Current Protocols in Molecular Biology

CHAPTER 1 Escherichia coli, Plasmids, and Bacteriophages INTRODUCTION M astery of current DNA technology requires fami...

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CHAPTER 1 Escherichia coli, Plasmids, and Bacteriophages INTRODUCTION

M

astery of current DNA technology requires familiarity with a small number of basic concepts and techniques. The goal of this chapter is to present this information concisely, yet in enough detail to be useful when a procedure goes wrong. Section I is devoted to Escherichia coli. Recipes are provided for media that support E. coli growth, as well as instructions for making the simple tools needed to work with bacterial cells. Growth of E. coli in liquid and solid media is then detailed. The final unit in Section I describes a few detailed aspects of E. coli biology learned from classical bacterial genetic studies, the understanding of which is especially relevant to the techniques used in modern DNA work. The remainder of the chapter discusses vectors used to introduce foreign DNA into E. coli. For the purposes of this chapter, vectors are said to be derived from plasmids, from bacteriophage lambda and related phages, or from filamentous phages. (Many modern vectors incorporate elements from more than one of these classes, and it is likely that this classification scheme will be hopelessly outdated by the time this chapter is revised.) Section II is concerned with plasmid vectors. Following a brief introduction to plasmid biology, procedures are described for purifying small and large amounts of plasmid DNA (“minipreps” and large preps). Finally, procedures for reintroducing plasmid DNA into bacterial cells are described. Section III covers vectors derived from bacteriophages. The biology of bacteriophage lambda is first introduced, followed by detailed aspects of biology that are especially significant when lambda derivatives are used as cloning vectors. Protocols in this section describe techniques for manipulating lambda-derived vectors, making single plaques, making and titering phage stocks, and isolating phage DNA. Finally, Section IV covers the biology and manipulation of vectors derived from filamentous phages. This chapter will be meaningful primarily to readers with some knowledge of the principles of molecular biology. Several books on molecular biology are recommended in the preface. For further advanced reading in the topics of this chapter, we recommend five books, all from Cold Spring Harbor Laboratory: Methods in Molecular Genetics (Miller, 1972), Advanced Bacterial Genetics (Davis et al., 1980), The Bacteriophage Lambda (Hershey, 1971), Lambda II (Hendrix et al., 1983), and Experiments with Gene Fusions (Silhavy et al., 1984). Many terms and jargon used by molecular biologists are introduced in this chapter. These terms are italicized at their first mention, and are defined below.

alpha fragment peptide containing the amino terminus of β-galactosidase, the lacZ gene product. Alpha fragments lack enzymatic activity, but can associate with omega fragments (see below) to form proteins whose β-galactosidase activity has been restored. alpha-complementation restoration of β-galactosidase activity to omega fragments by association with alpha fragments. Current Protocols in Molecular Biology (2002) 1.0.1-1.0.3 Copyright © 2002 by John Wiley & Sons, Inc.

amplification increase in copy number of some plasmids that occurs when host protein synthesis is inhibited. cloning site site on a vector into which foreign DNA is inserted. competent state in which bacterial or yeast cells are able to take up foreign DNA (for example, as the result of calcium treatment).

Escherichia coli, Plasmids, and Bacteriophages

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Introduction

cos site of action of phage lambda ter function. Cos site is cut by ter to yield two cohesive ends (cos ends). dilution, 10x-fold a solution or suspension that contains 1⁄10x as much (10−x as much) of the dissolved or suspended species as does the starting liquid. For example, to “do a 102-fold dilution” is to dilute a solution 100×. early-log phase period during the growth of a culture after the lag period. During early log phase growth, cells have begun exponential growth. efficiency of plating (EOP) titer of bacterial colonies or phage plaques under some experimental conditions divided by the titer of bacteria or phage obtained by growth on some reference medium. exponential growth period during which the number of cells in the culture increases as an exponential function of time, that is, during which cell number = ket. F factor genetic element found in some strains of E. coli and related species. F encodes proteins used in formation of sex pili which allow its transfer from bacterium to bacterium. female strain strain that does not contain the F factor and that receives genetic information when crossed with a strain containing F. helper phages bacteriophages that encode essential proteins and that allow other phages which do not encode these essential proteins to grow. incompatible phenomenon in which two plasmids cannot replicate in the same cell without continual selection for both of them. incompatibility group consists of plasmids that cannot be maintained together in the same cell. Compatible plasmids belong to different incompatibility groups. induction the onset of transcription of a new gene or operon, usually in response to some environmental stimulus. Phage induction or lysogenic induction describe the process in which prophage excise from the chromosome of bacteria that harbor them and begin to grow lytically. inoculation introduction of cells into a container of sterile growth medium. lag period period just after inoculation of a culture when cells have not yet begun to grow exponentially. late-log phase last period of exponential growth of a culture, after which growth slows and then stops altogether due to nutrient exhaustion or accumulation of waste products. lawn uniform layer of bacteria that covers the surface of a plate.

log phase period during growth of a culture in which cells are growing exponentially. low-copy-number plasmids plasmids found in less than about 20 copies per cell when cells containing them are grown in rich medium. lysogen E. coli cell or strain that harbors a dormant bacteriophage. male strain strain of bacteria that contains the F factor. male-specific phages bacteriophages that only grow on male strains because they adsorb to sex pili. marker detectable genetic difference between one organism and another (usually wild-type) organism of the same species. minimal medium growth medium for cells that contains only salts, vitamins, trace elements, and simple compounds which serve as carbon, nitrogen, and phosphorous sources. miniprep small-scale preparation or purification of some desired species, usually of plasmid or phage DNA. mobilization transfer of DNA from one cell to another caused by a mobile genetic element such as the F factor. multiplicity of infection (MOI) ratio of infecting bacteriophage to host cells. nonsense suppression the insertion of amino acids into proteins at positions where translation would normally not occur because the mRNA contains a UAG (amber), UAA (ochre) or UGA nonsense codon. nonsense suppressor tRNA that inserts amino acids at nonsense codons. The term is sometimes used for the genes encoding these tRNAs. omega fragment protein containing the carboxy-terminal fragment of β-galactosidase. This protein lacks enzymatic activity, but βgalactosidase activity can be restored when the peptide is complexed with an alpha fragment. ori (origin) site on genome at which DNA replication begins. outgrowth the growth of freshly transformed cells under nonselective conditions for enough time to allow proteins encoded by the foreign DNA to be expressed. overnight a small, freshly saturated liquid culture of bacteria. packaging extract extract from special strains of E. coli that contains bacteriophage lambda head proteins, tail proteins, and packaging proteins. Phage DNA added to such an extract is assembled into phage particles. par site on some plasmids which ensures that each daughter cell receives a plasmid copy. pilot protein protein in the coat of filamentous phages that helps phage DNA enter the cell.

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Current Protocols in Molecular Biology

plates petri dishes filled with solid medium, used to grow separated bacterial colonies or plaques. The term is sometimes used to refer to 96-well microtiter dishes. plating out the placement of bacteria or phage on plates so that colonies or plaques are formed. polylinker stretch of DNA that contains contiguous restriction sites. prophage dormant bacteriophage, usually integrated into the host chromosome, that replicates with the host bacterium. relaxed control applies to plasmids whose replication does not depend on the bacterial cell cycle. replicative form double-stranded circular filamentous phage DNA found inside infected cells. replicator stretch of DNA on a phage or plasmid that enables the phage or plasmid to replicate. rich medium growth medium that contains complex organic molecules (peptides, nucleotides, etc.). Typical components of rich media include tryptone (made from beef) and yeast extract (made from yeast). rolling-circle replication mechanism of replication sometimes used by circular molecules in which DNA polymerase continually circumnavigates the template, and thus synthesizes a long tail. satellite colonies small colonies that grow around a large colony on a plate containing selective medium. These are usually composed of cells unable to grow on selective medium, but which are able to grow near the large colony because the cells in the large colony neutralize the selective agent. saturated culture culture of cells in liquid medium that has stopped growing because nutri-

ents are exhausted or because waste products have accumulated. SOS response response of E. coli to DNA damage or other treatments that inhibit DNA replication. Lambda-derived phages are induced during this response. stringent control applies to plasmids whose replication is synchronized with the E. coli cell cycle. temperate describes bacteriophages capable of lysogenic growth. transfection introduction of bacteriophage DNA into competent E. coli cells. Also describes the introduction of any DNA (including plasmid DNA) into cells of higher eukaryotes. transformation introduction of plasmid DNA into E. coli or yeast. Also used to denote any of a number of changes in cultured higher eukaryotic cells to characteristics more typical of cancer cells (immortal growth, loss of contact inhibition, etc.).

LITERATURE CITED Davis, R., Botstein, D., and Roth, J.R. 1980. Advanced Bacterial Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Hendrix, R., Roberts, J., Stahl, F., and Weisberg, R. 1983. Lambda II. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Hershey, A.D. 1971. The Bacteriophage Lambda. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Miller, J. 1972. Methods in Molecular Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Silhavy, T., Berman, M.L., and Enquist, L.W. 1984. Experiments with Gene Fusions. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.


1.0.3 Current Protocols in Molecular Biology

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ESCHERICHIA COLI

SECTION I

Escherichia coli is a rod-shaped bacterium with a circular chromosome about 3 million base pairs (bp) long. It can grow rapidly on minimal medium that contains a carbon compound such as glucose (which serves both as a carbon source and an energy source) and salts which supply nitrogen, phosphorus, and trace metals. E. coli grows more rapidly, however, on a rich medium that provides the cells with amino acids, nucleotide precursors, vitamins, and other metabolites that the cell would otherwise have to synthesize. The purpose of this first section is to provide basic information necessary to grow E. coli. A more detailed introduction to certain aspects of E. coli biology may be found in UNIT 1.4. When E. coli is grown in liquid culture, a small number of cells are first inoculated into a container of sterile medium. After a period of time, called the lag period, the bacteria begin to divide. In rich medium a culture of a typical strain will double in number every 20 or 30 min. This phase of exponential growth of the cells in the culture is called log phase (sometimes subdivided into early-log, middle-log, and late-log phases). Eventually the cell density increases to a point at which nutrients or oxygen become depleted from the medium, or at which waste products (such as acids) from the cells have built up to a concentration that inhibits rapid growth. At this point, which, under normal laboratory conditions, occurs when the culture reaches a density of 1–2 × 109 cells/ml, the cells stop dividing rapidly. This phase is called saturation and a culture that has just reached this density is said to be freshly saturated. With very few exceptions, bacterial strains used in recombinant DNA work are derivatives of E. coli strain K-12. Most advances in molecular biology until the end of the 1960s came from studies of this organism and of bacteriophages and plasmids that use it as a host. Much of the cloning technology in current use exploits facts learned during this period.

Media Preparation and Bacteriological Tools

UNIT 1.1

Recipes are provided below for minimal liquid media, rich liquid media, solid media, top agar, and stab agar. Tryptone, yeast extract, agar (Bacto-agar), nutrient broth, and Casamino Acids are from Difco. NZ Amine A is from Hunko Sheffield (Kraft). MINIMAL MEDIA Ingredients for these media should be added to water in a 2-liter flask and heated with stirring until dissolved. The medium should then be poured into separate bottles with loosened caps and autoclaved at 15 lb/in2 for 15 min. Do not add nutritional supplements or antibiotics to any medium until it has cooled to 15 kb, do not transform well and frequently give lower DNA yields. Consider the final size of the vector plus insert when planning an experiment and wherever possible use smaller vectors. The higher the copy number the more vector DNA is produced, but high-copy-number vectors may not be applicable to all situations (see Mechanism of Replication and Copy-Number Control). When choosing a plasmid vector, consider both what sites are present in the polylinker and the order of the sites. If it will be necessary to manipulate the cloned sequence subsequent to insertion into the polylinker, plan ahead to ensure that the necessary sites will remain available in the polylinker. Selections or screens for identification of recombinant clones are useful for experiments where cloning efficiency is expected

to be low or when generation of a large number of clones is necessary. However, for routine subcloning experiments the advent of PCR (UNIT 15.1) has made it possible to rapidly screen through a large number of transformants to identify potential recombinant molecules, obviating the need for histochemical and genetic screening methods. The primary factor in choosing a plasmid vector is to understand and anticipate the experiments for which the recombinant clone will be used. The specialized functions of plasmid vectors are generally the key to selecting the correct vector for an experiment. For example, completely different types of vectors would be used for generating large quantities of DNA, expressing a fusion protein in bacteria, or for undertaking a two-hybrid screen in yeast. Once the type of vector required is determined then deciding upon a particular vector is dependent on both the details of the ancillary vector features, for example the type of promoter used to express a recombinant protein, and the properties of the replicator, polylinker, and selectable marker.

PLASMID VECTORS FOR PRODUCTION OF SINGLE-STRANDED DNA Plasmids have been developed that contain a filamentous phage origin of replication in addition to a plasmid ori. These “phagemid” vectors (UNIT 1.14) can be grown and propagated as plasmids. However, upon super-infection of a plasmid-containing cell with a wild-type helper phage, the phage ori becomes active, and single-stranded DNA (ssDNA) is produced and secreted. There are usually (+) and (−) versions of these vectors where the phage ori is in opposite orientation so that it is possible to produce ssDNA from either DNA strand. For many years, ssDNA was the substrate of choice for DNA sequencing (UNIT 7.4) and oligonucleotide-directed mutagenesis (UNIT 8.1). Development of sequencing and mutagenic protocols that use double-stranded templates has made the production of ssDNA a less frequently utilized featu re of plasmid vectors. The pBluescriptI, pBluescriptII, and pBS phagemid vectors derived from the general cloning vector pUC19 (Fig. 1.5.2) are examples of phagemids that incorporate the f1 filamentous phage ori.

PLASMID VECTORS FOR CLONING LARGE INSERTS Cosmid vectors, plasmids carrying a lambda phage cos site (e.g., pWE15, Fig. 1.5.3), were



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Eco O 109 2674

396 EcoRI SacI KpnI SmaI XmaI BamHI XbaI HincII Pst I SphI HindIII 447

Aat lI 2617 Nde l 183 Nar l 235 Ssp l 2501 Xmn l 2294 lac Z′ Scal 2177 Plac r

Ap

′lac l

pUC19 2686 bp ori

AfI III 806

Gsu l 1784 Cfr10I 1779 PpaI 1766

400

420

440

460

AGTGAATTCGAGCTCGGTACCCGGGGATCCTCTAGAGTCGACCTGCAGGCATGCAAGCTTGGCGTAATCATGGTCAT

EcoRI

Sac I

KpnI

SmaI XmaI

BamHI XbaI

HincII Acc I Sal I

Pst I

SphI

HindIII lac Z′

1 ThrlleMetThr(Met)

T3

EcoRI

NotI

Bam HI

NotI

EcoRI

Figure 1.5.2 Map of pUC19.

T7

Apr cos

ori

pWE15 8.2 kb

SV40 ori Neor

Introduction to Plasmid Biology

Figure 1.5.3 Map of pWE15 (adapted from Wahl et al., 1987, with permission).

1.5.6 Supplement 41


T7 SaII

NotI cosN IoxP

SP6 HB

NotI

IacZ

par C

EcoRI CMr

par B SnaBI pBeloBAC11 7.3 kb

XhoI

par A ori S XbaI rep E EcoRV EcoRV Figure 1.5.4 Map of pBeloBAC11 (adapted from Shizuya et al., 1992, with permission). Abbreviation: CM, chloramphenicol.

developed to facilitate cloning of large DNA fragments (UNIT 1.10). Cosmids can be transformed into cells like plasmids and once in the cells, replicate using their plasmid ori. The ColE1 type replicators are the most commonly used in these vectors, and cosmids can generally be maintained at high-copy-number in E. coli. Cosmids can also be packaged into lambda phage heads. In order for packaging to occur the cos sites must be separated by 40 to 50 kb, the approximate length of the wild-type lambda genome. Many cosmid vectors are between 5 to 10 kb in size and therefore can accept inserts of 30 to 45 kb. Because of the size of inserts they can accept, the high efficiency of packaging recombinant molecules into phage, and the efficiency of infection of cosmid-containing phage heads, cosmid vectors are frequently used for making genomic libraries. Unfortunately, propagation of insert-containing cosmid vectors in E. coli sometimes results in the deletion of all or a portion of the insert. To address this problem, a new set of cosmid vectors have been developed that replace the ColE1 replica-

tor with the F factor replicator. These “fosmid” vectors are maintained at low-copy-number, 1 to 2 copies per cell, and are more stable than higher-copy-number cosmid vectors when grown in E. coli (Kim et al., 1992). Another type of plasmid cloning vector, called bacterial artificial chromosome (BAC; Fig. 1.5.4), has been developed using the F factor replicator for propagation of very large pieces of DNA (100 to 500 kb). The vectors are used in a similar manner to yeast artificial chromosome (YAC) vectors but have the advantage of being manipulated solely in E. coli.

PLASMID VECTORS FOR EXPRESSION OF LARGE QUANTITIES OF RECOMBINANT PROTEINS There are a wide variety of vectors for expressing high levels of recombinant proteins in E. coli, insect, and mammalian cells (Chapter 16). The general goal of expressing proteins in any of these systems is to produce large quantities of a particular protein upon demand.



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While the features of the protein expression systems vary considerably, the basic properties outlined for expressing proteins in E. coli are common to all of them. Expression vectors are usually designed such that production of the foreign protein is tightly regulated. This is necessary because the host cellular machinery is co-opted to produce large quantities of the foreign protein, the shear amount of which may be toxic to the cell, and/or the foreign protein may encode a function that will inhibit cell growth or kill the host cell. Generally, expression vectors are configured so the polylinker cloning site is downstream of an inducible promoter. One of the promoters commonly used in E. coli expression vectors is the hybrid trp/lac promoter (trc) which contains the lacO operator site. This promoter is turned off in the presence of the lacIq repressor; the repressor gene is either carried by the bacterial host or is encoded on the expression vector itself. Expression of the foreign protein from the trc promoter is induced by the addition of IPTG. The quantity of foreign protein produced will be determined by both the rate of transcription of the gene and the efficiency of translation of the mRNA. Therefore, in addition to regulated highly inducible promoters, many E. coli protein expression vectors are designed to optimize translation of the foreign protein in bacterial cells. These expression vectors include an efficient ribosome binding site and an ATG start codon uptsream of the polylinker cloning site. Usually the cloning sites in the polylinker are designed so that it is possible to make an inframe fusion to the protein of interest in all three reading frames. For many experiments, high levels of pure recombinant protein are required, and some expression vectors are designed to create tagged or fusion proteins that facilitate purification of the recombinant protein. Tag sequences may be located 5′ or 3′ to the polylinker, creating either amino- or carboxy-terminal-tagged fusion proteins. Six polyhistidine residues (UNIT 10.11B), the FLAG epitope, and glutathione-S-transferase protein (UNIT 16.7) are some of the sequences that are appended to proteins to assist in purification. Tagged or fusion proteins can be rapidly and efficiently purified using an appropriate affinity column designed to tightly bind the tag or fusion region. Many protein expression vectors are created with specific protein cleavage sites adjacent to the tag or fusion sequences to allow for removal of these sequences from the purified protein. This feature may prove to be essential if the tag

or fusion sequence impairs the function of the protein in the relevant assays.

PLASMID VECTORS FOR REPORTER GENE FUSIONS Plasmid vectors have been designed to simplify the construction and manipulation of reporter gene fusions, where a promoter of interest is used to drive an easily scored marker gene (UNIT 9.6). Gene fusions provide a rapid and simple method for following the expression pattern conferred by a particular promoter. There are generally two types of reporter fusions, transcriptional and translational fusions. For transcriptional fusions, the ATG start codon is provided by the marker gene. In translational fusions, the 5′ untranslated region and ATG are provided by the gene of interest; in fact these constructs may fuse a large portion, or even entire coding region, to the amino terminus of a marker. In the reporter vector, the polylinker cloning site is located directly upstream of the reporter gene for insertion of the promoter fragment. In vectors that are designed for expression in eukaryotic cells, a polyadenylation signal is located downstream of the reporter gene. There are a variety of reporter genes used including chloramphenicol acetyltransferase, luciferase, β-galactosidase, secreted alkaline phosphatase, human growth hormone, β-glucuronidase, and green fluorescent protein (also see UNITS 9.6-9.7C). An example of a reporter vector pEGFP, is shown in Figure 1.5.5. The choice of reporter gene will depend on the cell type or organism, whether the assay will be done in vivo or in vitro, and whether quantitative or qualitative data is desired. The replicator, selectable marker, and other elements of the reporter vector will also have to be compatible with the system.

PLASMID VECTORS FOR YEAST Yeast plasmid vectors contain the same basic features as E. coli vectors—replicator, selectable marker, and cloning site (UNIT 13.4). There are two primary types of replicators used in yeast plasmid vectors, autonomously replicating sequences (ARS) derived from the yeast chromosomes and the natural yeast 2µm plasmid replicator. ARS-containing plasmids frequently contain yeast centromeric sequences to ensure their stable maintenance in the population of cells. Both ARS and 2µm vectors average 10 to 30 copies per cell. Many yeast vectors are “shuttle” vectors that can be maintained in both S. cerevisae and E. coli (e.g., pRS303; see Fig. 1.5.6). These vectors have an E. coli plas-

1.5.8 Supplement 41


tions in a biosynthetic pathway. For example, the URA3 marker carried on a plasmid is used to restore the ability to grow in the absence of uracil to a ura3− yeast. Therefore, yeast selectable markers, unlike most bacterial selectable markers, are strain dependent. There are a wide variety of yeast vectors available that have a range of specialized func-

P

f1 ori

SV40 poly A

NotI

Figure 1.5.5

HSV TK poly A

Map of pEGFP-1.

Kanr/ Neor

pEGFP-1 4.2 kb

e

SV40 ori PSV40

EGFP pUC ori

MCS

EGFP MCS

TA GCG CTA CCG GAC TCA GAT CTC GAG CTC AAG CTT CGA ATT CTG CAG TCG ACG GTA CCG CGG GCC CGG GAT CCA CCG GTC GCC ACC ATG GTG Eco47III XmaI BamHI AgeI SalI KpnI SacII ApaI EcoRI PstI XhoI SacI HindIII BglII Bsp1201 SmaI Ecl136II AccI Asp7181

mid replicator, frequently pMB1 derived, and a yeast replicator. Conversely, the yeast integrating plasmid vectors, used for introduction of genes into the yeast chromosome, have a bacterial replicator but no yeast replicator. The selectable markers in yeast are for the most part recessive markers, usually cloned yeast genes that are used to complement muta-



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CEN6 pRS 313 4967

ARSH4

AatII 4384 ScaI 3944 PvuI 3833

0

HindIII 808

4

HIS3

Apr

Bst XI 886 1

pRS303 4453 bp

HindIII 995 KpnI 1122 PstI 1183

3

ori

f1(+) lacZ 2

PvuII 2395

2073

2175 KpnI

NaeI 1748 PvuI 1920 PvuII1950

XhoI ClaI EcoRV PstI BamHI XbaI EagI Bst XI

ApaI SaII HindIII EcoRI SmaI SpeI NotI SacII SacI

T3

T7

Figure 1.5.6 Map of pRS303 (adapted from Sikorski and Heiter, 1989, with permission).

tions—e.g., expression of recombinant proteins in yeast, integration of sequences into the yeast genome, and cloning of very large fragments (hundreds of kilobases) of genomic DNA.

PLASMID VECTORS FOR EXPRESSION IN CULTURED MAMMALIAN CELLS


The type of vector that is used in mammalian cells depends on whether the experiment involves transient transfection into mammalian cells or the generation of stable mammalian cell lines carrying the construct of interest (UNITS 9.1-9.5). Virtually any plasmid vector that contains an appropriate construct for expression in mammalian cells can be used in transient assays. In transient assays, the plasmid vector carrying the DNA of interest is transfected into mammalian cells, the cells are harvested some time later (24 to 96 hr), and the pertinent assay

is performed. It is not necessary for plasmid vectors used in these assays to carry a mammalian selectable marker or to replicate in mammalian cells; therefore easily manipulatable bacterial plasmids, like pUC, are usually the vectors of choice. Plasmid vectors carrying a selectable marker that functions in mammalian cells are necessary for the generation of stable transgenic lines (UNITS 9.5; e.g., pcDNA3.1, Fig. 1.5.7). In order to generate a stable mammalian cell line, the plasmid DNA is transfected into the mammalian cells, and over a period of several weeks the DNA of interest is selected for based on expression of the vector-borne marker. Many mammalian vectors cannot replicate in mammalian cells, and the only way to maintain the DNA of interest and the selectable marker is for the vector to randomly integrate into the mammalian genome. However, there are some plasmid vectors that carry the simian

1.5.10 Supplement 41


T7

NheI PmeI AflII HindIII Asp718I KpnI BamHI Bst XI EcoRI EcoRV Bst XI NotI XhoI XbaI ApaI PmeI

pcDNA3.1(+)

T7

NheI PmeI ApaI XbaI XhoI NotI Bst XI EcoRV EcoRI Bst XI BamHI Asp718I KpnI HindIII AflII PmeI

pcDNA3.1(–)

PCMV

BGH polyA f1 ori

SV40 ori

Apr

pcDNA3.1 Vectors Neor

or

Zeor

SV40 polyA ColE1 Figure 1.5.7 Map of pcDNA3.1.

virus (SV40) or bovine papilloma virus (BPV) ori that can replicate in mammalian cells if the necessary viral replication proteins are provided; in most cases the viral replication proteins must be provided by the host cell line, limiting the range of cell types in which these vectors are useful. The most critical general feature of plasmid vectors (and also viral vectors) for creation of stable mammalian cell lines is the selectable marker (see UNIT 9.5). Unlike transient assays, formation and maintenance of stable cell lines requires selection. The number of selectable markers available for mammalian cells is limited; resistance to hygromycin, puromycin, G418, and neomycin are the predominant markers used. Since expression of foreign proteins or assaying the expression of a mammalian gene may require multiple plasmids and/or integration of constructs into the mammalian chromosome, careful planning must take place to ensure that all constructs can be selected for with the limited number markers. Further, since selection may be necessary over a long period

of time—weeks for generation of the lines and years for their maintenance—it is important to recognize that some of the antibiotics used in selection are very expensive and thus cost may be a factor in experimental design. Specialized plasmid vectors are also used for production of viruses that can infect mammalian cells. Plasmid vectors have been designed to produce infectious retroviral particles when transfected into the appropriate packaging cell line (UNIT 9.9). These retroviral vectors are then used to create stable transgenic lines in mammalian cell types not amenable to transfection. In addition, plasmid vectors have been designed to allow easy insertion of DNA sequences into vaccinia virus for the purpose of creating recombinant viruses that overexpress recombinant proteins (UNIT 16.15).

PLASMID VECTORS FOR NON–E. COLI BACTERIA Three features required of all bacterial plasmid vectors are that they replicate (unless they are suicide vectors), carry a selectable marker,



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trfA oriV

pRR54 8.1 kb bla par

ori T

H3 SphI XbaI BamHI Figure 1.5.8 Map of pRR54 (adapted from Roberts et al., 1990, with permission).


and can be easily introduced into host cells. The first thing to consider when selecting a plasmid vector for use in a non-E. coli host is whether or not it can replicate and be stably maintained in the particular strain. Different plasmid replicators have different host ranges, some have a narrow host range and can only replicate in a specific strain while others are promiscuous and can replicate in a wide variety of host (e.g., pRR54, Fig. 1.5.8). Unfortunately, the ColE1type replicators have a narrow host range, thus many standard E. coli vectors cannot be maintained in other bacteria. However, a number of broad-host-range replicators, such as RK2 and RSF1010, have been well characterized and used to construct vectors that can replicate in many gram-negative bacterial species (and in the case of RSF1010, some gram-positive species as well). Antibiotic resistance genes are used as selectable markers in non-E. coli bacterial hosts; however, the quantity of an antibiotic used to select against non-plasmid-containing cells is usually higher than the quantity used for E. coli selection. Furthermore, some bacterial strains are inherently resistant to particular antibiotics, thus it is important to determine whether the selectable marker carried on a plasmid vector is functional in a particular strain.

The favored method for introduction of plasmid DNA into bacterial host cells varies widely with the bacterial strain. Unlike E. coli, many bacteria cannot be efficiently transformed by chemical procedures or electroporation. In these cases bacterial mating is used to introduce plasmid DNA into the desired bacterial host. Mating to transfer a plasmid vector from an E. coli host where the vector is maintained and manipulated to a recipient bacterium requires both cis- and trans-acting functions. The tra (or mob) genes encode the trans-acting proteins necessary to transfer the plasmid DNA from one bacterium to another, and they are usually located on a helper plasmid that is distinct from the plasmid vector. Any plasmid vector that is to be mobilized must contain the cis-acting site called oriT where the DNA is cleaved and transfer is initiated.

MAPS OF PLASMIDS Figures 1.5.2-1.5.11 present maps of plasmids that are in widespread use or are examples of plasmids whose special functions make them useful for particular techniques described in this manual. Note that the trend in development of vectors is to include multiple features on a single vector, and many of these examples span the vector categories described.



EcoRI 4361 ClaI 23 HindIII 29 AatII 4286 EcoRV 185 SspI 4170 NheI 229 BamHI 375 SphI 562 Sal I 651

ScaI 3846 PvuI 3735 PstI 3609 r

r

Ap

PpaI 3435

Tet

EagI 939 NruI 972 BspMI 1063

pBR322 4363bp

ori

BsmI 1353 Sty I 1369 AvaI 1425 Bal I 1444 BspMII 1664

Afl lII 2475

PvuII 2066

Ndel 2297

Tth111I 2219 Figure 1.5.9 Map of pBR322 (Bolivar et al. 1977; sequence in Sutcliffe, 1978).

pBR322 is one of the classic cloning vectors from which many other vectors are derived. It contains an amplifiable pMB1 replicator and genes encoding resistance to ampicillin and tetracycline. Insertion of DNA into a restriction site in either drug-resistance gene usually inactivates it and allows colonies bearing plasmids with such insertions to be identified by their inability to grow on medium with that antibiotic (see Fig. 1.5.9; Bolivar et al., 1977; sequence in Sutcliffe, 1978). pUC19 belongs to a family of plasmid vectors that contains a polylinker inserted within the alpha region of the lacZ gene. The polylinkers are the same as those used in the m13mp series (Fig. 1.14.2). pUC19 and pUC18 have the same polylinker but in opposite orientations. Under appropriate conditions (see UNIT 1.4 for a description), colonies that bear plasmids containing a fragment inserted into the polylinker form white colonies instead of blue ones. These pMB1-derived plasmids (see Fig. 1.5.2) maintain a very high-copy-number (1000 to 3000 per genome). Wild-type and recombinant plasmids confer ampicillin resistance and can be amplified with chloramphenicol (Norrander et al., 1983). In addition wildtype plasmids confer a LacZ+ phenotype to appropriate cells (e.g., JM101 cells, UNIT 1.4). pBluescript is a commonly used phagemid cloning vector that contains a polylinker in-

serted into the alpha region of the lacZ gene and T3 and T7 promoter sequences flanking the cloning sites. The f1 (+) filamentous phage origin of replication in pBluescript SK+ allows for the recovery of the sense strand of the lacZ gene as ssDNA; the pBluescript SK(−) vector (Fig. 1.5.10) with the f1 origin in the opposite orientation, f1(−), facilitates recovery of the other strand. The position of the polylinker in the alpha region of the lacZ gene allows for identification of inserts based on a blue/white color screen under the appropriate conditions. The T3 and T7 promoters are recognized by bacteriophage RNA polymerases. Transcription from these promoters reads into the polylinker from either side. RNA transcripts of any DNA cloned into the polylinker can thus be produced by run-off transcription in vitro. pWE15 is an example of a cosmid vector used for cloning DNA fragments ∼35 to 45 kb (see Fig. 1.5.3; Wahl et al., 1987). The cos sites allow the DNA to be cut and packaged into phage heads by the appropriate lambda proteins. There is a single unique BamHI cloning site flanked by T3 and T7 promoter sequences. These promoters are particularly useful for production of labeled RNA probes corresponding to the ends of the insert DNA, and these can be used to identify overlapping cosmids for chromosome walking and construction of cosmid contigs. Not1 sites flanking the cloning site can



Supplement 41

1.5.14

Supplement 41


lacZ

T3 primer 5′–AATTAACCCTCACTAAAGGG– 3′

ColE1ori

2958 bp

AflIII 1153

pBluescript SK (+/–) MCS

Apr

f1(+)ori

SK primer 5′–CGCTCTAGAACTAGTGGATC– 3′

PvuII 977

SacI 759

KpnI 657

PvuI 503 PvuII 532

NaeI 333

SspI 445

Figure 1.5.10

759 Bsp106I HindII Eco 0109I ClaI AccI DraII ApaI XhoI Eco RV HindIII Sal I KpnI AATTCGATATCA AGCTTATCGATACCGTCGACC TCGAGGGGGGGCCCGGTACC CCAAT TCGCCCTATAGTGAGTCGTAT TACAAT TCACTGGCCGTCGT T T TACAA – 3′ (+) GCTATAGT TCGAATAGCTATGGCAGCTGGAGCTCCCCCCC GGGCCATGG GGTTAAGCGGGATAT CACTCAGCATAATGTTAAGTGACCGGCAGCAAAATGT T – 5′ (–) +1 T 7 promoter 657 3′–CGGGATATCACTCAGCATA A T G – 5′3′–TGACCGGCAGCAAAATG– 5′ 3′– C TATGGCAGCTGGAGCT– 5′ T7 primer M1320 primer KS primer

Map of pBluescript SK (+/−).

816 β-Galactosidase

SpeI SmaI Bst XI Eag I EcoRI MET T3 promoter +1 XbaI Sac I Sac II Not I BamHI Pst I 5′– GGAAACAGCTATGACCATGAT TACGCCAAGCTCGAAAT TAACCCTCACTAAAGGGAACAA AAGCTGGAGCTCCACCGCGGTGGCGGC CGCTCTAGAACTAGTGGATCCCCCGGGCTGCAGG 3′–CCTT TGTCGATACTGGTACTA ATGCGGT TCGAGCTTTAATTGGGAGTGATT TCCCTTGT T T TCGACCTCGAGGTGGCGCCAC CGCCGGCGAGATCTTGATCACCTAGGGGGCCCGACGTC CT TAA

reverse primer 5′– GGAAACAGCTATGACCATG–3′

PvuI 2416

ScaI 2526

XmnI 2645

SspI 2850 SspI 22

NaeI 134

4.0

3.5

lacI q

NcoI EcoRI SstI KpnI SmaI 4.37/0 BamHI XbaI 0.5 SalI PstI Ptrc 5S SphI T1 HindIII T2

1.0

pTrc99A,B,C 3.0

ori

Apr PvuI 1.5

2.5 2.0 pTrc99A

1

2

Met Glu

3

4

Phe Glu

5

6

7

8

9

Leu

Gly

Thr

Arg

Gly

11

12

13

14

15

16

17

18

19

Ser Ser

10

Arg

Val

Asp

Leu

Gln

Ala

Cys

Lys Leu

20

rrnB RBS TCACACAGGAAACAGACC ATG GAA TTC GAG CTC GGT ACC CGG GGA TCC TCT AGA GTC GAC CTG CAG GCA TGC AAG CTT GGCTGTTTTGGC

Nco l

pTrc99B

EcoRI

Sst I

XmaI SmaI

KpnI

1

2

3

4

5

6

Met

Gly

Ile

Arg

AIa

Arg

7 Tyr

8 Pro

BamHI

SaI I Acc I HincII

XbaI

9

10

11

Gly

IIe

Leu

Pst I

SphI

HindIII

rrnB RBS TCACACAGGAAACAGACC ATG GGA ATT CGA GCT CGG TAC CCG GGG ATC CTC TAG A GTCGACCTGCAGGCATGCAAGCTTGGCTGTTTTGGC Nco l

pTrc99C

EcoRI

1

2

3

Met

Gly

Asn

Sst I

KpnI

4

5

6

Ser

Ser

Ser Val

7

XmaI SmaI

BamHI

8

9

Pro

Gly

10

SaI I Acc I HincII

XbaI

11

Pst I

Sph I

HindIII

12

13

14

15

16

17

18

19

20

21

Asp Pro Leu

Glu

Ser

Thr

Cys

Arg

His

Ala

Ser

Leu

rrnB RBS TCACACAGGAAACAGACC ATG GGG AAT TCG AGC TCG GTA CCC GGG GAT CCT CTA GAG TCG ACC TGC AGG CAT GCA AGC TTG GCTGTTTTGGC Nco l

EcoRI

Sst I

KpnI

XmaI SmaI

BamHI

XbaI

SaI I Acc I HincII

Pst I

SphI

HindIII

Figure 1.5.11 Map of pTrc99A,B,C.

potentially be used to excise an intact insert fragment from the vector. The ColE1-derived ori and ampicillin resistance gene allow for replication and selection in bacteria. The SV40 promoter (included in SV40 ori) which drives the neomycin phosphotransferase gene enables selection in eukaryotic cells. pBeloBAC11 is an example of the family of bacterial-artificial-chromosome vectors based on the low-copy-number F factor replicator (see Fig. 1.5.4; Shizuya et. al., 1992). BAC vectors are used for cloning large DNA fragments (100 to 500 kb) in E. coli and are used commonly in genome-mapping strategies. oriS, repE, parA, parB, and parC genes are the

essential genes that compose the F factor replicator. oriS and repE genes are required for unidirectional replication of the plasmid, and parABC loci stably maintain the copy number at one to two per E. coli genome. There are two unique cloning sites (HindIII and BamHI) inserted into the lacZ alpha region. Other useful features of the cloning region are (1) T7 and SP6 promoter sequences flanking the cloning sites, (2) NotI restriction sites flanking the cloning sites for potential excision of the insert, and (3) and presence of the loxP and cosN sites that can be cleaved by specific enzymes. The ends generated by cleavage at loxP or cosN can be used as fixed reference points in building an



Supplement 41


ordered restriction map by end labeling and partial restriction digestion. pEGFP-1 is a selectable vector for monitoring promoter activity in mammalian cells via fluorescence of a green fluorescent protein (GFP) derivative (Clontech; see Fig. 1.5.5; Yang et al., 1996). The vector contains a neomycin resistance gene downstream of the SV40 early promoter for selection of stably transformed mammalian cells. It has a polylinker located upstream of the EGFP gene, so that the function of promoter sequences introduced into the polylinker can be assessed based on EGFP activity. The EGFP gene is modified from wildtype GFP to ensure expression in mammalian cells, it has silent base mutations that correspond to human codon-usage preferences, and sequences flanking the coding region have been converted to a Kozak consensus translation initiation signal. The vector backbone contains an f1 ori for production of ssDNA, a pUC-derived ori for propagation in E. coli and a kanamycin resistance gene for selection in bacteria. A series of yeast shuttle vectors (pRS304, 305, and 306) has been created to facilitate manipulation of DNA in Saccharomyces cerevisiae (see Fig. 1.5.6; Sikorski and Hieter, 1989). These vectors have a backbone derived from pBLUESCRIPT into which the features necessary for replication and maintenance in yeast have been introduced. The members of this series of plasmids differ only in the yeast selectable marker incorporated; pRS303 carries the HIS3 marker that complements a nonreverting his3 chromosomal mutation in specific yeast strains. These plasmids contain an autonomously replicating sequence as well as a centromere sequence, CEN6, that ensures stable maintenance in yeast cells. pcDNA3.1 is a selectable cloning and expression vector for use in mammalian cells. The features of this vector include a neomycin resistance gene driven by the SV40 early promoter (contained within the SV40 ori) and terminated by an SV40 polyadenylation signal for selection in mammalian cells (see Fig. 1.5.7). In addition, due to the inclusion of the SV40 ori, the vector can replicate as an episome in cells expressing the SV40 large T antigen. The polylinker cloning site is located downstream of strong cytomegalovirus enhancerpromoter sequences and upstream of the bovine growth hormone gene termination signals for high-level expression of protein-coding sequences cloned into this vector. This vector also contains some of the more standard features of

other plasmid vectors, including a ColE1 replicator for propagation in E. coli, the ampicillin resistance gene for selection in E. coli, the f1 ori for production of ssDNA, and the T7 promoter sequence for in vitro transcription of DNA inserted into the polylinker. pRR54 is an example of a broad-host-range mobilizable plasmid vector. This vector contains replicator and stablilization sequences derived from the natural RK2 broad-host-range plasmid (see Fig. 1.5.8; Roberts et al., 1990). oriV is the vegetative origin of replication, trfA encodes trans-acting functions necessary for replication, and par encodes a locus that enhances stability of the plasmid. This plasmid can be mated into diverse gram-negative species as long as the appropriate mobilization machinery is provided in trans because it contains the origin of conjugal transfer, oriT. The plasmid carries the β-lactamase gene, allowing for ampicillin/carbenecillin selection of plasmid containing bacteria. The pTrc series of plasmid expression vectors facilitates regulated expression of genes in E. coli. These vectors carry the strong hybrid trp/lac promoter, the lacZ ribosome-binding site (RBS), the MCS of pUC18 that allows insertion in three reading frames, and the rrnB transcription terminators (see polylinker sequences given below the vector diagram in Fig. 1.5.11). These vectors are equally useful for expression of unfused proteins (resulting from insertion in the NcoI site) or for expression of fusion proteins (using one of the cloning sites in the correct translational frame). The presence of the lacIq allele on the plasmid ensures complete repression of the hybrid trp/lac promoter during cloning and growth in any host strain (see Amann et al., 1988, for further details).

LITERATURE CITED Amann, E., Ochs, B., and Abel, K.-J. 1988. Tightly regulated tac promoter vectors useful for the expression of unfused and fused proteins in Escherichia coli. Gene 69:301-315. Bolivar, F., Rodriguez, R.L., Greene, P.J., Betlach, M.C., Heynecker, H.L, and Boyer, H.W. 1977. Construction of useful cloning vectors. Gene 2:95-113. Chang, A.C.Y. and Cohen, S.N. 1978. Construction and characterization of amplifiable mulitcopy DNA cloning vehicles derived from the P15A cryptic miniplasmid. J. Bacteriol. 134:11411156. Gerhart, E., Wagner, H., and Simmons, R.W. 1994. Antisense RNA control in bacteria, phages, and plasmids. Annu. Rev. Microbiol. 48:713-742.



Jackson, D.A., Symons, R.M., and Berg, P. 1972. Biochemical method for inserting new genetic information into DNA of Simian Virus 40 circular DNA molecules containing lambda phage genes and the galactose operon of Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 69:2904-2909. Kahn, M., Kolter, R., Thomas, C., Figurski, D., Meyer, R., Remaut, E., and Helinski, D.R. 1979. Plasmid cloning vehicles derived from plasmids ColE1, F, R6K, RK2. Methods Enzymol. 68:268280.

Sikorski, R.S. and Hieter, P.I. 1989. A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122:19-27. Stoker, N.G., Fairweather, N., and Spratt, B.G. 1982. Versatile low-copy-number plasmid vectors for cloning in Escherichia coli. Gene 18:335-341. Sutcliffe, J.G. 1978. Complete nucleotide sequence of the Escherichia coli plasmid pBR322. Cold Spring Harbor Symp. Quant. Biol. 43:77-90.

Kim, U.J., Shizuya, H., de Jong, P.J., Birren, B., and Simon, M.I. 1992. Stable propagation of cosmidsized human DNA inserts in an F-factor based vector. NAR 20:1083-1085.

Uhlin, B.E., Schweickart, V., and Clark, A.J. 1983. New runaway-replication-plasmid cloning vectors and suppression of runaway replication by novobiocin. Gene 22:255-265.

Norrander, J., Kempe, T., and Messing, J. 1983. Construction of improved M13 vectors using oligonucleotide-directed mutagenesis. Gene 26:101-106.

Wahl, G.M., Lewis, K.A., Ruiz, J.C., Rothenberg, B., Zhao, J., and Evans, G.A. 1987. Cosmid vectors for rapid genomic walking, restriction mapping, and gene transfer. Proc. Natl. Acad. Sci. U.S.A. 84:2160-2164.

Roberts, R.C., Burioni, R., and Helinski, D.R. 1990. Genetic characterization of the stabilizing functions of a region of broad-host-range plasmid RK2. J. Bacteriol. 172:6204-6216. Shizuya, H., Birren, B., Kim U.J., Mancin, V., Slepak, T., Tachiiri, Y., and Simon, M.I. 1992. A bacterial system for cloning large human DNA fragments. Proc. Natl. Acad. Sci. U.S.A. 89:8794-8797.

Yang, T., Cheng, L., and Kain, S.R. 1996. Optimized codon usage and chromophore mutations provide enhanced sensitivity with the green fluorescent protein. Nucl. Acids Res. 24:4592-4593.

Contributed by Rhonda Feinbaum Massachusetts General Hospital Boston, Massachusetts



Supplement 41

Minipreps of Plasmid DNA

UNIT 1.6

Although there are a large number of protocols for the isolation of small quantities of plasmid DNA from bacterial cells (minipreps), this unit presents four procedures based on their speed and success: the alkaline lysis prep, a modification of the alkaline lysis prep that is performed in 1.5-ml tubes or 96-well microtiter dishes, the boiling method, and a lithium-based procedure. A support protocol provides information on storing plasmid DNA. ALKALINE LYSIS MINIPREP The alkaline lysis procedure (Birnboim and Doly, 1979, and Birnboim, 1983) is the most commonly used miniprep. Plasmid DNA is prepared from small amounts of many different cultures (1 to 24) of plasmid-containing bacteria. Bacteria are lysed by treatment with a solution containing sodium dodecyl sulfate (SDS) and NaOH (SDS denatures bacterial proteins, and NaOH denatures chromosomal and plasmid DNA). The mixture is neutralized with potassium acetate, causing the covalently closed plasmid DNA to reanneal rapidly. Most of the chromosomal DNA and bacterial proteins precipitate—as does the SDS, which forms a complex with potassium—and are removed by centrifugation. The reannealed plasmid DNA from the supernatant is then concentrated by ethanol precipitation.

BASIC PROTOCOL

Materials LB medium (UNIT 1.1) containing appropriate antibiotic (Table 1.4.1) Glucose/Tris/EDTA (GTE) solution TE buffer (APPENDIX 2) NaOH/SDS solution Potassium acetate solution 95% and 70% ethanol 10 mg/ml DNase-free RNase (optional; UNIT 3.13) 1.5-ml disposable microcentrifuge tubes 1. Inoculate 5 ml sterile LB medium with a single bacterial colony. Grow to saturation (overnight). 2. Spin 1.5 ml of cells 20 sec in a microcentrifuge at maximum speed to pellet. Remove the supernatant with a Pasteur pipet. The spins in steps 2 and 6 can be performed at 4°C or at room temperature. Longer spins make it difficult to resuspend cells.

3. Resuspend pellet in 100 µl GTE solution and let sit 5 min at room temperature. Be sure cells are completely resuspended.

4. Add 200 µl NaOH/SDS solution, mix by tapping tube with finger, and place on ice for 5 min. 5. Add 150 µl potassium acetate solution and vortex at maximum speed for 2 sec to mix. Place on ice for 5 min. Be sure mixing is complete.

6. Spin 3 min as in step 2 to pellet cell debris and chromosomal DNA. 7. Transfer supernatant to a fresh tube, mix it with 0.8 ml of 95% ethanol, and let sit 2 min at room temperature to precipitate nucleic acids. Contributed by JoAnne Engebrecht, Roger Brent, and Mustak A. Kaderbhai Current Protocols in Molecular Biology (1991) 1.6.1-1.6.10 Copyright © 2000 by John Wiley & Sons, Inc.


1.6.1 Supplement 15

8. Spin 1 min at room temperature to pellet plasmid DNA and RNA. 9. Remove supernatant, wash the pellet with 1 ml of 70% ethanol, and dry pellet under vacuum. 10. Resuspend the pellet in 30 µl TE buffer and store as in support protocol. Use 2.5 to 5 µl of the resuspended DNA for a restriction digest. Contaminating RNA may interfere with detection of DNA fragments on the agarose gel; it can be destroyed by adding 1 ìl of a 10 mg/ml RNase solution (DNase-free) to the digestion mixture. ALTERNATE PROTOCOL

ALKALINE LYSIS IN 96-WELL MICROTITER DISHES Escherichia coli cells that contain plasmids are grown and lysed, and the plasmid DNA is precipitated—all in the wells of 96-well microtiter dishes. This procedure makes it possible to perform hundreds of rapid plasmid preps in a day. It is based on an unpublished procedure by Brian Seed of Massachusetts General Hospital. Additional Materials TYGPN medium (UNIT 1.1) 70% ethanol, ice-cold Isopropanol 96-well microtiter plates (Dynatech PS plates or equivalent) Multichannel pipetting device (8-prong Costar; 12-prong Titer Tek) Multitube vortexer Sorvall RT-6000 low-speed centrifuge, or equivalent, with microplate carrier in H-1000B rotor 1. Add 0.3 ml sterile TYGPN medium to each well of a 96-well microtiter plate (see sketch 1.6A). Inoculate each well with a single plasmid-containing colony. The 96-well microtiter plates used must have U-shaped bottoms. To take full advantage of this protocol, one should perform all pipetting steps with a multichannel pipetting device.

adding solution to 96-well microtiter dish

Minipreps of Plasmid DNA

Sketch 1.6A

1.6.2 Supplement 15


2. Grow bacteria to saturation at 37°C (∼48 hr). All subsequent steps are performed at room temperature unless otherwise noted. Potassium nitrate in the TYGPN medium presumably acts as a terminal electron acceptor when the bacteria in the wells are growing anaerobically, resulting in high cell densities.

3. Spin saturated cultures in H-1000B rotor with microplate carrier for 10 min at 2000 rpm (600 × g), 4°C. Decant supernatant with brief flick. A microplate carrier is available for the Beckman JS-4.2 rotor. The same rpm values can be used for the JS-4.2 as those given here for the Sorvall H-1000B (see APPENDIX 1 for rotor conversion values).

4. Resuspend cells in the well bottoms by clamping the plate in a multitube vortexer and running it 20 sec at setting 4. 5. Add 50 µl GTE solution to each well. 6. Add 100 µl NaOH/SDS solution to each well. Wait 2 min. 7. Add 50 µl potassium acetate solution to each well. 8. Cover with plate tape or parafilm. Agitate vigorously in vortexer 20 sec at setting 4. Spin 5 min at 2000 rpm (600 × g), 4°C. 9. Insert a pipet tip just at the edge of the U in the bottom of the well. Remove 200 µl from each well and transfer to a new plate. Do not try to recover all the fluid in each well.

10. Add 150 µl isopropanol to each well of the new plate. Cover with plate tape, agitate, and chill 30 min at −20°C. 11. Spin 25 min at 2000 rpm, 4°C, and decant supernatant. Wash pellets with cold 70% ethanol, gently decant supernatant, wash with 95% ethanol, and again gently decant supernatant. Pellets often shrink visibly during the 70% ethanol wash, as impurities in them are dissolved. Restriction enzymes will not cut well if the DNA is contaminated with even very small amounts of NaOH/SDS or potassium acetate solutions. It is therefore very important to decant the supernatants from the isopropanol precipitation and ethanol washes thoroughly, but not so vigorously that the pellets are flung out of the well bottoms. If pellets become detached during either washing step, the plate should be respun at 2500 rpm for 5 min to bring the pellets to the bottom of the wells again.

12. Air dry pellets for 30 min, then resuspend in 50 µl TE buffer. Store as in support protocol and use 10-µl aliquots for digestion.



Supplement 15 CPMB

BASIC PROTOCOL

BOILING MINIPREP Bacteria that contain plasmid DNA are broken open by treatment with lysozyme, Triton (a nonionic detergent), and heat. The chromosomal DNA remains attached to the bacterial membrane and is pelleted to the bottom of a centrifuge tube during a brief spin. Plasmid DNA is precipitated from the supernatant with isopropanol (Holmes and Quigley, 1981). This procedure is recommended for preparing small amounts of plasmid DNA from 1 to 24 cultures. It is extremely quick, but the quality of DNA produced is lower than that from the alkaline lysis miniprep. Materials LB medium (UNIT 1.1) containing appropriate antibiotic (Table 1.4.1) STET solution Hen egg white lysozyme Isopropanol, ice-cold TE buffer (APPENDIX 2) 10 mg/ml DNase-free RNase (optional; UNIT 3.13) 1.5-ml disposable microcentrifuge tubes Boiling water bath (100°C) 1. Inoculate 5 ml sterile LB medium with a single bacterial colony. Grow at 37°C at least until mid-log phase ∼6 hr, (a freshly saturated overnight culture works even better; see UNIT 1.2). 2. Transfer 1.5 ml of the saturated culture to a 1.5-ml microcentrifuge tube and pellet the cells by spinning 20 sec in microcentrifuge at maximum speed. Discard supernatant with a Pasteur pipet. The spins in steps 2, 6, and 7 can be performed at 4°C or room temperature. Longer spins make it difficult to resuspend cells.

3. Resuspend the bacteria in 300 µl of STET solution containing 200 µg lysozyme. Vortex to achieve complete suspension. Be sure cells are completely resuspended in order to maximize the number of cells exposed to the lysozyme and consequently the yield of plasmid DNA.

4. Place tube on ice for 30 sec to 10 min. The time required for this step can vary between the limits indicated without affecting the yield or quality of the plasmid DNA.

5. Place tube in a boiling water bath (100°C) 1 to 2 min. Heat and detergents cause the weakened cell walls to break, releasing plasmid DNA and RNA, but not the larger bacterial chromosome which remains attached to or trapped inside the lysed cells.

6. Spin in microcentrifuge 15 to 30 min at maximum speed. The pellet, which should be fairly gummy, contains bacterial debris as well as chromosomal DNA. The supernatant contains plasmid DNA and RNA.

7. Pipet off supernatant into a new tube, carefully, without dislodging pellet. Mix with 200 µl (an equal volume) of cold isopropanol. Place at −20°C for 15 to 30 min. Spin 5 min in microcentrifuge at maximum speed. The cold isopropanol precipitates the plasmid DNA and cellular RNA. Considerably shorter incubation periods (e.g., 2 to 5 min) may be sufficient for precipitation. Minipreps of Plasmid DNA

8. Remove the supernatant by inverting the tube and flicking it several times. Dry the

1.6.4 Supplement 15


pellet by placing under a vacuum until it looks flaky. If a vacuum source is unavailable, the pellet can be air dried.

9. Resuspend the pellet in 50 µl TE buffer and store as in the support protocol. Use 5 µl of the resuspended DNA for a restriction digest. Contaminating RNA may interfere with detection of DNA fragments on the agarose gel; it can be destroyed by adding 1 ìl of a 10 mg/ml RNase solution (DNase-free) to the digestion mixture.

LITHIUM MINIPREP Plasmid DNA is obtained from E. coli grown on plates as colonies or in liquid cultures. Bacterial cells harboring plasmid DNA are sequentially treated with Triton X-100/LiCl and phenol/chloroform. These steps solubilize plasmid DNA while precipitating chromosomal DNA with cellular debris. The debris is removed by centrifugation. This isolation procedure yields preparations of plasmid DNA that are virtually devoid of chromosomal DNA.

BASIC PROTOCOL

The procedure described here (originally presented by He et al., 1990) for small-scale isolation of plasmid DNA can also be readily extended for large-scale preparations as described in the annotation to the final step. The merit of the approach is that it is extremely reliable and rapid—requiring no more than 20 min of simple and economical operations for a preparation. The final plasmid DNA preparations are of a purity and quality usable for most biological applications. Materials TELT solution LB medium (UNIT 1.1) containing appropriate antibiotic (Table 1.4.1) l:l (w/v) phenol/chloroform (UNIT 2.1) 100% ethanol, prechilled to −20°C TE buffer (APPENDIX 2) 10 mg/ml DNase-free RNase A (optional; UNIT 3.13) 1.5-ml disposable microcentrifuge tubes NOTE: All steps are performed at room temperature. 1. To isolate plasmid DNA from transformant colonies grown on agar plates, prepare the cells as follows: a. Using a microspatula, scoop out an entire bacterial colony grown to 2- to 5-mm diameter on an LB agar plate. Transfer the colony to a 1.5-ml microcentrifuge tube containing 100 µl TELT solution. b. Vortex thoroughly to suspend the cells. Proceed to step 3. 2. To isolate plasmid DNA from liquid cultures, prepare the cells as follows: a. Inoculate a colony of bacteria into 1.8 ml of sterile LB medium supplemented with appropriate antibiotic. Grow to saturation with shaking for 18 to 24 hr at 30°C (see UNIT 1.2). Glass test tubes with plastic caps are suitable. Place the tubes at a suitably inclined angle to achieve good agitation.

b. Carefully transfer the entire culture volume into a 1.5-ml microcentrifuge tube. At this stage the volume of the liquid culture will have been reduced to ∼1.5 ml.



Supplement 24

c. Pellet the cells by spinning in a microcentrifuge (10,000 × g) for 20 sec. The spins in steps 2, 4, 7, and 9 can be performed at 4°C or room temperature. Centrifugation for longer periods or at higher speeds makes it difficult to resuspend the cells in the following step.

d. With the tube held in a vertical position, aspirate the supernatant using a longtipped Pasteur pipet connected to a vacuum line. e. Add 100 µl of TELT solution to the pellet and resuspend by vortexing. Ensure that the cells are thoroughly suspended. See annotation to step 11 for scaled-up DNA preparations.

3. Add 100 µl of 1:1 phenol/chloroform and thoroughly vortex for 5 sec. This mixture may be left at room temperature for ≤15 min. Plasmid yield will elevate with increasing duration of incubation; however, incubation periods >15 min may result in phenol-mediated modification of DNA.

4. Microcentrifuge 1 min at 15,000 × g (maximum speed). 5. Using a pipettor or similar device, carefully withdraw 75 µl of the upper aqueous phase and transfer the contents into a clean microcentrifuge tube. Do not agitate the resolved phases. If mixing occurs, recentrifuge. When collecting the top layer avoid picking the debris at the interface.

6. To the supernatant, add 150 µl of chilled 100% ethanol. Mix the contents well to precipitate the plasmid DNA. 7. Pellet the nucleic acids by microcentrifuging 5 min at maximum speed. 8. Discard the supernatant by inverting the tube. When all the supernatant has drained, hold the tube in the same position for a few seconds and wipe off the last droplet from the rim of the tube by touching the edge of a Kleenex paper tissue. 9. Wash the pellet with 1 ml of cold 100% ethanol and harvest the nucleic acid pellet as in steps 6 and 7. After centrifugation, decant the supernatant carefully as the pellet may be loose.

10. Cap the tube. Stab a small hole in the cap with a thumbtack or a syringe needle. Place the tube in a vacuum desiccator (without desiccant). Apply vacuum until the nucleic acid pellet appears completely dry. A water-pumped vacuum line suffices for the purpose and usually takes ≤15 min.

11. Dissolve the pellet in 30 µl TE buffer. Vortex the contents well, capturing most of the DNA around the inner surface of the microcentrifuge tube. Store as in the support protocol and use 2 to 5 µl of DNA solution in a final 20-µl reaction volume for restriction digestion. Contaminating RNA may interfere with detection of DNA fragments on the agarose gel; it can be destroyed by adding 1 ìl of a 10 mg/ml RNase solution (DNase-free) to the digestion mixture. For scaled-up plasmid DNA preparations (He et al., 1991), increase the amounts of TELT solution and 1:1 phenol/chloroform in direct proportion to the culture volume used. For cultures ≤5 ml, transfer the cells after suspension in TELT buffer into a microcentrifuge tube. Wash the final nucleic acids pellet twice with 1 ml of 100% ethanol. For cultures between 5 and 100 ml, use Corex glass tubes for treatment with TELT and phenol/chloroform and for centrifugations (Sorvall RC-5C centrifuge at 6000 × g). Minipreps of Plasmid DNA

1.6.6 Supplement 24


STORAGE OF PLASMID DNA Plasmids can be maintained for a short period (up to 1 month) in bacterial strains simply by growing on selective plates and storing at 4°C. For permanent storage, bacteria harboring the plasmid should be grown to saturation in the presence of the appropriate selective agent. An equal volume (∼1 to 2 ml) of bacteria should be added to sterile 100% glycerol or a DMSO-based solution (recipe in UNIT 1.3) and frozen at −70°C in sterile vials. Cells taken from storage should again be grown on a selective plate (UNIT 1.1), and the plasmid DNA should be checked by restriction analysis (UNIT 3.1).

SUPPORT PROTOCOL

Plasmid DNA can be stored in TE buffer at 4°C for several weeks or preserved for several years by storing at −20° or −70°C. Most investigators prefer to store plasmids as frozen DNA, due to the widely held belief that plasmids stored in bacteria are sometimes lost, are rearranged, or accumulate insertion sequences and transposons during storage or on revival. Although such rearrangements certainly occur during storage of plasmids in bacteria in stab vials, we are unaware of any report of rearrangements affecting plasmids stored in frozen cells. REAGENTS AND SOLUTIONS Glucose/Tris/EDTA (GTE) solution 50 mM glucose 25 mM Tris⋅Cl, pH 8.0 10 mM EDTA Autoclave and store at 4°C NaOH/SDS solution 0.2 N NaOH 1% (wt/vol) sodium dodecyl sulfate (SDS) Prepare immediately before use 5 M potassium acetate solution, pH 4.8 29.5 ml glacial acetic acid KOH pellets to pH 4.8 (several) H2O to 100 ml Store at room temperature (do not autoclave) STET solution 8% (wt/vol) sucrose 5% (wt/vol) Triton X-100 50 mM EDTA 50 mM Tris⋅Cl, pH 8.0 Filter sterilize and store at 4°C TELT solution 2.5 M LiCl 50 mM Tris⋅Cl, pH 8.0 62.5 mM Na2 EDTA 4% (wt/vol) Triton X-100 Store as 1- to 5-ml aliquots frozen at −20°C (do not filter sterilize or autoclave)



Supplement 24

COMMENTARY Background Information Isolation of small quantities of plasmid DNA from bacterial cells is essential for the analysis of recombinant clones (UNITS 3.2 & 6.3). A myriad of plasmid DNA miniprep methods now exist and investigators are generally remarkably loyal to his or her own particular protocol. This unit presents three of the most widely used and reliable methods—alkaline lysis, the boiling method, and a lithium-based miniprep. The success of any of these procedures is largely a function of the expertise of the investigator; choice of method is therefore determined by personal preference as well as the size and type of the plasmid and the host strain of E. coli. With practice, all three protocols yield plasmid DNA of sufficient quantity and quality for use in most enzymatic manipulations (Chapters 3 & 7), and in most bacterial (UNIT 1.8) and yeast (UNIT 13.7) transformation procedures. The procedures presented in this unit allow for the preferential recovery of circular plasmid DNA over linear chromosomal DNA. Treatments with either base or detergent (used in all three procedures) disrupt base pairing and cause the linear chromosomal DNA to denature and separate. In contrast, because of its supercoiled configuration, covalently closed circular plasmid DNA is unable to separate and readily reforms a correctly paired superhelical structure under renaturing conditions. In the alkaline lysis miniprep, treatment with SDS and NaOH breaks open bacterial cells. Subsequent addition of potassium acetate preferentially reanneals covalently closed plasmid DNA, while chromosomal DNA and proteins are trapped in a complex formed between the potassium and SDS. The lysis treatment in the boiling miniprep causes chromosomal DNA to remain attached to the bacterial membrane, while plasmid DNA remains in the supernatant. In the lithium method, treatment of bacterial cells with Triton X-100/LiCl results in dissolution of the inner bacterial plasma membrane. However, this treatment has no effect on the overall morphology of the cells as observed microscopically, nor does it lead to release of plasmid DNA from the cells. Subsequent addition of phenol/chloroform leads to denaturation and precipitation of intracellular proteins. The concomitant rapid shrinkage of the cells preferentially expels soluble, super-

coiled plasmid DNA into the medium while retaining chromosomal DNA and denatured cellular protein with the bulk of the cell mass. Bacterial morphology, particularly of the cell wall envelope, is preserved under these conditions for at least 30 min, at which point cell lysis ensues. In all preparations, chromosomal DNA is removed with cellular debris by centrifugation, and soluble, supercoiled plasmid DNA is concentrated by ethanol precipitation. In general, all three methods provide plasmid DNA of comparable yield and quality suitable for most biological applications. Yield of DNA is determined more by the type of plasmid than by method of isolation. Plasmids derived from pBR322 generally give lower (but, for most applications, sufficient) yields compared with the more recently derived pUC-like vectors, which contain a lesion in the plasmid-encoded rop gene, causing the plasmid to be maintained in high copy number in the cell (see UNIT 1.5). All three methods are successful for the isolation of small (10 µg/ml. This is not a concern when applied to plasmid DNA purification, where the concentrations should be orders of magnitude greater than that figure. Plasmid DNA purification by anion-exchange or size-exclusion chromatography. The columns available in kit form from a number of manufacturers are quite reliable. Almost all necessary reagents are provided, including common buffers such as TE buffer. Because the composition of the matrix is undisclosed, it is impossible to evaluate the procedure carefully and attempt to optimize it. Therefore, users of kits are strongly encouraged to follow the manufacturer’s recommended procedures. Additional discussion of one type of anion-exchange procedure (Qiagen) can be found in the Commentary of UNIT 2.1B. A major drawback to some prepared columns is the limited capacity of the matrix. A 500-ml culture of plasmid-containing bacteria can often yield 2 to 8 mg of plasmid DNA in the crude lysate. Some columns routinely yield only 500 to 1000 µg purified plasmid DNA, which is adequate for most purposes. Recovering the column flowthrough and rechromatographing it is the most practical method of increasing recovery of plasmid DNA. Not all columns can be reused, however, and recovery may require use of additional columns. Alter-

natively, smaller culture volumes can be used as suggested. If optimal recovery of DNA is desired, CsCl/ethidium bromide centrifugation or PEG precipitation should be used. DNA obtained from chromatographic purification of plasmid DNA is comparable in quality to that prepared by the other methods. It is of sufficient purity for virtually any procedure for which it can be used. For example, plasmid DNA prepared using Qiagen-tips is at least equivalent in purity to that obtained following two rounds of CsCl-gradient centrifugation. It is suitable for all applications from cloning to transfection, radioactive and fluorescent sequencing, and gene therapy research. Qiagen also offers an endotoxin-free plasmid DNA kit which yields plasmid DNA containing lower levels of endotoxins than DNA purified by two rounds of CsCl centrifugation, which is essential in gene therapy studies. The two most frequent contaminants are chromosomal DNA and high-molecularweight RNA. These contaminants may be detected by the presence of large, diffuse ethidium bromide–binding material in agarose gel electrophoresis of purified plasmid DNA. To prevent such contamination, follow the manufacturer’s suggestions for the use of RNase. Only common laboratory equipment is required for chromatographic purification of plasmid DNA. Another consideration with commercial kits is the large amount of packaging material and waste. In addition to the plastic columns and excess packaging, kits contain standard reagents supplied in plastic bottles. These reagents are solutions of buffers, salts, ethanol, and detergent—all of which can be, and usually are, prepared in the lab. Most suppliers will provide the column without the reagents. However, the major advantage of using prepared anion-exchange column kits includes the absence of organic extractions and exposure to toxic chemicals such as phenol, chloroform, ethidium bromide, and CsCl. The toxicity of DNA prepared by several methods has been assessed by performing a biological assay. Crude lysate from a 1-liter culture of plasmid-containing bacteria was prepared by the alkaline lysis procedure and divided into four equal aliquots. The aliquots were then subjected to purification by CsCl/ ethidium bromide centrifugation, PEG precipitation, or chromatography on Qiagen and pZ523 columns. Purified plasmid DNA was injected into Drosophila embryos, and the frequency of germline transformation and killing



of injected embryos was determined. No significant differences attributable to the method of purification were observed.

Anticipated Results Most plasmids currently used are derivatives of the pUC series (Fig. 1.5.2). These plasmids contain an origin of replication significantly more efficient than that of the previous generation of pBR322-derived plasmids. This allows recovery of 1 to 5 mg of plasmid DNA (free of contaminating bacterial products) from a 500ml culture following any of the crude lysate preparation methods or PEG precipitation. Purification by CsCl/ethidium bromide density gradient centrifugation yields 75% to 90% of the amount of plasmid DNA obtained using PEG precipitation. Yields obtained from column chromatography are limited by the capacity of the column and are generally 1 ìg of DNA in the ligation reaction, or if the ligation reaction is from low gelling/melting temperature agarose, it is wise to dilute the ligation mix (see UNIT 3.16).

12. Rapidly thaw competent cells by warming between hands and dispense 100 µl immediately into test tubes containing DNA. Gently swirl tubes to mix, then place on ice for 10 min. Introduction of Plasmid DNA into Cells

Competent cells should be used immediately after thawing. Remaining cells should be discarded rather than refrozen.

1.8.2 Supplement 37


13. Heat shock cells by placing tubes into a 42°C water bath for 2 min. Alternatively, incubate at 37°C for 5 min.

14. Add 1 ml LB medium to each tube. Place each tube on a roller drum at 250 rpm for 1 hr at 37°C. 15. Plate aliquots of transformation culture on LB/ampicillin or other appropriate antibiotic-containing plates. It is convenient to plate several different dilutions of each transformation culture. The remainder of the mixture can be stored at 4°C for subsequent platings.

16. When plates are dry, incubate 12 to 16 hr at 37°C. ONE-STEP PREPARATION AND TRANSFORMATION OF COMPETENT CELLS

ALTERNATE PROTOCOL 1

This procedure is considerably easier than Basic Protocol 1 because it eliminates the need for centrifugation, washing, heat shock, and long incubation periods (Chung et al., 1989). Moreover, competent cells made by this simple procedure can be directly frozen at −70°C for long-term storage. A variety of strains can be made competent by this procedure, and the transformation frequency can be as high as that achieved by Basic Protocol 1. However, frequency is considerably lower than can be obtained by electroporation. Additional Materials (also see Basic Protocol 1) 2× transformation and storage solution (TSS; see recipe), ice cold LB medium (UNIT 1.1) containing 20 mM glucose 1. Dilute a fresh overnight culture of bacteria 1:100 into LB medium and incubate at 37°C until the cells reach an OD600 of 0.3 to 0.4. The procedure will work if cells are harvested at other stages of the growth cycle (including stationary phase), but with reduced efficiency.

2. Add a volume of ice-cold 2× TSS equal to that of the cell suspension, and gently mix on ice. For long-term storage, freeze small aliquots of the suspension in a dry ice/ethanol bath and store at −70°C. To use frozen cells for transformation, thaw slowly and then use immediately. Cells can also be used if pelleted by centrifugation 10 min at 1000 × g, 4°C, and this may increase the frequency of transformation (according to Chung et al., 1989). Discard supernatant and resuspend cell pellet at one-tenth of original volume in 1× TSS (prepared by diluting 2× TSS). Proceed with transformation as in step 3.

3. Add 100 µl competent cells and 1 to 5 µl DNA (0.1 to 100 ng) to an ice-cold polypropylene or glass tube. Incubate 5 to 60 min at 4°C. As is the case for related procedures, the transformation frequency as measured by transformants/ìg DNA is relatively constant at amounts of DNA 3-fold.

Transform the cells 10. Set the electroporation apparatus to 2.5 kV, 25 µF. Set the pulse controller to 200 or 400 ohms. The pulse controller is necessary when high-voltage pulses are applied over short gaps in high-resistance samples (see Background Information).

11. Add 5 pg to 0.5 µg plasmid DNA in 1 µl to tubes containing fresh or thawed cells (on ice). Mix by tapping the tube or by swirling the cells with the pipettor. 12. Transfer the DNA and cells into a cuvette that has been chilled 5 min on ice, shake slightly to settle the cells to the bottom, and wipe the ice and water from the cuvette with a Kimwipe. The volume of DNA added to the cells should be kept small. Adding DNA up to one-tenth of the cell volume will decrease the transformation efficiency 2- to 3-fold. Also, since the resistance of the sample should be high, make sure that addition of the DNA to the cells does not increase the total salt concentration in the cuvette by >1 mM.

13. Place the cuvette into the sample chamber. If using a homemade apparatus, connect the electrodes to the cuvette.

14. Apply the pulse by pushing the button or flipping the switch. 15. Remove the cuvette. Immediately add 1 ml SOC medium and transfer to a sterile culture tube with a Pasteur pipet. Incubate 30 to 60 min with moderate shaking at 37°C. If the actual voltage and time constant of the pulse are displayed on the electroporation apparatus, check this information. Verify that the set voltage was actually delivered, and record the time constant of the pulse so that you may vary it later if necessary (see Critical Parameters).

16. Plate aliquots of the transformation culture on LB plates containing antibiotics.



Supplement 37


DIRECT ELECTROPORETIC TRANSFER OF PLASMID DNA FROM YEAST INTO E. COLI The use of “shuttle vectors”—plasmids that can be grown successfully in at least two different organisms—facilitates the transfer of DNA between, for example, yeast and E. coli. In this adaptation of the electroporation protocol, plasmid DNA from a shuttle vector is transformed directly from yeast into E. coli. Components of the interaction trap/twohybrid system (UNIT 20.1) are used as an example in this protocol. The transfer and selection of a “prey” plasmid from the yeast strain EGY48 into the E. coli strain KC8 is described here, but the approach can be adapted for use with other yeast and E. coli strains. Additional Materials (also see Basic Protocol 2) Single colony of E. coli KC8 cells (UNIT 20.1) Streak colony of Trp− plasmid–harboring EGY48 yeast cells on Gal/Raff/Xgal/CM plates (UNIT 20.1), no older than 2 weeks M9 minimal medium and plates (UNIT 1.1) containing 100 µg/ml ampicillin (Table 1.4.1) and standard concentrations of leucine, histidine, and uracil Additional reagents and equipment for growth and manipulation of yeast (UNIT 13.2) and for plasmid DNA miniprep (UNIT 1.6) or PCR (UNIT 15.1) 1. Prepare electrocompetent KC8 cells (see Basic Protocol 2, steps 1 to 9a), resuspending the final cell pellet in ice-cold water to obtain an OD600 of 100. Fresh KC8 cells work better in this electroporation method than frozen ones. To measure OD, dilute 5 ìl of the cell suspension with water to 1 ml and measure the OD600. If necessary add more water to the suspension to get an OD600 of 100.

2. Distribute 65-µl aliquots of the electrocompetent E. coli KC8 cells into ice-cold microcentrifuge tubes. 3. With a sterile wooden or plastic stick, scrape off ∼10 µl of yeast from a streak colony of EGY48 harboring the respective “prey” plasmid derivative of pJG4-5 and grown on Gal/Raff/Xgal/CM plates. Resuspend the yeast cells in the KC8 suspension by swirling the stick used for scraping off the cells. Avoid scraping off plate medium when collecting the yeast streak cells. Keep the microcentrifuge tube on ice as much as possible. Try to get an even distribution of the two cell types but do not vortex. Yeast grown on plates other than Gal/Raff/Xgal/CM will probably work as well; do not worry if the yeast colony used is blue.

4. Set the electroporation apparatus to 1.5 kV, 25 µF, and the pulse controller to 100 ohms. Transfer the cell suspension into a 0.2-cm cuvette that has been chilled 5 min on ice, shake slightly to settle the cells to the bottom, and wipe the ice and water from the cuvette with a Kimwipe. The use of Pasteur pipettes will facilitate placing the cell suspension at the bottom of the cuvette. Avoid any air bubbles.

5. Place the cuvette in the sample chamber of the apparatus and pulse. Take the cuvette out and place it on ice for ≥45 sec. Meanwhile, change the settings in preparation for the second pulse. The expected time constant for the first pulse is 2.2 to 2.4 msec.

6. Set the electroporation apparatus to 2.5 kV, 25 µF, and the pulse controller to 200 ohms. Wipe the cuvette again, place it in the sample chamber, and pulse. Introduction of Plasmid DNA into Cells

The expected time constant for the second pulse is 4.2 to 4.8 msec.

1.8.6 Supplement 37


7. Remove the cuvette, immediately add 1 ml LB medium, and transfer the suspension into a microcentrifuge tube. Incubate 45 min at room temperature. Incubation of the suspension after electroporation for 1 hr at 37°C decreases the yield of transformants, probably due to prolonged adhesion of the E. coli cells to the yeast cell debris.

8. Spread 150 µl of the suspension evenly onto M9 minimal medium plates containing 100 µg/ml ampicillin and leucine, histidine, and uracil. Incubate ≥24 hr at 37°C. A slight yeast background might appear on the plates but single E. coli colonies are easy to pick. Between 50 and 200 KC8 colonies have been obtained per plate employing 150 ìl out of the 1 ml LB suspension.

9. Pick a single KC8 colony, inoculate it into 1.5 to 5 ml M9 minimal medium (Leu+, His+, Ura+, 100 µg/ml Amp) or LB (100 µg/ml Amp) and grow at 37°C. Harvest at an appropiate OD to prepare miniprep DNA (UNIT 1.6) or perform PCR analysis (UNIT 15.1). Using M9 minimal medium to grow KC8 in liquid culture is an additional safety measure but not absolutely necessary. It ensures isolation of the plasmid whose marker complements the auxotrophic defect in KC8; in addition, slightly increased plasmid copy number and improved DNA quality have been reported.

REAGENTS AND SOLUTIONS CaCl2 solution 60 mM CaCl2 15% glycerol 10 mM PIPES [piperazine-N,N′-bis(2-hydroxypropanesulfonic acid)], pH 7 Filter sterilize using a disposable filter unit, or autoclave Store at room temperature (stable for years) SOC medium 0.5% yeast extract 2% tryptone 10 mM NaCl 2.5 mM KCl 10 mM MgCl2 10 mM MgSO4 20 mM glucose Store at room temperature (stable for years) Transformation and storage solution (TSS), 2× Dilute sterile (autoclaved) 40% (w/v) polyethylene glycol (PEG) 3350 to 20% PEG in sterile LB medium containing 100 mM MgCl2. Add dimethyl sulfoxide (DMSO) to 10% (v/v) and adjust to pH 6.5. COMMENTARY Background Information Calcium and one-step transformation Transformation of E. coli was first described by Mandel and Higa (1970). Subsequent modifications to improve transformation efficiencies have included prolonged exposure of cells to CaCl2 (Dagert and Ehrlich, 1974), substitution of calcium with other cations such as Rb+

(Kushner, 1978), Mn2+, and K+, and addition of other compounds such as dimethyl sulfoxide, dithiothreitol, and cobalt hexammine chloride (Hanahan, 1983). Basic Protocol 1 given here provides good transformation efficiencies, permits long-term storage of competent cells, and is relatively uncomplicated to perform. Variations on this protocol can be obtained from the references provided. Alternate Proto-



Supplement 37

col 1, a one-step preparation and transformation procedure, is considerably faster. Transformation by electroporation Electroporation has become a valuable technique for transfer of nucleic acids into eukaryotic and prokaryotic cells. The method can be applied to many different E. coli strains and to other gram-negative and gram-positive bacteria. In this technique, a high-voltage electric field is applied briefly to cells, apparently producing transient holes in the cell membrane through which plasmid DNA enters (Shigekawa and Dower, 1988). The field strength used for mammalian cell and plant protoplast electroporation is usually 0.5 to 1 kV/cm. The field strength needed for high-efficiency transformation of E. coli is much greater, usually ∼12.5 kV/cm. Under these conditions up to 1010 transformants/µg plasmid DNA have been reported (Calvin and Hanawalt, 1988; Miller et al., 1988; Dower et al., 1988). Recently, field strength up to 8 kV/cm was also used successfully to electroporate both mammalian cells and plant protoplasts. The capacitor discharge circuit of the electroporation apparatus typically generates an electrical pulse with an exponential decay waveform. The voltage across the electrodes rises rapidly to a peak voltage, which then declines over time as follows: −t/T

Vt = V0 [e

Introduction of Plasmid DNA into Cells

]

where Vt = voltage at a given time t after the time of V0, V0 = initial voltage, t = time (sec), T = pulse time constant = RC, R = resistance of circuit (ohms); and C = capacitance of circuit (Farads). The pulse time constant is ∼5 to 10 msec for electroporating E. coli cells and ranges from 5 µsec to 50 msec for higher eukaryotic cells. The pulse controller contains a number of different-sized resistors, any one of which is placed in parallel with the sample, and one resistor of fixed (20-ohm) resistance, which is placed in series with the sample. The resistor placed in parallel with the sample (usually 200 or 400 ohms) swamps out the effect of changes in the resistance of the sample on the total resistance of the circuit, thus determining the total resistance across the capacitor, and controlling the time constant (T) of the capacitor discharge. The 20-ohm resistor in series with the sample protects the circuitry by limiting the current should a short circuit (arc) occur and the capacitor discharge instantly.

The pulse controller is required when highvoltage electroporation pulses are delivered to high-resistance samples across narrow electrode gaps. In this procedure, the resistance of washed E. coli in the 0.2-cm cuvette is ∼5000 ohms. A pulse controller is not necessary when samples of low (500 to 1000 bp (UNITS 2.5A & 2.6) and smaller-pore acrylamide or sieving agarose gels (UNIT 2.7) are used for fragments 120 kb) genomic libraries. Two main precautions should be taken to maximize molecular weight: (1) minimize shearing forces by gentle (but thorough) mixing during extraction steps, and (2) after the extraction, remove organic solvents and salt from the DNA by dialysis, rather than by ethanol precipitation. Additional precautions must be taken to prepare very high-molecular-weight DNA for the construction of P1 or BAC libraries. The absence of both cellular proteins and proteinase K in the final DNA solution is important for susceptibility of the genomic DNA to restriction enzyme action; therefore, care should be exercised in deproteination. To remove protein completely it may be necessary to repeat the proteinase K digestion. In general, highly pure DNA has an A260/A280 ratio >1.8, while 50% protein/50% DNA mixtures have A260/A280 ratios of ∼1.5.

Approximately 2 mg DNA should be obtained from 1 g tissue or 109 cells. The DNA should be at least 100 kb long and should be digestible with restriction enzymes.

Time Considerations This protocol involves effort on 2 days: tissue preparation on the first day followed by overnight lysis, and extraction/precipitation on the second day. Actual time spent on the procedure, however, will be less than 1 hr each day. The DNA can be stored indefinitely in the presence of ethanol at 4° or in TE buffer at −20°C.

Literature Cited Enrietto, P.J., Payne, L.N., and Hayman, M.J. 1983. A recovered avian myelocytomatosis virus that induces lymphomas in chickens. Pathogenic properties and their molecular basis. Cell 35:369-379. Gross-Bellard, M., Oudet, P., and Chambon, P. 1973. Isolation of high-molecular-weight DNA from mammalian cells. Eur. J. Biochem. 36:3238.

Contributed by William M. Strauss Harvard Medical School and Beth Israel Deaconess Medical Center Boston, Massachusetts

Preparation and Analysis of DNA


Supplement 42

Preparation of Genomic DNA from Plant Tissue

UNIT 2.3

This unit describes two methods for preparing genomic DNA from plant tissue. In the first method, plant cells are lysed with ionic detergent, treated with protease, and subsequently purified by cesium chloride (CsCl) density gradient centrifugation. The second method is based upon a series of treatments with the nonionic detergent cetyltrimethylammonium bromide (CTAB) to lyse cells and purify nucleic acid. Nucleic acid is recovered from the final CTAB solution by isopropanol or ethanol precipitation. The first method, although somewhat more lengthy, results in highly purified nucleic acid. The second method requires fewer manipulations, results in very high yields (∼10-fold higher per gram fresh tissue depending on species and condition of starting material), and produces DNA that is less pure but nonetheless suitable in quality for use in many molecular biology manipulations. PREPARATION OF PLANT DNA USING CSCL CENTRIFUGATION Plant cells are lysed by the detergent N-lauroylsarcosine (Sarkosyl), and the lysate is digested with proteinase K. After clearing insoluble debris from the lysate, nucleic acids are precipitated and DNA is purified on a cesium chloride (CsCl) density gradient.

BASIC PROTOCOL

Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Plant tissue, fresh Liquid nitrogen Extraction buffer (see recipe) 10% (w/v) N-lauroylsarcosine (Sarkosyl) Isopropanol TE buffer, pH 8.0 Cesium chloride 10 mg/ml ethidium bromide CsCl-saturated isopropanol (equilibrate over a CsCl-saturated aqueous phase) Ethanol 3 M sodium acetate, pH 5.2 250-ml centrifuge bottle 55°C water bath Beckman JA-14, JA-20 or JA-21, and VTi80 rotors (or equivalents) 5-ml quick-seal ultracentrifuge tubes 15-G needle and 1-ml syringe Prepare plant tissue 1. Harvest 10 to 50 g fresh plant tissue. Plants may be placed in the dark for 1 to 2 days prior to harvest to reduce the starch content in the tissues. Younger plants are the preferred source of tissue because they have a lower polysaccharide content.

2. Rinse tissue with deionized water to remove adhering debris and blot dry. 3. Freeze tissue with liquid nitrogen and grind to a fine powder in a mortar and pestle. Keep the tissue frozen throughout this procedure by occasionally adding liquid nitrogen. Contributed by Eric Richards, Mark Reichardt, and Sharon Rogers Current Protocols in Molecular Biology (1994) 2.3.1-2.3.7 Copyright © 2000 by John Wiley & Sons, Inc.


2.3.1 Supplement 27

Lyse and digest cells 4. Transfer frozen powder to a 250-ml centrifuge bottle and immediately add 5 to 10 ml extraction buffer per gram fresh plant tissue. Stir gently to disperse tissue. 5. Add 10% N-lauroylsarcosine to a final concentration of 1%. Incubate 1 to 2 hr at 55°C. It is important to add N-lauroylsarcosine after the tissue is resuspended in extraction buffer. If N-lauroylsarcosine is included in extraction buffer, premature lysis of the plant cells will interfere with tissue dispersal and lead to unwanted shearing of DNA. The lysate should be clear, green, and slightly viscous. From this point on solutions should be handled gently to reduce shearing of the DNA—use a wide-bore pipet and do not vortex or mix vigorously.

6. Centrifuge lysate 10 min at 5500 × g (6000 rpm in a Beckman JA-14 rotor), 4°C, to pellet debris. Save the supernatant and centrifuge again if necessary to remove undigested debris. Precipite the DNA 7. Add 0.6 vol isopropanol to the supernatant and gently mix. A nucleic acid precipitate should be visible; if not, incubate 30 min at −20°C. 8. Centrifuge 15 min at 7500 × g (8000 rpm in a Beckman JA-14 rotor), 4°C. Discard supernatant. Do not let the nucleic acid pellet dry or it will become extremely difficult to dissolve.

Carry out CsCl centrifugation 9. Resuspend pellet in 9 ml TE buffer. If necessary, incubate at 55°C to aid resuspension. Add 9.7 g of solid CsCl and mix gently until dissolved. To minimize depurination, limit 55°C incubation to ≤2 hr.

10. Incubate 30 min on ice. Centrifuge 10 min at 7500 × g (8000 rpm in a JA-20 rotor), 4°C, and save supernatant. This clearing spin removes some of the insoluble debris remaining in the lysate. In addition, a small separate phase may form on the top of the solution after centrifugation; this is due to residual Sarkosyl in the lysate. The Sarkosyl phase can be removed by filtering the supernatant through two layers of cheesecloth. Collect the supernatant but discard the Sarkosyl phase.

11. Add 0.5 ml of 10 mg/ml ethidium bromide and incubate 30 min on ice. CAUTION: Ethidium bromide is a mutagen. Be careful and wear gloves.

12. Centrifuge 10 min at 7500 × g, 4°C. A large RNA pellet should form. At this point much of the unwanted constituents in the lysate—RNA, protein, and carbohydrates—have been removed.

13. Transfer the supernatant to two 5-ml quick-seal ultracentrifuge tubes and seal tubes. Make sure tubes are full, balanced, and well-sealed.

14. Centrifuge 4 hr at 525,000 × g (80,000 rpm in a Beckman VTi80 rotor), 20°C, or overnight at 300,000 × g (60,000 rpm in VTi80 rotor), 20°C. Preparation of Genomic DNA from Plant Tissue

2.3.2 Supplement 27


Collect and purify DNA 15. Gently remove the tube. Punch a hole in the top (to provide an air inlet) with a large-bore (15-G) collecting needle. Recover the DNA band by inserting needle, attached to a 1-ml syringe, through tube wall directly below the band (see Fig. 1.7.1). This operation is identical to that used during plasmid purification, except that only one band should be visible. CAUTION: If UV illumination is used to visualize the DNA, wear UV protective glasses or a face shield. Minimize exposure of gradient to visible light to reducing nicking of DNA caused by ethidium bromide.

16. Remove the ethidium bromide by repeatedly extracting the collected DNA with CsCl-saturated isopropanol. 17. Add 2 vol water and 6 vol ethanol to the DNA solution and mix. Incubate 1 hr at −20°C. DNA may precipitate immediately as a single white mass; it can be collected using a Pasteur pipet with a hook introduced at the tip or by brief centrifugation.

18. Centrifuge 10 min at 7500 × g, 4°C. 19. Resuspend pellet in TE buffer and reprecipitate DNA by adding 1⁄10 vol of 3 M sodium acetate and 2 vol ethanol. Incubate at −20°C if precipitate is not visible and collect DNA by centrifugation. 20. Briefly air dry the final pellet and resuspend in 0.5 to 2 ml TE buffer. A DNA concentration of 100 ng/ìl is generally convenient for most purposes.

PREPARATION OF PLANT DNA USING CTAB Alternatively, the nonionic detergent cetyltrimethylammonium bromide (CTAB) is used to liberate and complex with total cellular nucleic acids. This general procedure has been used on a wide array of plant genera and tissue types. Many modifications have been published to optimize yields from particular species. The protocol is relatively simple, fast, and easily scaled from milligram to grams of tissue; it requires no cesium chloride density gradient centrifugation.

ALTERNATE PROTOCOL

Additional Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

CTAB extraction solution (see recipe) CTAB/NaCl solution (UNIT 2.4) CTAB precipitation solution (see recipe) 2% (v/v) 2-mercaptoethanol (2-ME) High-salt TE buffer (see recipe) 24:1 (v/v) chloroform/octanol or chloroform/isoamyl alcohol 80% ethanol Pulverizer/homogenizer: mortar and pestle, blender, Polytron (Brinkmann), or coffee grinder Organic solvent–resistant test tube or beaker 65°C water bath Beckman JA-20 rotor or equivalent or microcentrifuge Preparation and Analysis of DNA


Supplement 27

Extract nucleic acids 1. Add 2-ME to the required amount of CTAB extraction solution to give a final concentration of 2% (v/v). Heat this solution and CTAB/NaCl solution to 65°C. Approximately 4 ml of 2-ME/CTAB extraction solution and 0.4 to 0.5 ml CTAB/NaCl solution are required for each gram of fresh leaf tissue. With lyophilized, dehydrated, or dry tissues such as seeds, 2-ME/CTAB extraction solution should be diluted 1:1 with sterile water. 2-ME should be used in a fume hood.

2. Chill a pulverizer/homogenizer with liquid nitrogen (−196°C) or dry ice (−78°C). Pulverize plant tissue to a fine powder and transfer the frozen tissue to an organic solvent–resistant test tube or beaker. Use young tissue and avoid larger stems and veins to achieve the highest DNA yield with minimal polysaccharide contamination.

3. Add warm 2-ME/CTAB extraction solution to the pulverized tissue and mix to wet thoroughly. Incubate 10 to 60 min at 65°C with occasional mixing. A 60-min incubation results in larger DNA yields. If maximum yield is not important, 10 min should be adequate. If the tissue contains large amounts of phenolic compounds, 1% (v/v) polyvinylpyrrolidone (mol. wt. = 40,000) may be added to absorb them.

4. Extract the homogenate with an equal volume of 24:1 chloroform/octanol or chloroform/isoamyl alcohol. Mix well by inversion. Centrifuge 5 min at 7500 × g (8000 rpm in JA-20; ∼10,000 rpm in a microcentrifuge, for smaller samples), 4°C. Recover the top (aqueous) phase. Octanol, rather than isoamyl alcohol, is used because it may enhance isolation of nuclei. Slower centrifugation speeds are possible if centrifugation time is increased accordingly; a microcentrifuge may be used for small-scale preparations (≤150 mg starting tissue). After centrifugation, two phases should be evident with tissue debris at the interface.

5. Add 1⁄10 vol 65°C CTAB/NaCl solution to the recovered aqueous phase and mix well by inversion. 6. Extract with an equal volume of chloroform/octanol. Mix, centrifuge, and recover as in step 4 above. The aqueous phase may still be light yellow-brown in color.

Precipitate nucleic acids 7. Add exactly 1 vol CTAB precipitation solution. Mix well by inversion. If precipitate is visible, proceed to step 8. If not, incubate mixture 30 min at 65°C. 8. Centrifuge 5 min at 500 × g (2000 rpm in JA-20; ∼2700 rpm in microcentrifuge), 4°C. Do not increase the speed or time of centrifugation as the pellet may become very difficult to resuspend. If there is no pellet, add more CTAB precipitation solution (up to 1⁄10 the total volume). Incubate 1 hr to overnight at 37°C. Centrifuge 5 min at 500 × g, 4°C.

9. Remove but do not discard the supernatant and resuspend pellet in high-salt TE buffer (0.5 to 1 ml per gram of starting material). If the pellet is difficult to resuspend, incubate 30 min at 65°C. Repeat until all or most of pellet is dissolved. Polysaccharide contamination may make it excessively difficult to resuspend the pellet. Read the A260 of the supernatant and discard pellet if nucleic acids are present in the supernatant. Preparation of Genomic DNA from Plant Tissue

2.3.4 Supplement 27


10. Precipitate the nucleic acids by adding 0.6 vol isopropanol. Mix well and centrifuge 15 min at 7500 × g, 4°C. Ethanol can be used for the precipitation, but isopropanol may yield cleaner pellets.

11. Wash the pellet with 80% ethanol, dry, and resuspend in a minimal volume of TE (0.1 to 0.5 ml per gram of starting material). Residual CTAB is soluble and is removed by the 80% ethanol wash. Further purification of the DNA with RNase A and proteinase K may be done using standard methods (UNITS 2.4 & 3.13).

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

CTAB extraction solution 2% (w/v) CTAB 100 mM Tris⋅Cl, pH 8.0 20 mM EDTA, pH 8.0 1.4 M NaCl Store at room temperature (stable several years) CTAB precipitation solution 1% (w/v) CTAB 50 mM Tris⋅Cl, pH 8.0 10 mM EDTA, pH 8.0 Store at room temperature (stable several years) Extraction buffer 100 mM Tris⋅Cl, pH 8.0 100 mM EDTA, pH 8.0 250 mM NaCl 100 µg/ml proteinase K (add fresh before use) Store indefinitely at room temperature without proteinase K High-salt TE buffer 10 mM Tris⋅Cl, pH 8.0 0.1 mM EDTA, pH 8.0 1 M NaCl Store at room temperature (stable for several years) COMMENTARY Background Information CsCl gradient purification This protocol is an adaptation of common DNA isolation procedures—cell lysis by detergent, protease treatment, and CsCl gradient purification. Because whole cells are lysed, DNA purified using this protocol will correspond to both the nuclear genome and cytoplasmic (mitochondrial and chloroplast) genomes. Methods for purifying nuclear DNA—free of plastid and mitochondrial DNA contamination—have been described by Watson and Thompson (1986). In addition, a miniprep pro-

tocol for isolation of total plant DNA has been described by Dellaporta et al. (1983); the miniprep protocol is similar to the protocol described here, except that it omits the CsCl gradient centrifugation. CTAB purification The alternate protocol, cetyltrimethylammonium bromide (CTAB) DNA isolation, was initially used in bacteria (Jones, 1953; UNIT 2.4) and later modified to obtain DNA from plants (Murray and Thompson, 1980). CTAB forms an insoluble complex with nucleic acids when the initial NaCl concentration is lowered to



Supplement 45

∼0.5 M (Rogers and Bendich, 1985). Polysaccharides, phenolic compounds, and other enzyme-inhibiting contaminants found in plant cells are efficiently removed in the supernatant because most do not precipitate under the conditions described. The nucleic acid–CTAB complex is only soluble in high salt; detergent is removed by raising the NaCl concentration and precipitating the nucleic acids. The residual CTAB is removed by washing the nucleic acid pellet with 80% ethanol; CTAB is more soluble in ethanol and is discarded with the wash solution. The CTAB method of DNA isolation is widely used on plants because of its versatility. Total genomic DNA has been isolated from many genera of monocotyledons and dicotyledons (Murray and Thompson, 1980; Rogers and Bendich, 1985). Various types of tissue can be used, including whole seedlings, leaves, cotyledons, seeds/grains, endosperm, embryos, tissue culture callus, and pollen. In addition, milligram amounts of tissue can be used when sample size is limiting. The starting material can be lyophilized, dehydrated (even mummified), frozen, or fresh. The authors’ (M.R. and S.R.) laboratory has successfully used this protocol to isolate DNA from Arabidopsis thaliana, Zea mays, Gossypium hirsutum, Flaveria sp., Linum usitatissimum, Petunia hybrida, Glycine sp., Nicotiana tabacum, and Lycopersicum esculentum. In addition, each year The Plant Molecular Biology Reporter publishes several modifications to this general protocol optimized for species from which nucleic acid extraction is difficult.

Critical Parameters

Preparation of Genomic DNA from Plant Tissue

The aim of any genomic DNA preparation technique is to isolate high-molecular-weight DNA of sufficient purity. Two factors affect the size of the DNA isolated: shear forces and nuclease activity. As noted in the protocols, lysates should be treated gently to minimize shear forces. Plant cells are rich in nucleases. To reduce nuclease activity, the tissue should be frozen quickly and thawed only in the presence of an extraction buffer that contains detergent and a high concentration of EDTA. Plant DNA isolated using the basic protocol should be in the range of 50 kb in length, which is quite acceptable for most applications. Arabidopsis DNA isolated using this protocol can be digested with restriction enzymes and ligated efficiently into cloning vectors. However, in some cases it may be necessary to modify the steps in order to reduce contamination by poly-

saccharides, phenolics, and other compounds that interfere with DNA isolation. Polysaccharides pose the most common problem affecting plant DNA purity. These carbohydrates are difficult to separate from the DNA itself, and they inhibit many enzymes commonly used in cloning procedures. The recommended procedure for polysaccharide removal is chloroform extraction of lysates in the presence of 1% CTAB and 0.7 M NaCl, as described by Murray and Thompson (1980) and in the alternate protocol. If the DNA pellet obtained using the alternate protocol is excessively difficult to resuspend, it may be due to the presence of polysaccharides that were not removed during CTAB precipitation. The DNA should be soluble in TE buffer; passage of the solution over an anion-exchange column should remove much of the contamination (Fang et al., 1992; UNIT 2.1B). If phenolic compounds are a problem, 1% polyvinylpyrollidone (mol. wt. = 40,000; Sigma) can be included during tissue homogenization to absorb them. Chloroform/octanol is preferred over chloroform/isoamyl alcohol for organic extractions because it has been reported to isolate n uclei mo re efficiently (Watson and Thompson, 1986). Finally, RNase A and proteinase K digestion followed by phenol extraction can be performed if further purification is required.

Anticipated Results CsCl gradient purification. Yields should be in the range of 10 to 40 µg DNA (50-kb length) per gram of fresh plant tissue. Isolated DNA should digest well with restriction enzymes and be ligated efficiently into cloning vectors. CTAB purification. Yields should be in the range of 100 to 500 µg DNA per gram of fresh plant tissue. The greatest yields will always be obtained using the youngest, freshest tissue available. DNA ≥50 kb can be obtained if care is taken not to shear it (by using wide bore pipettes and gentle mixing) and if nucleases are avoided (by keeping tissue frozen or lyophilized and thawing or rehydrating only in the presence of CTAB extraction solution). In addition to DNA, RNA is also efficiently liberated and purified by this method and can be separated from the DNA if desired (e.g., see UNIT 4.3 and Taylor and Powell, 1982).

Time Considerations CsCl gradient purification. Approximately 4 to 6 hr are required to work through the protocol to the point where the lysate is loaded

2.3.6 Supplement 45


onto the CsCl gradient. The gradient can be centrifuged overnight at 300,000 × g or 4 hr at 525,000 × g. Approximately 3 to 4 hr are required to process the banded DNA. CTAB purification. This procedure should take between 2 and 6 hr depending on the quantity of starting material, desired purity, and yield.

Rogers, S.O. and Bendich, A.J. 1985. Extraction of DNA from milligram amounts of fresh, herbarium and mummified plant tissues. Plant Mol. Biol. 5:69-76. Taylor, B. and Powell, A. 1982. Isolation of plant DNA and RNA. BRL Focus 4(3):4-6. Watson, J.C. and Thompson, W.F. 1986. Purification and restriction endonuclease analysis of plant nuclear DNA. Methods Enzymol. 118:57-75.

Literature Cited Dellaporta, S.L., Wood, J., and Hicks, J.B. 1983. A plant DNA minipreparation: Version II. Plant Mol. Biol. Rep. 1:19-21. Fang, G., Hammar, S., and Grumet, R. 1992. A quick and inexpensive method for removing polysaccharides from plant genomic DNA. BioTechniques 13:52-56. Jones, A.S. 1953. The isolation of bacterial nucleic acids using cetyltrimethlyammonium bromide (cetavlon). Biochim. Biophys. Acta. 10:607-612.

Contributed by Eric Richards (CsCl preparation) Washington University St. Louis, Missouri Mark Reichardt and Sharon Rogers (CTAB preparation) Lakeside Biotechnology Chicago, Illinois

Murray, M.G. and Thompson, W.F. 1980. Rapid isolation of high-molecular-weight plant DNA. Nucl. Acids Res. 8:4321-4325.



Supplement 27

UNIT 2.4 BASIC PROTOCOL

Preparation of Genomic DNA from Bacteria MINIPREP OF BACTERIAL GENOMIC DNA Bacteria from a saturated liquid culture are lysed and proteins removed by digestion with proteinase K. Cell wall debris, polysaccharides, and remaining proteins are removed by selective precipitation with CTAB, and high-molecular-weight DNA is recovered from the resulting supernatant by isopropanol precipitation. Materials TE buffer (APPENDIX 2) 10% sodium dodecyl sulfate (SDS) 20 mg/ml proteinase K (stored in small single-use aliquots at −20°C) 5 M NaCl CTAB/NaCl solution 24:1 chloroform/isoamyl alcohol 25:24:1 phenol/chloroform/isoamyl alcohol (UNIT 2.1) Isopropanol 70% ethanol 1. Inoculate a 5-ml liquid culture with the bacterial strain of interest. Grow in conditions appropriate for that strain (i.e., appropriate medium, drug selection, temperature) until the culture is saturated. This may take several hours to several days, depending on the growth rate. 2. Spin 1.5 ml of the culture in a microcentrifuge for 2 min, or until a compact pellet forms. Discard the supernatant. 3. Resuspend pellet in 567 µl TE buffer by repeated pipetting. Add 30 µl of 10% SDS and 3 µl of 20 mg/ml proteinase K to give a final concentration of 100 µg/ml proteinase K in 0.5% SDS. Mix thoroughly and incubate 1 hr at 37°C. The solution should become viscous as the detergent lyses the bacterial cell walls. There should be no need to predigest the bacterial cell wall with lysozyme.

4. Add 100 µl of 5 M NaCl and mix thoroughly. This step is very important since a CTAB–nucleic acid precipitate will form if salt concentration drops below about 0.5 M at room temperature (Murray and Thompson, 1980). The aim here is to remove cell wall debris, denatured protein, and polysaccharides complexed to CTAB, while retaining the nucleic acids in solution.

5. Add 80 µl of CTAB/NaCl solution. Mix thoroughly and incubate 10 min at 65°C. 6. Add an approximately equal volume (0.7 to 0.8 ml) of chloroform/isoamyl alcohol, mix thoroughly, and spin 4 to 5 min in a microcentrifuge. This extraction removes CTAB–protein/polysaccharide complexes. A white interface should be visible after centrifugation.

7. Remove aqueous, viscous supernatant to a fresh microcentrifuge tube, leaving the interface behind. Add an equal volume of phenol/chloroform/isoamyl alcohol, extract thoroughly, and spin in a microcentrifuge for 5 min. With some bacterial strains the interface formed after chloroform extraction is not compact enough to allow easy removal of the supernatant. In such cases, most of the interface can be fished out with a sterile toothpick before removal of any supernatant. Remaining CTAB precipitate is then removed in the phenol/chloroform extraction. Preparation of Genomic DNA from Bacteria

2.4.1 Supplement 27

8. Transfer the supernatant to a fresh tube. Add 0.6 vol isopropanol to precipitate the nucleic acids (there is no need to add salt since the NaCl concentration is already Contributed by Kate Wilson Current Protocols in Molecular Biology (1997) 2.4.1-2.4.5 Copyright © 1997 by John Wiley & Sons, Inc.

high). Shake the tube back and forth until a stringy white DNA precipitate becomes clearly visible. At this point it is possible to transfer the pellet to a fresh tube containing 70% ethanol by hooking it onto the end of a micropipet that has been heat-sealed and bent in a Bunsen flame. Alternatively, the precipitate can be pelleted by spinning briefly at room temperature. If no stringy DNA precipitate forms in the above step, this implies that the DNA has sheared into relatively low-molecular-weight pieces. If this is acceptable, i.e., if DNA is to be digested to completion with restriction endonucleases for Southern blot analysis, chromosomal DNA can often still be recovered simply by pelleting the precipitate in a microcentrifuge.

9. Wash the DNA with 70% ethanol to remove residual CTAB and respin 5 min at room temperature to repellet it. Carefully remove the supernatant and briefly dry the pellet in a lyophilizer. 10. Redissolve the pellet in 100 µl TE buffer. This may take some time (up to 1 hr) since the DNA is of high molecular weight. 15 ìl of this DNA will typically digest to completion with 10 U EcoRI in 1 hr, which is sufficient to be clearly visible on an agarose gel, or to give a good signal during Southern hybridization.

REMOVAL OF POLYSACCHARIDES FROM EXISTING GENOMIC DNA PREPS

SUPPORT PROTOCOL

Steps 4 through 10 of the basic protocol can be adapted for removing polysaccharides and other contaminating macromolecules from existing bacterial chromosomal DNA preparations. Simply adjust the NaCl concentration of the DNA solution to 0.7 M and add 0.1 vol CTAB/NaCl solution. A white interface after the chloroform/isoamyl extraction indicates that contaminating macromolecules have been removed. The CTAB extraction step (steps 5 and 6) can be repeated several times until no interface is visible.

MINIPREP OF BACTERIAL GENOMIC DNA 1. Grow bacterial strain to saturation.

SHORT PROTOCOL

2. Spin 1.5 ml for 2 min in microcentrifuge. 3. Resuspend in 567 µl TE buffer, 3 µl of 10% SDS, 3 µl of 20 mg/ml proteinase K. Mix and incubate 1 hr at 37°C. 4. Add 100 µl of 5 M NaCl. Mix thoroughly. 5. Add 80 µl of CTAB/NaCl solution. Mix. Incubate 10 min at 65°C. 6. Extract with an equal volume of chloroform/isoamyl alcohol. Spin 5 min in microcentrifuge. 7. Transfer aqueous phase to a fresh tube. Extract DNA with phenol/chloroform/ isoamyl alcohol. Spin 5 min in microcentrifuge. 8. Transfer aqueous phase to a fresh tube. Extract DNA with 0.6 vol isopropanol. Wash precipitate with 70% ethanol. Remove supernatant and briefly dry pellet in lyophilizer. 9. Resuspend pellet in 100 µl TE buffer.



Supplement 56

ALTERNATE PROTOCOL

LARGE-SCALE CsCl PREP OF BACTERIAL GENOMIC DNA This procedure is essentially a scale-up of the chromosomal miniprep described in the basic protocol, followed by additional purification on a cesium chloride gradient. This procedure may be used if large amounts of exceptionally clean genomic DNA are required, e.g., for the construction of genomic libraries. Additional Materials Cesium chloride 10 mg/ml ethidium bromide CsCl-saturated isopropanol or H2O-saturated butanol 3 M sodium acetate, pH 5.2 Beckman JA-20 rotor or equivalent 50-ml Oak Ridge centrifuge tubes Wide-bored pipet 4-ml sealable centrifuge tubes Beckman VTi80 rotor 3-ml plastic syringe with 15-G needle Preparation and lysis of cells 1. Grow 100 ml culture of bacterial strain to saturation. 2. Pellet cells for 10 min at 4000 × g (e.g., in a Beckman JA-20 rotor at 6000 rpm). Discard supernatant. This, and the following steps, can be conveniently carried out using 50-ml Oak Ridge centrifuge tubes.

3. Resuspend cells gently in 9.5 ml TE buffer. Add 0.5 ml of 10% SDS and 50 µl of 20 mg/ml proteinase K. Mix thoroughly and incubate 1 hr at 37°C. Precipitation and purification of DNA 4. Add 1.8 ml of 5 M NaCl and mix thoroughly. 5. Add 1.5 ml CTAB/NaCl solution. Mix thoroughly and incubate 20 min at 65°C. 6. Add an equal volume of chloroform/isoamyl alcohol. Extract thoroughly. Spin 10 min at 6000 × g (JA-20 rotor at 7000 rpm), room temperature, to separate phases. 7. Transfer aqueous supernatant to a fresh tube using a wide-bored pipet. The supernatant will probably be very viscous if the yield is high. An additional chloroform/isoamyl alcohol extraction, or a phenol/chloroform/isoamyl alcohol extraction, is optional but should not be necessary if the material is to be purified on a cesium chloride gradient.

8. Add 0.6 vol isopropanol and mix until a stringy white DNA pellet precipitates out of solution and condenses into a tight mass. Transfer the precipitate to 1 ml of 70% ethanol in a fresh tube, by hooking it on the end of a Pasteur pipet that has been bent and sealed in a Bunsen flame. 9. Spin the pellet 5 min at 10,000 × g (JA-20 rotor at 9900 rpm). Remove supernatant and redissolve the pellet in 4 ml TE buffer. This may take several hours to overnight— the DNA can be placed at 60°C to hasten the process.

Preparation of Genomic DNA from Bacteria

10. Measure the DNA concentration on a spectrophotometer. Adjust concentration to 50 to 100 µg/ml. This will give 200 to 400 µg chromosomal DNA per 4 ml gradient. It is not advisable to spin larger quantities of chromosomal DNA on such a small gradient.

2.4.3 Supplement 56


11. Add 4.3 g CsCl per 4 ml TE buffer. Dissolve. Add 200 µl of 10 mg/ml ethidium bromide. Transfer to 4-ml sealable centrifuge tubes. Adjust volume and balance tubes with CsCl in TE buffer (1.05 g/ml). Seal tubes. Spin 4 hr in a Beckman VTi80 rotor at 70,000 rpm, 15°C, or overnight at 55,000 rpm, 15°C. 12. Visualize gradient under longwave UV lamp. A single band should be visible. Remove band using a 15-g needle and a 3-ml plastic syringe. If the DNA is intact high-molecular-weight chromosomal DNA it will appear very viscous as the band is withdrawn from the gradient; hence, it is important to use a wide-bore needle to avoid mechanical shearing of the DNA. If the band appears right at the top of the gradient, then the gradient is too dense. Reduce the amount of CsCl added in step 11.

13. Remove the ethidium bromide by sequential extractions with CsCl-saturated isopropanol or water-saturated butanol, as described in UNIT 1.7. 14. Dialyze overnight against 2 liters TE buffer to remove CsCl. 15. Transfer DNA solution to a fresh tube. If required, precipitate chromosomal DNA as described above (steps 8 and 9) by adding 1⁄10 vol of 3 M sodium acetate and 0.6 vol isopropanol, and resuspend at desired concentration.

LARGE-SCALE CsCl PREP OF BACTERIAL GENOMIC DNA 1. Grow 100 ml culture of bacterial strain to saturation.

SHORT PROTOCOL

2. Spin 10 min at 4000 × g. 3. Resuspend pellet in 9.5 ml TE buffer, 0.5 ml of 10% SDS, and 50 µl of 20 mg/ml proteinase K. Mix and incubate 1 hr at 37°C. 4. Add 1.8 ml of 5 M NaCl. Mix thoroughly. 5. Add 1.5 ml CTAB/NaCl solution and mix. Incubate 20 min at 65°C. 6. Extract with an equal volume of chloroform/isoamyl alcohol. Spin 10 min at 6000 × g, room temperature. 7. Transfer aqueous phase to a fresh tube. Extract with phenol/chloroform/isoamyl alcohol if necessary. Spin as in step 6. 8. Transfer aqueous phase to a fresh tube. Precipitate DNA with 0.6 vol isopropanol. Wash precipitate with 70% ethanol. Remove supernatant and resuspend pellet in 4 ml TE buffer. 9. Measure DNA concentration. Adjust concentration to give 50 to 100 µg/ml. Add 4.3 g CsCl per 4 ml TE buffer. Add 200 µl of 10 mg/ml ethidium bromide. Transfer to sealable centrifuge tubes. Spin 4 hr at 70,000 rpm, 15°C. 10. Visualize gradient with UV light. Remove band. 11. Extract ethidium bromide with CsCl-saturated isopropanol. 12. Dialyze overnight against 2 liters TE buffer.



Supplement 56

REAGENTS AND SOLUTIONS CTAB/NaCl solution (10% CTAB in 0.7 M NaCl) Dissolve 4.1 g NaCl in 80 ml water and slowly add 10 g CTAB (hexadecyltrimethyl ammonium bromide) while heating and stirring. If necessary, heat to 65°C to dissolve. Adjust final volume to 100 ml. COMMENTARY Background Information Most commonly used protocols for the preparation of bacterial genomic DNA consist of lysozyme/detergent lysis, followed by incubation with a nonspecific protease and a series of phenol/chloroform/isoamyl alcohol extractions prior to alcohol precipitation of the nucleic acids (Meade et al., 1984; Silhavy et al., 1982). Such procedures effectively remove contaminating proteins, but are not effective in removing the copious amounts of exopolysaccharides that are produced by many bacterial genera, and which can interfere with the activity of molecular biological enzymes such as restriction endonucleases and ligases. In this procedure, however, the protease incubation is followed by a CTAB extraction whereby CTAB complexes both with polysaccharides and with residual protein; both groups of contaminating molecules are effectively removed in the subsequent emulsification and extraction with chloroform/isoamyl alcohol. This procedure is effective in producing digestible chromosomal DNA from a variety of gram-negative bacteria, including those of the genera Pseudomonas, Agrobacterium, Rhizobium, and Bradyrhizobium, all of which normally produce large amounts of polysaccharides. If large amounts of exceptionally clean DNA are required, the procedure can be scaled up and the DNA purified on a cesium chloride gradient, as described in the alternate protocol. The method can also be used to extract high-molecular-weight DNA from plant tissue (Murray and Thompson, 1980).

Critical Parameters The most critical parameter is the salt (NaCl) concentration of the solution containing the

lysed bacteria prior to adding CTAB. If the NaCl concentration is 500 bp) is the silica membrane spin column method (see Alternate Protocol 2). This is a very rapid method that has the advantage of concentrating purified DNA from agarose gels without the need for ethanol precipitation. Commercial kits for this procedure are now widely available (e.g., Qiagen, Promega, Invitrogen, Novagen).

Critical Parameters and Troubleshooting The quality of the preparative DNA digest and the resolution of the preparative gel are critical for obtaining optimal yields of fragments from all protocols. If the preparative gel is overloaded, cross-contamination will occur as a small amount of each fragment becomes “trapped” in the other bands. This low level of contamination can be significant if the fragment is to be used for some cloning procedures, or as a probe in some hybridizations. If it is

necessary to obtain a large amount of highly pure fragment, the first preparation should be re-electrophoresed and repurified. For the Elutip purification used in the electroelution or low gelling/melting temperature agarose protocols, it is essential that the DNA solution be passed through the column slowly to adsorb it completely. For the phenol extraction purification from low gelling/melting temperature agarose, it is essential to use straight buffered phenol and not phenol/chloroform/isoamyl alcohol mixtures. Transfer to NA-45 paper works well as long as the paper is carefully introduced in front of the band of interest, so that air bubbles are not trapped between the paper and the gel. It is convenient to examine the gel under UV illumination with a hand-held lamp prior to removing the NA-45 paper, to ensure that the fragment has migrated into the paper. If the fragment of interest is 500 bp; smaller-length fragments apparently bind tightly and irreversibly to the silica membrane.

Time Considerations The gel slice containing the target band should be cut out as soon as possible after the preparative gel has been electrophoresed—this will minimize diffusion of the DNA in the band. The gel slice can be stored for hours to days by wrapping the damp gel slice in a piece of plastic wrap and refrigerating. Depending on the size of the fragment to be isolated, electroelution should take 2 to 4 hr. After electroelution, the DNA solution can be stored for days at 4°C before purifying. Once Elutip purification has been started, however, it should be carried to the stage of ethanol precipitation. The Elutip-d column itself is completed in ∼15 min. NA-45 elution takes ∼1 hr for fragments 1500

bp. Purification of fragments separated on low gelling/melting temperature agarose, using either phenol extraction or β-agarase digestion, will take ∼1 hr. Purification via silica membrane spin column requires 20 to 40 min. Linker removal using Sephacryl S-300 takes ∼30 min.

Literature Cited Blin, N., Gabain, A.V., and Bujard, H. 1975. Isolation of large molecular DNA from agarose gels for further digestion by restriction enzymes. FEBS Lett. 53:84-86. Thuring, R.W., Sanders, J.B., and Borst, P.A. 1975. Freeze-squeeze method for recovering long DNA from agarose gels. Anal. Biochem. 66:213220. Vogelstein, B. and Gillespie, D. 1979. Preparative and analytical purification of DNA from agarose. Proc. Nat. Acad. Sci. U.S.A. 76:615-619. Wienand, U., Schwarz, Z., and Felix, G. 1978. Electrophoretic elution of nucleic acids from gels adapted for subsequent biological tests: Application for analysis of mRNAs from maize endosperm. FEBS Lett. 98:319-323.

Contributed by David Moore and Dennis Dowhan Baylor College of Medicine Houston, Texas Joanne Chory The Salk Institute La Jolla, California Randall K. Ribaudo (NA-45 paper) National Institute of Allergy and Infectious Diseases Bethesda, Maryland

Purification of Large DNA Restriction Fragments



RESOLUTION AND RECOVERY OF SMALL DNA FRAGMENTS

SECTION III

This section describes the use of polyacrylamide gels and sieving agarose gels for analytical or preparative separation of small double-stranded DNA fragments. Polyacrylamide gels provide somewhat better resolution as well as significantly higher capacity. Sieving agarose gels are much easier to pour and run, however, and are particularly useful for simple analytical applications. The use of denaturing polyacrylamide gels to separate single-stranded polynucleotides is described in UNITS 2.12 & 7.6.

Separation of Small DNA Fragments by Conventional Gel Electrophoresis

UNIT 2.7

Large amounts of small (200 mM. One unit is defined as the amount of enzyme that produces 1 µg of acid-soluble material in 1 min at 37°C using single-stranded salmon sperm DNA as the substrate. Enzymatic Manipulation of DNA and RNA


Supplement 8

Reaction Conditions For 100-ìl reaction: 30 mM sodium acetate, pH 5.0 50 mM NaCl 1 mM zinc acetate 5% (vol/vol) glycerol 1 µg DNA 50 µg/ml BSA 15 U mung bean nuclease Incubate 30 min at 37°C. Stop reaction by adding 1 µl of 0.5 M EDTA. The volume of reaction, amount of DNA, units of enzyme, temperature, and time of reaction will vary depending on the application. Applications Mung bean nuclease is used for most of the same purposes as described for S1 nuclease. However, mung bean nuclease is more precise than S1 nuclease for cleaving immediately adjacent to the last hybridized base pair without removing any of the base-paired nucleotides. For this reason, mung bean nuclease will typically generate single bands in transcript mapping experiments (UNIT 4.6), whereas S1 often results in multiple bands. In addition, mung bean nuclease is preferred over S1 for precisely deleting overhanging bases that result from restriction endonuclease cleavage. Unlike S1 nuclease, mung bean nuclease will not cleave the strand opposite a nick in duplex DNA. The disadvantages of mung bean nuclease are that it is more expensive than S1, and that its activity is more sensitive to reaction conditions. ENZYME

MICROCOCCAL NUCLEASE Micrococcal nuclease from Staphylococcus aureus is a relatively nonspecific nuclease that cleaves single- and double-stranded DNA and RNA to oligo- and mononucleotides with 3′ phosphates (Alexander et al., 1961). The enzyme is more active on single- stranded nucleic acids. Cleavage of DNA or RNA occurs preferentially at AT- or AU-rich regions although all sequences are ultimately cleavable. The enzyme is strictly dependent on calcium for activity and hence can be inactivated by Ca++-specific chelating agents such as EGTA. One unit is defined as the amount of enzyme that hydrolyzes 1 µmol of acid-soluble oligonucleotides from native DNA per minute at 37°C, pH 8.8. Reaction Conditions Typical micrococcal nuclease digestions are performed in solutions containing 10 mM Tris⋅Cl, pH 8.0, and 1 mM CaCl2. However, the enzyme is somewhat more active at higher pH. The volume of reaction, amount and source of nucleic acid, units of enzyme, temperature, ionic strength, and time of reaction will vary greatly depending on the application. The reactions can be stopped by EDTA or the Ca++-specific chelator EGTA. Applications 1. Studies of chromatin structure (Kornberg, 1977).

Endonucleases

2. Removing nucleic acid from crude cell-free extracts without destroying enzyme activities. Digestion of nucleic acid is performed under mild buffer and ionic conditions in the presence of CaCl2. When digestion is complete, micrococcal nuclease is inactivated with EGTA (Pelham and Jackson, 1976). Such treated extracts are generally active for all processes except those requiring calcium.

3.12.4 Supplement 8


DEOXYRIBONUCLEASE I (DNase I)

ENZYME

DNase I from bovine pancreas is an endonuclease that degrades double-stranded DNA to produce 3′-hydroxyl oligonucleotides (Moore, 1981; Fig. 3.12.3). The enzyme requires divalent cations; the specificity of the reaction differs depending upon which divalent cation is present (Campbell and Jackson, 1980). In the presence of Mg++, DNase I produces nicks in duplex DNA, while in the presence of Mn++ the enzyme produces double-stranded breaks in the DNA. I. Activity in the presence of Mg++:

P 5′ 3′

3′ 5′

5′ 3′

P

P

P 3′ 5′

P P

II. Activity in the presence of Mn++:

5′ 3′

3′ 5′

5′ 3′

P 5′ OH 3′

3′ OH 5′ P

3′ OH 5′ P

P 5′ OH 3′

P 5′ OH 3′

3′ OH 5′ P

3′ 5′

Figure 3.12.3 DNase I activities.

Reaction Conditions For 100-ìl reaction: 50 mM Tris⋅Cl, pH 7.5 10 mM MgCl2 (for single-strand nicks; to create double-strand breaks, replace with 10 mM MnCl2) 2 µg DNA 50 µg/ml BSA 1 µl DNase I (the concentration depends on the application) Incubate at 37°C for 1 to 30 min, depending upon the degree of digestion desired. Stop the reaction by adding 5 µl of 0.5 M EDTA. For nick translations, the DNase I reaction is carried out simultaneously with the DNA polymerase I reaction (UNIT 3.5). Preparation and Storage of DNase I Solutions DNase I is usually purchased as a lyophilized powder at a concentration of 2000 to 3000 U/mg protein. For almost all applications, it is useful to make up a solution containing DNase I that can be stored for long periods of time without loss of enzyme activity. One method is to dissolve 1 mg DNase I in 1 ml of a 50% (wt/vol) solution of glycerol containing 20 mM Tris⋅Cl, pH 7.5, plus 1 mM MgCl2; this solution can be stored in liquid form at −20°C. Alternatively, 1 mg DNase I can be dissolved in 1 ml of a solution containing 20 mM Tris⋅Cl, pH 7.5, plus 1 mM MgCl2, then aliquotted into small microcentrifuge tubes (100 10-µl aliquots are convenient). The aliquots should be quick-frozen on dry ice, then stored at −80°C. For use, an aliquot of DNase I is thawed on ice and an appropriate amount is added to the reaction mixture (for nick translations, the enzyme must first be diluted; see UNIT 3.5). Aliquots are not refrozen; they are simply discarded after use. DNase I solutions prepared by either method are stable for at least 1 year. It is important that DNase I be dissolved without vortexing (to minimize denaturation) and at a concentration of at least 1 mg/ml (to improve long-term stability).

Enzymatic Manipulation of DNA and RNA


Supplement 8

RNase-Free DNase I For many purposes, DNase I should be free of RNase. High-grade commercial preparations such as Worthington grade DPRF can be satisfactory. Alternatively, DNase I can be dissolved at 1 mg/ml in 0.1 M iodoacetic acid plus 0.15 M sodium acetate at a final pH of 5.3. The solution is then heated 40 min at 55°C and cooled. Finally, 1 M CaCl2 is added to the solution to 5 mM. This homemade preparation of RNase-free DNase I should be stored frozen in small aliquots. Applications 1. Nick translation (UNIT 3.5). Introduces nicks in duplex DNA which then serve as primer sites to initiate DNA synthesis by E. coli DNA polymerase I. 2. Cloning random DNA fragments by catalyzing double-stranded cleavage of DNA in the presence of Mn++ (Anderson, 1981). LITERATURE CITED Alexander, M., Heppel, L.A., and Hurwitz, J. 1961. The purification and properties of micrococcal nuclease. J. Biol. Chem. 236:3014-3019. Anderson, S. 1981. Shotgun DNA sequencing using cloned DNase I-generated fragments. Nucl. Acids Res. 9:3015-3027. Campbell, V.W. and Jackson, D.A. 1980. The effect of divalent cations on the mode of action of DNase I. The initial reaction products produced from covalently closed circular DNA. J. Biol. Chem. 255:37263735. Gray, H.B., Ostrander, D.A., Hodnett, J.L., Legerski, R.J., and Robberson, D.L. 1975. Extracellular nucleases of Pseudomonas Bal 31. I. Characterization of single strand-specific deoxyriboendonuclease and doublestrand deoxyriboexonuclease activites. Nucl. Acids Res. 2:1459-1492. Gray, H.B., Winston, T.P., Hodnett, J.L., Legerski, R.J., Nees, D.W., Wei, C.-F., and Robberson, D.L. 1981. The extracellular nuclease from Alteromonas espejiana: An enzyme highly specific for nonduplex structure in nominally duplex DNAs. In Gene Amplification and Analysis, Vol. 2: Structural analysis of nucleic acids (J.G. Chirikjian and T.S. Papas, eds.) pp. 169-203. Elsevier/North Holland, New York, Amsterdam, Oxford. Kornberg, R.D. 1977. Structure of chromatin. Ann. Rev. Biochem. 46:931-954. Kroeker, W.D., Kowalski, D., and Laskowski, M. 1976. Mung bean nuclease I. Terminally directed hydrolysis of native DNA. Biochemistry 15:4463-4467. Lau, P.P. and Gray, H.B. 1979. Extracellular nucleases of Alteromonas espejiana Bal 31. IV. The single strand-specific deoxyriboendonuclease activity as a probe for regions of staltered secondary structure in negatively and positively supercoiled closed circular DNA. Nucl. Acids Res. 6:331-357. Legerski, R.J., Gray, H.B. and Robberson, D.L. 1977. A sensitive endonuclease probe for lesions in DNA helix structure produced by carcinogenic or mutagenic agents J. Biol. Chem. 252:8740-8746. Legerski, R.J., Hodnett, J.L., and Gray, H.B. 1978. Extracellular nucleases of Pseudomonas Bal 31. III. Use of the double-strand deoxyriboexonuclease activity as a basis of a convenient method for the mapping of fragments of DNA produced by cleavage with restriction enzymes. Nucl. Acids Res. 5:1445-1464. Moore, S. 1981. Pancreatic DNase. In The Enzymes, Vol. 14A (P.D. Boyer, ed.) pp. 281-298. Academic Press, San Diego. Pelham, H.R.B. and Jackson, R.J. 1976. An efficient mRNA-dependent translation system from reticulocyte lysates. Eur. J. Biochem. 67:247-256. Vogt, V.M. 1980. Purification and properties of S1 nuclease from Aspergillus. Methods Enzymol. 65:248254. Wei, C.-F., Alianell, G.A., Bencen, G.H., and Gray, H.B. 1983. Isolation and comparison of two molecular species of the Bal 31 nuclease from Alteromonas espejiana with distinct kinetic properties. J. Biol. Chem. 258:13506-13512.

Contributed by Stanley Tabor and Kevin Struhl Harvard Medical School Boston, Massachusetts Endonucleases

3.12.6 Supplement 8


Ribonucleases

UNIT 3.13

Ribonucleases (RNases) with different sequence specificities are used for a variety of analytical purposes, including RNA sequencing, mapping, and quantitation. One very common application for RNase A (see below) is hydrolyzing RNA that contaminates DNA preparations. Two other commonly used RNases, RNase H and RNase T1, are also described below. Many commercially available RNases are sequence-specific endoribonucleases. This property has been used for enzymatic sequencing of RNA. For example, a combination of three different RNases and Staphylococcus aureus nuclease can be used in RNA sequence determination (see Table 3.13.1). Table 3.13.1

RNases Used in RNA Sequencinga

RNase

Sequence specificity

T1 U2 CL3 S. aureus nucleaseb

Gp↓N Ap↓N C(A/G)p↓N Np↓A/U

aAdapted from Boehringer Mannheim. bStaphylococcus aureus nuclease cleaves both RNA and DNA.

RIBONUCLEASE A

ENZYME

Ribonuclease A (RNase A) from bovine pancreas is an endoribonuclease that specifically hydrolyzes RNA after C and U residues (Richards and Wyckoff, 1971). Cleavage occurs between the 3′-phosphate group of a pyrimidine ribonucleotide and the 5′- hydroxyl of the adjacent nucleotide. The reaction generates a 2′:3′ cyclic phosphate which then is hydrolyzed to the corresponding 3′-nucleoside phosphates. RNase A activity can be inhibited specifically by an RNase inhibitor (e.g., RNasin from Promega), a protein isolated from human placenta. Reaction Conditions RNase A is active under an extraordinarily wide range of reaction conditions, and it is extremely difficult to inactivate. At low salt concentrations (0 to 100 mM NaCl), RNase A cleaves single-stranded and double-stranded RNA as well the RNA strand in RNA:DNA duplexes. However, at NaCl concentrations of 0.3 M or above, RNase A becomes specific for cleavage of single-stranded RNA. Removal of RNase A from a reaction solution generally requires treatment with proteinase K followed by multiple phenol extractions and ethanol precipitation. DNase-Free RNase A To prepare RNase A free of DNase, dissolve RNase A in TE buffer at 1 mg/ml, and boil 10 to 30 min. Store aliquots at −20°C to prevent microbial growth. Applications 1. Mapping and quantitating RNA species using the ribonuclease protection assay (UNIT 4.7). It is used in conjunction with RNase T1. 2. Hydrolyzing RNA that contaminates DNA preparations (UNITS 1.6 & 1.7). 3. RNA sequencing. Contributed by Kevin Struhl Current Protocols in Molecular Biology (1989) 3.13.1-3.13.3 Copyright © 2000 by John Wiley & Sons, Inc.


3.13.1 Supplement 8

4. Blunt-ending double-stranded cDNA (UNIT 5.5). It is used in conjunction with RNase H. ENZYME

RIBONUCLEASE H Ribonuclease H (RNase H) from E. coli is an endoribonuclease that specifically hydrolyzes the phosphodiester bonds of RNA in RNA:DNA duplexes to generate products with 3′ hydroxyl and 5′ phosphate ends (Berkower et al., 1973). It will not degrade singlestranded or double-stranded DNA or RNA. RNase H cleavage can be directed to specific sites by hybridizing short deoxyoligonucleotides to the RNA (Donis-Keller, 1979). One unit is defined as the amount of enzyme that produces 1 nmol of acid-soluble ribonucleotides from poly(A)⋅poly(dT) in 20 min at 37°C. Reaction Conditions For 100-ìl reaction: 20 mM HEPES⋅KOH, pH 8.0 50 mM KCl 4 mM MgCl2 1 mM DTT 2 µg RNA:DNA duplex 50 µg/ml BSA 1 U ribonuclease H Incubate 20 min at 37°C. Stop reaction by adding 1 µl of 0.5 M EDTA. The volume of reaction, amount of DNA, units of enzyme, temperature, time, and method of stopping the reaction will vary depending on the application. Applications 1. Facilitating the synthesis of double-stranded cDNA by removing the mRNA strand of the RNA:DNA duplex produced during first strand synthesis of cDNA (UNIT 5.5). It is also used later in the protocol to degrade residual RNA in conjunction with RNase A. 2. Creating specific cleavages in RNA molecules by using synthetic deoxyoligonucleotides to create local regions of RNA:DNA duplexes (Donis-Keller, 1979).

ENZYME

RIBONUCLEASE T1 Ribonuclease T1 (RNase T1) from Aspergillus oryzae is an endoribonuclease that specifically hydrolyzes RNA after G residues (Uchida and Egami, 1971). Cleavage occurs between the 3′-phosphate group of a guanine ribonucleotide and the 5′-hydroxyl of the adjacent nucleotide. The reaction generates a 2′:3′ cyclic phosphate which then is hydrolyzed to the corresponding 3′-nucleoside phosphates. Reaction Conditions The enzyme is active under a wide range of reaction conditions, and it is difficult to inactivate. At low salt concentrations (0 to 100 mM NaCl), it cleaves single-stranded and double-stranded RNA as well as the RNA strand in RNA:DNA duplexes. However, at NaCl concentrations of 0.3 M or above, it becomes specific for cleavage of single-stranded RNA. Removal of RNase T1 from a reaction solution generally requires treatment with proteinase K followed by multiple phenol extractions and ethanol precipitation.

Ribonucleases

3.13.2 Supplement 8


Applications 1. Mapping and quantitating RNA species using the ribonuclease protection assay (UNITS 4.7 & 9.8). It is used in conjunction with RNase A. 2. RNA sequencing. 3. Determining the level of RNA transcripts synthesized in vitro from DNA templates containing a “G-less cassette.” Because accurately initiated transcripts contain no G residues, treatment of the reaction products with RNase T1 preferentially destroys all nonspecific transcripts (Sawadogo and Roeder, 1985). LITERATURE CITED Berkower, I., Leis, J., and Hurwitz, J. 1973. Isolation and characterization of an endonuclease from Escherichia coli specific for ribonucleic acid in ribonucleic acid⋅deoxyribonucleic acid hybrid structures. J. Biol. Chem. 248:5914-5921. Boehringer Mannheim Biochemicals. Biochemicals for Molecular Biology (catalog). Indianapolis. Donis-Keller, H. 1979. Site specific enzymatic cleavage of RNA. Nucl. Acids Res. 7:179-192. Richards, F.M. and Wyckoff, H.W. 1971. Bovine pancreatic ribonuclease. In The Enzymes, Vol. IV (P.D. Boyer, ed.) pp. 647-806. Academic Press, San Diego. Sawadogo, M. and Roeder, R.G. 1985. Factors involved in specific transcription by human RNA polymerase II: Analysis by a rapid and quantitative in vitro assay. Proc. Natl. Acad. Sci. U.S.A. 82:4394-4398. Uchida, T. and Egami, F. 1971. Microbial ribonucleases with special reference to RNases T1, T2, N1, and U2. In The Enzymes, Vol. IV (P.D. Boyer, ed.) pp. 205-250. Academic Press, San Diego.

Contributed by Kevin Struhl Harvard Medical School Boston, Massachusetts



Supplement 8

UNIT 3.14

DNA Ligases DNA ligases catalyze the formation of phosphodiester bonds between juxtaposed 5′ phosphate and a 3′-hydroxyl terminus in duplex DNA. This activity can repair singlestranded nicks in duplex DNA (Fig. 3.14.1) and join duplex DNA restriction fragments having either blunt ends (Fig. 3.14.2) or homologous cohesive ends (Fig. 3.14.3).Two ligases are used for nucleic acid research—E. coli ligase and T4 ligase. These enzymes differ in two important properties. One is the source of energy: T4 ligase uses ATP, while E. coli ligase uses NAD. Another important difference is their ability to ligate blunt ends; under normal reaction conditions, only T4 DNA ligase will ligate blunt ends.

3′ Example:

3′ 5′

5′ 3′ ATP (T4, T7 ligase)

5′

P

A

G

C

T

A

G

G

A

T

C

G

A

T

C

P

P

P

P

P

P

5′

5′

3′

P

NAD

(T4, T7 ligase) AMP + PP i 3′ 5′

3′

P

T

ATP

NAD (E. coli ligase)

P

P

C

3′ 5′ 3′

5′ 3′

5′

P OH P

P

P

(E. coli ligase) AMP + NMN

P

P

P

P

P

C

T

A

G

C

T

A

G

G

A

T

C

G

A

T

C

P

P

P

P

P

P

3′

P

P

5′

Figure 3.14.1 DNA ligase activity at a nick.

ENZYME

T4 DNA LIGASE T4 DNA ligase, the product of gene 30 of phage T4, was originally purified from phage-infected cells of E. coli. The phage T4 gene 30 has been cloned, and the enzyme is now prepared from overproducing strains. Using ATP as a cofactor, T4 DNA ligase catalyzes the repair of single-stranded nicks in duplex DNA and joins duplex DNA restriction fragments having either blunt or cohesive ends. It is the only ligase that efficiently joins blunt-end termini under normal reaction conditions. It appears that T4 DNA ligase activity may be stimulated by T4 RNA ligase (UNIT 3.15). See UNIT 3.16 for a detailed ligation protocol.

DNA Ligases

3.14.1 Supplement 8

Contributed by Stanley Tabor Current Protocols in Molecular Biology (1987) 3.14.1-3.14.4 Copyright © 2000 by John Wiley & Sons, Inc.

Reaction Conditions For 50-ìl reaction: 40 mM Tris⋅Cl, pH 7.5 10 mM MgCl2 10 mM DTT 1 µg DNA 0.5 mM ATP 50 µg/ml BSA 1 “Weiss” U T4 DNA ligase Incubate at 12° to 30°C for 1 to 16 hr. Stop reaction by adding 2 µl of 0.5 M EDTA or by heating to 75°C for 10 min. The volume of reaction, concentration of DNA, and the temperature and time of the reaction will vary, depending upon the individual application. One Weiss unit is equivalent to 60 cohesive-end units. Ligation of cohesive ends is usually carried out at 12° to 15°C to maintain a good balance between annealing of the ends and activity of the enzyme. Higher temperatures make it difficult for the ends to anneal, whereas lower temperatures diminish ligase activity. Blunt-end ligations are typically performed at room temperature since annealing is not a factor (the enzyme is not particularly stable above 30°C). Blunt-end ligations require about 10 to 100 times more enzyme than cohesive-end ligations to achieve an equal efficiency. T4 DNA ligase is not inhibited by tRNA, but it is strongly inhibited by NaCl concentrations >150 mM. Macromolecular exclusion molecules (e.g., PEG 8000) have been shown to greatly increase the rate of both cohesive-end and blunt-end joining by T4 DNA ligase (Pfeiffer and Zimmerman, 1983). An inherent consequence of macromolecular crowding is that all ligations are intermolecular; thus, this technique is not suitable for the ligation and circularization of inserts and vectors that are required for most cloning experiments.

3′ 5′ Example:

5′ 3′

5′

P

P

OH P

P

3′

P

C

T

A

G

C

T

A

G

G

A

T

C

G

A

T

C

3′ 5′ 5′ 3′

3′ 5′ 3′

P

P

P

P OH

3′ 5′

P

P

P

5′

5′ 3′ ATP

ATP (T4 ligase) 5′ 3′

P

P

(T4 ligase)

AMP +PP i

5′

P

P

P

P

P

P

P

C

T

A

G

C

T

A

G

G

A

T

C

G

A

T

C

P

P

3′

Figure 3.14.2 DNA ligase activity at blunt ends.

P

P

P

P

3′

P

5′



Applications T4 DNA ligase is by far the most commonly used DNA ligase. It can be used for virtually any application requiring a DNA ligase. Importantly, it efficiently ligates blunt-end termini, a reaction that other ligases do not carry out in the absence of macromolecular exclusion molecules. ENZYME

ESCHERICHIA COLI DNA LIGASE DNA ligase from E. coli is the product of the lig gene. The lig gene has been cloned, and the enzyme is obtained from an overproducing strain. E. coli DNA ligase catalyzes the repair of single-stranded nicks in duplex DNA and joins restriction fragments having homologous cohesive ends. E. coli DNA ligase does not join termini with blunt ends under normal reaction conditions. Unlike the other ligases, it uses NAD as a cofactor. 3′ 5′

Example:

P OH P

5′

5′ 3′

3′ 5′

P

P

P

P

3′

P

C

T

A

G

C

T

A

G

G

A

T

C

G

A

T

C

3′ 5′

5′ 3′ P P P (T4, T7 ligase) (E. coli ligase) 3′ ATP ATP NA (T4, T7 ligase) D AMP +PP i 5′ 3′ 3′ 5′ 5′ P P P

P

P

OH P

P

5′ NAD5′ 3′ (E. coli ligase) AMP + NMN

P

P

P

C

T

A

G

C

T

A

G

G

A

T

C

G

A

T

C

P

P

P

3′

P

P

P

3′

P

P

5′

Figure 3.14.3 DNA ligase activity at cohesive ends.

Reaction Conditions For 50-ìl reaction: 40 mM Tris⋅Cl, pH 8 10 mM MgCl2 5 mM DTT 1 µg DNA 0.1 mM NAD 50 µg/ml BSA 10 “Modrich-Lehman” U E. coli DNA ligase Incubate at 10° to 25°C for 2 to 16 hr. Stop reaction by adding 2 µl of 0.5 M EDTA or by heating to 75°C for 10 min. The volume of reaction, concentration of DNA, and temperature and time of reaction will vary, depending upon the individual application. DNA Ligases

E. coli DNA ligase, in contrast to T4 DNA ligase, does not require reducing agents


(e.g., DTT) in the reaction. PEG 8000 greatly increases the rate of cohesive end joining by E. coli DNA ligase (Harrison and Zimmerman, 1983). Interestingly, the presence of macromolecular exclusion molecules also enables E. coli DNA ligase to efficiently join blunt-end termini, a reaction it is unable to carry out in their absence. Modrich-Lehman units measure the ability to form poly d(A-T) circles. One ModrichLehman unit is equivalent to 6 Weiss units (Modrich and Lehman, 1975). Applications E. coli DNA ligase can be used as an alternative to T4 DNA ligase when blunt-end ligations are not required. Transformation using DNA ligated with E. coli DNA ligase has a lower background that results from aberrant ligations compared with T4 DNA ligase, since T4 DNA ligase has a much lower specificity for the structure of the termini. LITERATURE CITED Harrison, B. and Zimmerman, S.B. 1983. Macromolecular crowding allows blunt-end ligation by DNA ligases from rat liver or Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 80:5852-5856. Modrich, T. and Lehman, I.R. 1975. Enzymatic joining of polynucleotides. J. Biol. Chem. 245:3626-3631. Pfeiffer, B.H. and Zimmerman, S.B. 1983. Polymer-stimulated ligation: Enhanced blunt- or cohesive-end ligation of DNA or deoxyribonucleotides by T4 DNA ligase in polymer solutions. Nucl. Acids Res. 11: 7853-7871.

KEY REFERENCE Engler, M.J. and Richardson, D.C. 1982. DNA ligases. In The Enzymes, Vol. 15B (P.D. Boyer, ed.) pp. 3-30. Academic Press, San Diego.

Contributed by Stanley Tabor Harvard Medical School Boston, Massachusetts



UNIT 3.15 ENZYME

RNA Ligases T4 RNA LIGASE (Uhlenbeck and Gumport, 1982) T4 RNA ligase, the product of the phage gene 63, is purified from phage-infected cells. It catalyzes the ATP-dependent covalent joining of single-stranded 5′-phosphoryl termini of DNA or RNA to single-stranded 3′-hydroxyl termini of DNA or RNA (Fig. 3.15.1).

Example: 3′ RNA [or DNA] 5′

RNA [or DNA] 3′

5′

5′

P

P

5′

OH

P

P

P

P

3′

P

3′

C

T

A

G

C

T

A

G

ATP ATP 5′

AMP + PP i

3′ 5′

P

C

T

P

A

P

G

P

C

P

T

P

A

P

3′

G

Figure 3.15.1 RNA ligase activity.

Reaction Conditions For 50-ìl reaction: 50 mM HEPES, pH 8.3 10 mM MgCl2 5 mM DTT 2 µg single-stranded DNA or RNA 2 mM ATP 50 µg/ml BSA 1 U T4 RNA ligase Incubate at 17°C for 10 hr. Stop the reaction by adding 2 µl of 0.5 M EDTA. Applications 1. Radioactive labeling of 3′ termini of RNA (Uhlenbeck and Gumport, 1982). The reaction contains the RNA molecule as the acceptor, a [5′-32P]nucleoside 3′, 5′bis(phosphate) (e.g., 5′-32P-Cp) as the donor, and ATP as the energy source. The product formed by this reaction is the RNA molecule containing one additional nucleotide at the 3′ terminus and a 32P-labeled phosphate in the last internucleotide linkage. 2. Circularizing deoxy- and ribo-oligonucleotides (Brennan et al., 1983). 3. Ligating oligomers for oligonucleotide synthesis (Romaniuk and Uhlenbeck, 1983). One such use is to synthesize oligomers that contain internally labeled oligomers at specific residues. RNA Ligases

3.15.1

4. Stimulating the blunt-end ligation activity of T4 DNA ligase (Sugino et al., 1977). Contributed by Stanley Tabor Current Protocols in Molecular Biology (1987) 3.15.1-3.15.2 Copyright © 2000 by John Wiley & Sons, Inc.

LITERATURE CITED Brennan, C.A., Manthey, A.E., and Gumport, R.I. 1983. Using T4 RNA ligase with DNA substrates. Meth. Enzymol. 100:38-52. Romaniuk, P.J. and Uhlenbeck, O.C. 1983. Joining of RNA molecules with RNA ligase. Meth. Enzymol. 100:52-59. Sugino, A., Goodman, H.M., Heyneker, H.L., Shine, J., Boyer, H.W., and Cozzarelli, N.R. 1977. Interaction of bacteriophage T4 RNA and DNA ligases in joining of duplex DNA at base-paired ends. J. Biol. Chem. 252:3987-3994.

KEY REFERENCE Uhlenbeck, O.C. and Gumport, R.I. 1982. T4 RNA ligase. In The Enzymes, Vol. 15B (P.D. Boyer, ed.) pp. 31-60. Academic Press, NY.




Supplement 13

SECTION IV

CONSTRUCTION OF HYBRID DNA MOLECULES Given the large number of restriction endonucleases and other enzymes for manipulating DNA, it is possible to create any DNA molecule of interest. As it is impossible to cover all possible ways of manipulating DNA molecules, this section will discuss the general principles involved along with several specific techniques that are frequently used. Chapter 8 describes methods for creating point mutations in DNA.


Subcloning of DNA Fragments In order to construct new DNA molecules, the starting DNAs are treated with appropriate restriction endonucleases and other enzymes if necessary. The individual components of the desired DNA molecule are purified by agarose or polyacrylamide gel electrophoresis, combined, and treated with DNA ligase. The products of the ligation mixture (along with control mixtures) are introduced into competent E. coli cells, and transformants are identified by an appropriate genetic selection. DNA is prepared from the colonies or plaques and subjected to restriction endonuclease mapping in order to determine if the desired DNA molecule was created. All cloning experiments follow the steps outlined below. Materials Calf intestine phosphatase (CIP) and buffer (optional; UNIT 3.10) dNTP mix (0.5 mM each; UNIT 3.4) Klenow fragment of E. coli DNA polymerase I or T4 DNA polymerase (optional; UNIT 3.5) Oligonucleotide linkers (optional) 10 mM ATP 0.2 mM dithiothreitol (DTT) T4 DNA ligase (measured in cohesive-end units; UNIT 3.14) 2× T4 DNA ligase buffer Additional reagents and equipment for restriction endonuclease digestions (UNIT 3.1), transformation of E. coli cells (UNIT 1.8), DNA minipreps (UNIT 1.6), and agarose or polyacrylamide gel electrophoresis (UNIT 2.5 or 2.7). 1. In a 20-µl reaction mixture, cleave the individual DNA components with appropriate restriction endonuclease. After the reaction is complete, inactivate the enzymes by heating 15 min to 75°C. If no further enzymatic treatments are necessary, proceed to step 6. Reaction mixtures can be done in any volume; 20 ìl is convenient for gel electrophoresis (step 6). Many of the subsequent enzymatic manipulations (steps 2 to 5) can be carried out sequentially without further buffer changes.

2. If the 5′ phosphates of one of the DNAs are to be removed (see Example 3.16.1), add 2 µl of 10× CIP buffer and 1 U CIP; incubate 30 to 60 min at 37°C as described in UNIT 3.10. After the reaction is complete, inactivate CIP by heating 15 min to 75°C. If no further enzymatic treatments are necessary, proceed to step 6.

Subcloning of DNA Fragments


3. If one or both ends generated by a restriction endonuclease must be converted to blunt ends (see Example 3.16.4), add 1 µl of a solution containing all 4 dNTPs (0.5 mM each) and an appropriate amount of the Klenow fragment of E. coli DNA polymerase I or T4 DNA polymerase; carry out the filling-in or trimming reaction as described in UNIT 3.5. After the reaction is complete, inactivate the enzymes by Contributed by Kevin Struhl Current Protocols in Molecular Biology (1987) 3.16.1-3.16.2 Copyright © 2000 by John Wiley & Sons, Inc.

heating 15 min to 75°C. If oligonucleotide linkers are to be added (see Example 3.16.6), proceed to step 4. If a DNA fragment containing only one blunt end is desired, cleave the reaction products with an appropriate restriction endonuclease. If no further enzymatic treatments are necessary, proceed to step 6. 4. Add 0.1 to 1.0 µg of an appropriate oligonucleotide linker, 1 µl of 10 mM ATP, 1 µl of 0.2 M DTT, and 20 to 100 cohesive-end units of T4 DNA ligase; incubate overnight at 15°C. Inactivate the ligase by heating 15 min to 75°C. In general, the oligonucleotide linkers are phosphorylated prior to ligation (UNIT 3.10). However, the linkers do not have to be phosphorylated, in which case the products will contain only one linker on each end.

5. Cleave the products from step 4 with the restriction endonuclease recognizing the oligonucleotide linker, adjusting the buffer conditions if necessary. If only one of the two ends is to contain a linker, cleave the products with an additional restriction enzyme (see Examples 3.16.2 and 3.16.6). 6. Isolate the desired DNA segments by gel electrophoresis, or by other methods if appropriate. Electrophoresis in agarose gels is the most common method. Purification is not essential for many cloning experiments, but it is usually very helpful. See critical parameters.

7. Using longwave UV light for visualization of the DNA, cut out the desired band(s) and purify the DNA away from the gel material using the procedures described in UNIT 2.6. It is critical to use longwave UV light sources to prevent damage to the DNA. If low gelling/melting temperature agarose is used, ligation reactions can usually be performed directly in the gel slice (see alternate protocol).

8. Set up the following ligation reaction: 9 µl component DNAs (0.1 to 5 µg) 10 µl 2× ligase buffer 1 µl 10 mM ATP 20 to 500 U (cohesive end) T4 DNA ligase Incubate 1 to 24 hr at 15°C. Simple ligations with two fragments having 4-bp 3′ or 5′ overhanging ends require much less ligase than more complex ligations or blunt-end ligations. The quality of the DNA will also affect the amount of ligase needed. Controlled cloning experiments usually involve several different ligation reactions. For simplicity, appropriate DNAs should be added to each tube (adding water if necessary) so that the volume is 9 ìl. Immediately prior to use, premix on ice the remaining ingredients (2× buffer, ATP, enzyme) in sufficient amounts for all the reactions. To start the ligation reactions, add 11-ìl aliquots of the premix to each tube containing DNA. The amounts and ratios of the component DNAs and the controls for these ligation reactions are discussed below.

9. Introduce 1 to 10 µl of the ligated products into competent E. coli cells and select for transformants by virtue of the genetic marker present on the vector. 10. From individual E. coli transformants, purify plasmid or phage DNAs by miniprep procedures and determine their structures by restriction mapping. If the frequency of transformants containing the desired hybrid DNA molecule is too low, initial screening by filter hybridization to immobilized colonies (UNIT 6.2) or plaques (UNIT 6.1) or by genetic techniques is necessary.



Supplement 9

ALTERNATE PROTOCOL

LIGATION OF DNA FRAGMENTS IN GEL SLICES This alternate protocol saves considerable time in comparison to the basic protocol because it eliminates purification of the DNA fragments away from the gel matrix (Struhl, 1985). It is suitable for most simple cloning experiments and is particularly valuable for carrying out a set of hybrid constructions involving a variety of different DNA fragments. However, as the cloning efficiency may be reduced, this method should be employed only when it is desired to make one (or relatively few) specific molecules. Additional Materials Low gelling/melting temperature agarose (SeaPlaque, FMC Marine Colloids) TAE buffer (APPENDIX 2) 1. Treat the starting DNAs with appropriate restriction endonucleases and other enzymes, as described in steps 1 to 6 of the basic protocol. 2. Subject the treated DNAs to electrophoresis in low gelling/melting temperature agarose using TAE buffer. It is critical that the agarose be of high quality. SeaPlaque is a good choice. The agarose concentration should be kept as low as possible (0.7% is suitable for most applications).

3. Cut out the desired band(s) in the smallest possible volume (20 to 50 µl) using a clean razor blade, and place the gel slice in a microcentrifuge tube. 4. Melt the gel slices containing DNA at 70°C for at least 10 min. This temperature is hot enough to melt the agarose without denaturing the DNA.

5. In separate tubes for each ligation reaction, combine the gel slices containing appropriate DNAs (and water if necessary) for a total volume of 9 µl. Place the tubes at 37°C for a few minutes. The gel slices should remain molten at 37°C.

6. To each tube containing DNA, add 11 µl of an ice-cold mixture containing 2× buffer, ATP, and T4 DNA ligase. Mix immediately by flicking the tube and place on ice. Then incubate the reaction mixtures 1 to 48 hr at 15°C. DNA fragments can still be ligated even though the reaction mixture has resolidified into a gel.

7. After the ligation reaction is complete, remelt the gel slices 5 to 10 min at 73°C and add 5 µl of the ligated products to 200 µl of competent E. coli cells (UNIT 1.8). The remelted gel must be diluted at least 30-fold so that it does not resolidify when the cells are placed on ice.

8. Carry out steps 9 and 10 of the basic protocol. Ligation in gel slices is suitable for most cloning applications. If the method does not work, DNA fragments can be purified from the gel by various techniques discussed in UNIT 2.6. Ligations are then carried out by the basic protocol.

REAGENTS AND SOLUTIONS


2× T4 DNA ligase buffer 100 mM Tris⋅Cl, pH 7.5 20 mM MgCl2 20 mM DTT

3.16.3 Supplement 9


COMMENTARY Background Information The essence of recombinant DNA technology is the joining of two or more separate segments of DNA to generate a single DNA molecule that is capable of autonomous replication in a given host. The simplest constructions of hybrid DNA molecules involve the cloning of insert sequences into plasmid or bacteriophage cloning vectors. The insert sequences can derive from essentially any organism, and they may be isolated directly from the genome (UNITS 5.3 and 5.4), from mRNA (UNIT 5.5), or from previously cloned DNA segments (in which case, the procedure is termed subcloning). Alternatively, insert DNAs can be created directly by DNA synthesis.

Critical Parameters Many different factors must be considered before embarking on any specific DNA manipulation. These include the number, type, and concentration of DNA fragments, the preparation and purification of DNA fragments, and the selection or screening systems that might be necessary for identifying the DNA molecule of interest. Another practical consideration is the number of hybrid DNA molecules that are desired for any particular experiment. For example, when generating genomic or cDNA libraries, it is critical to obtain the maximum number of recombinants. Strategies for producing large libraries are discussed elsewhere (UNITS 5.1 and 5.2). On the other hand, when it is desired to make one (or relatively few) specific DNA molecule, it is important to maximize the frequency of generating the “correct” molecule rather than the absolute cloning efficiency. In other words, it is better to obtain a low number of colonies, mostly containing the desired DNA rather than a large number of colonies few of which contain the desired DNA. In this way, screening for the correct colonies or plaques is unnecessary, and preparation and analysis of DNA from E. coli transformants is minimized; these represent the major time-consuming steps in the overall cloning process. Although it is obvious that the experimental conditions should be designed to favor the desired ligation events, the crucial factor for optimizing the frequency of correct molecules is to reduce the “background” of undesired events that will result in transformable DNA molecules. One major cause of background involves

“errors” in the preparation of the starting DNA components (e.g., incomplete digestion by restriction endonucleases). Such errors can be significantly reduced, but never entirely eliminated. The other major source of background is undesired ligation between molecules in the reaction mixture. This problem is addressed by considering all possible ways in which the starting components (and probable contaminants) can be joined to produce DNA molecules that can transform E. coli via the genetic selection being imposed. By appropriate “tricks,” the major “side reactions” can often be minimized; these are discussed below. DNA preparation and the purification of specific fragments. The efficiency of obtaining the desired recombinant DNA molecule is greatly improved by purifying the individual components by electrophoresis in agarose gels. The extra effort involved is more than compensated by the elimination (in most cases) of the need to screen the recombinants and the resulting preparation and analysis of fewer DNAs. Equally importantly, gel purification makes it possible to use crude miniprep DNAs (UNIT 1.6) as the initial starting material instead of CsClpurified DNAs, thus eliminating time, expense, and tedium. Purified DNA fragments are relatively free of minor contaminants that can cause major background problems. For example, if a circular plasmid vector is cleaved at 99% efficiency with a single restriction enzyme, the uncleaved 1% of the molecules are fully infectious even though they cannot be visualized by ethidium bromide staining. This background, which is unacceptably high for many experiments, can be greatly reduced by gel purification of the linearized DNA. Gel purification is extremely useful in situations where only one of several fragments is needed or where fragments are generated by partial digestion. It is also useful for removal of small oligonucleotide linkers or other low-molecular-weight contaminants. Electrophoretically separated DNA fragments should be isolated in the smallest possible volume of gel slice. Purity is more important than yield. DNA fragments can be purified away from the agarose (or acrylamide) by any of the procedures described in UNIT 2.6. It is important to use high-quality agarose such as SeaKem or SeaPlaque, since other preparations may contain inhibitors of DNA ligase that are hard to remove. DNA purified from acrylamide




gels does not usually contain such inhibitors. Finally, for most applications, ligation reactions can be performed directly in slices of low gelling/melting temperature agarose (Struhl, 1985; see alternate protocol on p. 3.16.3). By eliminating the need to purify the DNA away from the agarose, considerable time and effort is saved. Intramolecular versus intermolecular ligation of DNA fragments. The absolute and relative concentrations of the input DNA fragments influence the frequencies of specific ligation events. The first consideration is whether the end of a particular DNA molecule will be joined to the other end of the same molecule (intramolecular ligation) or to a separate DNA molecule (intermolecular ligation). The second consideration is whether intermolecular events will be between molecules of the same type (vector–vector or insert–insert) or between molecules of a different type (vector–insert). Most hybrid constructions involve one or more intermolecular events followed by a final intramolecular event to generate a circular plasmid molecule. Intramolecular events. Intramolecular events are a major source of background in many cloning experiments primarily because self-ligation of vector molecules produces undesired E. coli transformants. However, since intramolecular ligation can occur only if the ends of the molecule are cohesive or blunt, it can be almost completely eliminated by using molecules with heterologous ends. This is achieved by cleaving the vector with two restriction endonucleases that produce incompatible ends. For situations where the vector must be cleaved with a single endonuclease, intramolecular joining can be prevented by removing both 5′ phosphates of each molecule with calf intestinal phosphatase. These dephosphorylated vector molecules can be ligated to phosphorylated insert DNAs even though at each end only 1 of the 2 strands will be covalently joined in vitro. Escherichia coli can repair the single-stranded nicks in vivo to generate the desired DNA molecules. Intramolecular ligation of insert fragments does not generally affect the background of unwanted colonies. However, it strongly influences the frequency of obtaining the correct colonies because self-ligated insert DNA cannot be joined to the vector. Thus, in situations where the insert DNA fragments can cyclize, relatively high concentrations of DNA (20 to 100 µg/ml) should be used in order to favor

intermolecular ligation events. This is particularly true for short DNA fragments which are much more likely to cyclize than large fragments (for a theoretical discussion, see Dugaiczyk et al., 1975). Intermolecular events. Almost all cloning experiments using plasmid vectors require the joining of two or more separate DNA molecules followed by circularization of the product. Obviously, intermolecular ligation events become increasingly favored over self-joining as the total concentration of the relevant ends becomes higher. Thus, optimal cloning efficiency occurs at DNA concentrations that are high enough to permit sufficient intermolecular joining, but not so high as to reduce intramolecular ligation. For situations when the background is low, DNA concentrations between 1 and 50 µg/ml are acceptable. If we exclude self-ligation events, the frequency of joining different DNA segments is directly related to the molarity of each component in terms of specific ends to be joined. For example, if equimolar amounts of vector (V) and insert (I) are present, 50% of the bimolecular products will be V–I, 25% will be V–V, and 25% will be I–I. However, because in most cases only molecules containing vector sequences will generate E. coli transformants, it is preferable to use higher molar concentrations of insert fragment(s) because this will favor the desired V–I products over the V–V background. This consideration becomes less important if V–V events are precluded by phosphatase treatment or if V–V events cannot generate viable molecules. Amount of DNA. In general, it is not necessary to use very much DNA. For simple hybrid constructions, roughly 1 to 10% of the vector DNA molecules are converted to the recombinant DNA of interest. With normal transformation efficiencies, 106 colonies per microgram of pure vector DNA, ligated products containing as little as 1 ng of treated vector DNA should generate 10 to 100 transformants. This is more than enough if the background is not a problem. In practical terms, if starting DNA fragments are easily visualized by conventional ethidium bromide staining, there is enough for most experiments. This is true even when performing ligation in gel slices (see alternate protocol) where only a fraction of the electrophoretically separated DNA is used. More difficult constructions such as those involving blunt ends, three fragments, or Bal 31 nuclease usually require more DNA. How-


ever, even in these cases, there is sufficient DNA in a fraction of a gel slice such that the alternate protocol can be used. In general, 25 to 100 ng of each DNA component (in a standard 20-µl reaction) is usually enough to obtain the desired recombinants. Of course, the cloning efficiency will be increased by using higher concentrations of the components. If this proves to be necessary, the DNA can be concentrated easily after purifying it away from the agarose (UNIT 2.6). Selection, screening, and enrichment procedures. For some hybrid constructions, colonies containing the desired DNA molecule can be distinguished from background colonies by genetic methods. In such cases, techniques for background reduction become less important, which often means that time can be saved. As an extreme but relatively infrequent example, if one fragment contains the gene for ampicillin resistance and another fragment contains the gene for tetracycline resistance, the recombinant containing both fragments can be selected directly on plates containing both drugs; there is no need to purify the component DNAs prior to ligation. Cloning vectors have been designed to select or screen for recombinants. Perhaps those most commonly used are the pUC or M13mp series of vectors which contain the α-complementation region of the E. coli lacZ gene and produce blue plaques or colonies on appropriate indicator plates (UNIT 1.4). Insertion of a DNA fragment into the lacZ region generally causes a white colony or plaque, which is easily seen among a background of thousands of blue transformants. Another common vector, pBR322, contains two selectable markers, ampicillin and tetracycline resistance. Insertion into one of these marker genes renders the colony sensitive to that drug; such colonies can be screened by replica plating (UNIT 1.5). In many cases, the desired recombinant contains (or lacks) specific regions of DNA that are present in either or both of the starting molecules. Such recombinants can be identified by colony or plaque filter hybridization (UNITS 6.1 and 6.2) using an appropriate 32P-labeled probe (UNIT 3.5). These screening procedures can identify the correct recombinant from a background of hundreds or thousands of transformants. However, they take several days to perform and are unnecessary if the background is acceptably low. Finally, the proportion of correct colonies can be increased by appropriate enzymatic

treatment after the ligation reaction. For example, if the desired product lacks an EcoRI site, but undesired products contain such a site, the background can be reduced simply by EcoRI cleavage after the ligation reaction. Most restriction endonucleases are active in ligase buffer. Controls. It is critical to perform appropriate control experiments simultaneously with the reaction of interest. In this way, it is often possible to determine the relative success of the hybrid construction. Such knowledge is particularly important because preparation and analysis of prospective recombinant DNAs is the most time-consuming part of the overall process. Obviously, it is worth minimizing the number of colonies to be examined from successful experiments as well as avoiding further analysis from cloning attempts that were clearly unsuccessful. NOTE: The basic controls for any experiment are to set up and analyze parallel ligation reactions, each lacking a single DNA component. In order to keep the volume of the ligation reactions constant, the omitted DNA should be replaced by an equal volume of water. In a successful hybrid construction, the number of colonies obtained when all DNA components are present should be higher than any of the control reactions. If this is the case, many or most of the colonies should contain the molecule of interest. Two additional controls should be carried out as part of the transformation procedure. First, an aliquot of competent cells should be incubated in the absence of DNA. The appearance of transformants is indicative of contamination in the cells or transformation buffers. Second, another aliquot of competent cells should be incubated with 1 ng of a control DNA (either plasmid or phage depending on the experiment) to measure the transformation efficiency; 1000 colonies or plaques should be achieved routinely.

Anticipated Results For simple hybrid constructions, approximately 100 to 10,000 colonies or plaques should be generated, most of which should be the desired molecule. For more complex situations, the expected number of colonies should be between 1 and 1000; depending on the individual situation 5 to 80% of the molecules should contain the desired structure. For many cloning experiments, one correct colony is enough.



Time Considerations The enzyme reactions can usually be performed in 1 to 4 hr (except in experiments involving ligation of oligonucleotide linkers). Gel electrophoresis and DNA fragment purification should take an additional 1 to 4 hr; shorter times are obtained by using minigels (UNIT 2.5) and by using the alternate protocol. Thus, in one day, it should be possible to go from uncleaved starting DNAs to the ligation reaction. On day 2, the ligated products can be introduced into E. coli, and the resulting transformants will be obtained on day 3. After colony or plaque purification, DNA from the transformants can be prepared and analyzed on day 4.

EXAMPLES Example 3.16.1: Subcloning DNA Fragments with Homologous and Cohesive Ends Consider the cloning of a 2-kb EcoRI fragment into a vector containing a single EcoRI site. The major source of background is uncleaved or self-ligated vector DNA. In most cases, self-ligation should be reduced by treating the EcoRI cleaved vector DNA with calf intestinal alkaline phosphatase (see Fig. 3.16.1 below and UNIT 3.10). Although phosphatase treatment does lower the absolute efficiency of obtaining the desired DNA molecule, the great

reduction in background is necessary to obtain a decent relative frequency of the desired molecule. The concentration of insert DNA(s) should be relatively high in order to facilitate ligation to the vector. It should also be noted that the insert DNA can contribute to background transformants by self-ligation, especially when it is difficult to purify away from “vector” fragments derived from the original plasmid. Because all EcoRI ends are equivalent, the 2-kb insert can be cloned in either of the two possible orientations with respect to the vector sequences. These two outcomes can be distinguished only by restriction mapping using an enzyme that cleaves asymmetrically within the insert DNA. In addition, it is possible to insert multiple copies of the fragment into the vector. Generally, multiple insertion is infrequent unless the DNA concentrations are very high. When multiple insertions occur, the inserts are essentially always oriented in the same direction. Molecules containing tandem copies oriented in the opposite direction are incapable of replication in E. coli unless the insert fragment is very short (less than 30 bp). For all the reasons mentioned above, cloning of fragments with homologous and cohesive ends is best avoided when possible. Directional cloning, discussed below, is preferable.

vector 5′ pAATTC 3′ G

insert 3′ G CTTAAp 5′

5′ pAATTC 3′ G

G 3′ CTTAAp 5′

phosphatase ligase AATTC G

G CTTAA ligase

GAATTC CTTAAG

AA C TT G

G CTTAA

GAATTC CTTAAG

recombinant DNA (2 orientations)

no recircularization


Figure 3.16.1 Treatment of cleaved vector with CIP in order to reduce background.


Example 3.16.2: Directional Cloning Using Fragments with Heterologous Ends This is by far the most efficient method for cloning, and it should be used wherever possible. In the simplest example, consider the joining of two fragments, each produced by cleavage with enzymes A and B. Self-ligation of either fragment is theoretically eliminated, and ligation of two vector fragments produces a “perfect inverted repeat” molecule that cannot be stably maintained in E. coli. Thus, the background is very low, and if enzymes A and B produce 5′ or 3′ overhangs, the efficiency is very high. Moreover, almost all the transformants will contain recombinant molecules in which one copy of the insert DNA is oriented in a defined direction with respect to the vector. Molecules containing two copies of the insert cannot be formed, and those with three copies will be very rare. The major problems associated with directional cloning come from errors in generating the DNA segments, particularly incomplete cleavage of the vector by one of the restriction endonucleases. Vector molecules that are singly cleaved can self-ligate and generate background. Ideally, molecules cleaved by both enzymes can be purified away from those that are incompletely cleaved by gel electrophoresis. However, this is not possible when the sites for cleavage by enzymes A and B are close together. Moreover, if the two sites are extremely close together, as is the case for the polylinkers in the pUC and M13mp vectors, it can be difficult to cleave the molecules with both enzymes. The high efficiency of directional cloning makes it possible to create molecules composed of three or more segments. A typical three-piece ligation will consist of fragments generated by enzymes A + B, B + C, and A + C. The desired recombinant will occur less frequently than in the case for two-fragment ligation. However, the low background makes it possible to obtain the desired molecule at high enough frequency to permit individual analysis of colonies.

Example 3.16.3: Blunt-End Ligation Although T4 DNA ligase can join fragments with blunt ends, ligation efficiency is greatly reduced as compared to fragments with cohesive ends. Blunt-end ligation is facilitated by using higher concentrations of DNA and ∼10 times more ligase. Unlike the ligation of cohesive ends which requires compatible termini, blunt fragments are equally ligatible no matter which endonuclease is used to produce the

ends. The joining of blunt ends produced by different enzymes almost always results in a product that cannot be cleaved by either enzyme. The most difficult ligations are those in which all the relevant ends are blunt (e.g., cloning an SmaI fragment into an SmaI cleaved vector). The problems are similar to those described in Example 3.16.1 and are compounded by the inefficiency of blunt-end ligation. Cloning involving blunt ends is considerably easier if one of the ligation events involves cohesive ends. For example, two fragments generated by SmaI and EcoRI are much more easily joined than two SmaI-generated segments. In addition, such heterologous ends permit directional cloning and the background is very low (see Example 3.16.2).

Example 3.16.4: Joining DNA Fragments with Incompatible Ends Consider the case of joining an EcoRI end (overhang 5′ AATT) to an SacI end (overhang 3′ AGCT). To join such incompatible termini, it is necessary to convert them to blunt ends. For 5′ overhangs, this is accomplished by “filling in” the ends with the Klenow fragment of E. coli DNA polymerase I in the presence of all 4 dNTPs (see Fig. 3.16.2 below and UNIT 3.5). This reaction should be carried out in the presence of all 4 dNTPs in order to prevent more extensive exonuclease action. For 3′ overhangs, the 3′ to 5′ exonuclease activity of T4 DNA polymerase or Klenow enzyme is used to remove the protruding nucleotides (UNIT 3.5). The T4 enzyme is preferred because it has a much more active exonuclease. However, the Klenow enzyme will do the job, and it is considerably less expensive than T4 DNA polymerase. Both polymerases are active in all restriction endonuclease buffers and hence can be added (along with the 4 dNTPs) directly to the reaction mixture after cleavage. Once the relevant ends have been blunted, the ligation procedure proceeds as in Example 3.16.3. However, the additional step for generating the blunt ends usually results in a lowered efficiency of ligation; the more ends to be blunted, the worse the problem. The fill-in reaction for 5′ overhangs is usually more efficient than the trimming reaction for 3′ extensions. Thus, when incompatible ends must be joined, it is preferable if some of the required ligation reactions involve joining of cohesive ends; e.g., the joining of a BamHI–EcoRI (filled) fragment to a BamHI–SacI (trimmed) fragment.



Supplement 10

EcoRI end 5′ 3′

Klenow fragment DNA polymerase I

SacI end 5′ C 3′ TCGAG

3′ G CTTAA 5′

3′ 5′

T4 DNA

4 dNTPs

polymerase

4 dNTPs

C G

GAATT CTTAA ligase

GAATT C CTTAA G regenerated EcoRI site

Figure 3.16.2 Joining DNA fragments with incompatible ends.

In many cases, the joining of incompatible ends generates a product that cannot be cleaved by either of the restriction enzymes used to produce the ends. In such cases, the structure of the molecule at the fusion point can be determined only by DNA sequencing. Sometimes, however, one of enzyme sites should be regenerated, thus making it possible to analyze the prospective molecules by restriction mapping. For example, a filled in EcoRI site has the terminal sequence GAATT. If this is joined to a fragment whose 5′ terminal nucleotide is C, the EcoRI site will be restored. In principle, a SacI site (GAGCTC) that has been trimmed will regenerate an EcoRI site that has been filled in. Products where the EcoRI and SacI ends have been joined aberrantly will not result in the regeneration of the EcoRI site.

Example 3.16.5: Ligation of Fragments Generated by Partial Cleavage


Sometimes a given DNA fragment will contain internal restriction sites that are the same as those on one (or both) of the ends. In this case, the fragment can be obtained only by partial restriction endonuclease digestion. Contrary to popular opinion, it is not difficult to obtain hybrid DNA molecules when one or more of the components is generated by partial cleavage by a restriction endonuclease(s). By gel purifying the desired partial cleavage product(s), the ligation reactions can be set up in the normal way, and they should be equally efficient. The main difficulty with partial cleavage,

obtaining enough of the specific fragment, can usually be overcome by starting with more DNA (1 to 5 µg should be enough) prior to gel electrophoresis. To ensure the proper degree of partial digestion, the method of serial enzyme dilution should be performed (UNIT 3.1). In some cases, it will be difficult to purify the desired partial cleavage product from undesired products. This is especially true if the starting molecule contains many cleavage sites or if cleavage sites are close together. However, if the ligation reaction has a low background, such impurities are only a minor problem. For example, if the “band” that is cut from the gel contains four different clonable fragments, roughly 25% of the transformants should be the ones of interest. Moreover, if the fragment is generated by partial cleavage at one end and complete cleavage by another enzyme at the other end, many of the contaminating fragments cannot be cloned into the vector and hence will not contribute to the background.

Example 3.16.6: Ligations Involving Oligonucleotide Linkers Synthetic linkers are self-complementary oligonucleotides, typically 8 to 12 bases in length, that anneal to form blunt-ended, double-stranded DNA which contains a site for restriction endonuclease cleavage. For example, GGAATTCC is an 8-bp EcoRI linker (the underlined nucleotides represent the recognition site). Oligonucleotide linkers facilitate



the cloning of blunt-ended DNA fragments, and they are valuable for “introducing” new restriction sites at desired positions. A wide variety of linkers, each containing a specific recognition sequence, is available from commercial suppliers. Linkers are usually obtained in the nonphosphorylated form. For most applications, they are phosphorylated using ATP and T4 polynucleotide kinase (UNIT 3.10). After the kinase reaction, the phosphorylated linkers can be added directly to the ligation reaction. Ligations involving linkers are performed in three stages. First, the DNA of interest, which must contain blunt ends, is joined to the linker, as illustrated in Figure 3.16.3. This reaction is performed with 0.1 to 1 µg of linker, which represents a 100- to 1000-fold molar excess of linker over fragment. The high concentration of ends makes this blunt-end ligation reaction more efficient than normal. Generally, the linker is phosphorylated prior to ligation. In this case, the ligation products will typically contain several linkers on each end of the fragment. Some experiments involve ligation of nonphosphorylated linkers, in which case the products will contain only one linker on each end. Second, after heat treatment to inactivate the ligase, the products of this reaction are cleaved with the restriction enzyme that recognizes the linker, as illustrated in Figure 3.16.3. Because of the high concentration of restriction sites due to the linker, this step usually requires incubation for several hours with a large amount of restriction enzyme (20 to 50 U). It is important that the fragment does not contain an internal site that is recognized by the restriction enzyme (unless the fragment has been methylated previously by the methylase corresponding to the

restriction enzyme). If possible, the ligation products should be cleaved with a second restriction enzyme to produce a fragment with heterologous ends suitable for cloning (see Example 3.16.2). Third, the fragment is purified by gel electrophoresis and then ligated by conventional methods. The gel purification step also removes unligated linkers, which would interfere with this second ligation step. Linkers can also be removed by other methods (chromatography on Sepharose CL-4B or Sephacryl S-300; see UNITS 2.6 and 5.6), but electrophoresis is more efficient and allows for purification of the desired fragment away from undesired fragments.

Example 3.16.7: Cloning Synthetic Oligonucleotides Double-stranded oligonucleotides suitable for cloning can be obtained by annealing two single-stranded oligonucleotides that have been synthesized chemically. Generally, such oligonucleotides are phosphorylated with T4 kinase (UNIT 3.10) prior to annealing. Doublestranded and phosphorylated oligonucleotides can also be generated by “mutually primed synthesis” of appropriate single-stranded oligonucleotides (UNIT 8.2). Oligonucleotides obtained by either method can be cloned by standard ligation procedures using cohesive or blunt ends. The main problem associated with cloning oligonucleotides is that of multiple insertion. This is because small amounts of oligonucleotide represent a large molar excess. This problem is best dealt with by making a series of 10-fold serial dilutions of oligonucleotide and setting up parallel ligation reactions using

blunt-ended DNA

EcoRI linker pGGAATTCC CCTTAAGGp

p p ligase

pGGAATTCCGGAATTCCGGAATTCC CCTTAAGGCCTTAAGGCCTTAAGG

GGAATTCCGGAATTCCGGAATTCC CCTTAAGGCCTTAAGGCCTTAAGGp

EcoRI AATTCC GG

GG CCTTAA

Figure 3.16.3 Joining blunt-ended DNA to EcoRI linker.



Supplement 26

these various concentrations. If single insertions are desired, transformants should be picked off plates that represent the lowest amount of oligonucleotide needed to increase the number of colonies significantly above the background (i.e., the absence of oligonucleotide). In general, optimal conditions for single insertion occur when the oligonucleotide is present in a 5- to 20-fold molar excess over the other fragments in the ligation mixture.

Literature Cited Dugaiczyk, A., Boyer, H.W., and Goodman, H.M. 1975. Ligation of EcoRI endonuclease-generated DNA fragments into linear and circular structures. J. Mol. Biol. 96:174-184. Struhl, K. 1985. A rapid method for creating recombinant DNA molecules. Biotechniques 3:452453.

Contributed by Kevin Struhl Harvard Medical School Boston, Massachusetts




Constructing Recombinant DNA Molecules by the Polymerase Chain Reaction

UNIT 3.17

Any two segments of DNA can be ligated together into a new recombinant molecule using the polymerase chain reaction (PCR). The DNA can be joined in any configuration, with any desired junction-point reading frame or restriction site, by incorporating extra nonhomologous nucleotides within the PCR primers. Cloning by PCR is often more rapid and versatile than cloning with standard techniques that rely on the availability of naturally occurring restriction sites and require microgram quantities of DNA. It is not necessary to know the nucleotide sequence of the DNA being subcloned by this technique, other than the two short flanking regions (∼20 bp) that serve as anchors for the two oligonucleotide primers used in the amplification process. Moreover, PCR can be performed on low-abundance or even degraded DNA (or RNA) sources. This unit describes using PCR to construct hybrid DNA molecules. The main objective is to give an overview of how PCR can be exploited to accomplish numerous cloning strategies; it is assumed that the reader is already familiar with basic molecular biology techniques including PCR amplification (UNIT 15.1) and subcloning (UNIT 3.16). The basic protocol outlines the PCR amplification and cloning strategies. A troubleshooting guide for problems most frequently encountered in PCR cloning, and three specific examples of this technique—for creating (1) in-frame fusion proteins, (2) recombinant DNA products, and (3) deletions and inversions by inverse PCR—are presented in the Commentary. SUBCLONING DNA FRAGMENTS In this protocol, synthetic oligonucleotides incorporating new unique restriction sites are used to amplify a region of DNA to be subcloned into a vector containing compatible restriction sites. The amplified DNA fragment is purified, subjected to enzymatic digestion at the new restriction sites, and then ligated into the vector. Individual subclones are analyzed by restriction endonuclease digestion and either sequenced or tested in a functional assay. The procedure is summarized in Figure 3.17.1.

BASIC PROTOCOL

Materials Template DNA (1 to 10 ng of plasmid or phage DNA; 20 to 300 ng of genomic or cDNA) Oligonucleotide primers (0.6 to 1.0 mM; UNIT 8.5) Mineral oil TE-buffered phenol (UNIT 2.1) and chloroform 100% ethanol TE buffer, pH 8.0 (APPENDIX 2) Klenow fragment of E. coli DNA polymerase I (UNIT 3.5) Vector DNA Calf intestinal phosphatase (UNIT 3.10) Additional reagents and equipment for phosphorylating synthetic oligonucleotides (UNIT 3.10), enzymatic amplification of DNA by PCR (UNIT 15.1), agarose and polyacrylamide gel electrophoresis (UNITS 2.5 & 2.7), DNA extraction and precipitation (UNIT 2.1), purification of DNA by glass beads, electroelution from agarose gels, or from low-gelling/melting temperature agarose gels (UNIT 2.6), restriction endonuclease digestion (UNIT 3.1), ligation of DNA fragments (UNIT 3.16), transformation of E. coli (UNIT 1.8), plasmid DNA minipreps (UNIT 1.6), and DNA sequence analysis (UNIT 7.4) Contributed by Elaine A. Elion Current Protocols in Molecular Biology (1993) 3.17.1-3.17.10 Copyright © 2000 by John Wiley & Sons, Inc.



Amplify the target DNA 1. Prepare the template DNA. If using an impure DNA preparation (i.e., not purified by CsCl gradients), heat sample 10 min at 100°C to inactivate nucleases. Plasmid, phage, genomic, or cDNA, obtained from either rapid preparations or purified on CsCl gradients, can be used as the source of target DNA.

2. Prepare the oligonucleotide primers. If the PCR product is to be cloned by blunt-end ligation, phosphorylate the 5′ hydroxyl of the oligonucleotide primers. A 5′ phosphate on the ends of the PCR products will be needed to form the phosphoester linkage to the 3′OH of the vector during ligation. This step is essential if the vector has been treated with a phosphatase. Because the purity of the oligonucleotides does not seem to affect the PCR reaction, primer purification (as detailed in UNIT 8.5) may not be necessary.

3. Set up a standard amplification reaction and overlay with mineral oil as described in UNIT 15.1. Carry out PCR in an automated thermal cycler for 20 to 25 cycles under the following conditions: denature 60 sec at 94°C, hybridize 1 min at 50°C, and extend 3 min at 72°C. Extend an additional 10 min at 72°C in the last cycle to make products as complete as possible. Include negative controls of no template DNA and each oligonucleotide alone, as well as several different oligonucleotide:template ratios. For a discussion of optimization of amplification conditions see UNIT 15.1. A thermostable DNA polymerase with 3′→5′ exonuclease proofreading activity can be used instead of Taq DNA polymerase to reduce the amount of nucleotide misincorporation during amplification. Pfu DNA polymerase (Stratagene) and Vent DNA polymerase (New England Biolabs) have this activity (follow manufacturers’ instructions).

Recover the amplified fragment 4. Analyze an aliquot (e.g., 4 to 8 µl) of each reaction mix by agarose or polyacrylamide gel electrophoresis to verify that the amplification has yielded the expected product. 5. Recover amplified DNA from PCR reaction mix. Remove mineral oil overlay from each sample, then extract sample once with buffered chloroform to remove residual mineral oil. Extract once with buffered phenol and then precipitate DNA with 100% ethanol. Carrier tRNA may be added during precipitation if desired.

6. Microcentrifuge DNA 10 min at high speed, 4°C. Dissolve pellet in 20 µl TE buffer. Purify desired PCR product from unincorporated nucleotides, oligonucleotide primers, unwanted PCR products, and template DNA using glass beads, electroelution, or phenol extraction of low gelling/melting temperature agarose. Unused oligonucleotide primers can inhibit the ability of the restriction enzymes to digest the amplified PCR product. Amplified DNA that is greater than 100,000 Da (>150 bp) can be rapidly separated from the primers using a Centricon 100 microconcentrator unit from Amicon (follow manufacturers’ instructions). If the gel analysis in step 4 shows that amplification yielded only the desired PCR product, this microconcentrated DNA can be used directly for cloning.

Constructing Recombinant DNA Molecules by PCR

Prepare amplified fragment and vector for ligation 7a. If the PCR fragment is to be cloned by blunt-end ligation, repair the 3′ ends with DNA polymerase I (Klenow fragment). This step is necessary because Taq DNA polymerase adds a nontemplated nucleotide (usually dA) to the 3′ ends of PCR fragments.



7b. If primers contain unique restriction sites, digest half the amplified DNA in 20 µl with the appropriate restriction enzyme(s). Use an excess of enzyme, and digest for several hours. Reserve the undigested half for future use, if necessary.

8. Prepare the recipient vector for cloning by digesting 0.2 to 2 µg in 20 µl with compatible restriction enzymes. If necessary, treat vector DNA with calf intestinal phosphatase (UNIT 3.1) to prevent recircularization during ligation. 9. Separate the linearized vector from uncut vector by agarose or low-gelling/melting temperature gel electrophoresis. Recover linearized vector from the gel by adsorption to glass beads, electroelution, or phenol extraction of low-gelling/melting temperature agarose. Ligate amplified fragment and vector 10. Ligate the PCR fragment into the digested vector following the procedure outlined in UNIT 3.16. 11. Transform an aliquot of each ligation into E. coli. Prepare plasmid miniprep DNA from a subset of transformants. Analyze recombinant plasmids 12. Digest the plasmid DNA of the selected transformants with the appropriate restriction endonuclease. Analyze the digestions by agarose gel electrophoresis to confirm fragment incorporation. 13. Sequence the amplified fragment portion of the plasmid DNA to check for mutations. Alternatively, screen the subset of transformants using a biochemical or genetic functional assay if available. This analysis is critical because the Taq DNA polymerase can introduce mutations into the amplified fragment.

COMMENTARY Background Information The main benefit of cloning by PCR is that unique restriction sites can be introduced on either side of any segment region of amplified DNA to allow its ligation into a recipient vector (Mullis and Faloona, 1987; Chapter 15) in any configuration. The incorporation of additional nucleotides at the 5′ ends of the oligonucleotide primers permits the creation of novel restriction sites or changes in reading frame and coding sequence. The oligonucleotide primers can also be designed to contain mismatches, deletions, or insertions in the region of homology (UNIT 8.5). However, it is not necessary to always incorporate a new restriction site in the primer. Amplified PCR fragments can also be subcloned by blunt-end or sticky-end ligation using preexisting restriction sites within the amplified DNA. For example, PCR might be used to amplify a target DNA that already contains appropriate restriction sites, but is available in limited quantities. Finally, sequential polym-

erase chain reactions can be used to generate more complex recombinant PCR products which can subsequently be subcloned into a recipient vector. The most obvious disadvantage of PCR cloning is the need to verify that the subcloned PCR product does not contain mutations generated during the polymerase chain reaction. In cases where longer DNA segments (i.e., >1 kb) are being amplified, it may be more advantageous to use a polymerase which has a 3′→5′ exonuclease activity (e.g., Pfu polymerase, Stratagene) to reduce the chances of generating mutations. After subcloning, several independent PCR products should be analyzed by DNA sequencing to be sure that the recombinant DNA molecule is not mutated. Sequencing can be laborious when a large fragment of DNA is subcloned. However, subcloned PCR products can be prescreened by either biological or biochemical functional assays if they are available. In some instances, it may be more desir-



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able to break down the cloning into several steps that might involve the introduction of a needed restriction site within a short piece of DNA first.

Critical Parameters


In general, the DNA preparation, purification, and ligation guidelines outlined in UNIT 3.16 should be applied to PCR cloning to ensure recovery of the desired ligation products. However, the following points deserve special consideration. Design of oligonucleotide primers. Primers should only hybridize to the sequence of interest. This can be predicted in instances where sequence information is available. In general, primers with homology of 16 to 20 nucleotides to the target DNA and a GC content of ∼50% should be chosen. A longer oligonucleotide of ∼25 nucleotides should be used for AT-rich regions. In instances where genomic DNA is the source of the target DNA, the oligonucleotide primers should contain at least 20 nucleotides of homology to the target DNA to ensure that they anneal specifically (Arnheim and Erlich, 1992). When using primers to introduce a specific restriction site, a sequence within the target DNA should be selected that requires the addition of the fewest noncomplementary nucleotides to create the new site, if possible. Special consideration should be given to the choice of site itself, as restriction endonucleases vary in their ability to cleave recognition sequences within ten nucleotides of the end of a DNA duplex (consult Table 8.5.1 for the efficacies of different restriction enzymes in cleaving terminal recognition sequences). It is also recommended that four to five additional nucleotides be added on the 5′ side of the restriction site in the primer. Because DNA duplexes “breathe” at termini, potentially interfering with the ability of a restriction enzyme to cleave (Innis et al., 1990), it is useful to use the GCGC “clamp” sequence that is most thermostable (Sheffield et al., 1989). Finally, the sequence of the primer should be checked for internal complementarity to avoid secondary structure formation that will interfere with hybridization of the primer to the target DNA. The 3′ ends of the two primers being used must not be complementary, so that the formation of primer-dimers that will compete with the synthesis of the desired PCR product will be avoided. Additional details on primer design are discussed in UNIT 15.1.

DNA polymerase. Commercially available Taq DNA polymerase (Perkin-Elmer Cetus) lacks the 3′→5′ proofreading exonuclease activity used by DNA polymerase I Klenow fragment and T4 DNA polymerase to reduce error frequency (Kornberg, 1992). This absence of proofreading activity in Taq DNA polymerase is thought to result in a heightened error frequency. Old estimates indicate that the average rate of misincorporation is 8.5 × 10−6 nucleotides per cycle (Goodenow et al., 1989; Fucharoen et al., 1989). Two other thermostable DNA polymerases possessing proofreading 3′→5′ exonuclease activity have recently become commercially available: Pfu DNA polymerase, purified from Pyrococcus furiosus (Stratagene) and Vent DNA polymerase, purified from Thermococcus litoralis (New England Biolabs and Promega). Both are more thermostable than Taq DNA polymerase. Pfu DNA polymerase is 12-fold more accurate than Taq DNA polymerase, as assayed by the method of Kohler et al. (1991). Vent DNA polymerase is 4-fold more accurate than Taq DNA polymerase (Cariello et al., 1991). Although it is difficult to compare the relative error frequencies of three enzymes because they were assayed by different methods, the use of either Vent or Pfu DNA polymerases may reduce the amount of misincorporation. Removal of unincorporated nucleotide triphosphates. It is recommended that the amplified PCR fragment be purified from unincorporated nucleotides and primers. Any method of purification that involves electrophoresis can also separate the desired PCR product from any undesired DNA species produced during amplification. Typical methods of DNA purification include electrophoresis through lowgelling/melting temperature agarose or electrophoresis through agarose followed by DNA purification by electroelution or adsorption to glass beads (UNIT 2.6). However, amplified DNA can be more rapidly purified from unincorporated nucleotide triphosphates and primers using a Centricon microconcentration unit (Amicon). The disadvantage of using the microconcentrator is that undesirable PCR products and the starting template DNA will copurify with the amplified PCR fragment.

Troubleshooting PCR amplification. The use of appropriately designed primers should allow the amplification of the DNA segment of interest. Occasionally, however, primers may not be specific, leading to the amplification of undesired DNA



A

homologous to target DNA

primer 1: 5 ′ GCGCGAAT TCAATGNNNNNNNNNNN

EcoRI

homologous to target DNA

primer 1: 5 ′ GCGCGGATCC NNNNNNNNNNNNNNN

Bam HI

B target DNA primer 1

E

vector DNA primer 2

B

amplify

E ••••••••••••••••••••• ••••••••••••••••••••• B

E

B

open reading frame digest with Eco RI and Bam HI

E

digest with Eco RI and Bam HI B

••••••••••••••• •••••••••••••••

B purify digested fragment and vector; ligate and transform E. coli

E

original DNA

chimeric open reading frame

• • • • • • PCR DNA vector DNA

E

B

restriction site overhang

Figure 3.17.1 Introducing unique restriction sites and creating an in-frame fusion protein by PCR. Abbreviations: E, EcoRI; B, BamHI. For a full description, see Example 3.17.1 in Commentary.



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segments. The specificity of primer to template hybridization will depend upon temperature and salt (see UNITS 15.1 & 6.4 for a thorough discussion). The highest annealing temperature possible should be used to reduce nonspecific associations. Some nonspecific amplifications can be avoided by employing a “hot-start” technique—i.e., adding the DNA polymerase to a prewarmed sample (D’Aquila et al., 1991). In addition, purifying the PCR product by gel electrophoresis will help ensure that the proper DNA fragment is subcloned. If the synthesized primer does not bind with specificity, it may be simplest to have another one synthesized. In setting up the amplification cycle, keep in mind that for cloning, fidelity is more important than yield, so it is better to keep a low cycle number and not to raise the MgCl2 concentration too much. 1.5 mM MgCl2 in the amplification buffer should be sufficient for most primers. Cloning of the amplified fragment. “Stickyend” ligation of the amplified DNA can sometimes be difficult, due to poor cutting of the terminal restriction site by the desired restriction endonuclease. Careful choice of restriction sites and the addition of extra nucleotides to the 5′ end of the primer (see critical parameters) will facilitate digestion. Other common explanations for poor cutting by restriction endonucleases include: (1) Blockage of the duplex terminus by bound Taq DNA polymerase. If this is the case, the amplified DNA can be treated with proteinase K to remove associated protein. This is done by adding 50 µg/ml proteinase K in 10 mM Tris⋅Cl (pH 7.8)/5 mM EDTA/0.5% (v/v) SDS to the sample and incubating 30 min at 37°C. The sample must be extracted with phenol/chloroform (UNIT 2.1) to remove the proteinase K. (2) Inefficient extension by Taq DNA polymerase. The nonduplex ends generated in this fashion can be repaired by filling in with Klenow fragment (UNIT 3.5). (3) An insufficient number of extra nucleotides 5′ to the restriction site in the primer. In this case, the primer should be resynthesized. Prolonged (typically overnight) DNA digestion should also be tried. (4) 5′ terminal breathing of duplex DNA. Duplex formation can be stabilized by the inclusion of 0.1 mM spermidine in the digestion reaction. An alternative approach is to internalize the restriction site by concatemerizing the PCR products prior to restriction endonuclease di-

gestion (Jung et al., 1990). To do this, 5′ phosphorylated oligonucleotide primers are used in the PCR amplification, or following amplification, the DNA fragment is phosphorylated using T4 polynucleotide kinase (UNIT 3.10). The amplified fragments are then concatemerized by ligation using T4 DNA ligase (UNIT 3.16) prior to restriction endonuclease treatment. Aliquots of the amplified DNA before and after concatemerization, and after digestion, can be compared on an agarose gel to confirm that the procedure worked. Blunt-end ligations are often inhibited by nonflushed ends in the PCR fragment, due to the presence of a nontemplate-directed nucleotide (usually dATP) added by Taq DNA polymerase (Clark, 1988). Treatment of the PCR fragment with Klenow polymerase in the presence of dNTPs (UNIT 3.5) to make the ends flush should circumvent this problem. In the event that this simple approach does not work, the PCR fragment can be subcloned into a MstII (or Bsu36 I) site (CC↓TNAGG) which leaves a 5′ dT overhang. This approach requires a recipient vector with a unique MstII site.

Anticipated Results All of the cloning approaches outlined are reliable and should result in efficient recovery of the desired recombinant molecules.

Time Considerations Once the oligonucleotides have been synthesized, the PCR amplification, purification, ligation, and transformation steps can all be done within 2 days. The appropriate subclones can then be sequenced or tested in a functional assay immediately thereafter.

EXAMPLES Example 3.17.1: Creating In–Frame Fusion Proteins by PCR PCR cloning is particularly useful for creating in-frame fusions between two open reading frames, as is often done for synthesizing fusion proteins with E. coli expression vectors (Chapter 16). The essence of this type of subcloning involves incorporating additional noncomplementary nucleotides within the oligonucleotide primer that will encode the junction sequences of the amplified PCR fragment. Consider the introduction of a unique EcoRI site into a piece of target DNA that is to be fused with an open reading frame in the recipient vector, as depicted in Figure 3.17.1. For this experiment, the primers should be



designed as indicated in panel A. Each primer is designed to contain a unique restriction site not present within the target DNA. The primer carrying the EcoRI site contains an additional nucleotide (shown in bold) to allow the ATG of the amplified target DNA to be in-frame with the open reading frame in the vector (bold bracket). The second primer contains a unique BamHI site. Both oligonucleotide primers are designed to be homologous to and anneal with ∼20 nucleotides of DNA flanking the target DNA. They are oriented such that their 3′ hydroxyl ends point toward the target DNA. These unique restriction sites are 5′ to the region of the primer that is homologous to the target DNA. Each new restriction site is separated from the 5′ end of the oligonucleotide by four additional nucleotides to facilitate enzymatic digestion of the amplified DNA. Shown in this example is a GC clamp (Myers et al., 1985) which favors duplex formation at the ends of the amplified fragment. Panel B depicts the sequence of events in this experiment. Following primer annealing and PCR amplification, the amplified DNA is first digested with restriction endonucleases that cleave at the new restriction sites, then purified by gel electrophoresis. The recipient vector DNA is digested with either the same or compatible restriction endonucleases and is purified before being ligated to the recipient vector. In this example, one oligonucleotide contains additional bases to create an in-frame fusion with the plasmid-borne open reading frame. More elaborate primers can be designed to include additional restriction sites in different reading frames to allow subcloning of the same PCR fragment into multiple recipient fusion vectors that may have nonidentical cloning sites. This is efficient both in terms of labor and the cost of having to synthesize a new oligonucleotide primer. Note that an ATG can also be incorporated into the 5′ oligonucleotide primer to create an open reading frame with a new translational start (e.g., in the construction of a promoter-exon fusion).

Example 3.17.2: Creating a Recombinant DNA Molecule by Sequential PCR Amplifications Consider creating a chimeric DNA molecule by sequential polymerase chain reactions rather than by ligation. This technique is useful for complex cloning schemes that involve fusing together more than two pieces of DNA, as depicted in the creation of the gene fusion

shown in Figure 3.17.2. In this example, two PCR products are made from noncontiguous regions of DNA (that are also nonhomologous) in separate reactions. Two of the first-round amplification primers are designed to contain 5′ extensions that are homologous to a portion of the other target gene (see primers 1b and 1c). In this example, the primers for target gene 1 are labeled as 1a and 1b. Primer 1a contains a unique EcoRI site; primer 1b contains a 5′ extension that is homologous to a region in target gene 2 that will be amplified (thin line of arrow). Primers 1c and 1d are for amplifying target gene 2: primer 1c contains a 5′ extension that is homologous to a portion of target gene 1 that will be amplified (bold line of arrow), and primer 1d contains a unique BamHI restriction site. Because primers 1b and 1c contain complementary 5′ extensions, two PCR products containing a region of overlapping homology are generated. The two PCR fragments are purified away from the primers, then mixed together and annealed by denaturation and renaturation. Four DNA species are generated in this reaction: two heteroduplexes associating at the region of overlapping homology and two parental homoduplexes. The recessed 3′ ends of the heteroduplexes are extended by Taq DNA polymerase to produce a single fragment that is equal in length to the sum of the two overlapping fragments. In a second round of amplification, the combined heteroduplex DNA species is amplified by adding the outside set of primers (1b and 1d) to the PCR assay. These primers will now have complete homology to the amplified heteroduplex DNA species. (Note that the parental homoduplexes will not be amplified because only one of each outside primers will anneal to each parental homoduplex.) The complementary primers used in the first polymerase chain reaction step can be designed to either insert a restriction site at the junction between the joined PCR products, or alter a reading frame.

Example 3.17.3: Inserting a Restriction Site by Inverse PCR Consider deleting a segment of DNA from a plasmid by inserting a unique restriction site by inverse PCR as depicted in Figure 3.17.3. Divergent primers containing the same novel restriction site (R) are annealed to two portions of the plasmid. (Note that R can be any restriction site not found in the plasmid.) The primers are oriented with their 3′ ends facing away from



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target gene 2

target gene 1

E

gene 1 homology

primer 1a

primer 1c

gene 2 homology

primer 1b

amplify each target DNA in separate reactions

E • • • • • • • • • • • • • • • ••• • • • • • • • • • • • • • • • •••

B

primer 1d • • ••••••••••••••••••••••••••• • • ••••••••••••••••••••••••••• B

remove primers mix amplified DNA together denature and anneal

E • • • • • • • • • • • • • • • • • • ••• B • • • ••••••••••••••••••••••••••••••• B • • • ••••••••••••••••••••••••••••••• • • • • • • • • • • • • • • • • • • ••• E E

not shown in rest of reactions

B extend with Taq DNA polymerase

E B • • • • • • • • • • • • • • • • • • ••••••••••••••••••••••••••••••• • • • • • • • • • • • • • • • • • • ••••••••••••••••••••••••••••••• E B • • • • • • • • • • • • • • • • • • ••••••••••••••••••••••••••••••• • • • • • • • • • • • • • • • • • • ••••••••••••••••••••••••••••••• anneal outside primers (1a,1d)

1a

E ••••••••••••••••••• •••••••••••••••••••

B ••••••••••••••••••••••••••••••• •••••••••••••••••••••••••••••••

1d

amplify hybrid PCR products

E

gene 1

gene 2

B

original target gene 1 original target gene 2 • • • • • • • • • • • PCR DNA, target gene 1 • • • • • • • • • • • • • • • • • • • • • PCR DNA, target gene 2 PCR DNA, hybrid gene


Figure 3.17.2 Creating a recombinant DNA molecule by sequential PCR amplifications. Primer 1b has a region of homology to target gene 2 (open box); primer 1c has a region of homology to target gene 1 (closed box). Abbreviations: E, EcoRI; B, BamHI. For a complete description, see Example 3.17.2 in Commentary.



5′

R

deletion/ fusion region

R

5′

amplify target DNA

R

R

remove primers; digest DNA with R; circularize by ligation

R

Figure 3.17.3 Inserting a restriction site by inverse PCR. Abbreviation: R, restriction enzyme not found in target plasmid. For a complete description, see Example 3.17.3 in Commentary.

each other so that sequences flanking the region to be deleted will be amplified. The full-length PCR product is purified and digested at the new restriction site and the ends are ligated in a unimolecular reaction. The amount of spacing between the two primers will determine whether the final product contains all of the original sequence or a deletion. Insertions can also be made at specific sites by including 5′ extensions in the primers. Mismatches within the body of the primer can also be used to introduce mutations. This method is useful for rapid introduction of desired restriction sites, and is limited only by the size of the plasmid and the ability of

DNA polymerase to synthesize complete products. This methodology allows the amplification of DNA flanking a region of known sequence. It is also useful for cloning DNA that has not yet been sequenced and for making hybridization probes (Ochman et al., 1990).

Literature Cited Arnheim, N. and Erlich, H. 1992. Polymerase chain reaction strategy. Annu. Rev. Biochem. 61:131156. Cariello, N.F., Swenberg, J.A., and Skopek, T.R. 1991. Fidelity of Thermococcus litoralis DNA polymerase (Vent) in PCR determined by denaturing gradient gel electrophoresis. Nucl. Acids Res. 19:4193-4198.



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Clark, J.M. 1988. Novel nontemplated nucleotide addition reactions catalyzed by procaryotic and eucaryotic DNA polymerases. Nucl. Acids Res. 16:9677-9689. D’Aquila, R.T., Bechtel, L.J., Videler, J.A., Eron, J.J., Gorczyca, P., and Kaplan, J.C. 1991. Maximizing sensitivity and specificity of PCR by pre-amplification heating. Nucl. Acids Res. 19:3749. Fucharoen, S., Fucharoen, G., Fucharoen, P., and Fukamaki, Y. 1989. A novel ochre mutation in the beta-thalassemia gene of a Thai identified by direct cloning of the entire beta-globin gene amplified using polymerase chain reactions. J. Biol. Chem. 264:7780-7783. Goodenow, M., Huet, T., Saurin, W., Kwok, S., Sninsky, J., and Wain-Hobson, S. 1989. HIV-1 isolates are rapidly evolving quasi-species: Evidence for viral mixtures and preferred nucleotide substitutions. J. Acquired Immunol. Defic. Syndr. 2:344-352. Innis, M.A., Gelfand, D.H., Sninsky, J.J., and White, T.J. (eds.) 1990. PCR Protocols. Academic Press, San Diego. Jung, V., Pestka, S.B., and Pestka, S. 1990. Efficient cloning of PCR-generated DNA containing terminal restriction endonuclease sites. Nucl. Acids Res. 18:6156.

Kornberg, A. and Baker, T.A. 1992. DNA Replication, 2nd ed. W.H. Freeman, New York. Mullis, K.B. and Faloona, F.A. 1987. Specific synthesis of DNA in vitro via a polymerase-catalyzed chain reaction. Methods Enzymol. 155:335-350. Ochman, H., Medhora, M.M., Garza, D., and Hartl, D.L. 1990. Amplification of flanking sequences by inverse PCR. In PCR Protocols. (M.A. Innis, D.H. Gelfand, J.J Sninsky, and T.J. White, eds.) pp. 219-227. Academic Press, San Diego, Calif. Sheffield, V.C., Cox, D.R., Lerman, L.S., and Myers, R.M. 1989. Attachment of a 40 base pair G+Crich sequence (GC clamp) to genomic DNA fragments by the polymerase chain reaction results in improved detection of single-base changes. Proc. Natl. Acad. Sci. U.S.A. 86:232-236.

Key Reference Innis et al., 1990. See above. Provides an in-depth analysis of PCR methods and techniques.

Contributed by Elaine A. Elion Harvard Medical School Boston, Massachusetts




SPECIALIZED APPLICATIONS

SECTION V

Labeling and Colorimetric Detection of Nonisotopic Probes

UNIT 3.18

Although a number of different nonisotopic labels have been described in the literature, biotin and digoxigenin are used most frequently and are commercially available. Either label can be easily incorporated into DNA probes and be detected colorimetrically; a number of fluorochromes, as well as alkaline phosphatase and horseradish peroxidase (which produce colored precipitates) are available directly conjugated to anti-digoxigenin antibodies and to avidin. Chemiluminescent detection methods (UNIT 3.19) and indirect immunofluorescent techniques (UNIT 14.6) also provide sensitive alternatives for many molecular biology applications. Because one of the advantages of nonisotopically labeled probes is their long shelf life (≥2 years), many micrograms of DNA can be labeled in one reaction to provide probes of constant quality for multiple experiments. The two basic protocols describe incorporating biotinylated nucleotides into DNA probes by nick translation and random-primed synthesis. The support protocol describes colorimetric detection of the probes, which also serves to check the extent of nucleotide incorporation. An alternate protocol describes adaptations of the basic protocols for incorporation of digoxigenin-modified nucleotides. PREPARATION OF BIOTINYLATED PROBES BY NICK TRANSLATION Nick translation is frequently used to label DNA probes for nonisotopic detection procedures. The DNA to be labeled can be an isolated fragment or—as is often the case for in situ hybridization probes—the intact phage, cosmid, or plasmid clone. The biotinavidin system is the most widely used nonisotopic labeling method and the biotinylated deoxynucleotides and detection reagents required are commercially available from several sources.

BASIC PROTOCOL

The protocol is quite similar to the standard nick translation procedure (UNIT 3.5) except for some modifications—the biotin-11-dUTP is substituted for dTTP in a standard nick translation reaction mixture and the DNase I concentration is adjusted to ensure a size range of 100 to 500 nucleotides. A minimum of 2 hr incubation time is necessary for optimal incorporation of the modified deoxynucleotide. The reaction provides sufficient probe for 15 to 50 in situ hybridizations or 2 to 5 Southern blots. Materials E. coli DNA polymerase I (UNIT 3.5) and 10× buffer (UNIT 3.4) 0.5 mM 3dNTP mix (minus dTTP; UNIT 3.4) 0.5 mM biotin-11-dUTP stock 100 mM 2-mercaptoethanol (2-ME) Test DNA 1 mg/ml DNase I stock (UNIT 3.12) prepared in 0.15 M NaCl/50% glycerol DNA molecular weight markers (UNIT 2.5A) 0.5 M EDTA, pH 8.0 (APPENDIX 2) 10% (w/v) SDS 100% ethanol SDS column buffer 1-ml syringe Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5) and removal of unincorporated nucleotides by Sephadex G-50 spin columns (UNIT 3.4) Contributed by Ann Boyle and Heather Perry-O’Keefe Current Protocols in Molecular Biology (1992) 3.18.1-3.18.9 Copyright © 2000 by John Wiley & Sons, Inc.



1. Prepare a 100-µl reaction mix as follows: 10 µl 10× E. coli DNA polymerase I buffer 10 µl 0.5 mM 3dNTP mix 10 µl 0.5 mM biotin-11-dUTP stock 10 µl 100 mM 2-ME 2 µg DNA 20 U E. coli DNA polymerase I DNase I stock diluted 1:1000 in cold H2O immediately before use H2O to 100 µl. Incubate reaction 2 to 2.5 hr at 15°C. Other biotinylated nucleotides such as biotin-14-dATP and biotin-16-dUTP can be used. Adjust the mixture of unsubstituted deoxynucleotides accordingly. The quantity of DNase I depends on the individual lot used. The enzyme can be diluted in water as described, or in a buffer that is compatible with the nick translation reaction. See notes on DNase I titration in critical parameters.

2. Place reaction on ice. Remove a 6-µl aliquot, boil it 3 min and place on ice 2 min. 3. Load aliquot on an agarose minigel, along with suitable size markers (0.1- to 10-kb range). Run gel quickly (15 V/cm) in case additional incubation is necessary. 4. If the digested DNA is between 100 and 500 nucleotides, proceed to step 5. If the probe size is between 500 and 1000 nucleotides (or larger) add a second aliquot of DNase I and incubate further. Additional DNase I is added in a more concentrated form than in the initial reaction to avoid significant volume changes. For example, if 10ìl of a 1:1000 dilution of DNase I is added initially, the second addition might be 1 ìl of a 1:100 dilution incubated for ~1 hr. Monitor the additional incubation carefully to avoid complete digestion of the probe. If little or no digestion occurs, purify a new DNA sample by phenol extraction and ethanol precipitation (UNIT 2.1) and repeat the nick translation.

5. Add 2 µl of 0.5 M EDTA, pH 8.0 (10 mM final) and 1 µl of 10% SDS (0.1% final) to the reaction. Heat 10 min at 68°C to stop the reaction and inactivate the DNase I. 6. Prepare a Sephadex G-50 spin column in a 1-ml syringe. Wash syringe and silanized glass wool plug with 2 ml of 100% ethanol, then 4 ml water. Pack G-50 resin to the 1-ml mark. Wash column 3 to 4 times with 100 µl SDS column buffer before loading the sample. An additional wash after loading the sample is not necessary. Alternatively, the G-50 resin can be stored in SDS column buffer, reducing the number of times the column should be washed to 2 to 3 times. SDS prevents the biotinylated DNA from sticking nonspecifically to the resin and the glass wool plug.

7. Separate the biotinylated probe from unincorporated nucleotides. The eluted probe concentration should be ∼20 ng/µl (for 2 µg nick-translated DNA) and is ready to use without further treatment. Probe can be stored at −20°C for years without loss of activity. Biotinylated DNA should not be subjected to phenol extraction, as biotin causes the probe to partition to the phenol/water interface or completely into the phenol if heavily biotinylated.

Labeling and Colorimetric Detection of Nonisotopic Probes

8. Assess the extent of the biotinylation reaction and the probe quality by colorimetric (support protocol) or chemiluminescent (UNIT 3.19) detection. Chemiluminescent detection is 5 × 103 times more sensitive than colorimetric detection (Beck and Koster, 1990).



PREPARATION OF BIOTINYLATED PROBES BY RANDOM OLIGONUCLEOTIDE–PRIMED SYNTHESIS

BASIC PROTOCOL

This protocol outlines a priming reaction using random octamers that have been biotinylated at the 5′ end. Octamers are the optimal length for efficient hybridization of 5′-labeled oligonucleotides because the additional two bases act to minimize the steric hindrance from the 5′ end label. This ensures that every probe molecule generated contains at least one biotin, but biotin-16-dUTP is also included in the reaction so that additional biotin is incorporated into some of the probe molecules. Other biotinylated nucleotides (e.g., biotin-14-dATP and biotin-11-dUTP) can be used in place of the biotin-16-dUTP. The starting DNA template should be linear. Best results are obtained with isolated insert DNA that is ≥200 bp. The use of shorter templates will lead to ineffective probe synthesis because of the inability for many random primers to bind on a short target. The amount of template that can be labeled by this procedure ranges from 25 ng to 2 µg. Materials Linear template DNA Biotinylated random octamers dNTP/biotin mix 5 U/µl Klenow fragment (UNIT 3.5) TE buffer, pH 7.5 (APPENDIX 2) 0.5 M EDTA, pH 8.0 (APPENDIX 2) 4 M LiCl 100% and 70% ethanol, ice-cold 1. Place 500 ng to 1 µg template DNA in a 1.5-ml microcentrifuge tube. Add nucleasefree water to a total volume of 34 µl. Up to 2 ìg template DNA can be biotinylated without scaling up the reaction volumes.

2. Denature the DNA for 5 min in boiling water. Place on ice 5 min and microcentrifuge briefly. 3. Add the following to the sample in the order listed: 10 µl biotinylated random octamers 5 µl dNTP/biotin mix 1 µl (5 U) Klenow fragment. Incubate 1 hr at 37°C. If desired, an additional 2.5 U of Klenow fragment can be added after 30 min of incubation to “boost” the reaction. Continuing the incubation for up to 6 hr can increase the yield.

4. Terminate reaction by adding 3 µl of 0.5 M EDTA, pH 8.0. Precipitate the probe by adding 5 µl of 4 M LiCl and 150 µl ice-cold 100% ethanol. Place 30 min on dry ice. 5. Microcentrifuge 10 min at top speed, room temperature, and wash DNA pellet with ice-cold 70% ethanol. 6. Resuspend DNA pellet in 20 µl TE buffer, pH 7.5. Assess the quality of the biotinylation reaction by colorimetric (support protocol) or chemiluminescent (UNIT 3.19) detection. Chemiluminescent detection is 5 × 103 times more sensitive than colorimetric detection (Beck and Koster, 1990). Enzymatic Manipulation of DNA and RNA


Supplement 20

SUPPORT PROTOCOL

COLORIMETRIC DETECTION OF BIOTINYLATED PROBES This support protocol provides a method for checking the extent of the biotinylated incorporation into the labeled probe and serves as the basis for detecting biotinylated DNA probes hybridized to Southern blots. Because the detectability of biotinylated probes is a function of the number of biotin molecules per kilobase, rather than the specific activity of a corresponding radiolabeled probe, the extent of biotinylated deoxynucleotide incorporation is checked by a colorimetric assay (similar assessment can be made by chemiluminescent detection; UNIT 3.19). Under the standard nick translation conditions provided, ∼50 biotin molecules are incorporated per kilobase of DNA. In this procedure, biotinylated DNA is spotted on a nitrocellulose filter. The streptavidin– alkaline phosphatase conjugate binds strongly to the biotin and is visualized when the enzyme is provided with substrates that produce a colored precipitate. As little as 2 pg biotinylated DNA should be visible. If the protocol is being utilized as a method for detecting DNA probes by Southern analysis, follow standard Southern blot protocols (UNIT 2.9) substituting the biotinylated probe (∼100 ng/ml) for the radiolabeled probe and proceed to steps 3 to 8 below. Additional Materials Biotinylated standard DNA (basic protocols or GIBCO/BRL) and test DNA DNA dilution buffer Alkaline phosphatase pH 7.5 (AP 7.5) buffer Blocking buffer: 3% (w/v) BSA fraction V in AP 7.5 buffer 1 mg/ml streptavidin–alkaline phosphatase (AP) conjugate (GIBCO/BRL) Alkaline phosphatase pH 9.5 (AP 9.5) buffer 75 mg/ml nitroblue tetrazolium (NBT) 50 mg/ml 5-bromo-4-chloro-3-indoyl phosphate (BCIP) TE buffer, pH 8.0 (APPENDIX 2) Small piece of membrane (e.g., 5 × 3 cm2): nitrocellulose or uncharged nylon Sealable bags NOTE: To avoid nonspecific background, wear powder-free gloves when handling the membranes. 1. Prepare biotinylated standard DNA in concentrations of 0, 1, 2, 5, 10 and 20 pg/µl in dilution buffer. Dilute the biotinylated test DNA in a similar fashion. 6× SSC in the dilution buffer prevents the DNA spot from spreading on the nitrocellulose, which diminishes the color intensity of the spot.

2a. For nitrocellulose membrane: Spot 1 µl of each dilution on a small piece of nitrocellulose. Bake the membrane ∼1 hr at 80°C. Proceed directly to step 3. Nitrocellulose membranes cannot be used in chemiluminescent detection procedures (UNIT 3.19).

2b. For nylon membrane: Spot 1 µl of each dilution on the nylon membrane. Air-dry and cross-link DNA to the membrane by UV illumination (UNIT 3.19). Proceed to step 3 or develop membrane as described for chemiluminescent detection (UNIT 3.19). This cross-linking step is critical. If done incorrectly, ≤50% of the DNA can be washed off the membrane during the detection steps, resulting in inaccurate probe assessment. Labeling and Colorimetric Detection of Nonisotopic Probes

If chemiluminescent detection is used, the test DNA should be diluted in a series from 10−1 to 10−6, and must be visible at a 10−3 dilution. If it is not, the probe is not sufficiently biotinylated.



3. Float membrane in a small volume of AP 7.5 buffer for 1 min to rehydrate. Place membrane and 10 ml blocking buffer in a sealable bag (cut to size). Avoid trapping air bubbles when sealing. Block for 1 hr at 37°C. 4. Dilute 10 µl streptavidin–AP conjugate with 10 ml AP 7.5 buffer (1 µg/ml final). Cut corner of bag and squeeze out blocking buffer. Replace with the streptavidin–AP solution and reseal. Incubate 10 min at room temperature with agitation on a platform shaker. Do not allow the membrane to dry out at any stage.

5. Remove membrane from bag and transfer to a shallow dish. Wash in 200 ml AP 7.5 buffer (twice, 15 min each time) and in 200 ml AP 9.5 buffer (once, 10 min) with gentle agitation. 6. Add 33 µl of 75 mg/ml NBT to 7.5 ml AP 9.5 buffer and invert to mix. Add 25 µl of 50 mg/ml BCIP and mix gently. Be sure to add reagents in this order and mix gently to prevent precipitation of the reagents.

7. Incubate membrane with NBT/BCIP solution in a shallow dish in low light, checking periodically until color development is satisfactory (usually 15 to 60 min). 8. Stop reaction by washing with TE buffer, pH 8.0. Check incorporation of biotinylated dUTP by comparing the intensities of standard and test DNA. If the probe is at least half as intense as the standard DNA at the corresponding dilution, it should be suitable as an in situ hybridization probe. PREPARATION AND DETECTION OF DIGOXIGENIN-LABELED DNA PROBES

ALTERNATE PROTOCOL

The digoxigenin-based detection system is an alternative nonisotopic labeling method offered by Boehringer Mannheim. Detection is achieved by incubation with antidigoxigenin antibodies coupled directly to one of several fluorochromes or enzymes, or by indirect immunofluorescence (UNIT 14.6). The availability of uncoupled antibodies also permits signal-amplification protocols to be employed. Biotin- and digoxigenin-labeled probes can be visualized simultaneously using a different fluorochrome for each probe. Digoxigenin-11-dUTP can be incorporated into DNA by either of the nick translation or random oligonucleotide–primed synthesis protocols. Boehringer Mannheim advises that nick translation incorporation is not as efficient as random-priming; however, nick translation affords greater control over the final probe size. Nick translated probes perform well with in situ hybridizations, with no apparent sensitivity problems. The protocol is virtually identical to that for biotin-11-dUTP incorporation by nick translation. The standard 100-µl reaction is set up as described in the basic protocol, but 10 µl of a 10× digoxigenin-11-dUTP/dTTP stock solution (see reagents and solutions) is substituted for 10 µl of a 10× biotin-11-dUTP stock solution. The amount of DNase I added to the sample and the time of incubation remain the same and do not depend on the modified deoxynucleotide in the reaction. After incubation at 15°C for 2 hr, an aliquot is run on a minigel to check the probe size. When the correct size has been obtained, the reaction is stopped and the probe is separated from the unincorporated nucleotides using a G-50 spin column. It is advisable to avoid phenol extraction of digoxigenin-labeled probes. The support protocol for colorimetric detection of biotinylated probes can be modified for use with digoxigenin-labeled probes. If incorporation is to be checked, one can use as a control either digoxigenin-labeled standard DNA (obtained from Boehringer Mannheim as part of the Genius kit) or digoxigenin-labeled DNA that has already been used



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successfully. Anti-digoxigenin antibody conjugated to alkaline phosphatase is substituted for streptavidin–alkaline phosphatase. Boehringer Mannheim recommends a 1:5000 dilution. The other steps in the support protocol are unchanged and can be followed exactly. A protocol for chemiluminescent detection of digoxigenin-labeled probes is discussed in UNIT 3.19. REAGENTS AND SOLUTIONS Alkaline phosphatase pH 7.5 (AP 7.5) buffer 0.1 M Tris⋅Cl, pH 7.5 0.1 M NaCl 2 mM MgCl2 Autoclave or filter sterilize and store at room temperature Alkaline phosphatase pH 9.5 (AP 9.5) buffer 0.1 M Tris⋅Cl, pH 9.5 0.1 M NaCl 50 mM MgCl2 Autoclave or filter sterilize and store at room temperature ≤1 yr Biotin-11-dUTP, 0.5 mM Prepare a 0.5 mM stock of lyophilized biotin-11-dUTP (e.g., Sigma) in 20 mM Tris⋅Cl, pH 7.5 (at 289 nm, the molar extinction coefficient ε = 7100; see Table A.3D.1). Check the pH of the stock; if 5 kb is required from a gel that has an agarose concentration of >1.0% and is >5 mm thick.

11. Replace solution with 10 gel volumes of 20× SSC and soak for 45 min. This step is also optional but improves transfer efficiency with some brands of membrane.

Transfer RNA from gel to membrane 12. Place an oblong sponge slightly larger than the gel in a glass or plastic dish (if necessary, use two or more sponges placed side by side). Fill the dish with enough 20× SSC to leave the soaked sponge about half-submerged in buffer. Refer to Figure 2.9.1A for a diagram of the transfer setup. The sponge forms the support for the gel. Any commercial sponge will do, but before a sponge is used for the first time, it should be washed thoroughly with distilled water to remove any detergents that may be present. Two or more sponges can be placed side by side if necessary. As an alternative, a solid support with wicks made out of Whatman 3MM paper (Fig. 2.9.1B) may be substituted. Do not use an electrophoresis tank, as the high-salt transfer buffer will corrode the electrodes. If using a nylon membrane, a lower concentration of SSC (e.g., 10×) may improve transfer of molecules >4 kb; reduction of the SSC concentration is not recommended for a nitrocellulose membrane as the high salt is needed for retention of RNA.

13. Cut three pieces of Whatman 3MM paper to the same size as the sponge. Place them on the sponge and wet them with 20× SSC. 14. Place the gel on the filter paper and squeeze out air bubbles by rolling a glass pipet over the surface. 15. Cut four strips of plastic wrap and place over the edges of the gel. This is to prevent buffer from “short-circuiting” around the gel rather than passing through it.

16. Cut a piece of nylon or nitrocellulose membrane just large enough to cover the exposed surface of the gel. Pour distilled water ∼0.5 cm deep in an RNase-free glass dish and wet the membrane by placing it on the surface of the water. Allow the membrane to submerge. For nylon membrane, leave for 5 min; for nitrocellulose membrane, replace the water with 20× SSC and leave for 10 min. Avoid handling nitrocellulose and nylon membranes even with gloved hands—use clean blunt-ended forceps instead.

17. Place the wetted membrane on the surface of the gel. Try to avoid getting air bubbles under the membrane; remove any that appear by carefully rolling a glass pipet over the surface. 18. Flood the surface of the membrane with 20× SSC. Cut five sheets of Whatman 3MM paper to the same size as the membrane and place on top of the membrane. 19. Cut paper towels to the same size as the membrane and stack on top of the Whatman 3MM paper to a height of ∼4 cm. Preparation and Analysis of RNA


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20. Lay a glass plate on top of the structure and add a weight to hold everything in place. Leave overnight. The weight should be sufficient to compress the paper towels to ensure good contact throughout the stack. Excessive weight, however, will crush the gel and retard transfer. An overnight transfer is sufficient for most purposes. Make sure the reservoir of 20× SSC does not run dry during the transfer.

Prepare membrane for hybridization 21. Remove paper towels and filter papers and recover the membrane and flattened gel. Mark in pencil the position of the wells on the membrane and ensure that the up-down and back-front orientations are recognizable. Pencil is preferable to pen, as ink marks may wash off the membrane during hybridization. With a nylon membrane only, the positions of the wells can be marked by slits cut with a razor blade (do not do this before transfer or the buffer will short-circuit). The best way to record the orientation of the membrane is by making an asymmetric cut at one corner.

22. Rinse the membrane in 2× SSC, then place it on a sheet of Whatman 3MM paper and allow to dry. The rinse has two purposes: to remove agarose fragments that may adhere to the membrane and to leach out excess salt.

Immobilize the RNA and assess transfer efficiency 23a. For nitrocellulose membranes: Place between two sheets of Whatman 3MM filter paper and bake in a vacuum oven for 2 hr at 80°C. Baking results in noncovalent attachment of RNA to the membrane; the vacuum is needed to prevent the nitrocellulose from igniting.

23b. For nylon membranes: Bake as described above or wrap the dry membrane in UV-transparent plastic wrap, place RNA-side-down on a UV transilluminator (254-nm wavelength), and irradiate for the appropriate length of time (determined as described in UNIT 2.9A, Support Protocol). CAUTION: Exposure to UV irradiation is harmful to the eyes and skin. Wear suitable eye protection and avoid exposure of bare skin. UV cross-linking is recommended for a nylon membrane as it leads to covalent attachment and enables the membrane to be reprobed several times. The membrane must be completely dry before UV cross-linking; check the manufacturer’s recommendations, which may suggest baking for 30 min at 80°C prior to irradiation. The plastic wrap used during irradiation must be UV transparent—e.g., polyvinylidene (Saran Wrap). A UV light box (e.g., Stratagene Stratalinker) can be used instead of a transilluminator (follow manufacturer’s instructions).

24. If desired, check transfer efficiency by either staining the gel in ethidium bromide or acridine orange as in steps 7 and 8 or (if using nylon membrane) staining the membrane in 0.03% (w/v) methylene blue in 0.3 M sodium acetate, pH 5.2, for 45 sec and destaining in water for 2 min. If significant fluorescence is observed in the gel, not all the RNA has transferred. RNA bands on a nylon membrane will be stained by the methylene blue (Herrin and Schmidt, 1988).

Analysis of RNA by Northern and Slot Blot Hybridization

Membranes can be stored dry between sheets of Whatman 3MM filter paper for several months at room temperature. For long-term storage they should be placed in a desiccator at room temperature or 4°C.

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25. Prepare DNA or RNA probe labeled to a specific activity of >108 dpm/µg and with unincorporated nucleotides removed. Probes are ideally 100 to 1000 bp in length. DNA for a double-stranded probe is obtained as a cloned fragment (Chapter 1) and purified from the vector by restriction digestion (UNIT 3.1) followed by recovery from an agarose gel (UNIT 2.6). The DNA is labeled by nick translation or random oligonucleotide priming (UNIT 3.5) to create the radioactive probe. A single-stranded DNA probe is created in the same fashion but using a single-stranded vector; the probe should be antisense so it will hybridize to the sense RNA strands that are bound to the membrane. An RNA probe, which should also be antisense, is created by in vitro synthesis from a single-stranded sense DNA fragment (UNIT 2.10).

26. Wet the membrane carrying the immobilized RNA (from step 23) in 6× SSC. 27. Place the membrane RNA-side-up in a hybridization tube and add ∼1 ml formamide prehybridization/hybridization solution per 10 cm2 of membrane. Prehybridization and hybridization are usually carried out in glass tubes in a commercial hybridization oven. Alternatively, a heat-sealable polyethylene bag and heat-sealing apparatus can be used. The membrane should be placed in the bag, all edges sealed, and a corner cut off. Hybridization solution can then be pipetted into the bag through the cut corner and the bag resealed.

28. Place the tube in the hybridization oven and incubate with rotation 3 hr at 42°C (for DNA probe) or 60°C (for RNA probe). If using a bag, it can be shaken slowly in a suitable incubator or water bath. If using a nylon membrane, the prehybridization period can be reduced to 15 min.

29. If the probe is double-stranded, denature by heating in a water bath or incubator for 10 min at 100°C. Transfer to ice. 30. Pipet the desired volume of probe into the hybridization tube and continue to incubate with rotation overnight at 42°C (for DNA probe) or 60°C (for RNA probe). The probe concentration in the hybridization solution should be 10 ng/ml if the specific activity is 108 dpm/ìg or 2 ng/ml if the specific activity is 109 dpm/ìg. For denatured probe, add to hybridization tube as soon after denaturation as possible. If using a bag, a corner should be cut, the probe added, and the bag resealed. It is very difficult to do this without contaminating the bag sealer with radioactivity. Furthermore, the sealing element (the part that gets contaminated) is often difficult to clean. Hybridization in bags is therefore not recommended.

Wash membrane and perform autoradiography 31. Pour off hybridization solution and add an equal volume of 2× SSC/0.1% SDS. Incubate with rotation 5 min at room temperature, change wash solution, and repeat. CAUTION: Hybridization solution and all wash solutions must be treated as radioactive waste and disposed of appropriately. To reduce background, it may be beneficial to double the volume of the wash solutions. If using a bag, transfer the membrane to a plastic box for the washes.

32. Replace wash solution with an equal volume of 0.2× SSC/0.1% SDS and incubate 5 min with rotation at room temperature. Change wash solution and repeat (this is a low-stringency wash; see UNIT 2.10 Commentary). 33. If desired, carry out two further washes using prewarmed (42°C) 0.2× SSC/0.1% SDS for 15 min each at 42°C (moderate-stringency wash). Preparation and Analysis of RNA


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34. If desired, carry out two further washes using prewarmed (68°C) 0.1× SSC/0.1% SDS for 15 min each at 68°C (high-stringency wash). 35. Remove final wash solution and rinse membrane in 2× SSC at room temperature. Blot excess liquid and cover in UV-transparent plastic wrap. Do not allow membrane to dry out if it is to be reprobed.

36. Perform autoradiography. If the membrane is to be reprobed, the probe can be stripped from the hybridized membrane without removing the bound RNA (see Support Protocol). Do not add NaOH. The membrane must not be allowed to dry out between hybridization and stripping, as this may cause the probe to bind to the matrix. ALTERNATE PROTOCOL 1

NORTHERN HYBRIDIZATION OF RNA DENATURED BY GLYOXAL/DMSO TREATMENT In this procedure denaturation of the RNA is achieved by treating samples with a combination of glyoxal and DMSO prior to running in an agarose gel made with phosphate buffer. The glyoxal/DMSO method produces sharper bands after northern hybridization than do formaldehyde gels, but is more difficult to carry out as the running buffer must be recirculated during electrophoresis. Additional Materials (also see Basic Protocol 1) 10 mM and 100 mM sodium phosphate, pH 7.0 (see recipe) Dimethyl sulfoxide (DMSO) 6 M (40%) glyoxal, deionized immediately before use (see recipe) Glyoxal loading buffer (see recipe) 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2) Apparatus for recirculating running buffer during electrophoresis 50° and 65°C water baths NOTE: All solutions should be prepared with sterile deionized water that has been treated with DEPC as described in UNIT 4.1; see unit introduction for further instructions and precautions regarding establishment of an RNase-free environment. Denature and carry out agarose gel electrophoresis 1. Prepare a 1.0% agarose gel by dissolving 1.0 g agarose in 100 ml of 10 mM sodium phosphate, pH 7.0. Cool to 60°C in a water bath, pour gel, and allow to set. Remove comb, place gel in gel tank, and add 10 mM sodium phosphate (pH 7.0) until gel is submerged to a depth of ∼1 mm (see UNIT 2.5A). provides details on preparing, pouring, and running the agarose gel; vary as described here.

UNIT 2.5A

A 1.0% gel is suitable for RNA molecules 500 bp to 10 kb in size. A higher-percentage gel (1.0 to 2.0%) should be used to resolve smaller molecules or a lower percentage (0.7 to 1.0%) for longer molecules. The recipe may be scaled up or down depending on the size of gel desired; the gel should be 2 to 6 mm thick after it is poured and the wells large enough to hold 60 ìl of sample.

2. Adjust volume of each RNA sample to 11 µl with water, then add:


4.5 µl 100 mM sodium phosphate, pH 7.0 22.5 µl DMSO 6.6 µl 6 M glyoxal. Mix samples by vortexing, spin briefly (5 to 10 sec) in a microcentrifuge to collect the liquid, and incubate 1 hr at 50°C.

4.9.8 Supplement 67


3. Cool samples on ice and add 12 µl glyoxal loading buffer to each sample. Load samples onto gel. 0.5 to 10 ìg of RNA should be loaded per lane (see Commentary). Duplicate samples should be loaded at one side of gel for ethidium bromide staining.

4. Run the gel at 4 V/cm with constant recirculation of running buffer for ∼3 hr or until bromphenol blue dye has migrated one-half to two-thirds the length of the gel. Recirculation is needed to prevent an H+ gradient forming in the buffer. If a gradient forms, the pH in parts of the gel may rise to >8.0, resulting in dissociation of the glyoxal from the RNA followed by renaturation. If no recirculation apparatus is available, electrophoresis should be paused every 30 min and the tank shaken to remix the buffer.

5. Remove the gel, cut off lanes, and stain with ethidium bromide (see Basic Protocol 1, steps 7a and 8). The RNA transfer (using the remaining portion of the gel) should be set up as soon as the gel is cut, before starting the staining.

Carry out northern transfer and hybridization analysis 6. Transfer RNA (see Basic Protocol 1, steps 9 to 24). 7. Immediately before hybridization, soak the membrane in 20 mM Tris⋅Cl (pH 8.0) for 5 min at 65°C to remove glyoxal. 8. Continue with hybridization analysis (see Basic Protocol 1, steps 25 to 36). NORTHERN HYBRIDIZATION OF UNFRACTIONATED RNA IMMOBILIZED BY SLOT BLOTTING


RNA slot blotting is a simple technique that allows immobilization of unfractionated RNA on a nylon or nitrocellulose membrane. Hybridization analysis is then carried out to determine the relative abundance of target mRNA sequences in the blotted samples. The technique is based on the DNA dot- and slot-blotting procedure (UNIT 2.9B), the main difference being the way in which the samples are denatured prior to immobilization. RNA dot blots can be prepared by hand but slot blots constructed using a manifold apparatus are preferable because the slots make it easier to compare hybridization signals by densitometry scanning. Additional Materials (also see Basic Protocol 1) 0.1 M NaOH 10× SSC (APPENDIX 2) 20× SSC (APPENDIX 2), room temperature and ice-cold Denaturing solution (see recipe) 100 mM sodium phosphate, pH 7.0 (see recipe) Dimethyl sulfoxide (DMSO) 6 M (40%) glyoxal, deionized immediately before use (see recipe) Manifold apparatus with a filtration template for slot blots (e.g., Bio-Rad Bio-Dot SF, Schleicher and Schuell Minifold II) 50° and 60°C water baths NOTE: All solutions should be prepared with sterile deionized water that has been treated with DEPC as described in UNIT 4.1; see unit introduction for further instructions and precautions regarding establishment of an RNase-free environment. Preparation and Analysis of RNA


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Set up membrane for transfer 1. Clean the manifold with 0.1 M NaOH and rinse with distilled water. 2. Cut a piece of nylon or nitrocellulose membrane to the size of the manifold. Pour 10× SSC (for nylon membrane) or 20× SSC (for nitrocellulose membrane) into a glass dish; place membrane on top of liquid and allow to submerge. Leave for 10 min. Avoid handling nitrocellulose and nylon membranes even with gloved hands—use clean blunt-ended forceps instead.

3. Place the membrane in the manifold. Assemble the manifold according to manufacturer’s instructions and fill each slot with 10× SSC. Ensure there are no air leaks in the assembly. Denature RNA samples 4a. Add 3 vol denaturing solution to RNA sample. Incubate 15 min at 65°C, then place on ice. Up to 20 ìg of RNA can be applied per slot. Total cellular RNA (UNITS 4.1-4.4) or poly(A)+ RNA (UNIT 4.5) can be used, although the latter is preferable (see Commentary).

4b. Alternatively, mix: 11 µl RNA sample 4.5 µl 100 mM sodium phosphate, pH 7.0 22.5 µl DMSO 6.6 µl 6 M glyoxal. Mix by vortexing, spin briefly in a microcentrifuge to collect liquid, and incubate 1 hr at 50°C. 5. Add 2 vol ice-cold 20× SSC to each sample. Pass samples through manifold 6. Switch on the suction to the manifold device and allow the 10× SSC added in step 3 to filter through. Leave the suction on. The suction should be adjusted so that 500 ìl buffer takes ∼5 min to pass through the membrane. Higher suction may damage the membrane. Slots that are not being used can be blocked off by placing masking tape over them or by applying 500 ìl of 3% (w/v) gelatin to each one. The former method is preferable as use of gelatin may lead to a background signal after hybridization. Alternatively, keep all slots open and apply 10× SSC instead of sample to the slots not being used.

7. Load each sample to the slots and allow to filter through, being careful not to touch the membrane with the pipet tip. 8. Add 1 ml of 10× SSC to each slot and allow to filter through. Repeat. 9. Dismantle the apparatus, place the membrane on a sheet of Whatman 3MM paper, and allow to dry. Immobilize RNA and carry out hybridization 10. Immobilize the RNA (see Basic Protocol 1, step 23). If glyoxal/DMSO denaturation has been used, immediately before hybridization soak the membrane in 20 mM Tris⋅Cl (pH 8.0) for 5 min at 65°C to remove glyoxal. Analysis of RNA by Northern and Slot Blot Hybridization

11. Carry out hybridization analysis as described in steps 25 to 36 of Basic Protocol 1.



REMOVAL OF PROBES FROM NORTHERN BLOTS Hybridization probes can be removed from northern blots on nylon membranes without damage to the membrane or loss of the transferred RNA. Some probes (particularly RNA probes) are more resistant to stripping. In these cases, higher temperatures, longer incubation periods, or the inclusion of formamide may be necessary for complete probe removal. The following stripping procedures are appropriate for both radioactive and chemiluminescent probes. Begin with the mildest conditions (step 1a) and monitor results to determine the extent of stripping. If the hybridization signal is still evident, proceed with the more stringent treatments (steps 1b and 1c) until stripping is complete.

SUPPORT PROTOCOL

Materials Northern hybridization membrane containing probe (see Basic Protocol 1, Alternate Protocol 1, or Alternate Protocol 2) Stripping solution (see recipe) Hybridization bags 65°, 80°, or 100° (boiling) water bath UV-transparent plastic wrap (e.g., Saran Wrap or other polyvinylidene wrap) Additional reagents and equipment for autoradiography (APPENDIX 3A) CAUTION: If hybridization probes include a radioactive label, dispose of stripping solutions as radioactive waste. Observe appropriate caution when working with the toxic compound formamide. 1a. To remove probes at 80°C: Place membrane in a hybridization bag containing stripping solution without formamide. Place bag in water preheated to 80°C for 5 min. Pour out solution, then repeat this washing process three to four times. Add sufficient solution to cover the membrane completely when using a bag; alternatively, stripping can be done in an open container, again with sufficient solution to cover the membrane.

1b. To remove probes at 100°C: Place membrane in a hybridization bag containing stripping solution without formamide. Place bag in boiling water for 5 min. Pour out solution, then repeat this washing process three to four times. 1c. To remove probes with formamide: Place membrane in a hybridization bag containing stripping solution with formamide. Place bag in water preheated to 65°C for 5 min. Pour out solution, then repeat this washing process three times using stripping solution with formamide and once using stripping solution without formamide. 2. Place membrane on filter paper to remove excess solution. Wrap membrane in plastic wrap and perform autoradiography to verify probe removal. If a chemiluminscent probe was used, verify probe removal by chemiluminscent detection (UNIT 3.19). The membrane may be immediately rehybridized or air-dried and stored for future use.

NORTHERN HYBRIDIZATION OF SMALL RNA FRACTIONATED BY POLYACRYLAMIDE GEL ELECTROPHORESIS This protocol is adapted for analysis of small RNAs. The major differences between this procedure and the traditional northern hybridization procedure are the fractionation system and the transfer system applied. Fractionation using denaturing polyacrylamide gel electrophoresis (PAGE) allows better separation of small RNAs. The introduction of a semidry transfer system reduces the time of the experimental procedure to 2 days.

BASIC PROTOCOL 2

Preparation and Analysis of RNA


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Tissue or cell samples TRIzol reagent (Invitrogen) RNase-free H2O (UNIT 4.1) 15% denaturing polyacrylamide sequencing (urea/TBE) gel (UNIT 7.6) 0.5× TBE electrophoresis buffer (APPENDIX 2) Formamide loading dye (see recipe) 2× SSC (APPENDIX 2) 50 µM probe oligonucleotide (DNA or RNA; UNIT 2.11) in RNase-free H2O ≥10 mCi/ml [γ-32P]ATP (6000 Ci/mmol; ICN Biomedicals) 10× T4 polynucleotide kinase buffer (New England Biolabs) 200 U/µl T4 polynucleotide kinase (New England Biolabs) Prehybridization/hybridization solution (see recipe), prewarmed to 37°C 2× SSC (APPENDIX 2) containing 0.1× (w/v) SDS, prewarmed to 37°C 95°C heating block or water bath Hybond N+ Nylon Transfer Membrane (Amersham Biosciences) Extra-thick blotting paper (Bio-Rad), slightly larger than the gel being blotted Semi-dry transfer apparatus (e.g., Bio-Rad Trans-Blot SD cell) Sephadex G-25 spin column Hybridization oven with rotating glass hybridization bottles, 37°C Image-analysis software (also see UNIT 10.5): e.g., QuantityOne (Bio-Rad) or ImageGauge (Fuji) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (UNIT 7.6), phosphor imaging (APPENDIX 3A), and digital electrophoresis analysis (UNIT 10.5) Prepare RNA sample and run the gel 1. Isolate total RNA from tissue or cell samples with TRIzol reagent according to the manufacturer’s instructions. At the end of the procedure, dissolve the RNA to a final concentration of 10 µg/µl in RNase-free water. 2. Prerun 15% denaturing polyacrylamide sequencing (urea/TBE) gel for 15 min at 25 W in 0.5× TBE electrophoresis buffer. Mix 5 µl RNA sample (10 µg/µl) with equal volume of formamide loading dye. Heat 2 min at 95°C, and load onto gel (UNIT 7.6). 3. Run the gel at 25 W until the bromophenol blue dye has migrated to the bottom of the gel. Transfer RNA from gel to membrane 4. Cut a piece of Hybond N+ nylon membrane slightly larger than the gel. Soak the membrane and four pieces of blotting paper of appropriate size in 0.5× TBE buffer for 10 min. 5. Stack two pieces of blotting paper on the anode platform of the transfer cell. Avoid getting air bubbles under or between the papers; remove any that appear by carefully rolling a glass pipet over the surface. 6. Place the membrane on top of the blotting paper and squeeze out air bubbles by rolling a glass pipet over the surface. 7. Carefully transfer the gel from glass plate to the top of the membrane and squeeze out air bubbles. 8. Stack another two pieces of blotting paper on the gel and squeeze out air bubbles. Analysis of RNA by Northern and Slot Blot Hybridization

9. Set the cathode assembly and the safety lid on the sandwich. Transfer for 1 hr at 300 mA.



Prepare membrane for hybridization 10. Disassemble the transfer cell. Remove the paper and the gel. Rinse the membrane in 2× SSC, then place it on a sheet of filter paper and allow it to air dry. 11. Place membrane RNA-side-down on a UV transilluminator (254-nm wavelength) or in a UV light box for the appropriate length of time to covalently attach the RNA to the membrane. Prepare probe The authors typically use a chemically synthesized 21-22 nt DNA or RNA oligonucleotide (see UNIT 2.11 for oligonucleotide synthesis) perfectly complementary to the small RNA to be detected. 12. Set up the 5′-end-labeling reaction by combining the following reagents: 1 µl of 50 µM probe oligonucleotide (DNA or RNA) 1 µl of [γ- 32P]ATP (6000 Ci/mmol, ≥10 mCi/ml) 4 µl of 10× T4 polynucleotide kinase buffer H2O to a final volume of 40 µl 1 µl of 200 U/µl T4 polynucleotide kinase. Incubate reaction 1 hr at 37°C. 13. Pass reaction mixture through Sephadex G-25 spin column (centrifuging per manufacturer’s instructions) to remove unincorporated [γ- 32P]ATP. Perform prehybridization and hybridization 14. Place membrane (from step 11) RNA-side-up in a hybridization bottle and add ∼1 ml prewarmed (37°C) prehybridization/hybridization solution per 10 cm2 of membrane. Place the bottle in a hybridization oven and incubate with rotation for 30 min at 37°C. 15. Pipet the entire reaction mix (from step 13) into the hybridization bottle and continue to incubate with rotation overnight at 37°C. Wash membrane 16. Pour off hybridization solution and wash the membrane briefly with 2× SSC for 5 min. Add prewarmed (37°C) 2× SSC containing 0.1% SDS to the bottle. Incubate with rotation for 15 min at 37°C. Replace solution with fresh solution and repeat. Remove final wash solution, blot excess liquid, and wrap with plastic wrap. Do not allow membrane to dry out if it is to be reprobed.

Perform phosphor imaging and analyze the hybridization signals 17. Visualize the hybridization signals by phosphor imaging (APPENDIX 3A). Analyze hybridization result using appropriate software (UNIT 10.5). Subtract the background from the original signal to obtain the specific hybridization. 18. To compare the amount of small RNA in different samples, normalize the amount of small RNA detected to the nonspecific hybridization of the probe to 5S rRNA. Alternatively, the blot can be reprobed with a probe specific for 5S rRNA (for Drosophila 5′-CAA CAC GCG GTG TTC CCA AGC CG-3′) or, for Drosophila, the 2S RNA (5′-TAC AAC CCT CAA CCA TAT GTA GTC CAA GCA-3′). The precise amount (pmol or molecules) of small RNA can be determined if concentration standards of synthetic RNA are included on the blot. The assay is typically linear with respect to concentration over a 10,000-fold range.



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NORTHERN HYBRIDIZATION OF RNA USING CHURCH’S HYBRIDIZATION BUFFER Church’s hybridization buffer can be used as an alternative for the standard prehybrization/hybridization solution in this assay. It provides similar sensitivity. Additional Materials (also see Basic Protocol 2) Church’s hybridization buffer without BSA (see recipe) 1. Perform northern blotting and prepare probe (see Basic Protocol 2, steps 1 through 13). 2. Perform prehybridization and hybridization (see Basic Protocol 2, steps 14 to 15) using Church’s hybridization buffer (without BSA) in place of the prehybridization/hybridization solution. 3. Wash membrane and proceed with development and analysis (see Basic Protocol 2, steps 16 to 18). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Church’s hybridization buffer without BSA 0.5 M sodium phosphate buffer, pH 7.2 (APPENDIX 2) 1 mM EDTA, pH 8.0 (APPENDIX 2) 7% (w/v) SDS Store up to 1 year at room temperature Denaturing solution 500 µl formamide 162 µl 12.3 M (37%) formaldehyde 100 µl MOPS buffer (see recipe) Make fresh from stock solutions immediately before use If formamide has a yellow color, deionize as follows: add 5 g of mixed-bed ion-exchange resin (e.g., Bio-Rad AG 501-X8 or X8(D) resins) per 100 ml formamide, stir 1 hr at room temperature, and filter through Whatman #1 filter paper. CAUTION: Formamide is a teratogen. Handle with care.

Formaldehyde loading buffer 1 mM EDTA, pH 8.0 (APPENDIX 2) 0.25% (w/v) bromphenol blue 0.25% (w/v) xylene cyanol 50% (v/v) glycerol Store up to 3 months at room temperature Formamide loading dye 98% (v/v) deionized formamide 10 mM EDTA pH 8.0 (APPENDIX 2) 0.025% (w/v) xylene cyanol 0.025% (w/v) bromphenol blue Store indefinitely at −20°C


Glyoxal, 6 M, deionized Immediately before use, deionize glyoxal by passing through a small column of mixed-bed ion-exchange resin (e.g., Bio-Rad AG 501-X8 or X8(D) resins) until the pH is >5.0.



Glyoxal loading buffer 10 mM sodium phosphate, pH 7.0 (see recipe) 0.25% (w/v) bromphenol blue 0.25% (w/v) xylene cyanol 50% (v/v) glycerol Store up to 3 months at room temperature MOPS buffer 0.2 M MOPS [3-(N-morpholino)-propanesulfonic acid], pH 7.0 0.5 M sodium acetate 0.01 M EDTA Store up to 3 months at 4°C Store in the dark and discard if it turns yellow.

MOPS running buffer, 10× 0.4 M MOPS, pH 7.0 0.1 M sodium acetate 0.01 M EDTA Store up to 3 months at 4°C Prehybridization/hybridization solution 5× SSPE (see recipe) 5× Denhardt solution (APPENDIX 2) 50% (v/v) formamide 0.5% (w/v) SDS 72 µg/ml denatured herring sperm DNA (Promega) Make fresh from stock solutions immediately before use The herring sperm DNA is denatured by heating 10 min at 75°C just before it is added.

Sodium phosphate, pH 7.0, 100 mM and 10 mM 100 mM stock solution: 5.77 ml 1 M Na2HPO4 4.23 ml 1 M NaH2PO4 H2O to 100 ml Store up to 3 months at room temperature 10 mM solution: Dilute 100 mM stock 1/10 with H2O Store up to 3 months at room temperature SSPE, 10× 1.5 M NaCl 50 mM NaH2PO4⋅H2O 5 mM EDTA Store indefinitely at room temperature Stripping solution 1% (w/v) SDS 0.1× SSC (APPENDIX 2) 40 mM Tris⋅Cl, pH 7.5 to 7.8 (APPENDIX 2) Store up to 1 year at room temperature Where formamide stripping is desired, prepare the above solution and add an equal volume of formamide just before use. Preparation and Analysis of RNA


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COMMENTARY Background Information The development of Southern blotting (UNIT 2.9A; Southern, 1975) was quickly followed by an equivalent procedure for the immobilization of gel-fractionated RNA (Alwine et al., 1977). The term northern blotting, initially used in a humorous fashion, has become enshrined in molecular biology jargon. Northern hybridization is a standard procedure for identification and size analysis of RNA transcripts and RNA slot blotting is frequently used to assess the expression profiles of tissue-specific genes (Kafatos et al., 1979). Procedures for the removal of hybridization probes from northern blots are similar to those for Southern blots, except that NaOH is omitted to prevent hydrolysis of the RNA, and formamide may be included. The recent discovery of microRNAs (miRNAs) revealed an entire new class of molecules that regulate gene expression. miRNAs are small noncoding RNAs that range from 20 to 30 nucleotides, making traditional formaldehyde-agarose gel electrophoresis unsuitable for their size fractionation. The modified northern protocol here (Basic Protocol 2) combines denaturing polyacrylamide gel electrophoresis (PAGE), which is ideal for the separation of small RNAs, with standard blotting and hybridization procedures.

Critical Parameters


Gel electrophoresis and northern blotting The main distinction between northern and Southern blotting lies with the initial gel fractionation step. Because single-stranded RNA can form secondary structures, samples must be electrophoresed under denaturing conditions to ensure good separation. A variety of denaturants for RNA gels have been used, including formaldehyde (Basic Protocol 1; Lehrach et al., 1977), glyoxal/DMSO (Alternate Protocol 1; Thomas, 1980), and the highly toxic methylmercuric chloride (Bailey and Davidson, 1976). Because of the substantial health risks, use of methylmercuric chloride is not advised. Formaldehyde gels are recommended, as they are easy to run and reasonably reliable. The formaldehyde must be rinsed from the gel before the transfer is set up, but this is a minor inconvenience compared to assembling the buffer recircularization system required for electrophoresis of glyoxal-denatured RNA.

Total cellular RNA (UNITS 4.1-4.4) or poly(A)+ RNA (UNIT 4.5) can be used for northern transfers and slot blots. Total RNA is less satisfactory because nonspecific hybridization, however slight, to one or both of the highly abundant rRNA molecules will lead to a substantial hybridization signal. Any hybridizing band that appears in the vicinity of an rRNA should be treated with suspicion and its identity confirmed by blotting with poly(A)+ RNA. Under ideal conditions, a band that contains as little as 1 pg of RNA can be detected by northern hybridization with a probe labeled to a specific activity of 109 dpm/µg. In practice, the effective detection limit with an overnight exposure is ∼5 pg RNA. An mRNA is usually considered to be abundant if it constitutes >1% of the mRNA fraction. In a typical mammalian cell, the mRNA fraction makes up about 0.5% of total RNA, so >5 pg of an abundant mRNA should be present in just 100 ng of total RNA. If 10 µg of total RNA is transferred, abundant mRNAs should give strong hybridization signals and less abundant ones (down to 0.01% of the mRNA population) should be detectable with an overnight exposure. For rarer molecules, the poly(A)+ fraction must be prepared. In this sample 3 µg is sufficient for detecting an mRNA that makes up 0.0002% of the polyadenylated population. Unlike probing for mRNA, which often requires enrichment by poly(A) selection prior to analysis, total cellular RNA can always be used for the detection of miRNA, because individual miRNA species can be present in thousands to tens of thousands of copies per cell. The highly abundant rRNA and microRNAs are well separated in a 15% denaturing polyacrylamide gel; thus, the nonspecific hybridization to rRNA will not affect the interpretation of the desired hybridizing signal. RNA slot blots Although easy to perform, RNA slot-blot hybridization is one of the most problematic techniques in molecular biology. A number of criteria must be satisfied if slot blotting is to be used to make meaningful comparisons of mRNA abundance in different extracts. The first requirement is that equal amounts of RNA must be loaded in each slot. In practice this is difficult to achieve, especially if RNA concentrations are estimated by absorbance spectroscopy (APPENDIX 3D), which is subject to errors



due to the small quantities being measured and the presence of contaminants such as protein and DNA. Even if equal amounts of RNA are loaded, a difference in hybridization signal does not necessarily mean that the gene whose transcript is being studied is more active in a particular tissue. The analysis provides information on the abundance of an mRNA (i.e., the fraction of total RNA that it constitutes), not its absolute amount. To illustrate this point, consider a tissue in which a highly active gene is switched on at time t, where the transcripts of this gene constitute 0% of the mRNA at t − 1 but 20% of the mRNA at t + 1. If the slot blots of RNA from t − 1 and t + 1 are probed with the highly active gene, there will be a clear increase in hybridization signal after time t. In contrast, hybridization of the same slot blots with a second gene whose transcription rate is unchanged will show a decreased hybridization signal at t + 1. Transcripts of this gene are present in the same absolute amounts at t − 1 and t + 1, but their abundance decreases as the total mRNA population becomes larger due to activation of the highly expressed gene. To the unwary, the result of the hybridization analysis could appear to indicate down-regulation of a gene whose expression rate in fact remains constant. Choice of membrane and transfer system General information relating to the choice of membrane for a nucleic acid transfer is given in the Commentary to UNIT 2.9A. Because of the greater tensile strength of nylon, together with the fact that the RNA can be bound covalently by UV cross-linking, most transfers are now carried out using nylon rather than nitrocellulose. Nylon has the added advantage of being able to withstand the highly stringent conditions (50% formamide at 60°C) that may be required during hybridization with an RNA probe; nitrocellulose tends to disintegrate under these conditions. For DNA transfer, a major advantage of positively charged nylon is that nucleic acids become covalently bound to the membrane if the transfer is carried out with an alkaline buffer. RNA can also be immobilized on positively charged nylon by alkaline transfer, but the procedure is not recommended as the alkaline conditions result in partial hydrolytic degradation of the RNA. This hydrolysis is difficult to control and smaller molecules are easily broken down into fragments too short for efficient retention by the membrane (see Table 2.9.1). This results in a loss of signal after hybridiza-

tion, a problem that is exacerbated by the increased background caused by lengthy exposure of the membrane to the alkaline solution. Only if the signals are expected to be strong should an alkaline transfer be considered. In this case, Basic Protocol 1 should be modified as follows: omit the pre-transfer alkaline hydrolysis (step 10), use 8 mM NaOH rather than 20× SSC as the transfer buffer, do not transfer for more than 6 hr, and rinse the membrane in 2× SSC/0.1% (w/v) SDS rather than plain SSC immediately after transfer (step 22). If using Alternate Protocol 1, omit step 7 as well, as the alkaline transfer buffer removes glyoxal from the RNA. The standard northern transfer system can be modified as described for Southern blotting (UNIT 2.9A). Aqueous transfers onto nylon can be performed using a variety of buffers, although SSC is still most frequently used. Changes can be made to the transfer time and architecture of the blot (e.g., downward transfer; Chomczynski, 1992), and alternative methods such as electroblotting (Smith et al., 1984) and vacuum transfer (Peferoen et al., 1982) can be used. Hybridization procedures Hybridization analysis of an RNA blot is subject to the same considerations as DNA hybridization (see UNIT 2.10 Commentary). The factors that influence sensitivity and specificity are the same, and incubation times, hybridization solutions, probe length, and mechanics of hybridization all have similar effects. There are just two additional points that need to be made with respect to RNA blots. The first point is that formamide is almost always used in RNA hybridization solutions. The primary reason for this is to permit a lower hybridization temperature to be used, minimizing RNA degradation during the incubations. The second point concerns the stability of the hybrids formed between the immobilized RNA and the probe molecules. For a DNA probe the relevant equation is (Casey and Davidson, 1977): Tm = 79.8°C + 18.5(log M) + 0.58(%GC) + 11.8(%GC)2 − 0.50(%form) − 820⁄L

and for an RNA probe (Bodkin and Knudson, 1985): Tm = 79.8°C + 18.5(log M) + 0.58(%GC) + 11.8(%GC)2 − 0.35(%form) − 820⁄L

where Tm is the melting temperature, M is the molarity of monovalent cations, %GC is the



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percentage of guanosine and cytosine nucleotides in the DNA, %form is the percentage of formamide in the hybridization solution, and L is the length of the hybrid in base pairs. What these equations indicate is that an RNA-RNA hybrid is more stable than a DNA-RNA hybrid: if %form is 50%, the Tm for an RNA-RNA hybrid is 7.5°C higher than that for an equivalent RNA-DNA hybrid. The greater stability of the RNA-RNA hybrid means that an RNA probe requires a more stringent hybridization and washing regime than a DNA probe (e.g., hybridization at 60°C in 50% formamide and final wash at 68°C in 0.1× SSC/0.1% SDS). Reprobing conditions Nitrocellulose presents problems regarding both membrane integrity and RNA retention (see UNIT 2.10). UV cross-linking of RNA to a neutral nylon membrane presents optimal conditions for northern blot reprobing. Because of the sensitivity of RNA to alkaline hydrolysis, NaOH, which is included in protocols for probe removal from Southern blots, should not be used when removing probes from northern blots. It is recommended that probes be removed prior to membrane storage, because unstripped probes remain permanently attached if the blot dries.

Troubleshooting


The appearance of the agarose gel after staining gives a first indication of how successful a northern experiment is likely to be. If total RNA has been used, the rRNA bands should be clear and sharp (Fig. 4.9.1) with no “smearing” toward the positive electrode. The only exception is when the RNA has been prepared by the guanidinium isothiocyanate procedure (UNIT 4.2), in which case some smearing is normal. If the rRNA bands are not sharp, the RNA preparation may be of poor quality (usually because insufficient care has been taken in establishing an RNase-free environment) or the denaturing gel electrophoresis system may not have worked adequately. If the latter problem is suspected, make sure that the formaldehyde concentration in the gel is 2.2 M or, if glyoxal denaturation has been used, that the buffer recircularization is sufficient to maintain the gel pH at 7.0. Whatever the problem, if the rRNAs are not distinct, there is no point in proceeding with the transfer as the bands obtained after hybridization will also be fuzzy. In fact, even if the rRNA bands are clear there is no guarantee that the mRNAs are intact.

An indication of the efficiency of transfer onto nylon can be obtained by staining the membrane with methylene blue (see Basic Protocol 1, step 24), but often a problem with transfer is not recognized until after hybridization. If poor signals are obtained, the troubleshooting section of UNIT 2.10 (including Table 2.10.4) should be consulted to identify the likely cause. Note that it is relatively easy to detach RNA from a membrane before immobilization, so some loss may occur when the membrane is rinsed in 2× SSC to wash off agarose fragments and leach out salt (see Basic Protocol 1, step 22). If necessary, this rinse can be postponed until immediately before hybridization, after the RNA has been immobilized. Other problems, such as high backgrounds, extra bands, and difficulties with probe stripping, should be dealt with by referring to Table 2.10.4. To increase the sensitivity of the miRNA assay, RNA probes can be used instead of DNA probes. In some cases, RNA probes work better to detect miRNA precursors (∼60 to 70 nt long) which contain the immature miRNAs in a stemloop whose structure can prevent the hybridization of DNA probes.

Anticipated Results Using either a nylon or nitrocellulose membrane and a probe labeled to ≥5× 108 dpm/µg, it should be possible to detect transcripts that represent 0.01% of the mRNA population with a blot of 10 µg total mammalian RNA or 0.0002% of the population with a blot of 3 µg poly(A)+ RNA. It should be possible to detect small RNAs at levels as low as 0.3 fmol miRNA by following the Basic Protocol 2. Small RNAs that differ by as little as 1 nt (or even by a single phosphate group) can be separated on 15% denaturing polyacrylamide gel (50 to 100 cm), particularly when a long gel is used. For long gels, only the lower portion of the gel is used for transfer to the membrane.

Time Considerations Traditional blots A northern experiment can be completed in 3 days. The agarose gel is prepared and electrophoresed during the first day and the transfer carried out overnight. On the second day the blot is prehybridized and then hybridized overnight. Washes are completed early on the third day. A slot-blot experiment takes only two days, as the blot can be prepared and prehybridized



on the first day, hybridized overnight, and washed on the second day. The length of time needed for the autoradiography depends on the abundance of the target sequences in the blotted RNA. Adequate exposure can take anything from overnight to several days. With blots that are intended for reprobing, stripping procedures can be completed in ∼1 hr, not including the verification steps. miRNA blots Using a semidry transfer system, the northern experiment can be completed in 2 days. The length of time needed for the autoradiography depends on the abundance of the small RNA sequence and the detection system used. Optimal exposure can range from overnight to several days. Under ideal conditions, a band that contains as little as 0.3 fmol of small RNA can be detected by northern hybridization with a 1-day exposure to a phosphor imager plate when scanned at 25 µm resolution. The amount of RNA loaded and the exposure time may vary with the abundance of the individual miRNA species. For abundant miRNAs, loading of 5 µg total RNA and overnight exposure will give strong hybridization signals.

Literature Cited Alwine, J.C., Kemp., D.J., and Stark, G.R. 1977. Method for detection of specific RNAs in agarose gels by transfer to diazobenzyloxymethyl-paper and hybridization with DNA probes. Proc. Natl. Acad. Sci. U.S.A. 74:5350-5354. Bailey, J.M. and Davidson, N. 1976. Methylmercury as a reversible denaturing agent for agarose gel electrophoresis. Anal. Biochem. 70:75-85. Bodkin, D.K. and Knudson, D.L. 1985. Assessment of sequence relatedness of double-stranded RNA genes by RNA-RNA blot hybridization. J. Virol. Methods 10:45-52. Casey, J. and Davidson, N. 1977. Rates of formation and thermal stabilities of RNA:DNA and DNA:DNA duplexes at high concentrations of formamide. Nucl. Acids Res. 4:1539-1552.

Chomczynski, P. 1992. One-hour downward alkaline capillary transfer for blotting of DNA and RNA. Anal. Biochem. 201:134-139. Herrin, D.L. and Schmidt, G.W. 1988. Rapid, reversible staining of Northern blots prior to hybridization. BioTechniques 6:196-200. Kafatos, F.C., Jones, C.W., and Efstratiadis, A. 1979. Determination of nucleic acid sequence homologies and relative concentrations by a dot hybridization procedure. Nucl. Acids Res. 7:1541-1552. Lehrach, H., Diamond, D., Wozney, J.M., and Boedtker, H. 1977. RNA molecular weight determinations by gel electrophoresis under denaturing conditions: A critical reexamination. Biochemistry 16:4743-4751. Peferoen, M., Huybrechts, R., and De Loof, A. 1982. Vacuum-blotting: A new simple and efficient transfer of proteins from sodium dodecyl sulfate–polyacrylamide gels to nitrocellulose. FEBS Lett. 145:369-372. Smith, M.R., Devine, C.S., Cohn, S.M., and Lieberman, M.W. 1984. Quantitative electrophoretic transfer of DNA from polyacrylamide or agarose gels to nitrocellulose. Anal. Biochem. 137:120124. Southern, E.M. 1975. Detection of specific sequences among DNA fragments separated by gel electrophoresis. J. Mol. Biol. 98:503-517. Thomas, P.S. 1980. Hybridization of denatured RNA and small DNA fragments transferred to nitrocellulose. Proc. Natl. Acad. Sci. U.S.A. 77:5201-5205. Wilkinson, M. 1991. Purification of RNA. In Essential Molecular Biology: A Practical Approach, Vol. 1 (T.A. Brown, ed.) pp. 69-87. IRL Press, Oxford.

Contributed by Terry Brown University of Manchester Institute of Science and Technology Manchester, United Kingdom Karol Mackey (probe removal) Molecular Research Cincinnati, Ohio Tingting Du (miRNA blots) University of Massachusetts Medical School Worcester, Massachusetts



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UNIT 4.10

Identification of Newly Transcribed RNA Newly transcribed RNA can be identified using the nuclear runoff transcription assay. Isolated nuclei, free of membranes and cytoplasmic debris, are required for the assay. Cell lysis that does not allow the isolation of nuclei free of cell membranes and cytoplasmic material often results in poor incorporation of 32P-labeled UTP into nascent transcripts. Although there is no way to predict what cell types present this problem, many adherent cell lines and lymphocytes isolated from murine spleen or thymus do. Interestingly, very few nonadherent cell lines have posed this problem. Isolating nuclei by detergent lysis of cells (basic protocol) works well for many tissue culture cell lines but may not be appropriate for all cell lines and many tissues. Detergent lysis and Dounce homogenization (first alternate protocol) or cell lysis in an isoosmotic solution and centrifugation through a sucrose cushion (second alternate protocol) are alternative methods for preparing nuclei. The support protocol describes preparation of the cDNA nitrocellulose filter strips that are used to detect the presence of specific transcripts in the nuclear runoff transcription assay. NOTE: Experiments involving RNA require careful technique to prevent RNA degradation; see Chapter 4 Section I introduction.

BASIC PROTOCOL

NUCLEAR RUNOFF TRANSCRIPTION IN MAMMALIAN CELLS Nuclear runoff transcription is currently the most sensitive procedure for measuring specific gene transcription as a function of cell state. Nuclei are first isolated from cultured cells or tissues and frozen in liquid nitrogen. Thawed nuclei are incubated with 32P-labeled UTP and unlabeled NTPs to label nascent RNA transcripts. 32P-labeled RNA is purified and used to detect specific RNA transcripts by hybridization to cDNAs immobilized on nitrocellulose membranes. Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Identification of Newly Transcribed RNA


Cultures of mammalian cells (UNIT 9.0) or freshly isolated lymphoid cells Phosphate-buffered saline (PBS; APPENDIX 2), made fresh and ice cold Nonidet P-40 (NP-40) lysis buffer A (see recipe) Glycerol storage buffer (see recipe), ice cold 2× reaction buffer with and without nucleotides (see recipes) 10 mCi/ml [α-32P]UTP (760 Ci/mmol) 1 mg/ml DNase I (RNase-free; see recipe) HSB buffer (see recipe) SDS/Tris buffer (see recipe) 20 mg/ml proteinase K 25:24:1 (v/v/v) buffered phenol/chloroform/isoamyl alcohol (UNIT 2.1) 10% (v/v) trichloroacetic acid (TCA)/60 mM sodium pyrophosphate 10 mg/ml tRNA (UNIT 4.6) 5% (v/v) TCA/30 mM sodium pyrophosphate DNase I buffer (see recipe) 0.5 M EDTA, pH 8.0 20% (w/v) SDS Elution buffer (see recipe) 1 M NaOH 1 M HEPES (free acid) Contributed by Michael E. Greenberg and Timothy P. Bender Current Protocols in Molecular Biology (1997) 4.10.1-4.10.11 Copyright © 1997 by John Wiley & Sons, Inc.

3 M sodium acetate, pH 5.2 100% ethanol TES solution (see recipe) TES/NaCl solution (see recipe) cDNA plasmid(s) immobilized on nitrocellulose membrane (support protocol) 2× SSC (APPENDIX 2) 10 mg/ml heat-inactivated RNase A (UNIT 7.3) Rubber policeman 15- and 50-ml conical polypropylene centrifuge tubes Beckman JS-4.2 and JA-20 rotors or equivalent 30° and 65°C shaking water baths 42° and 65°C water baths 0.45-µm HA filters (Millipore) 30-ml Corex tube, silanized (APPENDIX 3) Whatman GF/F glass fiber filters 5-ml plastic scintillation vials Whatman 3MM filter paper NOTE: Keep cells and nuclei on ice until the nuclei are frozen. Isolate nuclei 1a. For cultures of adherent cells: Remove medium from monolayer cultures (5 × 107 cells per assay) and place cells on ice. Rinse twice with 5 ml ice-cold PBS. Scrape flask with a rubber policeman and collect cells in a 15-ml centrifuge tube. Centrifuge 5 min at 500 × g (1500 rpm in JS-4.2 rotor), 4°C. Remove supernatant. 1b. For cultures of nonadherent cells: Pipet up and down several times to resuspend cells (5 × 107 cells per assay) and transfer cells and medium to a 50-ml conical centrifuge tube. Centrifuge 5 min at 500 × g (1500 rpm in JS-4.2 rotor), 4°C, and remove supernatant. Wash by gently resuspending pellet in 5 ml ice-cold PBS, adding 45-ml ice-cold PBS, and collecting cells by centrifuging 5 min at 500 × g. Remove supernatant. Wash cells once more with PBS and remove supernatant. 1c. For freshly isolated lymphoid cells: Transfer lymphoid cells (5 × 107 cells per assay) removed directly from organ to a 50-ml conical centrifuge tube. Centrifuge 5 min at 500 × g (1500 rpm in JS-4.2 rotor), 4°C, and remove supernatant. Wash by resuspending pellet in 5 ml ice-cold PBS, adding 45 ml PBS, and collecting cells by centrifuging 5 min at 500 × g, 4°C. Remove supernatant. Wash cells once more with PBS and remove supernatant. Lymphoid cell nuclei are more fragile than other cell types, so a gentle procedure (e.g., second alternate protocol) may be required to isolate intact nuclei. It is not necessary to eliminate erythrocytes from lymphoid cells prior to preparation of nuclei. 5 × 107 cells are required for each nuclear runoff transcription assay.

2. Loosen cell pellet by gently vortexing 5 sec. Add 4 ml NP-40 lysis buffer A, continuing to vortex as buffer is added. After lysis buffer is completely added, vortex cells 10 sec at half maximal speed. Gentle vortexing (at a setting of six) uniformly resuspends cells and inhibits clumping. The same method is used to resuspend nuclei.

3. Incubate lysed cells 5 min on ice. Examine a few microliters of cell lysate on a hemacytometer with a phase-contrast microscope to ensure that cells have uniformly lysed and nuclei appear free of cytoplasmic material. Centrifuge 5 min at 500 × g, 4°C. Remove supernatant.



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Supernatant contains cytoplasmic RNA that can be purified as described in UNITS 4.1 & 4.5, if desired.

4. Resuspend the nuclear pellet in 4 ml NP-40 lysis buffer A by vortexing as described in step 2. Centrifuge 5 min at 500 × g, 4°C. Discard supernatant and resuspend nuclei in 100 to 200 µl glycerol storage buffer by gently vortexing. Freeze resuspended nuclei in liquid nitrogen. Nuclei are stable in liquid nitrogen for >1 year.

Perform nuclear runoff transcription 5. Thaw 200 µl frozen nuclei at room temperature and transfer to a 15-ml conical polypropylene centrifuge tube. Immediately add 200 µl of 2× reaction buffer with nucleotides plus 10 µl of 10 mCi/ml [α-32P]UTP. Incubate 30 min at 30°C with shaking. This reaction is done in a 15-ml polypropylene tube rather than a microcentrifuge tube to reduce the possibility of spilling radioactive materials.

6. Mix 40 µl of 1 mg/ml RNase-free DNase I and 1 ml HSB buffer. Add 0.6 ml of this solution to labeled nuclei and pipet up and down 10 to 15 times with a Pasteur pipet to mix thoroughly. Incubate 5 min at 30°C. 7. Add 200 µl SDS/Tris buffer and 10 µl of 20 mg/ml proteinase K. Incubate for 30 min at 42°C. DNA and protein should be well digested and a fairly uniform solution should be obtained. The presence of a substantial amount of particulate matter usually indicates that either DNase I or proteinase K treatment was not effective and should be repeated. It may be necessary first to ethanol precipitate the RNA, then to repeat the treatment with DNase I and proteinase K with fresh reagents.

Extract and precipitate RNA 8. Extract sample with 1 ml 25:24:1 buffered phenol/chloroform/isoamyl alcohol. Centrifuge 5 min at 800 × g (2000 rpm in JS-4.2 rotor), at or below room temperature. Transfer aqueous phase to a clean 15-ml polypropylene centrifuge tube. 9. Add 2 ml water, 3 ml of 10% TCA/60 mM sodium pyrophosphate, and 10 µl of 10 mg/ml E. coli tRNA carrier to aqueous phase. Incubate 30 min on ice. 10. Filter TCA precipitate onto 0.45-µm Millipore HA filter. Wash filter three times with 10 ml of 5% TCA/30 mM sodium pyrophosphate. If the HA filter clogs and filters very slowly, a Whatman GF/A glass fiber filter can be used instead.

11. Transfer filter to a glass scintillation vial. Incubate with 1.5 ml DNase I buffer and 37.5 µl of 1 mg/ml RNase-free DNase I for 30 min at 37°C. Quench the reaction by adding 45 µl of 0.5 M EDTA and 68 µl of 20% SDS. 12. Heat sample 10 min at 65°C to elute the RNA. Remove supernatant and save. Add 1.5 ml elution buffer to filter and incubate 10 min at 65°C. Remove supernatant and combine with original supernatant. This procedure removes >95% of the radioactivity from the filter. Identification of Newly Transcribed RNA

13. Add 4.5 µl of 20 mg/ml proteinase K to 3 ml supernatant containing 32P-labeled RNA. Incubate 30 min at 37°C.



14. Extract 3 ml RNA solution once with 3 ml of 25:24:1 buffered phenol/chloroform/isoamyl alcohol. 15. Remove aqueous phase to a silanized 30-ml Corex tube. Add 0.75 ml of 1 M NaOH to aqueous phase. Let stand 10 min on ice. Quench reaction by adding 1.5 ml of 1 M HEPES. 16. Precipitate RNA by adding 0.53 ml of 3 M sodium acetate and 14.5 ml of 100% ethanol. Incubate 30 min on dry ice or overnight at −20°C. 17. Centrifuge RNA 30 min at 10,000 × g (9000 rpm in JA-20 rotor), 4°C. Remove ethanol and resuspend pellet in 1 ml TES solution. Shake 30 min at room temperature. RNA should be completely dissolved.

18. Count a 5-µl aliquot of each sample in duplicate by spotting onto Whatman GF/F glass fiber filters. If necessary, dilute sample by adding TES solution to adjust 32 P-labeled RNA to ≥5 × 106 cpm/ml. Hybridize RNA to cDNA 19. Mix 1 ml RNA solution with 1 ml TES/NaCl solution. In a 5-ml plastic scintillation vial, hybridize to cDNA immobilized on nitrocellulose membrane strip for 36 hr at 65°C with shaking. Use a vial rather than a plastic bag for hybridization to ensure reproducible quantitative hybridization of the same number of cpm to each sample within a given experiment. Coil the nitrocellulose strip before inserting it into the scintillation vial and be sure the strip is completely immersed in hybridization solution.

20. After hybridization, transfer the strips to a 50-ml tube and wash filter in 25 ml of 2× SSC 1 hr at 65°C. Repeat wash once with fresh 2× SSC. 21. Remove filter to a glass scintillation vial containing 8 ml of 2× SSC and 8 µl of 10 mg/ml RNase A. Incubate without shaking 30 min at 37°C. 22. Wash filter once more in 25 ml of 2× SSC 1 hr at 37°C. Blot filter dry on Whatman 3MM filter paper. Unravel the strips, tape them to Whatman 3MM filter paper, and expose to X-ray film. Appropriate exposure time and conditions will vary depending on the experiment.

ISOLATION OF NUCLEI BY DOUNCE HOMOGENIZATION This protocol is used for isolation of nuclei from cell types that do not give clean nuclear preparations after lysis in NP-40 lysis buffer A. If cells are not lysed by treatment with NP-40 lysis buffer A (basic protocol), the addition of Dounce homogenization in NP-40 lysis buffer B is usually sufficient to lyse the cells.

ALTERNATE PROTOCOL


Lysis buffer (see recipe), ice cold Nonidet P-40 (NP-40) lysis buffer B (see recipe) Glycerol storage buffer (see recipe), ice cold Dounce homogenizer with type B pestle, ice cold 1.5-ml microcentrifuge tubes, chilled on dry ice NOTE: Keep cells and nuclei on ice until the nuclei are frozen.



Supplement 26

Harvest cells 1. Harvest and wash cells as in basic protocol step 1. 2. Remove supernatant and loosen cell pellet by gently vortexing 5 sec. Resuspend cell pellet to a single-cell suspension in 5 to 10 ml ice-cold lysis buffer. Add ice-cold lysis buffer to a total of 40 ml and rock tube back and forth for several seconds to distribute the cells. 3. Pellet cells at 500 × g (1500 rpm in JS-4.2 rotor), 4°C. Remove and discard supernatant. Resuspend pellet in 1 ml lysis buffer per 5 × 107 cells and vortex gently to mix. After centrifugation, the pellet should appear to be two to three times its initial size.

4. Add 1 ml NP-40 lysis buffer B per 5 × 107 cells and mix by gently rocking the tube. Break cells and collect nuclei 5. Transfer cells to an ice-cold Dounce homogenizer and break them with ten strokes of a B pestle or until nuclei appear free of membrane components by phase-contrast microscopy. 6. Transfer homogenized cells to a plastic 50-ml conical centrifuge tube and pellet nuclei by centrifugation 5 min at 500 × g, 4°C. Pellet should now be approximately one-third to one-half the starting volume and appear opaque white.

7. Carefully remove supernatant with a Pasteur pipet attached to a vacuum supply. Tilt the tube sideways so supernatant is pulled away from the pellet. Remove any bubbles or liquid that remain on the side of the tube. Return pellet to an ice bucket. 8. Loosen pelleted nuclei by gentle vortexing. Add 200 µl ice-cold glycerol storage buffer per 5 × 107 nuclei and resuspend pellet by pipetting up and down. Nuclei will be clumped at first but will disperse with continued pipetting. Pipetting should be steady but not hard enough to cause bubbles.

9. Aliquot 210 µl (∼5 × 107 nuclei) into chilled 1.5-ml microcentrifuge tube and immediately return tube to dry ice. Store nuclei at −70°C or in liquid nitrogen. Frozen nuclei are stable for at least 1 year.

10. Proceed with nuclear runoff transcription assay starting with basic protocol step 5.




ISOLATION OF NUCLEI BY SUCROSE GRADIENT CENTRIFUGATION Quality of nuclei used in nuclear runoff protocols is a major determinant in the success of the experiment. Normal lymphocytes in particular can present a problem because the nuclei are more fragile. In this protocol, cells are resuspended in an isoosmotic buffer containing nonionic detergent, then lysed by Dounce homogenization. Nuclei are collected by ultracentrifugation through a sucrose cushion and are quite clean and free of contaminating membranes and cytoplasmic components. Typically, 10% to 30% more [α-32P]UTP is incorporated into nascent transcripts of nuclei prepared by this method. The density of nuclei varies with cell type so a pilot experiment should be performed to verify that these conditions (which work well for murine splenic lymphocytes and other vertebrate cells) result in a nuclear pellet.

ALTERNATE PROTOCOL


Sucrose buffer I (see recipe), ice cold Sucrose buffer II (see recipe) Dounce homogenizer with B pestle, ice cold Polyallomer centrifuge tubes (9⁄16 × 33⁄4 in., Beckman) for SW 40.1 rotor Ultracentrifuge and SW 40.1 rotor or equivalent 1.5-ml microcentrifuge tubes, chilled on dry ice NOTE: Keep cells and nuclei on ice until the nuclei are frozen. Harvest and lyse cells 1. Harvest and wash cells as in basic protocol step 1. 2. Loosen cell pellet by gently vortexing 5 sec. Resuspend cell pellet in 4 ml ice-cold sucrose buffer I. Examine a small aliquot of cells for lysis with a phase-contrast microscope. Many cell types will lyse at this point and do not require Dounce homogenization. If cells have lysed, proceed directly to step 4.

3. Transfer cells to an ice-cold Dounce homogenizer and break the cells with five to ten strokes of a B pestle or until the nuclei appear free of cytoplasmic tags. Check a few microliters of cells with a phase-contrast microscope to be sure they are uniformly lysed. 4. Transfer nuclei to a clean 50-ml conical polypropylene centrifuge tube and add 4 ml sucrose buffer II. Mix by gentle pipetting and inversion. The final concentration of sucrose in cell homogenate should be sufficient to prevent a large buildup of debris at the interface between homogenate and sucrose cushion. The amount of sucrose buffer II added to cell homogenate may need to be adjusted.

Collect nuclei 5. Add 4.4 ml sucrose buffer II to polyallomer SW 40.1 tube. Sucrose buffer II serves as the sucrose cushion. Unlysed cells will not sediment through the sucrose cushion. If these conditions do not result in a nuclear pellet, adjust the concentration of sucrose in sucrose buffer II.

6. Carefully layer nuclei (from step 4) onto the sucrose cushion. Use sucrose buffer I to top off the gradient. Do not centrifuge more than 2 × 108 nuclei per tube.



Supplement 26

7. Centrifuge the gradient 45 min at 30,000 × g (15,500 rpm in SW 40.1 rotor), 4°C. 8. Remove supernatant by vacuum aspiration. Tilt the tube sideways so supernatant is pulled away from the pellet and remove any bubbles or liquid that remain on the side of the tube. Return tube to an ice bucket. Nuclei should form a tight pellet at the bottom of the tube and there may be some debris caught at the interface between sucrose buffers I and II. If the cells did not lyse during Dounce homogenization, nuclei will not pellet. Thus, it is important to be sure that the majority of the cells are clearly lysed in step 3. If the pellet appears as a gelatinous mass, nuclei have lysed and the pellet should be discarded.

9. Loosen nuclear pellet by gently vortexing 5 sec. Add 200 µl ice-cold glycerol storage buffer per 5 × 107 nuclei and resuspend nuclei by pipetting up and down. Nuclei will be clumped at first but will disperse with continued pipetting. Pipetting should be steady but should not create air bubbles.

10. Aliquot 210 µl (∼5 × 107 nuclei) into chilled microcentrifuge tube and immediately return tube to dry ice. Store frozen nuclei at −70°C or in liquid nitrogen. Frozen nuclei are stable for at least 1 year.

11. Proceed with nuclear runoff transcription assay starting at basic protocol step 5. SUPPORT PROTOCOL

PREPARATION OF NITROCELLULOSE FILTERS FOR NUCLEAR RUNOFF TRANSCRIPTION ASSAY cDNA plasmids are linearized and immobilized on nitrocellulose membrane filters for hybridization in the nuclear runoff transcription assay. Filters are prepared in advance and may be stored at least 6 months. Prior to their use for hybridization, filters are cut into strips that contain the cDNA plasmids of interest. Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

cDNA plasmid 1 M NaOH 6× SSC (APPENDIX 2) 0.45-µm nitrocellulose membrane Slot blot apparatus 80°C vacuum oven Additional reagents and equipment for restriction endonuclease digestion (UNIT 3.1) NOTE: Wear gloves and handle membranes with blunt-ended forceps. 1. Linearize 200 µg cDNA plasmid by digestion with an appropriate restriction enzyme. It is usually not necessary to phenol extract or ethanol precipitate the DNA after digestion if BSA is absent from the restriction enzyme digestion buffer. If the buffer contains BSA, extract plasmid DNAs with phenol/chloroform/isoamyl alcohol, ethanol precipitate, and resuspend in TE or similar buffer prior to denaturation (UNIT 2.1).


2. Add 49 µl of 1 M NaOH to linearized DNA (200 µg in 440 µl). Incubate 30 min at room temperature to denature DNA. 3. Add 4.9 ml of 6× SSC to DNA and place on ice to neutralize the sample.



4. Set up slot blot apparatus with 0.45-µm nitrocellulose membrane. Apply 125 µl of sample (∼5 µg cDNA plasmid) to each slot under a low vacuum provided by a water aspirator. Rinse each slot with 500 µl of 6× SSC. 5. Use a blue pencil to mark the location on the membrane of slots containing DNA. It is difficult to detect the location of slots once nitrocellulose has dried. Mark the edge of the slot so that the nitrocellulose strip can be trimmed very close to the edge of the slot. It is possible to minimize the volume of hybridization solution if a narrow filter strip is used.

6. Air dry nitrocellulose filter overnight. Bake filter 2 hr in an 80°C vacuum oven. Store filter in a vacuum desiccator at either room temperature or 4°C. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

DNase I, RNase-free (1 mg/ml) Adjust pH of 0.1 M iodoacetic acid/0.15 M sodium acetate to 5.3 and filter sterilize. Add sterile solution to lyophilized RNase-free DNase I (Worthington) to give a final concentration of 1 mg/ml. Heat 40 min at 55°C. Cool and add 1 M CaCl2 to a final concentration of 5 mM. Store 0.3-ml aliquots at −20°C. DNase I buffer 20 mM HEPES, pH 7.5 5 mM MgCl2 1 mM CaCl2 Sterilize by autoclaving Elution buffer 1% (w/v) SDS 10 mM Tris⋅Cl, pH 7.5 5 mM EDTA Sterilize by autoclaving Glycerol storage buffer 50 mM Tris⋅Cl, pH 8.3 40% (v/v) glycerol 5 mM MgCl2 0.1 mM EDTA HSB buffer 0.5 M NaCl 50 mM MgCl2 2 mM CaCl2 10 mM Tris⋅Cl, pH 7.4 Sterilize by autoclaving Lysis buffer 10 mM Tris⋅Cl, pH 7.4 3 mM CaCl2 2 mM MgCl2 Sterilize by autoclaving



Supplement 26

Nonidet P-40 (NP-40) lysis buffers Buffer A (basic protocol) 10 mM Tris⋅Cl, pH 7.4 10 mM NaCl 3 mM MgCl2 0.5% (v/v) NP-40

Buffer B (alternate protocol) 10 mM Tris⋅Cl, pH 7.4 3 mM CaCl2 2 mM MgCl2 1% (v/v) NP-40

Autoclave first three components and cool before adding NP-40. Reaction buffer, 2× 10 mM Tris⋅Cl, pH 8.0 5 mM MgCl2 0.3 M KCl Sterilize by autoclaving Reaction buffer with nucleotides, 2× 1 ml 2× reaction buffer (see recipe) 10 µl 100 mM ATP 10 µl 100 mM CTP 10 µl 100 mM GTP 5 µl 1 M DTT Prepare immediately prior to use Separate 100 mM solutions of each nucleotide should be prepared in 0.5 M EDTA (pH 8.0) and the pH of each one checked to be sure it is between 7.0 and 8.0. The solutions should be stored in aliquots at −20°C.

SDS/Tris buffer 5% (w/v) SDS 0.5 M Tris⋅Cl, pH 7.4 0.125 M EDTA Sterilize by autoclaving Sucrose buffer I 0.32 M sucrose 3 mM CaCl2 2 mM magnesium acetate 0.1 mM EDTA 10 mM Tris⋅Cl, pH 8.0 1 mM DTT 0.5% (v/v) Nonidet P-40 (NP-40) Prepare without DTT and NP-40. Autoclave and cool to room temperature. Add DTT and NP-40 just prior to use. Sucrose buffer II 2 M sucrose 5 mM magnesium acetate 0.1 mM EDTA 10 mM Tris⋅Cl, pH 8.0 1 mM DTT Prepare without DTT. Autoclave buffer and cool to room temperature. Add DTT just prior to use. Identification of Newly Transcribed RNA



TES solution 10 mM N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid (TES), pH 7.4 10 mM EDTA 0.2% (w/v) SDS Sterilize by autoclaving TES/NaCl solution 10 mM N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid (TES), pH 7.4 10 mM EDTA 0.2% (w/v) SDS 0.6 M NaCl Sterilize by autoclaving COMMENTARY Background Information The nuclear runoff transcription assay allows direct measurement and comparison of specific gene transcription in cells in various states of growth or differentiation. It takes advantage of the fact that newly synthesized RNA can be labeled to high specific activity in isolated nuclei, something that is difficult to accomplish in intact cells. The protocol described here has the advantage that it facilitates measurement of the level of transcription for many different genes in a single experiment. A point of controversy is whether some initiation of new RNA synthesis occurs in isolated nuclei during the runoff transcription reaction. What is clear is that transcripts that have initiated prior to cell lysis are faithfully elongated. Elongation of previously initiated transcripts most likely accounts for the bulk of the radioactivity incorporated into RNA in these reactions. Therefore, the method gives a reasonably accurate measure of the level of transcription occurring at the time of cell lysis. The runoff transcription assay is often used to assess whether changes in mRNA levels of a particular gene that occur as a function of cell state reflect a change in its synthesis as opposed to a change in mRNA degradation or transport from the nucleus to the cytoplasm. A variety of different nuclear runoff transcription protocols have been described. The procedures differ primarily in the method of isolation of 32P-labeled RNA. The protocol described here is advantageous because very low levels of background transcription are obtained. This is attributed in part to the TCA precipitation step, which allows effective removal of unincorporated 32P. Limited digestion of 32P-labeled RNA with NaOH appears to facilitate hybridization. Excellent reviews of

the nuclear runoff transcription method have been published by Marzluff (1978) and by Marzluff and Huang (1985).

Critical Parameters The most critical step in the nuclear runoff assay is isolation of nuclei. For many different types of tissue culture cells, the procedure described in the basic protocol works well. However, the isolation protocol may have to be altered somewhat for isolating nuclei from tissues and lymphocytes (Marzluff and Huang, 1985). Poor incorporation of 32P-labeled UTP into RNA in isolated nuclei may reflect damage to the nuclei during isolation or failure to isolate nuclei free of cytoplasmic and membrane contaminants. Lymphocyte nuclei are quite fragile, and it is necessary to employ an alternative isolation procedure to obtain nuclei that incorporate significant levels of radioactive isotope. In this alternative protocol, cells are lysed by Dounce homogenization in an isoosmotic buffer with nonionic detergent and nuclei are collected by centrifugation through a sucrose cushion. A minimum of 5 × 106 nuclei is required for a successful assay, but 5 × 107 is recommended. With fewer nuclei the level of incorporation of 32P-labeled UTP into RNA drops significantly, and the level of specific transcription is usually difficult to distinguish from background radioactivity. In this protocol the cpm/ml of 32P-labeled RNA is equalized in each sample prior to hybridization based on the assumption that the overall level of RNA synthesis is not changing as a function of the cell state. When analyzing gene transcription under a new set of conditions, it is critical to determine if this is a reasonable assumption. Particular care must be taken in these experiments to ensure that the



Supplement 26

same number of nuclei is used for each cell state analyzed. Incorporation of 32P-labeled UTP into RNA appears to increase when the number of nuclei is increased between 5 and 50 × 106 nuclei. Increasing the amount of 32P-labeled UTP beyond 100 µCi per sample appears to be less useful. Increasing the incubation time of nuclei with 32P-labeled UTP beyond 30 min is also ineffective for increasing the amount of 32P incorporated into RNA.

Anticipated Results

Beginning with 5 × 106 nuclei and 100 µCi of 32P-labeled UTP, the nuclear runoff transcription protocol allows incorporation of 1–10 × 106 cpm into total RNA. Good results have been obtained when as few as 1 × 106 cpm were used in the hybridization reaction.

Time Considerations Nuclei to be tested are isolated and stored in liquid nitrogen. NP-40 lysis and Dounce homogenization take 3 mm thick. Resolution in the Bull’s-eye apparatus is determined by several factors, including the concentration of the gel (Southern, 1979a,b). As the concentration of the gel increases, so does the time required for a DNA fragment of a certain size to elute. There is an upper limit to gel concentration, however, because with higher concentrations the time required for separation becomes unacceptably long, and the DNA concentration in each sample is reduced. Generally, a gel concentration of 1.2% is optimal for separation of fragments for cloning.

Literature Review The concept of simplifying cloning by gene enrichment has been reviewed by Edgell et al. (1979). They describe a two-step DNA fractionation procedure involving RPC5 column chromatography followed by preparative electrophoresis. Bott et al. (1980) have used preparative agarose electrophoresis to fractionate the Bacillus subtilis genome to produce DNA fragments for transformation analysis and cloning. The degree of enrichment achieved with preparative electrophoresis alone is adequate and allows one to start with a smaller quantity of DNA. Methods for constructing an apparatus that allows collection of multiple samples of sizefractionated DNA from a slab horizontal gel have been described (Polsky et al., 1978; Southern, 1979a,b). The resolution on a preparative agarose slab gel is adequate (although not as good as the Bull’s-eye apparatus). The primary difficulty with some preparative agarose gels is that the DNA that is eluted contains impurities that inhibit DNA ligase and prevent cloning. The Bull’s-eye apparatus is discussed here because it is commercially available, yields DNA that can be ligated, and produces large numbers

Troubleshooting Using the preparative agarose gel technique, fragments can generally be identified and eluted. Problems sometimes arise in that not enough clones can be obtained to make a subgenomic library (see below). Generally, if this occurs, it is best to start over with another fractionation technique. The Bull’s-eye apparatus is not easy for the beginner. Setting up the apparatus can be difficult, and instruction from someone skilled in the use of this machine is advised. In particular, placement of the dialysis tubing over the central anode is a tricky and crucial step in assembly. However, once assembled correctly the machine rarely malfunctions. Occasionally, bubbles will impede flow of buffer in one of the circuits. This is prevented by clearing the tubing of bubbles before starting a run. Failure of the tubing leading to the fraction collector to run dry in less than 7 min usually means that either the peristaltic pump in this circuit is too slow or that there is a leak into the sample collection chamber. Such a leak occurs through the dialysis tubing that covers the anode and is corrected by replacing the

Construction of Recombinant DNA Libraries


dialysis tubing. DNA fractionated in this fashion and cleaned by ethanol precipitation and chloroform extractions usually can be ligated efficiently into bacteriophage or plasmid vectors. If appropriate controls point to the fractionated DNA as the source of difficulty, it may help to repeat the phenol and chloroform extractions or to pass the DNA over an Elutip column (UNIT 2.6). If the DNA still cannot be ligated it probably means that the restriction fragment ends were damaged by exonuclease activity.

Anticipated Results Up to 80% of the DNA applied to a gel (either a slab gel or a Bull’s-eye gel) should be recovered. If 1 mg DNA is digested and loaded onto the gel, and 100 fractions collected, each fraction will contain several micrograms of DNA. This is enough DNA to produce a number of subgenomic libraries. The size distribution of the fragments will vary, depending on the enzyme used, but fractions around 4 kb will usually contain the most DNA.

Time Considerations Fractionation in a preparative agarose slab gel is rapid, generally 5 kb) cDNA.



Supplement 29

3. In a separate tube add in the following order (180 µl total): 20 µl 5 mM dNTPs (500 µM final each) 40 µl 5× RT buffer (1× final) 10 µl 200 mM DTT (10 mM final) 20 µl 0.5 mg/ml oligo(dT)12-18 (50 µg/ml final) 60 µl H2O 10 µl (10 U) RNasin (50 U/ml final). Mix by vortexing, briefly microcentrifuge, and add the mixture to the tube containing the RNA. Add 20 µl (200 U) AMV reverse transcriptase for a final concentration of 1000 U/ml in 200 µl. Mix as above and remove 10 µl to a separate tube containing 1 µl of [α-32P]dCTP. Leave both tubes at room temperature 5 min, then place both tubes at 42°C for 1.5 hr. The aliquot is removed to determine incorporation and permit an estimation of recovery. The remainder of the cDNA will be labeled during second-strand synthesis. Labeling cDNA during first-strand synthesis to a high enough specific activity to permit easy detection with a hand-held radiation monitor during all subsequent steps requires a relatively large amount of label, which may then interfere with reverse transcription due to buffer effects. Some investigators check the quality of the cDNA by fractionating the radiolabeled cDNA on an alkaline agarose gel and detecting it by autoradiography. Much of the cDNA should be >1000 bp long. For a specifically primed library, substitute an equal weight of antisense 15- to 40-mer primer for the oligo(dT)12-18. Expect a 100-fold enrichment of specific clones in the library. For a randomly primed library, substitute an equal weight of random-hexamer primers for oligo(dT)12-18 (or use a 50:50 mix), and perform the reverse transcription at 37° instead of 42°C.

4. Add 1 µl of 0.5 M EDTA, pH 8.0, to the radioactive reaction and freeze it at −20°C. It will be used later to estimate the amount of cDNA synthesized. 5. To the main reaction add 4 µl of 0.5 M EDTA, pH 8.0, and 200 µl buffered phenol. Vortex well, microcentrifuge at room temperature for 1 min to separate phases, and transfer the upper aqueous phase to new tube. Save the tube containing the phenol layer, too.

6. Add 100 µl TE buffer, pH 7.5, to the phenol layer and vortex and microcentrifuge as in step 5. Remove the aqueous layer and add it to the aqueous phase from the first extraction. The volume of aqueous phase is now about 300 ìl; the phenol may be discarded. Back extraction of the organic phase at each phenol extraction significantly improves the yield. See UNIT 2.1 for a discussion of phenol extraction and ethanol precipitation.

7. Add 1 ml diethyl ether, vortex, and microcentrifuge as in step 5. Remove and discard the upper (ether) layer with a glass pipet. Repeat the extraction with an additional 1 ml of ether. A single chloroform/isoamyl alcohol extraction followed by back extraction of the organic phase may be substituted for the two ether extractions; however, the yield is normally slightly lower.

Conversion of mRNA into Double-Stranded cDNA

8. Add 125 µl of 7.5 M ammonium acetate to the aqueous phase (final concentration 2.0 to 2.5 M) and 950 µl of 95% ethanol. Place in dry ice/ethanol bath 15 min, warm

5.5.4 Supplement 29


to 4°C, and microcentrifuge at 10 min at full speed, 4°C, to pellet nucleic acids. A small, yellow-white pellet may be visible. Precipitation from ammonium acetate leaves short oligonucleotides in the supernatant, thus removing the oligo(dT) primer and enriching the pellet in longer cDNAs. Do not substitute sodium acetate for ammonium acetate.

9. Remove the supernatant with a pipet, fill the tube with ice-cold 70% ethanol, and microcentrifuge 3 min at full speed, 4°C. Again remove the supernatant, then dry the tube containing the precipitated nucleic acids briefly in a vacuum desiccator. 10. Thaw the tube containing the radioactive aliquot of the first-strand synthesis reaction and spot the sample onto a nitrocellulose membrane filter. 11. Wash the membrane with ice-cold 10% TCA and determine the radioactivity bound to the filter with a fluor and scintillation counter. Use the specific activity of the radiolabel in the reaction, the amount of RNA used, the counts incorporated, and the efficiency of the β-counter to calculate the amount of cDNA synthesized (see Sample Calculation for Determining Amount of cDNA Synthesized, in Commentary). From 1 to 4 ìg is typical even though the theoretical maximum is 10 ìg.

Convert cDNA into double-stranded cDNA 12. Resuspend the pellet from the first-strand synthesis in 284 µl water and add to the tube in the following order (400 µl total): 4 µl 5 mM dNTPs (50 µM final each) 80 µl 5× second-strand buffer I (1× final) 12 µl 5 mM β-NAD+ (150 µM final) 2 µl 10 µCi/µl [α-32P]dCTP (50 µCi/ml final). Mix by vortexing, briefly microcentrifuge, and add: 4 µl (4 U) RNase H (10 U/ml final) 4 µl (20 U) E. coli DNA ligase, (not T4 DNA ligase; 50 U/ml final) 10 µl (100 U) E. coli DNA polymerase I (250 U/ml final). Mix by vortexing, briefly microcentrifuge, and incubate 12 to 16 hr at 14°C. Unrelated cDNA fragments are not ligated in this reaction because E. coli DNA ligase does not catalyze blunt ligations.

13. After second-strand synthesis, remove 4 µl of the reaction to a new tube and freeze at −20°C. Later, when time permits, determine the incorporation of radiolabel into acid-insoluble material as outlined in steps 10 and 11. Expect 1-10 × 106 cpm incorporated in the total reaction.

14. Phenol extract the second-strand synthesis reaction with 400 µl buffered phenol and back extract the phenol phase with 200 µl TE buffer, pH 7.5, as in steps 5 and 6. 15. Pool the two aqueous phases and extract twice with 900 µl ether, as in step 7. The volume of the aqueous phase is now ∼600 µl. 16. Divide the aqueous phase evenly between two tubes, add ammonium acetate, and ethanol precipitate as in steps 8 and 9. Unincorporated radioactive dCTP is removed by the ethanol precipitation and wash steps. From this point on, the cDNA may be followed with a hand-held radiation monitor. Construction of Recombinant DNA Libraries


Supplement 29

Create blunt ends on double-stranded cDNA 17. Complete second-strand synthesis and blunt the double-stranded cDNA by resuspending the pooled pellets in 42 µl water. Add in the following order (80 µl total): 5 µl 5 mM dNTPs (310 µM final each) 16 µl 5× TA buffer (1× final) 1 µl 5 mM β-NAD+ (62 µM final). Mix by vortexing, microcentrifuge briefly, and add: 4 µl of 2 µg/ml RNase A (100 ng/ml final) 4 µl (4 U) RNase H (50 U/ml final) 4 µl (20 U) E. coli DNA ligase (250 U/ml final) 4 µl (8 U) T4 DNA polymerase (100 U/ml final). Mix as above and incubate 45 min at 37°C. Volume of T4 DNA polymerase used to obtain 8 U may require adjustment depending on batch. If the library is to be screened with an antiserum, some investigators (see Tamkun et al., 1986) digest the cDNA at this point with AluI or HaeIII to prepare small inserts that may produce more stable fusion proteins in λgt11. Do this after the T4 polymerase step by diluting the 80-ìl reaction to 170 ìl with water; add 24 ìl of 5× TA buffer and 6 ìl of one of the above restriction enzymes. Incubate 1 hr at 37°C, then proceed to step 18, except do not add any TE buffer. An insert isolated from an immunological screen may then be used to screen a second library of full-length inserts.

18. Add 120 µl TE buffer, pH 7.5, and 1 µl of 10 mg/ml tRNA. Extract with 200 µl buffered phenol and back extract the phenol phase with 100 µl TE buffer as described in steps 5 and 6. 19. Pool the two aqueous phases and extract twice with 1 ml ether, as in step 7. 20. Ethanol precipitate the cDNA as in steps 8 and 9. The cDNA is now ready to be tailed or linkered to create compatible ends for subsequent cloning steps. ALTERNATE PROTOCOL

CONVERSION OF mRNA INTO DOUBLE-STRANDED cDNA FOR DIRECTIONAL CLONING Generation of cDNA that has unique ends for directional cloning is carried out in two parts, as in the basic protocol. First, the mRNA is hybridized to a linker-primer that incorporates a poly(dT) tract (at its 3′ end) as well as a restriction site for XhoI (Fig. 5.5.2). The linker-primer is extended using an RNase H− version of the Moloney murine leukemia virus reverse transcriptase (SuperScript) and a nucleotide mix in which dCTP is replaced with 5-methyl-dCTP. When first-strand synthesis is completed, the reaction mixture is transferred into a second tube that contains the prechilled second-strand mixture. The second strand is synthesized using RNase H and E. coli DNA polymerase I. Finally, a blunting step (consisting of treatments with mung bean nuclease and Klenow fragment) is carried out to prepare the cDNA for the EcoRI adaptor ligation.


Additional Materials (also see Basic Protocol) 5× SuperScript buffer (RNase-free; see recipe) 0.1 M DTT (RNase-free) 3dNTP/methyl-dCTP mix: 10 mM each dATP, dGTP, dTTP, and 5-methyl-dCTP (Pharmacia Biotech) continued

5.5.6 Supplement 29


AAAAAAAn TTT n GAGCTC (GA)10

reverse transcription AAAAAAA TTTTTTTGAGCTC (GA)10

Me

Me

second - strand synthesis AAAn TTT n GAGCTC (GA)10

Figure 5.5.2 Synthesis of cDNA with unique ends for directional cloning.

0.25 µg/µl oligonucleotide primer (UNIT 2.11) incorporating (from 5′ to 3′): (dGdA)10, XhoI restriction site, and (dT)18 200 U/µl SuperScript or SuperScript II (GIBCO/BRL) 5× second-strand buffer II (see recipe) 10 mM and 2 mM 4dNTP mix (Pharmacia Biotech) 10 µCi/µl [α-32P]dATP (3000 Ci/mmol) 0.8 U/µl RNase H (Pharmacia Biotech) 10 U/µl E. coli DNA polymerase I (New England Biolabs) 1:1 (w/v) phenol/chloroform 100% ethanol ice-cold 10× Klenow buffer (see recipe) 5 U/µl Klenow fragment of E. coli DNA polymerase I (New England Biolabs) 10× mung bean nuclease buffer (see recipe) 10 U/µl mung bean nuclease (New England Biolabs) 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2) 16°C incubator Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5) Synthesize cDNA 1. Prepare 5 to 7 µg poly(A)+ RNA. If mRNA is prepared as in UNIT 4.5, store under ethanol in a microcentrifuge tube until ready to proceed. If mRNA is in aqueous solution, add 3 M sodium acetate (RNase-free) to 0.3 M and 3 vol of 100% ethanol, chill ≥20 min at −20°C, and microcentrifuge 20 min at maximum speed, 4°C, to collect mRNA. Rinse the pellet with 70% ethanol (RNase-free), and invert the tube to air dry.

2. Prepare first-strand premix (45 µl final by mixing in the order listed): 10 µl 5× SuperScript buffer 5 µl 0.1 M DTT 2.5 µl 3dNTP/methyl-dCTP mix 1 µl RNasin 12 µl 0.25 µg/µl oligonucleotide primer 14.5 µl H2O. 3. Resuspend RNA pellet directly into premix. Incubate 15 min at room temperature, then add 5 µl of 200 U/µl SuperScript and incubate 1 hr at 42°C.



Supplement 29

4. Shortly before end of incubation, prepare second-strand premix (336 µl final) containing: 80 µl 5× second-strand buffer II 6 µl 10 mM 4dNTP mix 2.5 µl 10 µCi/µl [α-32P]dATP 247.5 µl H2O. Place on ice. 5. Continuing to work on ice, add the first-strand mix from step 3 (50 µl total) to prechilled second-strand premix from step 4. 6. In a second microcentrifuge tube combine 4 µl of 0.8 U/µl RNase H (3.2 U) and 10 µl of 10 U/µl E. coli DNA polymerase I (100 U). Add combined enzymes to the tube from step 5 containing the cDNA and mix by rapidly inverting a few times. In order to prevent formation of “snapbacks” during second-strand synthesis it is important to prechill the premix and add the RNase H and DNA polymerase simultaneously.

7. Microcentrifuge 5 sec at maximum speed, then incubate reaction 1 hr at 16°C and 1 hr at room temperature. At this step, second-strand synthesis is completed.

8. Electrophorese ∼2% to 5% of total reaction on a 0.7% agarose gel, and monitor size range of labeled cDNA by autoradiography. Optimally, a smear ranging from ∼200 bp to >8 kb will be seen, with a peak at 1 to 2 kb. The reaction can be frozen overnight (or longer) at −20°C, or one can immediately proceed to blunting and linker ligation.

Create blunt-ends on double-stranded cDNA 9. Add 500 µl of 1:1 phenol/chloroform to reaction. Microcentrifuge 5 min at maximum speed to separate phases. If reaction was frozen, thaw by vortexing with the phenol/chloroform.

10. Remove upper phase to a fresh tube, add 200 µl of 7.5 M ammonium acetate, and mix by inversion. 11. Divide sample evenly between two tubes (300 µl in each). Add 600 µl ice-cold 100% ethanol to each tube, place both tubes 5 min at −80°C, then microcentrifuge 20 min at maximum speed, 4°C. This and the following series of steps involve transfer of labeled cDNA from tube to tube, and require a number of extractions and precipitations. After this initial precipitation, the bulk of the radiolabel is incorporated in the cDNA. It is important to monitor using a Geiger counter to ensure that all cDNA is transferred from step to step and not lost during the extractions.

12. Pour off supernatants and rinse pellets with ice-cold 70% ethanol. Dry briefly in a Speedvac evaporator. Add 30 µl water to one of the two tubes, vortex to resuspend cDNA pellet, then transfer contents to the second tube. 13. Add an additional 10 µl water, 5 µl of 10× Klenow buffer, and 5 µl of 2 mM 4dNTP mix to the first tube. Vortex and transfer to the second tube. Add 5 µl of 10 U/µl Klenow fragment to the second tube and incubate 30 min at 37°C. Conversion of mRNA into Double-Stranded cDNA

The aim of the first Klenow treatment is to ensure that the 3′ end of the cDNA is

5.5.8 Supplement 29


completely blunted so that the XhoI restriction site will not be harmed by the subsequent enzymatic manipulations.

14. Add 50 µl of 1:1 phenol/chloroform to reaction, extract, and transfer upper phase to new tube. Back extract lower phase with an additional 50 µl water and combine it with the first extraction. 15. Add 50 µl of 7.5 M ammonium acetate and 300 µl ice-cold 100% ethanol. Place at −80°C for 5 min, then microcentrifuge 20 min at maximum speed, 4°C. 16. Pour off supernatant and rinse pellet with cold 70% ethanol. Dry pellet briefly in a Speedvac evaporator. Resuspend cDNA in 225 µl water. 17. Add 25 µl of 10× mung bean nuclease buffer and 1 µl of 1 U/µl mung bean nuclease. Incubate 15 min at 37°C. This mung bean exonuclease treatment removes any residual single-stranded nucleic acid extensions (i.e., single-stranded DNA, RNA overhangs, or small hairpins) from the 5′ end of the cDNA.

18. Add 25 µl of 1 M Tris⋅Cl, pH 8.0. Extract with 200 µl 1:1 phenol/chloroform and back extract with 50 µl water, pooling the upper phases in a new tube. 19. Add 175 µl of 7.5 M ammonium acetate and fill tube with ice-cold 100% ethanol. Place at −80°C for 5 min, then microcentrifuge 20 min at maximum speed, 4°C. 20. Rinse pellet with 70% ethanol and air dry briefly. Resuspend in 20 µl water, then add 2.5 µl of 10× Klenow buffer and 1 µl of 5 U/µl Klenow fragment. Incubate 5 min at 37°C. Add 2.5 µl of 2 mM dNTP mix to the reaction and incubate an additional 25 min at room temperature. Do not overdry pellet (i.e., if Speedvac evaporation is used, do not dry >5 min). The goal of this second Klenow fragment treatment is to effectively blunt any short single-stranded extensions that might remain after the mung bean exonuclease treatment or be caused by the “sloppiness” of this enzyme. This series of steps effectively causes essentially all the cDNA to become blunt-ended.

21. Add 25 µl of 1:1 phenol/chloroform to the reaction and extract, removing upper phase to a new tube. Back extract lower phase with an additional 25 µl water and pool with first upper phase. Store sample frozen at −20°C until ready for the EcoRI adaptor ligation step. EcoRI adaptor ligation is carried out as it is described in UNIT 5.6. Subsequently, XhoI digestion of the EcoRI adaptor–ligated cDNA results in a 5′ end that has an EcoRI-compatible overhang and a 3′ end with XhoI-compatible overhang.



Supplement 29


Buffer stock solutions Evaluate water and buffers for nuclease activity by incubating 5 µl solution with 3 µg of intact total cellular RNA or 0.5 µg of a DNA fragment, whichever is relevant, for 30 min. Check the RNA or DNA by agarose gel electrophoresis to be sure it was not degraded. Prepare at least 10 ml of each of the following 1 M stock solutions. Use RNA-grade water (see UNIT 4.1) and pass each solution through a sterile 0.45-µm filter. Store at room temperature unless otherwise indicated. 1 M Tris⋅Cl, pH 8.2, 42°C 1 M Tris⋅acetate: titrate aqueous Tris base to pH 7.8 with acetic acid 1 M KCl 1 M MgCl2 1 M (NH4)2SO4 1 M potassium acetate 1 M magnesium acetate 1 M DTT, −20°C, in tightly capped tube 5 mg/ml nuclease-free BSA, −20°C The pH of Tris buffers varies considerably with temperature. Be sure to measure the pH at the indicated temperature and use an electrode that accurately measures the pH of Tris buffers.

Enzymes Enzymes should be of the highest quality available. Before starting a cDNA cloning experiment with a rare or hard-to-obtain RNA prep, it is advisable to check the activities of all the enzymes used. Klenow buffer, 10× 0.5 M Tris⋅Cl, pH 7.6 0.1 M MgCl2 Prepare 1.0 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C. Mung bean nuclease buffer, 10× 0.3 M sodium acetate, pH 4.5 0.5 M NaCl 10 mM ZnCl2 50% (v/v) glycerol Prepare 1.0 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C.


Reverse transcriptase (RT) buffer, 5× 250 µl 1 M Tris⋅Cl, pH 8.2 250 µl 1 M KCl 30 µl 1 M MgCl2 470 µl H2O Prepare 1.0 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C.



RNase A (DNase-free) Dissolve 1 mg/ml RNase A (Sigma) in TE buffer (APPENDIX 2); boil 10 min to remove contaminating DNase and store in aliquots at −20°C. Dilute to 2 µg/ml in TE buffer when needed. Second-strand buffer I, 5× 100 µl 1 M Tris⋅Cl, pH 7.5 500 µl 1 M KCl 25 µl 1 M MgCl2 50 µl 1 M (NH4)2SO4 50 µl 1 M DTT 50 µl 5 mg/ml BSA 225 µl H2O Prepare 1.0 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C. Second-strand buffer II, 5× 94 mM Tris⋅Cl, pH 6.9 453 mM KCl 23 mM MgCl2 50 mM (NH4)2SO4 Prepare 1.0 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C. SuperScript buffer, 5× 250 mM Tris⋅Cl, pH 8.3 375 mM KCl 15 mM MgCl2 Prepare 1.0 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C. TA buffer, 5× 200 µl 1 M Tris⋅acetate, pH 7.8 400 µl 1 M potassium acetate 60 µl 1 M magnesium acetate 3 µl 1 M DTT 105 µl 5 mg/ml BSA 432 µl H2O Prepare 1.2 ml buffer using high-quality water and buffer stock solutions (see recipe), then check for nuclease activity as described in buffer stock solution recipe. Freeze in 200-µl aliquots in screw-cap microcentrifuge tubes at −80°C.



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COMMENTARY Background Information


cDNA synthesis and subsequent library preparation are necessary for the study of most mRNAs because no technology for conveniently manipulating and propagating RNA sequences is currently available. Of fundamental importance is that the cDNAs faithfully represent the sequence, size, and complexity of the mRNA population. The high quality of commercially available enzymes and reagents has dramatically changed the major concerns of the scientist preparing a cDNA library. In the past, considerable attention was paid to the source of reverse transcriptase, the quality of other enzymes, methods for their further purification, and conditions for optimum activity (Maniatis et al., 1982). These issues have been successfully addressed and full-length cDNAs of 5 to 8 kb mRNAs are now obtainable. The major focus currently is on the preparation of libraries of full-length cDNAs that will contain copies of even the rarest mRNA. There are several relatively recent innovations incorporated into these protocols. First, it is recommended that size fractionation of the mRNA be omitted in order to minimize the opportunities for degradation of the mRNA template. Size fractionation of the cDNA, which is much more stable, is discussed in UNITS 5.3 & 5.4. Second-strand synthesis employs RNase H, E. coli DNA ligase, and E. coli DNA polymerase I as described by Okayama and Berg (1982) and modified by Gubler and Hoffman (1983). More recently, Neve et al. (1986) introduced during the blunting process the additional second-strand synthesis step, which increases the yield and size of the cDNA. Using this modification, S1 nuclease cleavage of the 5′ “hairpin loop” generated by reverse transcriptase, a problematic step in the past, is avoided. The cDNA produced is ready to be prepared for insertion into an appropriate vector. Variations such as omitting the DNA ligase, (e.g., as per Gubler and Hoffman, 1983), omitting the additional second-strand synthesis, or substituting Klenow fragment for the intact E. coli DNA polymerase I have been successfully employed by others. Finally, the basic protocol uses AMV (avian myeloblastosis virus) reverse transcriptase (RT), but the alternate protocol uses SuperScript, a genetically engineered derivative of Moloney murine leukemia viral (MoMuLV) RT that has had its RNase H activity removed (Kotewicz et al., 1988) and its polymerase activity enhanced, resulting in a

greater size and yield of cDNA from first-strand synthesis. A number of variations of the protocols in this unit exist for particular applications. Random hexamers can be used to prime first-strand synthesis rather than oligo(dT)-containing primers: resulting cDNAs will be enriched for the 5′-end sequences of very long transcripts, although average cDNA length will be shorter than with the methods described here. Another method that has gained popularity for a variety of analytical techniques is to perform a standard first-strand synthesis from an mRNA of interest, and then amplify specific rare transcripts using PCR. Over the last several years, a number of kits for cDNA synthesis have become commercially available (e.g., from GIBCO/BRL, Stratagene, Promega, Amersham, and Pharmacia Biotech), and generally yield excellent results.

Critical Parameters RNA used to generate the cDNA must be undegraded and of the highest quality obtainable. It must also be DNA-free. Buffers and solutions must be nuclease-free. These may be divided into aliquots, frozen, and stored for long periods; thus, it is worthwhile to check each by incubating intact RNA or DNA, whichever is appropriate, with a sample of the solution and checking for degradation by agarose gel electrophoresis. Enzymes (and all reagents) should be the highest quality available. If experimenting with a rare or hard-to-isolate RNA preparation, it is advisable to check the activities of all enzymes used. Enzymes should be relatively fresh and stored at −20°C in a freezer that is not frost-free (because the warming and cooling cycles of a frost-free freezer cause the enzymes to lose activity). The RNasin added in the first-strand synthesis is a potent though reversible inhibitor of RNase and must be added after the DTT, as its activity is dependent on the presence of a reducing agent.

Troubleshooting Once high-quality mRNA has been obtained, the cDNA synthesis usually proceeds well. Difficulties, if any, only occur later— when the cDNA is ligated to the vector. If the yield of cDNA is low, likely causes are inactive reverse transcriptase, inactive E. coli polymerase I, or deteriorated dNTP mixes. If the calculated yield is low, double-check the result



Sample Calculation for Determining Amount of cDNA Synthesized 1. Assume 2 × 1012 cpm/Ci; thus, if [α-32P]dCTP is at a specific activity of 3000 Ci/mmol, there are 6 × 1015 cpm/mmol. 2. The second-strand synthesis aliquot representing 1% of the total contains 105 TCA-precipitable cpm, thus 105 cpm × 100 6 × 1015 cpm/mmol

= 1.67 × 10−9 mmol [alpha−32P]dCTP incorporated.

3. The initial concentration of [α-32P]dCTP was 50 µCi/ml = 1.67 × 10–2 µM. 3000 Ci/mmol

4. Thus, the total dCTP incorporated is 50 µM –2

1.67 × 10

µM

× 1.67 × 10–9 mmol ×

0.3 g dCMP = 1.5 µg. mmol

5. There were four deoxynucleotides present in the reaction, so the total cDNA synthesized in the second-strand reaction is 6 µg and since the cDNA is doublestranded, there are 12 µg of cDNA total.

by running 5% of the total sample on an agarose minigel and staining with ethidium bromide, or by determining the amount of DNA in the sample by optical density or DAPI fluorescence. If the radiolabel was old or deteriorated, incorporation will be low but the amount of cDNA synthesis will be correct. Very small cDNA is usually the result of DNase contamination. This is readily evaluated by adding a restriction fragment of DNA to the reaction mixes and checking the fragment after incubation by gel electrophoresis.

Anticipated Results The typical amount of poly(A)+ RNA used to generate a library is 10 µg; however, the protocol may be scaled down should RNA availability be limiting. The 10 µg of poly(A)+ RNA will yield 2 to 10 µg of double-stranded cDNA. An aliquot representing 2% to 5% of the total volume may be checked by agarose gel electrophoresis, ethidium bromide staining, and autoradiography. The aliquots removed from the first- and second-strand synthesis should contain 2 to 8 × 105 cpm and 2 to 10 × 104 cpm TCA-insoluble counts, respectively (basic protocol). 100 ng double-stranded cDNA should generate a full-complexity library (1 × 106 recombinants) if a phage vector such as λgt10 is employed.

To determine the amount of cDNA synthesized, see Sample Calculation for Determining Amount of cDNA Synthesized.

Time Considerations Either of these protocols can usually be completed in two days. In the basic protocol, first-strand synthesis should be started in the morning to permit the initial second-strand synthesis to proceed overnight. The final secondstrand synthesis should be carried out the following morning. In the alternate protocol, 4 hr should be allowed for synthesis of the doublestranded cDNA and 4 to 5 hr for generation of blunt ends. The cDNA may be linkered or tailed as described in UNIT 5.6 later the same day. If necessary, the protocol may be interrupted at any ethanol precipitation step.

Literature Cited Gubler, U. and Hoffman, B.J. 1983. A simple and very effective method for generating cDNA libraries. Gene 25:263-269. Kotewicz, M.L., Sampson, C.M., D’Alessio, J.M., and Gerard, G.F. 1988. Isolation of cloned Moloney murine leukemia virus reverse transcriptase lacking ribonuclease H activity. Nucl. Acids Res. 16:265-277. Maniatis, T., Fritsch, E.F., and Sambrook, J. 1982. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.



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Neve, R.L., Hanis, P., Kosik, K.S., Kurnit, D.M., and Donlon, T.A. 1986. Identification of the gene for the human microtubule-associated protein tau and chromosomal localization of the genes for tau and microtubule-associated protein 2. Brain Res. 387(3):271-80. Okayama, H. and Berg, P. 1982. High-efficiency cloning of full-length cDNA. Mol. Cell. Biol. 2:161-170. Tamkun, J.W., DeSimone, D.W., Fonda, D., Patel, R.S., Buck, D., Horwitz, A.F., and Hynes, R.O. 1986. Structure of integrin, a glycoprotein involved in the transmembrane linkage between fibronectin and actin. Cell 46:271-282.

Key References Gubler and Hoffman, 1983. See above. Huse, W.D. and Hansen, C. 1988. cDNA cloning redefined. Strategies (Stratagene) 1:1-3. First demonstrated use of linker primers and methylated nucleotides in cDNA synthesis.

Okayama and Berg, 1982. See above. These authors first developed the RNase H/E. coli DNA polymerase I alternative to S1 nuclease and Klenow fragment for second-strand synthesis.

Contributed by Lloyd B. Klickstein Brigham and Women’s Hospital Boston, Massachusetts Rachael L. Neve McLean Hospital Belmont, Massachusetts Erica A. Golemis Fox Chase Cancer Center Philadelphia, Pennsylvania Jeno Gyuris Mitotix, Inc. Cambridge, Massachusetts




Ligation of Linkers or Adapters to Double-Stranded cDNA

UNIT 5.6

Linkers or adapters can be ligated to double-stranded cDNA (UNIT 5.5) to provide restriction endonuclease sites used in the production of a cDNA library (UNIT 5.8). For cloning purposes, only one linker or adapter must be present on each end of the cDNA. However, multiple linkers are usually ligated to the cDNA because both ends are phosphorylated and contain cohesive sequences. As a result, cDNA must first be methylated to protect it from a subsequent restriction digest designed to remove the multiple linkers (basic protocol). The procedure for ligating adapters to the cDNA is much simpler than that for linkers because only one end is phosphorylated, resulting in the ligation of just one adapter (alternate protocol). Linkered or adapted cDNA is then passed over a Sepharose CL-4B column to remove unligated linkers or adapters and other low-molecular-weight material (1.5 kb. Since only 50 to 100 ng of cDNA are required to produce a full complexity library with a phage vector, it is recommended that a library be produced at this stage in any event. A

A

B

pooled cDNA fractions 6 -10 inclusive

(origin)

32

P cpm /2 x 10

-2

µl

28 24 20 16

tracking d ye begins to elute

12 8

λ

ds cDNA

kb 21.4 9.6 6.7 4.5 2.1 1.9

4 2

4

6

8

10 12 14 16

CL - 4B fraction no.

Figure 5.6.1 Fractionation of EcoRI-digested cDNA by (A) Sepharose CL-4B chromatography and (B) agarose gel electrophoresis.



Supplement 13

library may be stored for years and may be useful in the future. For example, sequences related to the gene of interest may be identified that were excluded from a size-fractionated library.

Size-select the cDNA to obtain long inserts 16. Pour a 0.8% TBE agarose minigel—the gel should be thick enough such that all of the cDNA will fit into a single well. Rinse the gel box, tray, and comb thoroughly, and use fresh TBE electrophoresis buffer. High-quality, nuclease-free agarose that does not inhibit ligation is essential. Most commercial agarose advertised as molecular biology grade is adequate. The agarose may be checked by first carrying an EcoRI fragment of a plasmid through the procedure and comparing its cloning efficiency in the cDNA vector, expressed as recombinants/ng insert, to the cloning efficiency of the same fragment prior to fractionation. Wash the gel box thoroughly afterward!

17. Add 10× loading buffer to 1× final to the cDNA and load it into a well near the center. Load DNA molecular weight standards (e.g., an HindIII digest of λ phage) two wells away from the sample. Electrophorese at 70 V until adequate resolution is achieved as determined by ethidium bromide fluorescence, usually 1 to 4 hr. Be sure not to use standards with EcoRI ends.

18. Elute double-stranded cDNA of the desired size as estimated by comparison with the comigrated standards. λgt10 and λgt11 have a maximum insert size of 7 kb, so collecting cDNA larger than this won’t be useful unless a plasmid vector or a phage vector such as Charon 4A or EMBL 4 will be used.

19. Add 10 mg/ml tRNA to 20 µg/ml final, 1⁄10 vol of 3 M sodium acetate, pH 5.2 (APPENDIX 2), and 2.5 vol ice-cold 95% ethanol and place 15 min on dry ice. Microcentrifuge, wash and dry as in step 14, and resuspend pellet in 20 µl TE buffer. Ethanol precipitation also extracts the ethidium from the DNA.

20. Determine radioactivity in 1 µl using a fluor and scintillation counter and then calculate the recovery of double-stranded cDNA (see commentary). Proceed to library construction protocols in UNIT 5.8. ALTERNATE PROTOCOL

LIGATION OF BstXI SYNTHETIC ADAPTERS Blunt-ended, double-stranded cDNA is ligated to phosphorylated BstXI adapters and then purified as described in the basic protocol. Alternatively, EcoRI or EcoRI-NotI adapters may be used for cDNA to be cloned in vectors with the EcoRI site (Fig. 5.6.2). This protocol is simpler than that for linkers because the methylation and restriction digestion steps are unnecessary. Additional Materials BstXI adapters (UNIT 2.11; Invitrogen), EcoRI adapters (New England Biolabs), or EcoRI-NotI adapters (Invitrogen) 1. Dissolve blunt-ended, double-stranded cDNA pellet in 23 µl water and add in the following order (30 µl final volume):


3 µl 10× T4 DNA ligase buffer containing 5 mM ATP (1× and 0.5 mM ATP final) 2 µl 1 µg/µl EcoRI, EcoRI-NotI, or phosphorylated BstXI adapters (67 µg/ml final).

5.6.4 Supplement 13


adapters

5′ CCATTGTG 3′ GGTA

CTCTAAAG

CTTTAGAGCACA

ACACGAGATTTC

GAAATCTC

ATGG 3′ GTGTTACC 5′

insert cDNA

vector cut with BstXI

Figure 5.6.2 Noncomplementary adapter strategy. The insert and vector ends are compatible with each other but cannot self-ligate because the cohesive ends are not self-complementary. The BstXI sites (underlined and bold) are not regenerated.

Mix gently by pipetting up and down with a pipettor. Add 2 µl (800 U) T4 DNA ligase to 27,000 U/ml final, mix as above, and incubate overnight at 4°C. It is helpful to use 32P-labeled cDNA to follow the DNA on the subsequent CL-4B column. If the cDNA is not 32P-labeled, it may be labeled at this step by using [32P]labeled adapters, prepared with [γ-32P]ATP as in the T4 polynucleotide kinase exchange reaction (UNIT 3.10). Alternatively, if the adapters are not yet phosphorylated, they may be labeled with [γ-32P]ATP as in the T4 polynucleotide kinase forward reaction (UNIT 3.10).

2. Add 100 µl TE buffer and remove excess adapters as in steps 10 to 15 of the basic protocol. If desired, size-select the cDNA as in steps 16 to 20 of the basic protocol. Resuspend purified cDNA pellet (obtained from either the CL-4B column or the gel) in 10 to 15 µl TE buffer. Proceed to library construction protocols in UNIT 5.8. Because adapter dimers formed during the ligation reaction will clone into the vector very efficiently, removal of the excess adapters is essential.

PREPARATION OF A CL-4B COLUMN The CL-4B column (Fig. 5.6.3) effectively removes linkers or adapters that would otherwise interfere in subsequent cloning steps; it also allows selection of cDNA ≥350 bp (see basic and alternate protocols). The column may be prepared while the cDNA is being digested with EcoRI, as described in steps 7 and 8 of the basic protocol.

SUPPORT PROTOCOL

Additional Materials Preswollen Sepharose CL-4B (Pharmacia), 4°C CL-4B column buffer Silanized glass wool Plastic tubing (new) with clamp



Supplement 13

disposable plastic pipet

Sepharose CL-4B

to ring stand clamp

silanized glass wool tubing clamp plastic tubing

Figure 5.6.3 CL-4B column used for removal of EcoRI-digested linkers and selection of cDNA ≥350 bp.

1. Transfer 10 ml of preswollen Sepharose CL-4B to a 50-ml polypropylene tube and fill the tube with CL-4B column buffer. Mix by inverting several times and let the Sepharose CL-4B settle by gravity for 10 to 15 min. Aspirate the buffer above the settled gel, removing also the unsettled “fines.” 2. Fill the tube two times with CL-4B column buffer—allow the Sepharose CL-4B to settle each time and remove the fines as in step 1. 3. Add 10 ml CL-4B column buffer and mix by inverting several times. Incubate the tube 10 min at 37°C, then proceed at room temperature. Outgassing may occur if the column is poured cold. The bubbles thus formed in the gel will interfere with the chromatography.

4. Break off the top of the 5-ml plastic pipet. Wearing gloves, use the 1-ml pipet to push a small piece (3- to 4-mm3) of silanized glass wool down to the tip of the 5-ml pipet. Push a 3-cm length of plastic tubing firmly onto the tip of the 5-ml pipet. Clamp the tubing and attach the column to the ring stand as shown in Figure 5.6.3. 5. With a pipet, carefully fill the column with the gel slurry from step 3. After a few minutes, release the clamp on the tubing and allow the column to flow. Periodically add more slurry to the column as the level drops until the volume of packed gel in the column is at the 5-ml mark. Ligation of Linkers or Adapters to Double-Stranded cDNA

6. Allow the level of buffer in the column to drop until it is just above the level of the gel and clamp the tubing to stop the flow. The column is ready to be loaded (see basic or alternate protocols).

5.6.6 Supplement 13


REAGENTS AND SOLUTIONS Buffer stock solutions Prepare ≥10 ml of each of the following stock solutions. Use autoclaved water and pass each solution through a sterile 0.45-µm filter. Store at room temperature unless otherwise indicated. 1 M Tris⋅Cl, pH 8.0 and pH 7.5 0.5 M EDTA, pH 8.0 3 M sodium acetate, pH 5.2 5 M NaCl (prepare 100 ml) 1 M MgCl2 7.5 M ammonium acetate 20% N-lauroylsarcosine (Sarkosyl) 1 M DTT; store at −20°C in tightly capped tube 0.1 M ATP, pH 7.0 (prepare 1.0 ml); neutralize as described in UNIT 3.4; store at −20°C From these stock solutions, prepare the following buffers, which should be checked for nuclease activity as described in UNIT 5.5. Enzyme buffers should be frozen in 200-µl aliquots at −80°C in screw-cap microcentrifuge tubes. Enzyme buffers prepared and stored as described will last for years. Several of these solutions are routine and may be already available. Nonetheless, to help ensure success, it is best to prepare separate stocks for critical applications such as library preparation.

CL-4B column buffer, 500 ml 5 ml 1 M Tris⋅Cl, pH 8.0 60 ml 5 M NaCl 1 ml 0.5 M EDTA, pH 8.0 2.5 ml 20% Sarkosyl 431.5 ml H2O Filter sterilize and store at room temperature 50× S-adenosylmethionine (SAM) 1 mg SAM 1.0 ml 50× SAM dilution buffer (see below) Prepare fresh just prior to use. Store dry SAM at −80°C for no longer than 2 months.

50× SAM dilution buffer, 7 ml 330 µl 3 M sodium acetate, pH 5.2 6.67 ml H2O Store in 1-ml aliquots Silanized glass wool Submerge the glass wool in a 1:100 dilution of a silanizing agent such as Prosil 28 (VWR) for 15 sec with shaking. Rinse the glass wool extensively with distilled H2O. Autoclave the glass wool for 10 min and store at room temperature.



Supplement 13



The most common method currently employed to create compatible ends on cDNA prior to cloning is the attachment of synthetic linkers (basic protocol). This has for the most part superseded homopolymeric tailing (Maniatis et al., 1982), since linkering is relatively efficient, and its use eliminates the need for the sometimes tricky procedure of titrating the tailing reaction conditions. Other methods, such as the sequential ligation of two different linkers (Maniatis et al., 1982), are now rarely used. The linkered cDNA may be cloned into a plasmid or a phage vector. The use of synthetic adapters that ligate at high efficiency instead of linkers eliminates the methylation and restriction digestion steps (alternate protocol). The recently developed noncomplementary adapter strategy (see below) may ultimately supersede all the above methods as vectors with appropriate sites become available. Size fractionation of cDNA is preferable to size fractionation of the initial mRNA primarily because DNA is considerably less susceptible to degradation than is RNA. In addition, the small fragments that are generated during the cDNA synthesis process (e.g., due to contaminating nucleases) are removed. These would otherwise be preferentially inserted and decrease the yield of long cDNA clones in the library. RNA enrichment procedures were undertaken in the past because cloning efficiencies then were not high enough to ensure the representation of very rare mRNAs in the library. Currently, the most common reason for using mRNA fractionation or enrichment is in preparation of subtracted libraries (UNIT 5.8; Hedrick et al., 1984). The strategy of noncomplementary adapters (Fig. 5.6.2), developed by Brian Seed and coworkers (Seed, 1987), overcomes the loss of library complexity due to self-ligation of the cDNA inserts or the vector. This enables a high efficiency for the ligation of the vector to the insert since there are no competing reactions and the vector ends are similarly noncomplementary and do not self-ligate. The much higher yield of desirable ligation products ultimately results in a greater number of clones in the library. Furthermore, the occasional “scrambles” produced as a consequence of two unrelated cDNAs ligating together during the vector ligation step are eliminated; however, “scrambles” at the stage of adapter ligation

remain possible, though infrequent. Adapters are favored over linkers in general because the methylation and restriction digestion steps are bypassed, thus simplifying the procedure. adapters are required in the alternate protocol procedure to ensure that all of the cDNA will have the proper, noncomplementary “sticky ends.” The use of EcoRI-NotI adapters for cloning with EcoRI compatible vectors introduces the rare NotI site at both ends of the insert, allowing the insert to be cut out of the vector as one fragment, which is not always possible with EcoRI linkers if there are internal EcoRI sites present. The development of noncomplementary adapters has enabled the production of highcomplexity cDNA libraries in multifunctional plasmid vectors such as CDM8 (Seed, 1987). These vectors permit library screening by functional expression in eukaryotic cells and production of single-stranded DNA for mutagenesis or subtraction, in addition to conventional hybridization methods. For some specialized applications and for selectable markers other than the supF present in CDM8, other vectors that employ the noncomplementary linker strategy (and use the same BstXI sequence) include several available from Invitrogen with different antibiotic resistances and eukaryotic selectable markers. Other available vectors include AprM8, an ampicillin-resistant version of CDM8 (L.B. Klickstein, unpublished results), and retroviral vectors in the pBabe series (Morgenstern and Land, 1990).

Critical Parameters In order to maximize the length and cloning efficiency of the cDNA, it is essential that contamination by endo- and exonucleases be avoided. EDTA, an inhibitor of most nucleases, should be present whenever feasible. Reagents, solutions, and enzymes should be of the highest quality obtainable. The SAM reagent in the methylation step is very unstable, must be freshly dissolved prior to use, and should not be kept >2 months, even if stored dry at −80°C and never thawed. Stabilized SAM is supplied free of charge with methylases purchased from New England Biolabs. Methylation conditions may be checked by methylating λ DNA under the conditions described in step 1 of the basic protocol, digesting methylated and unmethylated DNA with

5.6.8 Supplement 13


EcoRI, and comparing the two samples by agarose gel electrophoresis. Methylated λ DNA should be protected from EcoRI digestion. Digested linkers must be completely removed. Because of their small size, even as a small weight percentage of the total DNA, they will comprise a large mole fraction; thus, if not completely removed, most clones in the library will contain only linkers. Some investigators remove digested linkers by three sequential precipitations from 2 M ammonium acetate. However, column chromatography provides better resolution of the cDNA from the linkers. The CL-4B column also removes small doublestranded cDNA molecules. Purification of cDNA by gel electrophoresis after ligation of the linkers or adapters may be performed as an alternative to the CL-4B column, but the yield usually is not as high and the separation of linkers or adapters from cDNA is not as complete. If noncomplementary adapters are employed, both 5′ ends should be phosphorylated for optimum results. If complementary adapters are used, only the blunt end should have a 5′ phosphate. The other end is phosphylated after ligation to the cDNA. Because BstXI produces a cohesive end with a sequence not specified by the restriction site sequence, the adapters and vector must correspond—e.g., the cohesive ends of the adapters used for the cDNA must be complementary to that of the vector. The vectors CDM8, AprM8, pCDNAI, pCDNAII, and the pBabe series all use the same sticky ends (Fig. 5.6.2).

Troubleshooting Methylation and addition of linkers to double-stranded DNA are important to the successful construction of a cDNA library, but cannot be evaluated immediately. Unmethylated cDNA will clone efficiently and the problem will only be detected if the isolated inserts all end at an internal EcoRI site. This possibility may be minimized by prior evaluation of the quality of the SAM used in the methylation reaction (see reagents and solutions), or by use of the stabilized SAM that accompanies New England Biolab’s EcoRI methylase. Incomplete addition of linkers or adapters will be detected as a poor cloning efficiency in the next step of ligating the inserts to the vector. The problem may be any of the following: (1) cDNA was not blunted properly; (2) linkers or adapters were not kinased well or not annealed; (3) multiple linkers were not cut off with

EcoRI; (4) ligation reaction did not work. However, the problem is usually improperly blunted cDNA or linkers that do not ligate well. Evaluate the linkered cDNA (after the CL-4B column) by ligating 5% of the sample with no vector and running the reaction on a 1% agarose minigel next to an equal amount of unligated cDNA. Properly linkered cDNA should significantly increase in size as determined by ethidium bromide staining or autoradiography of the dried gel. Poorly linkered cDNA may often be salvaged by repeating the blunting and linkering steps with fresh reagents. Inadequate separation of linkers or adapters from cDNA will be detected as cloning efficiency that is too high and as clones with no inserts detectable by gel electrophoresis (white plaques with no inserts in λgt11 or plaques on C600hflA with λgt10 that have no insert). The remainder of this cDNA may be salvaged by repeating the CL-4B column chromatography.

Anticipated Results Approximately 50% to 70% of the starting radioactivity present in the blunt-ended, double-stranded cDNA should be collected after the CL-4B column. This typically represents 1 to 3 µg of cDNA. In a human tonsil library prepared in λgt11 where the inserts were thoroughly evaluated, the mean insert size was 1.4 kb and actin clones represented 0.34% of all recombinants. Three-quarters of the actin cDNAs were nearly full length (actin mRNA = 2.1 kb).

Time Considerations The methylation can be done on the same day as the blunting step from the cDNA synthesis protocol. The linker ligation is then set up overnight. EcoRI digestion and CL-4B chromatography are performed the following day, and the cDNA is either ligated to the vector overnight (UNIT 5.8) or stored as an ethanol precipitate overnight and further size selected by agarose gel electrophoresis the next day. At any ethanol precipitation in the procedure, the cDNA may be stored for several days as an ethanol precipitate. The use of adapters rather than linkers eliminates the need for the methylation and linker digestion steps. The adapted cDNA is immediately loaded onto the CL-4B column after the adapter ligation and is used directly from the column after ethanol precipitation. The subsequent ligation of the adapted cDNA and the vector is performed overnight, requiring the same amount of time with adapters as with linkers.



Supplement 13

Literature Cited

Key References

Hedrick, S.M., Cohen, D.I., Nielsen, E.A., and Davis, M.M. 1984. Isolation of cDNA clones encoding T cell-specific membrane-associated proteins. Nature (Lond.) 308:149-153.

Neve, R.L., Hanis, P., Kosik, K.S., Kurnit, D.M., and Donlon, T.A. 1986. Identification of the gene for the human microtubule-associated protein tau and chromosomal localization of the genes for tau and microtubule-associated protein 2. Brain Res. 387:271-80.

Maniatis, T., Fritsch, E.F., and Sambrook, J. 1982. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.

Seed, B. 1987. See above.

Morgenstern, J.P. and Land, H. 1990. Advanced mammalian gene transfer: high titre retroviral vectors with multiple drug selection markers and a complementary helper-free packaging cell line. Nucl. Acids Res. 18:3587-3596.

Contributed by Lloyd B. Klickstein Brigham and Women’s Hospital Boston, Massachusetts

Seed, B. 1987. An LFA-3 cDNA encodes a phospholipid-linked membrane protein homologous to its receptor CD2. Nature (Lond.) 329:840842.

Rachael L. Neve McLean Hospital Belmont, Massachusetts




PRODUCTION OF GENOMIC DNA AND cDNA LIBRARIES

SECTION IV

In this section the insert DNA (prepared in UNIT 5.2 or 5.3) is ligated to vector DNA and then the ligated DNA is propagated in Escherichia coli. The technical problems associated with joining DNA molecules together with ligase are described in UNIT 3.14. The creation of genomic and complete or subtracted cDNA libraries requires the production of large numbers of recombinant clones and this causes further complications. To recover large numbers of recombinant clones without using large amounts of packaging mixture, vector, or insert DNA, the optimal ratio of insert DNA to vector DNA must be determined prior to the production of the library. Without this optimization, the library will frequently not be large enough to contain the entire genome. After ligation of vector DNA to insert DNA the ligated DNA must be introduced into E. coli. The introduction of bacteriophage or cosmid DNA into E.coli is carried out most efficiently by packaging the DNA into bacteriophage particles then allowing these bacteriophage particles to infect E. coli (see UNIT 1.11 for a detailed description of this process). Unfortunately, plasmid DNAs cannot be packaged and thus must be introduced into E. coli by bacterial transformation (UNIT 1.8). Because of the high efficiency of introducing DNA into E. coli by the packaging procedure, we recommend producing genomic and complete or subtracted cDNA libraries using bacteriophage vectors whenever possible. In this section the protocols required to ligate vector DNA to insert DNA and to introduce these molecules into E. coli are described. The protocols are divided into two parts—one for genomic DNA libraries and the other for complete or subtracted cDNA libraries—because the amount of insert DNA required for genomic libraries is usually considerably more than for cDNA libraries.

Production of a Genomic DNA Library A number of small-scale ligations are performed using a set amount of vector and varying amounts of insert. Test ligations are transformed into bacteria (plasmid vectors) or packaged and plated on host bacteria (λ and cosmid vectors). The number of clones in the different ligations is compared, and the optimum ratio of vector to insert is indicated by the ligation with the most recombinant clones. A large-scale ligation is then set up using this optimum ratio. This protocol employs a bacteriophage vector; however, cosmid or plasmid vectors can be used with minor modifications (see commentary).


Determining the number of clones required to make a genomic DNA library is discussed in the chapter introduction. A library with a base of about 700,000 clones is required for a complete bacteriophage library of mammalian DNA. Materials Vector DNA (phage arms, cosmid arms, or linearized plasmid) Insert fragment (UNITS 5.3-5.6) 10× DNA ligase buffer (UNIT 3.4) T4 DNA ligase (measured in cohesive-end units; New England Biolabs; UNIT 3.14) Packaging extract (phage and cosmid) or competent E. coli (plasmid) LB or LB/ampicillin plates (UNIT 1.1) Top agarose containing 10 mM MgSO4 (UNIT 1.1) Additional reagents and equipment for plating, packaging, and titering bacteriophage (UNIT 1.11) Contributed by Thomas Quertermous Current Protocols in Molecular Biology (1988) 5.7.1-5.7.4 Copyright © 2000 by John Wiley & Sons, Inc.


5.7.1 Supplement 13

Ligating insert DNA to vector DNA 1. Perform a series of test ligations that bracket equimolar concentrations of vector and insert, i.e., insert/vector molar ratios of 5:1, 2:1, 1:1, 0.5:1, and 0.2:1. (Keep the vector constant and vary the insert DNA.) Since only the relative quantities of recombinants obtained are important, a small quantity of vector can be used for each reaction (40 to 100 ng). A control tube containing only ligated vector (tube no. 6 in Table 5.7.1.) is important to assess the yield of recombinants (see below). Conditions of the ligation reaction are as described in UNIT 3.14 (1× ligase buffer, 100 U T4 DNA ligase) and should be in a total volume of 5 to 10 µl. See Table 5.7.1 for a sample set of reactions. Vectors should be chosen as described in UNIT 5.1. Genomic DNA libraries should be constructed only with vectors that provide a selection for insert DNA. Table 5.7.1

Sample Set of Reactions for Ligating λ Vector to Insert DNA

Reaction component (µl) Vector DNA (100 ng/tube) Insert DNA (20 kb, 100 ng/µl) Insert DNA (20 kb, 50 ng/µl) Insert DNA (20 kb, 10 ng/µl) 10× ligation buffer Water T4 DNA ligase

Tube number 3 4

1

2

5

6

1

1

1

1

1

1

2

—

—

—

—

—

—

2

1

0.5

—

—

—

—

—

—

1

—

0.5 1 0.5

0.5 1 0.5

0.5 2 0.5

0.5 2.5 0.5

0.5 2 0.5

0.5 3 0.5

2. Package the phage recombinants as described in UNIT 1.11. Normally, commercially prepared packaging extract is designed for approximately 1 µg of λ DNA, so it is possible to divide such an aliquot of packaging extract among several test ligations. 3. Dilute packaged extract to 0.5 ml and plate 1 µl and 10 µl of packaged bacteriophage per plate. Grow overnight. 4. Compare the number of plaques on the plates to determine the optimum relative concentrations of arms and insert. The yield of plaques with insert DNA (from ligation tubes 1 to 5) should be at least twice and preferably 5 times the yield of colonies on the control plate (from tube 6) containing vector DNA alone. 5. Prepare a large-scale ligation reaction using the optimum ratio of insert DNA to vector DNA determined in step 4. The amount of DNA is determined by the size of the library that is desired. See UNIT 5.1 to determine total number of desired recombinants. If the expected number of recombinants that can be made uses too much insert DNA or vector DNA, either the insert DNA or the vector DNA is probably defective. The quality of the vector or insert DNA can be tested by ligating them together and fractionating the ligated DNA on a gel (see UNITS 3.14, 2.5A, & 2.5B) using commercially prepared extracts.

Production of a Genomic DNA Library

Introducing ligated DNA into E. coli 6. Package the ligated DNA. Remember that bacteriophage packaging extracts can be saturated by too much DNA.

5.7.2 Supplement 13


Because producing extracts that efficiently package DNA can be quite difficult, we recommend the use of commercially available packaging kits (see UNIT 1.11). The number of recombinants that can be obtained with 1 ìg λ DNA from large-scale ligation will depend on the efficiency of the packaging extract or competent cells, but can be as high as 108 for phage vectors. For cloning from size-fractionated DNA, this represents a vast excess of recombinants. It is frequently possible to obtain over 106 recombinant phage from the test ligations, and this is often enough recombinants to clone a single-copy gene. Thus, it is useful to save the test ligations.

7. Package and titer the large-scale ligation and store at 4°C. Plate the bacteriophage for amplification or screening as soon as possible after the titer is known, as the titer will drop 3- to 10-fold within 48 hr of packaging.

COMMENTARY Background Information The theory of the ligation reaction is presented in UNIT 3.16 and is well outlined by Williams and Blattner (1980). From these theoretical considerations it is possible to predict the optimum concentration of vector and insert. However, because of the difficulty of estimating small quantities of DNA and the inability of some ends to ligate, it is helpful to conduct a series of test ligations as outlined here to determine the ratio of reagents that will result in the maximum number of clones. The theory of ligation reactions suggests that the optimum concentration for a ligation reaction varies for different sized molecules. However, the conditions suggested here are optimal for most vectors and inserts used for making genomic and cDNA libraries. The production of a cosmid library is identical to the procedure described here except that a cosmid vector is used rather than a bacteriophage vector. Different amounts of cosmid vector are used because cosmids are smaller (about 5 kb) than bacteriophages (50 kb). Thus the weight of vector DNA should be reduced 10fold. Production of a plasmid library involves the same procedure except that ligated DNA is introduced into E. coli cells by transformation (see UNIT 1.8) rather than by packaging.

Critical Parameters The quality of the insert DNA and vector DNA are critical to the success of these experiments. The most frequent cause of failure is that one of these DNAs has defective ends and thus the molecules are not able to ligate efficiently. Quality of packaging extracts is also critical. Normally, commercial packaging extracts are of high quality and yield large numbers of bacteriophage. However, an occasional bad lot

of packaging extract is obtained.

Troubleshooting A large number of clones in all the test ligations, or a large number of clones with extremely small quantities of insert, may indicate a high background of nonrecombinant clones. Background is determined by ligating the vector in the absence of insert. A large number of clones in this control ligation indicates inadequate dephosphorylation, inadequate purification from stuffer, or possibly that the bacterial strain being used for genetic selection is not correct. If few recombinants are obtained, there are a number of possible explanations. The packaging extract should be evaluated by appropriate control experiments. Usually, the commercially available packaging extract kits contain a suitable control fragment. At the same time a test ligation can be run on an agarose gel and compared to the unligated vector and insert. If both the insert and vector DNA appear intact (i.e., not degraded), either the sticky ends of the vector or insert have been damaged or one of the DNA samples (usually the insert) contains an inhibitor of the ligase reaction. Clonability of these two reagents can be evaluated by determining the ability of insert or vector DNA to ligate to itself or to an appropriate control. Inhibiting contaminants can sometimes be removed by phenol/ chloroform extraction followed by ethanol precipitation, or by passing the DNA over an Elutip column (UNIT 2.6).

Anticipated Results The number of clones obtained will depend on a number of factors including the efficiency of ligation, transformation, and packaging. At best, in a λ vector, over 108 recombinants per µg of insert DNA can be achieved. Cloning into



plasmid usually produces fewer recombinants per quantity of insert because transformation into competent E. coli is 10 to 100 times less efficient than the in vitro packaging of λ phage. In a plasmid, it may be necessary to use 1 to 2 µg insert DNA to achieve 106 recombinant clones. Cosmid clones usually are obtained at 105 to 106 colonies per µg insert DNA.

Time Considerations Test ligations are incubated overnight at 13°C (alternatively, incubate 2 hr at room temperature). The ligation reactions are packaged and then titered on the second day. Results are available late on the second day or the morning of the third day.

Literature Cited Kaiser, K. and Murray, N.E. 1984. The use of phage lambda replacement vectors in the construction of representative genomic DNA libraries. In DNA Cloning, Vol. 1: A Practical Approach (D. Glover, ed.) pp. 1-47. IRL Press, Oxford. Rodriguez, R.L. and Tait, R.C. 1983. Recombinant Techniques: An Introduction. Addison-Wesley, Reading, MA. Williams, B.G. and Blattner, F.R. 1980. Bacteriophage lambda vectors for DNA cloning. In Genetic Engineering, Vol. 2 (J.K. Setlow and A. Mullander, eds.) p. 201. Plenum, NY. Describes the ligation reaction as applied to cloning with λ vectors.

Contributed by Thomas Quertermous Massachusetts General Hospital Boston, Massachusetts

Production of a Genomic DNA Library


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Construction of Bacterial Artificial Chromosome (BAC/PAC) Libraries

UNIT 5.9

Large-insert genomic libraries are important reagents for physical mapping of large chromosomal regions, for isolation of complete genes including all regulatory sequences, and for use as intermediates in DNA sequencing of entire genomes. There is a need for libraries of large insert clones in which rearrangements and cloning artifacts are minimal. With recent developments in host/vector systems, it is now possible to efficiently create large-insert libraries using low-copy-number bacterial plasmid vectors. Suitable vector systems have been developed using Escherichia coli F factor or P1 phage replicons, which are capable of producing large insert clones by bacterial transformation. Construction of BAC and PAC libraries is detailed first (see Basic Protocol). Two vectors, pCYPAC2 (Ioannou et al., 1994) and pPAC4 (E. Frengen et al., unpub. observ.) have been used for preparing PAC libraries, and a new BAC vector pBACe3.6 (Frengen et al., 1999) has been developed for construction of BAC libraries. All vectors are illustrated in Figure 5.9.1. Support Protocol 1 describes preparation of PAC or BAC vector DNA for cloning by digestion with BamHI or EcoRI, simultaneous dephosphorylation with alkaline phosphatase, and subsequent purification through pulsed-field gel electrophoresis (PFGE). For the preparation of high-molecular weight DNA for cloning, Support Protocols 2 and 3 provide procedures for embedding total genomic DNA from lymphocytes or animal tissue cells, respectively, in InCert agarose. Support Protocol 4 details the next steps for the genomic DNA: partial digestion with MboI or with a combination of EcoRI endonuclease and EcoRI methylase (including appropriate methods for optimizing the extent of digestion), and subsequent size fractionation by preparative PFGE. Finally, Support Protocol 5 covers the isolation of BAC and PAC plasmid DNA for analyzing clones. Isolation of BAC and PAC plasmid DNA is somewhat more difficult than standard plasmid extraction, because it requires more attention to cell concentration during extraction and resuspension of ethanol-precipitated DNA. In order to construct, for example, a 10-fold redundant human library with 150-kb average insert size, 200,000 clones must be generated. The EcoRI-EcoRI ligation efficiency seems to be higher than the MboI-BamHI ligation. A reasonable transformation efficiency from EcoRI-EcoRI ligation is ∼2,000 colonies per transformation using 2 µl ligation mixture and 20 µl electrocompetent cells. In comparison, the efficiency of MboI-BamHI ligation is ∼800 colonies per transformation. Thus, it is necessary to perform at least 100 individual transformations for an EcoRI partially digested 10-fold redundant library, and 250 individual transformations for an MboI partially digested library. In order to perform 100 electroporations, 200 µl of concentrated ligation mixture is required, thus five individual 250-µl ligations must be prepared. For 250 transformations, 500 µl of ligation mixture is needed, so thirteen individual 250-µl ligations are required. Note that it will be necessary to repeat the Basic Protocol and Support Protocol 4 multiple times until enough clones are generated to complete the library. Steps 24 to 30 of the Basic Protocol are designed to handle the amount of size-fractionated genomic DNA that can reasonably and comfortably be generated from a single preparation as described in Support Protocol 4. It is advisable to prepare at least 20 DNA plugs (20 to 40 µg/plug) for the entire library construction procedure; up to four plugs may be needed to optimize the partial digestion, and as many as sixteen plugs may be needed for library construction. CAUTION: Radioactive, biological, and chemical substances require special handling; see APPENDIX 1F & 1H for guidelines.


Contributed by Kazutoyo Osoegawa, Pieter J. de Jong, Eirik Frengen, and Panayiotis A. Ioannou Current Protocols in Molecular Biology (2001) 5.9.1-5.9.33

5.9.1

Copyright © 2001 by John Wiley & Sons, Inc.

Supplement 55

A

BamHI (1)

NotI (18,720)

ScaI (21) ScaI (1789) T7 ScaI (2723) pUC-link BamHI (2739) P1 lytic SP6 NotI (2778) replicon sacBll pCYPAC2 loxP r 18.8 kb Km

P1 plasmid replicon

B

BamHI (1) SacII (8) EcoRI (10) SacI (20) MluI (22) NsiI (36) ApaLI (766)

NotI (11,456) Tn7att PI-SceI loxP 511

T7 Cmr

ApaLI (2012) pUC-link

pBACe3.6 11.5 kb F plasmid replicon

ApaLI (2509) Eco RI (2728) SacI (2738) NsiI (2784) MluI (2790) SacI (2800) EcoRI (2802) SacII (2810) BamHI (2811) NotI (2850)

SP6

sacBII

loxP

C

NotI (19,466) BamHI (1) Tn7 att PI-SceI ScaI (21) loxP 511 ScaI (1789) ScaI (2723) oriP T7 pUC-link BamHI (2739) SP6 NotI (2778) sacBII Kanr pPAC4 19.5 kb P1 plasmid replicon

bsrr cassette

cos hCMV loxG

Construction of BAC/PAC Libraries

5.9.2 Supplement 55


Figure 5.9.1 (at left) The (A) pCYPAC2, (B) pBACe3.6, and (C) pPAC4 vectors contain common features for positive selection of cloned inserts. The insert-containing E. coli cells can be grown in preference over cells containing plasmids without an insert by including sucrose in the medium. Sucrose is converted into toxic metabolites by levansucrase encoded by the sacBII gene present in all three vectors. The pUC-link segment contains a functional high-copy-number plasmid that includes the ampicillin resistance gene (Apr; not shown). The pUC-link sequence is removed in the cloning procedure and is not present in the PAC and BAC clones. The digested genomic DNA fragment replaces the fragment containing the pUC-link in the PAC or BAC vectors. The loxP recombination sites are not used in cloning. A detailed description of vector elements is given elsewhere (see, Background Information, vector elements). (A) The pCYPAC2 vector contains the P1 plasmid replicon, which maintains recombinant clones at around one copy per cell. Alternatively, the P1 lytic replicon can be induced to provide higher copy numbers by adding the lac inducer IPTG into the medium. The kanamycin resistance gene (Kmr) is also present in this vector. (B) The pBACe3.6 vector contains the chloramphenicol resistance gene (Cmr) and the F plasmid replicon, maintaining recombinant clones at one copy per cell. The multiple cloning sites flanking the pUC-link segment allow positive selection for cloned inserts using any of the restriction enzymes BamHI, SacII, EcoRI, SacI, MluI, or NsiI. The Tn7att sequence permits specific Tn7-based retrofitting of BAC clones. (C) The pPAC4 vector contains four elements from pCYPAC2: the P1 plasmid replicon, the kanamycin resistance gene, the sacBII gene, and the pUC-link. The P1 plasmid replicon ensures low-copy-number maintenance of recombinant clones; the P1 lytic replicon is removed. In addition, the oriP and bsrr cassette have been included to facilitate the use of PAC clones for functional analysis of genes carried on the cloned inserts. The Tn7att sequence is also present, enabling specific retrofitting of PACs.

PREPARATION OF BAC/PAC CLONES USING pCYPAC2, pPAC4, OR pBACe3.6 VECTOR

BASIC PROTOCOL

Preparation of large-insert bacterial and P1-derived artificial chromosome (BAC and PAC) clones is conceptually similar to procedures used for preparing small plasmid clones. A single vector fragment with dephosphorylated BamHI or EcoRI ends (see Support Protocol 1) is ligated to size-fractionated MboI- or EcoRI-digested genomic DNA fragments that are recovered from agarose slices using an electroelution procedure (see Support Protocol 4). Ligations are performed at low genomic DNA concentrations to favor the formation of circular molecules. A number of 120- to 300-kb fractions from the preparative gel are tested initially to identify the fraction giving the largest average insert size at an acceptable cloning efficiency. Once the optimal fraction is identified, a large-scale ligation is performed to make sufficient clones for a several-fold redundant library. Size fractionation of the partially digested DNA is important to select for the desired size fragments and to remove smaller digestion products, which otherwise would result in an overwhelming background of small BAC or PAC clones. Circular plasmid ligation products are introduced directly into host cells by electroporation. Although this procedure is conceptually easy, there are several aspects of the Basic and Support protocols that are critical for successful preparation of BAC and PAC libraries (see Critical Parameters). Materials ≥2 to 10 ng/µl size-fractionated MboI- or EcoRI-digested genomic DNA (see Support Protocol 4) 10 to 50 ng/µl pBACe3.6, pCYPAC2, or pPAC4 vector DNA prepared for cloning (see Support Protocol 1) 1 Weiss U/µl T4 DNA ligase (Life Technologies) and 5× buffer (see recipe) 0.5 M EDTA, pH 8.0 (APPENDIX 2) 10 mg/ml proteinase K



Supplement 55

100 mM phenylmethylsulfonyl fluoride solution (PMSF; see recipe) TE/PEG solution: 0.5× TE buffer, pH 8.0 (APPENDIX 2) containing 30% (w/v) polyethylene glycol 8000 (PEG 8000) Electrocompetent bacterial cells (ElectroMAX DH10B; Life Technologies) SOC medium (Life Technologies; also see UNIT 1.8, but reduce yeast extract to 0.5%) LB plates (UNIT 1.1) containing 5% (w/v) sucrose and either 25 µg/ml kanamycin (for PAC clones) or 20 µg/ml chloramphenicol (for BAC clones): 100 × 15–mm petri dishes for test transformation 22 × 22–cm trays for picking colonies LB medium (UNIT 1.1) containing 20 µg/ml chloramphenicol (BAC clones) or 25 µg/ml kanamycin (PAC clones) TE buffer, pH 8.0 (APPENDIX 2) NotI restriction endonuclease and buffer (New England Biolabs) 1% (w/v) ultrapure agarose solution (Life Technologies) 0.5× TBE buffer (APPENDIX 2) Low-range PFG markers in agarose containing a mixture of lambda HindIII fragments and lambda concatemers (New England Biolabs) 0.5 µg/ml ethidium bromide in 0.5× TBE buffer (APPENDIX 2) 80% (v/v) glycerol, sterile Dry ice/ethanol bath 16° and 37°C water baths 0.025-µm-pore-size microdialysis filters (Millipore): 25-mm diameter for small-scale test ligation and 47-mm diameter for large-scale ligation Wide-bore pipet tips, sterile Disposable microelectroporation cuvettes with a 0.15-cm gap (Life Technologies or equivalent) Electroporator (Cell Porator equipped with a voltage booster; Life Technologies or equivalent) 15-ml snap-cap polypropylene tubes, sterile Orbital shaker, 37°C Automated plasmid isolation system (AutoGen 740, Integrated Separation Systems, optional) Flexible plastic 96-well plate (Falcon or equivalent) Contour-clamped homogeneous electrical field (CHEF; UNIT 2.5B) apparatus (Bio-Rad) or field-inversion gel electrophoresis (FIGE; UNIT 2.5B) apparatus (Bio-Rad or equivalent) Digital imager (Alpha Innotech IS1000 or equivalent) 50-ml disposable centrifuge tube (Corning or equivalent) Additional reagents and equipment for modified alkaline lysis preparation of BAC or PAC clone DNA (see Support Protocol 5; optional) CAUTION: To prevent shearing, use sterile wide-bore pipet tips for all steps involving the handling of genomic DNA. Perform small-scale test ligation 1. Mix ∼50 ng of each 120- to 300-kb size fraction of MboI- or EcoRI-digested genomic DNA (see Support Protocol 4) with 25 ng BamHI- or EcoRI-digested and dephosphorylated pBACe3.6 vector DNA, or 50 ng BamHI-digested and dephosphorylated pCYPAC2 or pPAC4 vector DNA prepared for cloning (see Support Protocol 1). Construction of BAC/PAC Libraries

Suitable ligation controls include reactions with vector only or without ligase.

5.9.4 Supplement 55


The vector fragment is 8.7 kb for pBACe3.6, ∼15.5 kb for pCYPAC2, and 16.8 kb for pPAC4. The average size of fractionated genomic DNA is ∼150 to 200 kb. Thus, a 10/1 molar ratio of vector to insert is obtained by using approximately half quantity of pBACe3.6 vector to insert DNA in the ligation reaction, or equal quantities of vector and insert DNA for PAC cloning. The genomic DNA concentration in the final reaction is maintained at a low value (1 ng/ìl) to favor circle formation over concatemer formation. Ligation schedules will vary somewhat depending on the restriction endonuclease used in preparing the vector and insert DNA. For a typical EcoRI-EcoRI ligation schedule, perform step 1 through step 6 in a day and then dialyze the solution against TE/PEG solution at 4°C overnight. For BamHI-MboI cloning, incubate the ligation mixture in a thermal cycler at 16°C for 8 hr, then hold at 4°C overnight, and start step 4 the following day.

2. To each tube, add 10 µl of 5× T4 DNA ligase buffer (1× final) and sterile water to bring the total volume to 50 µl. Mix very gently. 3. Add 1 Weiss unit T4 DNA ligase and incubate at 16°C for 4 hr for EcoRI-EcoRI cloning or 8 hr for BamHI-MboI cloning. Keep in mind that the vector preparation can always contain a very low fraction of shortened fragments caused by BamHI or EcoRI degenerate (star) activities. Ligation could occur at those unspecific sites after long incubation, which could result in a small portion (200,000 clones. It is important to achieve as high a cloning efficiency as possible with the largest possible insert size, because this will reduce the number of electroporations that are subsequently required.

Isolate and characterize individual test clones 19. Pick 40 clones with a sterile toothpick and grow the cells overnight in 1.5 ml LB medium containing 20 µg/ml chloramphenicol (BAC clones) or 25 µg/ml kanamycin (PAC clones). 20. Extract DNA using an automated plasmid isolation system (AutoGen 740) or the modified alkaline lysis procedure (see Support Protocol 5). 21. Dissolve DNA (0.5 to 1 µg) in 100 µl TE buffer. Digest 5 to 10 µl DNA with 0.1 U NotI in a 20-µl volume to separate the vector and insert DNA fragments. It is convenient to perform the reaction in a flexible 96-well plate at 37°C for 2 hr.

22. Analyze digested DNA using a CHEF or a FIGE apparatus. Use a 1% agarose gel, 0.5× TBE buffer, low-range PFG markers, and the following conditions: For CHEF: 14°C, 6 V/cm, 16 hr, 0.1 to 40 sec pulse time, and 120° angle. For FIGE: Room temperature, 180 V forward voltage, 120 V reverse voltage, and 16 hr with 0.1 to 14 sec switch time linear shape. The FIGE system is much less expensive than CHEF, and is able to obtain enough resolution for this purpose.

23. Stain the gel with 0.5 µg/ml ethidium bromide solution and calculate the molecular weight of inserts using a digital imager. Most of the clones will give two bands—a vector fragment (8.7 kb for pBACe3.6, 15.5 kb for pCYPAC2, and 16.8 kb for pPAC4) and the insert DNA of variable size. If the insert contains internal NotI sites, additional bands will be seen. Incomplete digestion usually leads to a characteristic doublet, where the two bands differ by the size of the vector.

Create library 24. Repeat the ligation procedure using a size fraction of genomic DNA that gives the desired average insert size and cloning efficiency. Scale up the ligation reaction to 500 to 1000 µl using the entire remaining electroeluted insert DNA. The ligation reaction is usually performed in several microcentrifuge tubes at 250 ìl per tube. Otherwise, the increase in volume may cause the solution to be dispersed into the water during subsequent dialysis, even though a 47-mm-diameter filter is used.

25. Dialyze and concentrate the ligation mixture as described (steps 6 to 10), but use a 47-mm-diameter microdialysis filter for each 250-µl aliquot. After concentration, ∼40 ìl should be recovered from each 250-ìl ligation solution.

26. Add 12 µl concentrated DNA to tubes containing ∼110 µl each of electrocompetent bacterial cells. Mix well by gentle stirring using a wide-bore pipet tip. Thaw enough electrocompetent cells to complete all transformations using the entire recovered ligation mixture. Construction of Recombinant DNA Libraries


Supplement 55

27. Perform identical transformations at 20 µl per electroporation and collect the transformed cells from 10 electroporations into a 50-ml disposable centrifuge tube containing 5 ml SOC medium at room temperature. The transformation efficiency will drop if the medium containing transformed cells is kept at 4°C during the transformation experiment.

28. Incubate the cells on an oribital shaker at 200 rpm at 37°C for 1 hr. 29. Add 800 µl of 80% sterile glycerol solution into the bacterial culture and mix very well. Spread 200 µl on two individual 100 × 15–mm LB plates containing 5% sucrose and the appropriate antibiotic to examine the titer from each tube. The amount of culture spread per plate is determined based on the titer estimated from the test ligation and transformation (step 18). The number of colonies on a plate should be between 300 and 700 to estimate the accurate titer for colony picking. It is easier to homogeneously spread 200 ìl culture when 300 ìl SOC medium is also added to the plate.

30. Freeze the remaining cells in a dry ice/ethanol bath and keep at −80°C until colony picking can be scheduled. To pick colonies, spread culture at 1600 clones/plate on 22 × 22–cm LB plates containing 5% sucrose and the appropriate antibiotic. Grow overnight at 37°C. The colony picking is usually scheduled within a couple of months. The frozen cells should be able to be stored for at least 1 year. SUPPORT PROTOCOL 1

PREPARATION OF BAC/PAC VECTOR FOR CLONING The integrity of the vector’s cohesive ends and the absence of uncut vector molecules are prerequisites for success in preparing BAC and PAC libraries. This protocol describes steps for preparing the vectors to maximize cloning efficiency and to minimize the level of nonrecombinant clones. Preparation of pBACe3.6, pCYPAC2, and pPAC4 for cloning involves a single restriction endonuclease digestion with BamHI or EcoRI. These plasmids contain the pUC-link insert at the cloning site, thus inactivating the sacB gene and enabling clones containing the original plasmid to grow in the presence of sucrose. Clones containing uncut vectors cannot be distinguished from truly recombinant clones, and great care should be taken to ensure complete digestion of the original vector with BamHI or EcoRI during vector preparation. Depending on the source and activity of the different enzymes used, it is necessary to optimize the conditions for each digestion. The following conditions have been found to yield good vector preparations using the recommended enzymes.


Additional Materials (also see Basic Protocol) pBACe3.6, pCYPAC2, or pPAC4 stock in E. coli DH10B cells (P. de Jong; [email protected]) LB plates (UNIT 1.1) containing: 25 µg/ml kanamycin (for PAC) or 20 µg/ml chloramphenicol (for BAC) 5% (w/v) sucrose and either kanamycin or chloramphenicol 5% (w/v) sucrose, 100 µg/ml ampicillin, and either kanamycin or chloramphenicol BamHI and EcoRI restriction endonucleases and 10× buffers (New England Biolabs or equivalent) 0.7% (w/v) agarose gels (for standard electrophoresis) Calf intestine alkaline phosphatase (AP; Boehringer Mannheim) 10 mg/ml proteinase K (Boehringer Mannheim) stock solution

5.9.8 Supplement 55


95% (v/v) ethanol 1.0% (w/v) ultrapure agarose solution (Life Technologies; for CHEF system) 6× loading buffer (UNIT 2.5A), not containing xylene cyanol FF or SDS 1-kb ladder or lambda HindIII markers T4 polynucleotide kinase (New England Biolabs) 30% (w/v) polyethylene glycol (PEG) 8000 1.5-mm-thick electrophoresis comb for CHEF apparatus Dialysis tubing of 3⁄4 in. diameter, mol. wt. exclusion limit of 12,000 to 14,000 daltons (Life Technologies or equivalent) Dialysis clip Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5A), plasmid extraction (e.g., UNIT 1.7), CsCl/ethidium bromide equilibrium centrifugation (e.g., UNIT 1.8), electroelution (see Support Protocol 4), and ethanol precipitation (UNIT 2.1A) Test vector DNA 1. Streak recombinant DH10B cells harboring BAC or PAC vectors onto an LB plate containing the appropriate antibiotic (chloramphenicol or kanamycin, respectively). Incubate overnight at 37°C. 2. Isolate five single colonies and inoculate into separate 15-ml snap-cap polypropylene tubes containing 3 ml LB medium with the appropriate antibiotic. Grow cultures overnight at 37°C. 3. Use 1.5 ml of each culture to prepare DNA using an automated plasmid isolation system (AutoGen 740) or the modified alkaline lysis protocol (see Support Protocol 5). Store the remainder of each culture at 4°C. 4. Resuspend DNA in 100 µl TE buffer. 5. Digest 5 µl DNA from each preparation with NotI, 5 µl with BamHI, and 5 µl with EcoRI, using the manufacturers’ recommended conditions. 6. Analyze digested DNA in a 0.7% agarose gel (UNIT 2.5A). Discard clones that contain any rearrangements of the vector. PAC vectors do not contain EcoRI sites. Otherwise all enzymes should liberate pUC19-link from the vectors, producing fragments of ∼2.7 kb (the pUC19 stuffer fragment) and ∼8.7 kb (pBACe3.6), 15.5 kb (pCYPAC2), or 16.8 kb (pPAC4).

Prepare vector for large-scale ligation 7. Dilute the selected culture into 1 liter LB medium containing the appropriate antibiotic. Grow to saturation at 37°C. This culture is used for making a large-scale preparation of the PAC or BAC vectors.

8. Prepare a crude lysate of the culture by the alkaline lysis or cleared lysate method. Any standard protocol for plasmid extraction may be used here (UNIT 1.7).

9. Purify vector plasmid from crude lysate by CsCl/ethidium bromide equilibrium centrifugation. Any standard protocol for CsCl/ethidium bromide equilibrium centrifugation may be applied (see UNIT 1.8). Construction of Recombinant DNA Libraries


Supplement 55

Digest vector with BamHI or EcoRI 10. To establish the minimal amount of restriction enzyme required for complete digestion of the vector, first digest 50 ng vector DNA with 0.1, 0.2, 0.5, and 1 U of either BamHI or EcoRI for pBACe3.6, and BamHI for PAC vectors, in separate 10-µl reactions for 1 hr at 37°C. Perform conventional agarose gel electrophoresis and ethidium bromide staining to view the digestion. It is essential to establish the minimal amount of enzyme required for complete digestion. If too little enzyme is used, many nonrecombinant clones containing undigested vector can be expected. On the other hand, excess enzyme results in a very high ratio of noninsert clones, which are most likely induced by star activity.

11. Next, to establish the maximal amount of DNA in the reaction (and the minimum scale up volume), digest 100, 200, and 400 ng vector DNA with 2, 4, and 8 times the minimal amount of enzyme in separate 10-µl reactions (total 9 reactions) for 1 hr at 37°C. Perform conventional agarose gel electrophoresis and ethidium bromide staining to view the digestion. High-quality vector preparation is difficult. If necessary, the vector can be digested with ApaLI for pBACe3.6 or ScaI for pCYPAC2 and pPAC4 to destroy the pUC-stuffer fragment in the vectors nearly completely. Note that a precipitation step is needed in order to change the reaction buffer and that optimal enzyme concentration must be determined.

12. Incubate ∼30 µg vector DNA with the defined amount of the appropriate enzyme at 37°C for 15 min. 13. Add 1 U calf intestine AP and continue incubating at 37°C for 1 hr. 14. To stop the reaction, add 0.5 M EDTA to a final concentration of 15 mM, add 10 mg/ml proteinase K to 200 µg/ml final, and incubate at 37°C for 1 hr. 15. To inactivate proteinase K, add 100 mM PMSF solution to 2 mM final and incubate at room temperature for 1 hr. Perform PFGE 16. During step 15, clean a 1.5-mm-thick electrophoresis comb with 95% ethanol and cover teeth with autoclave tape to create a large preparative slot with sufficient space to load the reaction mixture. Leave enough empty wells for a 1-kb ladder or lambda-HindIII markers on both sides and at least one empty well between the markers and the preparative slot. 17. Prepare a 1% agarose gel in a CHEF mold using the preparative comb. While the gel is solidifying, add 2 liters of 0.5× TBE buffer to the CHEF apparatus tank and equilibrate the unit at 14°C. 18. Place the gel in the precooled unit. Add 6× gel loading buffer to the samples (final 1×) and load samples in the large preparative well. Load 1-kb ladder or lambda HindIII markers in both side wells. Perform electrophoresis at 14°C and 6 V/cm for 16 hr with 0.1 to 40 sec pulse time at 120° angle. Use of xylene cyanol FF in the gel loading buffer is not recommended because this dye migrates with the vector band under these electrophoresis conditions.


Recover vector DNA 19. Cut away two flanking gel slices that contain the markers as well as 1 to 2 mm from each side of the central vector lane. Wrap the remaining middle portion of the preparative lane in plastic wrap and store at 4°C. 20. Stain the flanking slices with ethidium bromide to detect the major vector fragment.



21. Cut linear vector DNA lacking the pUC-stuffer fragment from the central lane and recover by electroelution (see Support Protocol 4, steps 38 to 46). 22. Load sample into 3⁄4-in.-diameter dialysis tubing with a 12,000- to 14,000-Da mol. wt. exclusion limit, and dialyze once against 1 liter TE buffer at 4°C for >2 hr. 23. Recover solution from the dialysis tubing and precipitate DNA with sodium acetate and either ethanol or isopropanol (UNIT 2.1A). Dissolve DNA in 100 µl TE buffer. Check efficiency of dephosphorylation 24. Place four microcentrifuge tubes on ice and number them 1 to 4. Add 4 µl of 5× T4 DNA ligase buffer and 50 ng of digested and AP-treated vector DNA into the tubes, and adjust the volume to 20 µl with sterile distilled and deionized water. 25. Add 1 U T4 polynucleotide kinase to tubes 3 and 4. Incubate all four tubes at 37°C for 1 hr. T4 polynucleotide kinase will be active in 1× ligase buffer.

26. Heat at 65°C for 20 min, then place on ice to inactivate the enzyme. 27. Add 1 U T4 DNA ligase to tubes 2 and 4. Incubate all four tubes at 16°C for >4 hr. Thus, tube 1 contains no enzyme, tube 2 contains T4 DNA ligase, tube 3 contains T4 polynucleotide kinase, and tube 4 contains T4 polynucleotide kinase and T4 DNA ligase.

28. Run samples in a 0.7% agarose gel, stain the gel with ethidium bromide, and view. Closed circular DNA should not be seen in samples 1, 2, and 3. Sample 4 should have closed circular DNA but no linear DNA.

29. Perform a ligation using all the remaining vector DNA solution in a 200-µl reaction volume with 4 U T4 DNA ligase. 30. Purify the linear monomeric vector DNA away from the background of ligated vector by PFGE in a CHEF apparatus (see steps 16 through 23). Check ratio of background clones 31. Perform test ligations using vector DNA alone (25 ng pBACe3.6, 50 ng PAC vectors) and vector plus 50 ng control DNA (see Basic Protocol, steps 1 to 5). 32. Dialyze against water and 30% PEG 8000, and perform transformation as described (see Basic Protocol, steps 6 to 16). 33. Spread transformed cells on LB plates containing 5% sucrose and either kanamycin or chloramphenicol, as appropriate. Grow overnight at 37°C. Kanamycin/sucrose and chloramphenicol/sucrose plates are used to determine the level of recombinant clones and nonrecombinant clones. If the number of colonies obtained from vector DNA alone is high (>100), vector must be prepared again, beginning with enzyme titration (step 10). If the plasmid is isolated from these background clones and analyzed by PFGE after NotI digestion, smaller vector sizes will be observed. Presumably, star activity causes nonspecific digestion in the sacB gene (either promoter or coding sequence), thus inactivating the gene, and the nonspecifically digested, nondephosphorylated DNA is circularized by self-ligation. Thus, these colonies can grow on sucrose plates without insert DNA.

34. To examine the complete removal of stuffer fragment from the vector, spread the transformed cells with self-ligated vector on LB plates containing 5% sucrose, 100 µg/ml ampicillin, and either kanamycin or chloramphenicol, as appropriate. Grow overnight at 37°C. Kanamycin/ampicillin/sucrose and chloramphenicol/ampicillin/sucrose plates are used to determine the remaining level of undigested vector containing the pUC-link. Ideally, there should be zero AmpR colonies.



Supplement 55

SUPPORT PROTOCOL 2

PREPARATION OF HIGH-MOLECULAR-WEIGHT DNA FROM LYMPHOCYTES IN AGAROSE BLOCKS Construction of a total genomic library demands that some consideration be given to the source of DNA to be used. Although DNA from cultured cell lines has been extensively used in constructing cosmid libraries, it is well known that passage in tissue culture may result in chromosomal rearrangements; therefore, it is considered more desirable to use DNA obtained directly from human or animal sources. A convenient source of DNA for this purpose is whole blood, and more specifically, the circulating lymphocytes. Assuming an average of ∼5 × 106 lymphocytes per milliliter of blood and 6 pg DNA per cell, as much as 30 µg DNA can be recovered per ml of blood, making it a very convenient and inexpensive source of DNA. This method depends on preferential lysis of red blood cells (RBC) using a gentle procedure that does not affect lymphocytes. The method can be applied equally well to cultured cells by omitting the RBC-lysis step; however, the cells should be collected as quickly as possible, especially when using attached cells in culture. Other protocols for embedding high-molecular-weight mammalian DNA from tissues (see Support Protocol 3 and UNIT 2.5B) should also produce acceptable results. Materials Healthy human volunteer PBS (APPENDIX 2), ice cold 1× RBC lysis solution (see recipe) InCert agarose (FMC Bioproducts) Proteinase K lysis solution (see recipe) TE50 buffer: 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2)/50 mM EDTA 0.1 mM PMSF solution: 100 mM PMSF solution (see recipe) diluted 1/1000 in TE50 buffer immediately before use 0.5 M EDTA, pH 8.0 (APPENDIX 2) Blood-drawing equipment Blood collection tubes containing EDTA Automated hematology counter 50-ml conical screw-cap polypropylene tubes, sterile Refrigerated centrifuge with rotor/adapters for 50-ml tubes (e.g., Sorvall RI6000D centrifuge with H-1000B swinging-bucket rotor or equivalent) Roller mixer (Robbins Scientific or equivalent) 50°C water bath 10 × 5 × 1.5–mm disposable DNA plug mold (Bio-Rad) Collect blood and remove red blood cells 1. Use blood-drawing equipment to obtain 45 ml venous blood from a healthy human volunteer in blood collection tubes containing EDTA. Mix well to avoid clot formation. 2. Count the total number of lymphocytes using an automated hematology counter. The total number should be 2.25 to 3.3 × 108 in 45 ml.

3. Transfer blood to two 50-ml conical screw-cap polypropylene tubes, add 10 ml ice-cold PBS to each, and mix gently. 4. Centrifuge at 1876 × g for 5 min at 4°C. Discard the supernatant using a 10-ml disposable pipet, being careful not to remove any lymphocytes from the fuzzy coat layer. Construction of BAC/PAC Libraries

Centrifugation results in a large pellet of red blood cells at the bottom of the tube and a very thin layer of lymphocytes (the fuzzy coat layer) at the interface with the plasma layer.



5. Repeat wash with ice-cold PBS ten times. 6. Discard the final supernatant, mix the cell suspension well, and divide into four 50-ml tubes. 7. Add 25 ml of 1× RBC lysis solution to each tube, mix samples gently on a roller mixer, and incubate on mixer ∼20 min at room temperature. During this time, an exchange process catalyzed by carbonic anhydrase results in accumulation of ammonium chloride inside the cells; this accumulation increases the internal osmotic strength and causes swelling and bursting of RBCs. Lymphocytes have much lower carbonic anhydrase activity and do not undergo lysis.

8. Visually monitor the progress of RBC lysis. When it appears complete (usually 200 kb) is always observed. This results from at least three factors: (1) DNA fragments and circular ligation products being increasingly more fragile, subject to shearing or nuclease action; (2) a lower efficiency in the formation of circular ligation intermediates; and (3) a dropoff in transformation efficiency of plasmids with increasing size, expressed as a molecular efficiency, and becoming more pronounced when expressed per DNA weight. It is imperative that highly competent bacterial cells are used to ensure that reasonable numbers of clones are obtained per electroporation. As a consequence of the dropoff in cloning efficiency with size, it is difficult to prepare BAC and PAC libraries with insert sizes averaging in 200 to 250 kb, and nearly impossible to construct highly redundant libraries with an average size >250 kb using the presented technology. Once a provisional BAC or PAC library has been constructed, it is essential to characterize the clones and isolate plasmid DNA. The use of Support Protocol 5 or an automatic plasmid isolation machine (AutoGen 740) will result in ∼0.5 to 1 µg PAC DNA (170 kb) per 1.5 ml of overnight culture (not induced with IPTG). When the pCYPAC2, pPAC4, or pBACe3.6 vector is used for library construction, two NotI sites flank the insert in recombinant plasmids; therefore, most clones should show only two bands when analyzed by PFGE: the vector (at ∼15.5 kb for pCYPAC2, 16.8 kb for pPAC4, and 8.7 kb for pBACe3.6) and the insert of variable size from clone to clone. Additional bands may be seen if the insert contains internal NotI sites. Incomplete digestion usually leads to a very characteristic doublet, where the two bands differ by the size of the vector.

Time Considerations Anticipated Results Under optimal conditions, one can obtain as many as 1800 transformants per 2 µl dialyzed solution with an average insert size of 160 kb, and as many as 250 transformants per 2 µl at 230 kb for BAC. Due to the low DNA concentrations in the ligation, a typical 10-fold redundant mammalian BAC or PAC library requires anywhere from a few hundred up to 2000 electroporations. Each electroporation should result in 300 to 2000 colonies for fragments in

It takes several days to prepare agarose-embedded high-molecular-weight DNA starting from a blood or animal tissue sample and at least another week for testing various conditions of partial digestion and analyzing the results using pulsed-field gel electrophoresis (weeks 1 and 2). Ligations and transformations take 3 to 5 days, and another 3 to 4 days are required to characterize candidate colonies by pulsed-field gel electrophoresis of NotI-restricted plasmids (weeks 2 and 3). If the par-



Supplement 55

ticular ligation gives desirable results (good cloning efficiency, low nonrecombinant levels, and large inserts), another week (week 4) should be dedicated to the 200 to 2000 electroporations required for a 10-fold redundant mammalian BAC or PAC library. It is often convenient, perhaps necessary, and possibly desirable to take a break at this point. The arraying of the BAC or PAC libraries can be done at a different time and place, as it requires other facilities. Picking of colonies (e.g., 100,000) will take about one week and will require at least two people to prepare and label microtiter dishes and spread all the cells, provided that there is access to a facility equipped with microtiter plate fillers and a colony-picking robot. Hence, under the most optimal laboratory conditions and with the appropriate skills, a 10-fold redundant mammalian BAC or PAC library can be constructed in 1 to 3 months, requiring at least two people; however, it is more realistic to expect that most of the time will be invested in acquiring the skills and optimizing the various critical parameters. Anytime between 6 months and a year should be dedicated by a single person, who should be able to count on assistance from others from time to time.

Literature Cited Chalker, A.F., Leach, D.R., and Lloyd, R.G. 1988. Escherichia coli sbcC mutants permit stable propagation of DNA replicons containing a long palindrome. Gene 71:201-205. Chu, G., Vollrath, D., and Davis, R.W. 1986. Separation of large DNA molecules by contourclamped homogeneous electric field. Science 234:1582-1585. Donovan, J. and Brown, P. 1995. Euthanasia. In Current Protocols in Immunology. (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevock, and W. Strober, eds.) pp. 1.8.1-1.8.4. John Wiley & Sons, New York. Frengen, E., Weichenhan, D., Zhao, B., Osoegawa, K., van Geel, M., and de Jong, P.J. 1999. A modular, positive selection bacterial artificial chromosome vector with multiple cloning sites. Genomics. In press. Fukushige, S. and Sauer, B. 1992. Genomic targeting with a positive-selection lox integration vector allows highly reproducible gene expression in mammalian cells. Proc Natl Acad Sci U.S.A. 89:7905-7909.


Gibson, T.J., Coulson, A.R., Sulston, J.E., and Little, P.F. 1987. Lorist2, a cosmid with transcriptional terminators insulating vector genes from interference by promoters within the insert: Effect on DNA yield and cloned insert frequency. Gene 53:275-281.

Gibson, F.P., Leach, D.R., and Lloyd, R.G. 1992. Identification of sbcD mutations as cosuppressors of recBC that allow propagation of DNA palindromes in Escherichia coli K-12. J. Bacteriol. 174:1221-1228. Grant, S.G., Jessee, J., Bloom, F.R., and Hanahan, D. 1990. Differential plasmid rescue from transgenic mouse DNAs into Escherichia coli methylation-restriction mutants. Proc. Natl. Acad. Sci. U.S.A. 87:4645-4649. Hanahan, D., Jessee, J., and Bloom, F.R. 1991. Plasmid transformation of Escherichia coli and other bacteria. Methods Enzymol. 204:63-113. Ioannou, P.A., Amemiya, C.T., Garnes, J., Kroisel, P.M., Shizuya, H., Chen, C., Batzer, M.A., and de Jong, P.J. 1994. A new bacteriophage P1-derived vector for the propagation of large human DNA fragments. Nature Genet. 6:84-89. Osoegawa, K., Woon, P.-Y., Zhao, B., Frengen, E., Tateno, M., Catanese, J.J., and de Jong, P.J. 1998. An improved approach for construction of bacterial artificial chromosome libraries. Genomics 52:1-8. Pierce, J.C., Sauer, B., and Sternberg, N. 1992. A positive selection vector for cloning high molecular weight DNA by the bacteriophage P1 system: Improved cloning efficacy. Proc. Natl. Acad. Sci. U.S.A. 89:2056-2060. Raleigh, E.A. 1987. Restriction and modification in vivo by Escherichia coli K12. Methods Enzymol. 152:130-141. Schwartz, D.C., Li, X., Hernandez, L.I., Ramnarain, S.P., Huff, E.J., and Wang, Y.K. 1993. Ordered restriction maps of Saccharomyces cerevisiae chromosomes constructed by optical mapping. Science 262:110-114. Sheng, Y.L., Mancino, V., and Birren, B. 1995. Transformation of Escherichia coli with large DNA molecules by electroporation. Nucl. Acids Res. 23:1990-1996. Shizuya, H., Birren, B., Kim, U.-J., Mancino, V., Stepak, T., Tachiiri, Y., and Simon, M. 1992. Cloning and stable maintenance of 300-kilobase-pair fragments of human DNA in Escherichia coli using an f-factor-based vector. Proc. Natl. Acad. Sci. U.S.A. 89:8794-8797. Sternberg, N. 1990. Bacteriophage P1 cloning system for the isolation, amplification, and recovery of DNA fragments as large as 100 kilobase pairs. Proc. Natl. Acad. Sci. U.S.A. 87:103-107. Strong, S.J., Ohta, Y., Litman, G.W., and Amemiya C.T. 1997. Marked improvement of PAC and BAC cloning is achieved using electroelution of pulsed-field gel-separated partial digests of genomic DNA. Nucl. Acids Res. 25:3959-3961. Wang, M., Chen, X.N., Shouse, S., Manson, J., Wu, Q., Li, R., Wrestler, J., Noya, D., Sun, Z.G., Korenberg, J., and Lai, E. 1994. Construction and characterization of a human chromosome 2-specific BAC library. Genomics 24:527-534. Wang, G.L., Holsten, T.E., Song, W.Y., Wang, H.P., and Ronald, P.C. 1995. Construction of a rice bacterial artificial chromosome library and iden-



tification of clones linked to the Xa-21 disease resistance locus. Plant J. 7:525-533. Woo, S.S., Jiang, J., Gill, B.S., Paterson, A.H., and Wing, R.A. 1994. Construction and characterization of a bacterial artificial chromosome library of Sorghum bicolor. Nucl. Acids Res. 22:4922-4931.

Osoegawa et al., 1998. See above. An improved BAC/PAC cloning approach. Pierce et al., 1992. See above. Development of an elegant and efficient P1 cloning vector, a direct predecessor to the pCYPAC vectors.

Woon, P.-Y., Osoegawa, K., Kaisaki, P.J., Zhao, B., Catanese, J.J., Gauguier, D., Cox, R., Levy, E. R., Lathrop, G.M., Monaco, A.P., and de Jong, P.J. 1998. Construction and characterization of a 10fold genome equivalent rat P1-derived artificial chromosomal library. Genomics 50:306-316.

Shizuya et al., 1992. See above.

Wyman, A.R. and Wertman, K.F. 1987. Host strains that alleviate underrepresentation of specific sequences: Overview. Methods Enzymol. 152:173180.

The first large-insert P1 plasmids constructed using in vitro packaging and viral infection to transform E. coli.

Key References Albertsen, H.M., Abderrahim, H., Cann, H.M., Dausset, J., Le Paslier, D., and Cohen, D. 1990. Construction and characterization of a yeast artificial chromosome library containing seven haploid equivalents. Proc. Natl. Acad. Sci. U.S.A. 87:4256-4260. The first description of use of PFGE to size select DNA for ligation. Ioannou et al., 1994. See above. The first paper describing PAC cloning. O’Connor, M., Peifer, M., and Bender, W. 1989. Construction of large DNA segments in Escherichia coli. Science 244:1307-1312.

The development of efficient procedures for constructing large-insert plasmids by bacterial transformation. Sternberg, 1990. See above.

Contributed by Kazutoyo Osoegawa and Pieter J. de Jong Children’s Hospital Oakland Research Institute Oakland, California Eirik Frengen The Biotechnology Centre of Oslo University of Oslo Oslo, Norway Panayiotis A. Ioannou The Murdoch Institute for Research into Birth Defects Royal Children’s Hospital Melbourne, Australia

Describes first large BAC clone constructed by a combination of in vivo recombination and recombinant DNA technology.



Supplement 55

AMPLIFICATION OF TRANSFORMED OR PACKAGED LIBRARIES

SECTION V

After libraries are packaged (or transformed) they can be amplified for storage. Amplification of a library involves replicating each clone in the library. This is frequently a worthwhile procedure for genomic or cDNA libraries because the libraries can then be used many times to isolate clones corresponding to different sequences. Procedures for amplifying bacteriophage, cosmid, and plasmid libraries are presented in this section.

Amplification of a Bacteriophage Library This protocol may be used for genomic DNA or cDNA libraries. A freshly packaged and titered library is adsorbed to log phase plating bacteria. The mixture is then plated at high density and allowed to grow until the plaques are just subconfluent. The phage are eluted from the plate by overnight incubation with phage buffer and the library is titered and stored at both 4°C and −80°C.


Materials LB medium containing 0.2% maltose and 10 mM MgSO4 (UNIT 1.1) Suitable host (Table 5.10.1) In vitro packaged phage library (UNITS 5.7 & 5.8) Top agarose (UNIT 1.1), warmed to 47°C 150-mm H plates (UNIT 1.1), warmed to 37°C Suspension medium (SM; UNIT 1.11) Chloroform Dimethyl sulfoxide (DMSO) Additional reagents and equipment for titering bacteriophage (UNIT 1.11) 1. Inoculate 250 ml LB medium containing 0.2% maltose and 10 mM MgSO4 with 2.5 ml of a fresh overnight culture of host bacteria. Shake vigorously 2 to 4 hr at 37°C until the OD600 ≅ 0.5. 2. Prepare E. coli cells for plating as described in UNIT 1.11. The integrity of λ phage particles requires the presence of Mg2+. Maltose induces the expression of the λ phage receptor. To maximize the complexity of the library, it is important that the number of nonviable cells among the host bacteria be minimized; these represent “dead ends” for phage. Table 5.10.1 Suitable Escherichia coli Host Strains for Amplifying Lambda-Constructed Libraries

Vector

E. coli host

Relevant host genotype

λgt10 λgt11

C600hflA Y1088

EMBL 3 or 4

P2392, Q359, NM539 LE392

hflA SupF, lacIq (no antibiotics needed) P2 lysogen

Charon 4A

SupF Construction of Recombinant DNA Libraries

Contributed by Lloyd B. Klickstein Current Protocols in Molecular Biology (1996) 5.10.1-5.10.3 Copyright © 2000 by John Wiley & Sons, Inc.


3. Shake by hand and then briefly microcentrifuge the tube containing the library to separate out the chloroform in the packaged library. Add the packaged, titered phage (but no chloroform) to plating bacteria, using 0.25 ml of host cells for every 1 × 105 phage. Incubate 15 min at 37°C to allow phage to adsorb to bacteria. 4. Add 0.5 ml of the host/phage mixture to 8 ml of 47°C top agarose, mix by inverting a few times, and pour onto a fresh 150-mm H plate, warmed to 37°C. Use as many plates as necessary to plate the entire mixture. Spread the top agarose by gently rotating the plate on the benchtop, and allow the agarose to harden for 5 min. Invert and incubate 6 to 7 hr at 37°C (39° to 41°C for λgt11 libraries amplified in Y1088). A myriad of tiny plaques should barely be visible at ∼4 to 5 hr. These should expand and be just less than confluent at 6 hr. If the plaques are growing slowly, continue the incubation for up to 8 hr.

5. Remove plates from incubator, cover each lawn with 10 ml SM, and place on a level surface at 4°C for 2 to 16 hr to elute the phage. 6. Using a pipet, combine the SM from all plates into a single glass or polypropylene tube. Centrifuge 5 min at 3000 rpm (2800 × g) and transfer the supernatant to a new tube. Add 0.5 ml chloroform and mix by inverting several times. Titer the amplified library as described in UNIT 1.11. Expect a titer of 1010 to 1011/ml. The volume of SM recovered from each plate will be nearly 1 ml less than what was added. If the plates were not fresh, greater losses of volume will occur. The total volume of the library will, of course, depend on the number of plates used in the amplification and may approach 100 ml in the case of large libraries.

7. Store the library at 4°C in Teflon-capped glass tubes with 0.5 ml chloroform. The library is stable in this form for years. (Storage in plastic results in a 100- to 1000-fold drop in titer.) For added security, transfer 930 µl-aliquots of amplified library (no chloroform) to screw-cap microcentrifuge tubes, add 70 µl of DMSO (final concentration 7%), mix thoroughly by inverting several times, and place at −80°C. COMMENTARY Background Information

Amplification of a Bacteriophage Library

Amplification is not essential to the successful creation and screening of a λ phage library; however, an unamplified library must be used right away and the number of times the library may be screened is limited by the life of the nitrocellulose filters or the primary plates which dry out in a few weeks to a month. In contrast, amplification allows the library to be stored nearly indefinitely and to be screened as many times as necessary. The principle of amplification is to allow each in vitro packaged phage to produce thousands of identical clones by a limited infection of a host. A potential disadvantage is that the composition of the library may change as a consequence of differences in growth rate during the amplification step, some clones being overrepresented compared to the corresponding mRNA abundance in the total cellular mRNA and others being underrepresented. The preadsorption of the li-

brary to the bacteria, the high plating density, and the relatively short incubation period all help minimize changes in the composition. An alternative approach used by some is amplification in liquid culture. This method is not recommended, since rapidly growing phage are overrepresented to a greater degree than when plates are used. Furthermore, there are more steps involved, since the phage must be purified from the relatively large volume by PEG precipitation and CsCl-gradient centrifugation.

Critical Parameters Amplification should be carried out as soon as possible after the library is packaged. Phage particles stored in the in vitro packaging mixture are unstable because the phage particles adsorb to bacterial debris in the mixture and are inactivated. A decrease in titer at this point decreases the complexity of the library. Fresh,



log-phase host cells are important to ensure that each phage infects a viable cell and makes a phage burst and thus is successfully amplified. A phage that injects its DNA into a dead cell is lost from the library. For libraries constructed in vectors such as λgt10 or the EMBL series, which allow selection for recombinants, amplification should be done under selective conditions since this eliminates nonrecombinants. Subsequent platings of the amplified library may be done on either selective or nonselective hosts. The titer of a library stored as described in the basic protocol will usually drop only 2- to 3-fold over the years. If stored in plastic, then the titer may drop as much as 100- to 1000-fold.

that occurred during the elution of the plates. Storing the library in several tubes and freezing aliquots will circumvent loss of library due to this problem.

Anticipated Results The titer of an amplified phage library will be in the range of 1-10 × 1010 pfu/ml. The volume obtained will depend on the number of plates used, which in turn depends on the initial complexity of the library. For example, a library initially containing 2 × 106 clones in 1.5 ml requires ten 150-mm plates, so ∼100 ml of SM will be recovered in total. Thus, the library will have been amplified by a factor of roughly 105.

Time Considerations Troubleshooting The only difficulties that can occur here are a failure of the phage to infect the host or a failure of the host to grow. This may be avoided if the plates, media, and host strain are first tested with nonrecombinant phage prior to amplifying the library. Causes of problems include top agarose that is too hot, incorrect host strain, incorrect plates, incorrect incubator temperature, and so forth. Losing a library because a tube was dropped or contaminated may be avoided by splitting the packaged ligation into two tubes and amplifying in duplicate. A library stored over chloroform at 4°C may occasionally drop in titer due to fungal contamination

After in vitro packaging, the library is stored overnight at 4°C while a titer plate is incubating. The library should be amplified the next day by starting step 1 early in the day. The plaques should develop in 6 to 10 hr. Suspension medium is added and the plates elute overnight at 4°C. Alternatively, step 1 may be started in the late afternoon and the plates grown overnight for 7 hr.

Contributed by Lloyd B. Klickstein Brigham and Women’s Hospital Boston, Massachusetts



Supplement 34

Amplification of Cosmid and Plasmid Libraries Bacteria containing the recombinant clones are grown on agar plates, washed off the plates, and stored in glycerol. This procedure produces about a million-fold amplification of the library.


Materials LB plates containing appropriate antibiotic (UNIT 1.1) LB medium (UNIT 1.1) Sterile glycerol Nitrocellulose membrane filters (Millipore HATF) Sterile rubber policeman 1. Plate drug-resistant bacteria on nitrocellulose filters that have been placed on LB plates (containing antibiotic to which the colonies are resistant), as described in UNIT 6.2. The filters are used so that the colonies remain small and do not grow together. Be sure to use commercially available triton-free filters to ensure bacterial growth.

2. Grow bacteria until colonies are just confluent. 3. Flood each plate with LB medium. Use ∼2 ml/10-cm plate and 4 ml/15-cm plate. 4. Using a sterile rubber policeman, rub colonies off from nitrocellulose filter, making a bacterial suspension. 5. Pool the suspensions from the different plates into one 50-ml plastic tube. 6. Add sterile glycerol to give a final glycerol concentration of 15%. 7. Mix the solution thoroughly and dispense 500 µl into 1-ml tubes. 8. Freeze the aliquots of the library at −70°C. They should remain viable without appreciable loss of bacteria for over 1 year. 9. To screen the library, simply thaw an aliquot, titer the bacteria concentration (UNIT 1.3), and plate the appropriate numbers of cosmid- or plasmid-containing bacteria on the screening filters. COMMENTARY Background Information Amplification of phage libraries is an established procedure, but the effective amplification of plasmid and especially cosmid libraries has proven to be more difficult. The major concern in amplifying a plasmid or cosmid library is that the doubling times of bacteria are often altered, depending upon the type of insert included within the plasmid or cosmid vector. This disproportionate growth of recombinant bacteria can easily be visualized by plating out a portion of a library and noting the variable colony size after 16 hr at 37°C. However, there is ample incentive to amplify the library. The advantages of obtaining thousands of copies of a cosmid or plasmid library from a single liContributed by John H. Weis Current Protocols in Molecular Biology (1996) 5.11.1-5.11.2 Copyright © 2000 by John Wiley & Sons, Inc.

brary are obvious. When amplifying a cosmid or bacterial library the individual must weigh the concern of the possible underrepresentation of particular clones because of overgrowth of some colonies, against the time required to create a library de novo for each screening or for the dissemination of the library to other researchers.

Critical Parameters The major concern with any amplification step is that each original recombinant be equally represented. Differences in the duplication rate of any recombinant bacteria will result in over- or underrepresentation in the amplified library. This concern is less for sim-



ple plasmid libraries than for cosmid libraries, which contain bacteria that grow at markedly different rates.

Anticipated Results The purpose of amplification is to provide a reagent library that can be used many times. An amplified cosmid or plasmid library will contain one library’s equivalence in 10 to 100 µl. By producing 100 µl of such a library frozen in 100 1-ml tubes, over 1000 platings can be expected.

Time Considerations After the transformation or transfection step in the preparation of the plasmid or cosmid libraries, respectively, the plating of the outgrowths will require 1 to 2 hr. The plates are then incubated overnight at 30° to 37°C, depending upon the desired colony size and the density of the platings. The filter wash, glycerol mix, and aliquoting into freezer tubes require an additional 2 hr.

Contributed by John H. Weis Harvard Medical School Boston, Massachusetts

Amplification of Cosmid and Plasmid Libraries



CHAPTER 6 Screening of Recombinant DNA Libraries INTRODUCTION

T

he usual approach to isolating a recombinant DNA clone encoding a particular gene or mRNA sequence is to screen a recombinant DNA library. As described in Chapter 5, a recombinant DNA library consists of a large number of recombinant DNA clones, each one of which contains a different segment of foreign DNA. Since only a few of the thousands of clones in the library encode the desired nucleic acid sequence, the investigator must devise a procedure for identifying the desired clones. The optimal procedure for isolating the desired clone involves a positive selection for a particular nucleic acid sequence. If the desired gene confers a phenotype that can be selected in bacteria, then the desired clone can be isolated under selective conditions (UNIT 1.4). However, most eukaryotic genes and even many bacterial sequences do not encode a gene with a selectable function. Clones encoding nonselectable sequences are identified by screening libraries: the desired clone is identified either because (1) it hybridizes to a nucleic acid probe, (2) it expresses a segment of protein that can be recognized by an antibody, or (3) it promotes amplification of a sequence defined by a particular set of primers. Screening libraries involves the development of a rapid assay to determine whether a particular clone contains the desired nucleic acid sequence. This assay is used first to identify the recombinant DNA clone in the library and then to purify the clone (see Fig. 6.0.1). Normally, this screening procedure is performed on bacterial colonies containing plasmids or cosmids or on bacteriophage plaques. To test a large number of clones at one time, the library is spread out on agarose plates (UNIT 6.1), then the clones are transferred to filter membranes (UNIT 6.2). The clones can be simultaneously hybridized to a particular probe (UNITS 6.3 & 6.4) or bound to an antibody (UNITS 6.7 & 6.11). When the desired clone is

bacteriophage, cosmid, or plasmid libraries

plate library (consider library base and insert size)

screen library by: hybridization to nick-translated DNA and synthetic oligonucleotides or,

immunoreactivity or, hybrid selection of mRNA and translation

purify plaques or colonies

Figure 6.0.1

Flow chart for screening libraries.

Contributed by J.G. Seidman Current Protocols in Molecular Biology (1994) 6.0.3-6.0.4 Copyright © 2000 by John Wiley & Sons, Inc.

Screening of Recombinant DNA Libraries

6.0.3 Supplement 27

first identified, it is usually found among many undesirable clones; an important feature of library screening is the isolation of the desired clones (UNITS 6.5, 6.6 & 6.12). Another method for identifying the desired clone involves hybrid selection (UNIT 6.8), a procedure by which the clone is used to select its mRNA. This mRNA is characterized by its translation into the desired protein. Libraries consisting of large genomic DNA fragments (∼1 Mb) carried in yeast artificial chromosome (YAC) vectors have proven to be tremendously useful for genome analysis. In general, these libraries (which are usually produced by large “core” laboratories) are intially screened using a locus-specific PCR assay (UNIT 6.9); the clone resulting from the initial round of screening is subsequently analyzed by more conventional hybridization methods (UNIT 6.10). To screen a DNA library, one must first devise the screening procedure. The next important choice is the selection of a recombinant DNA library. When choosing which library to screen the investigator should consider whether he or she wants to isolate clones encoding the gene or the mRNA sequence. cDNA clones encode the mRNA sequence and allow prediction of the amino acid sequence, whereas genomic clones may contain regulatory as well as coding (exon) and noncoding (intron) sequences. The differences between genomic and cDNA libraries are discussed in Chapter 5. Another critical parameter to be determined before proceeding with a library screen is the number of clones in the library that must be screened in order to identify the desired clone. That is, what is the frequency of the desired clone in the library? This frequency is predicted differently for genomic and cDNA libraries, as described below. Screening a genomic library. In general, genomic libraries can be made from DNA derived from any tissue, because only two copies of the gene are present per cell or per diploid genome. The predicted frequency of any particular sequence should be identical to the predicted frequency for any other sequence in the same genome. The formula for predicting the number of clones that must be screened to have a given probability of success is presented in UNIT 5.1. This number is a function of the complexity of the genome and the average size of the inserts in the library clones. For amplified libraries, the base (see UNIT 5.1) must exceed this number. Usually about 1 million bacteriophage clones or 500,000 cosmid clones must be screened to identify a genomic clone from a mammalian DNA library. Many of the clones that are screened from an amplified library will be screened more than once; the total number of clones that must be screened is 30 to 40% greater than the number calculated by the formula. Screening a cDNA library. The optimal cDNA library is one made from a particular tissue or cell that expresses the desired mRNA sequence at high levels. In highly differentiated cells, a particular mRNA may comprise as many as 1 of 20 of the poly(A)+ mRNA molecules, while some mRNAs are either not present at all or comprise as low as 1 molecule in 100,000 poly(A)+ mRNA molecules. When choosing a cDNA library the investigator must make every effort to obtain a library from a cell where the mRNA is being expressed in large amounts. Of course, the number of clones that must be screened is determined by the abundance of the mRNA in the cell. The amount of protein that is found in the cell is frequently a good indicator of the abundance of the mRNA. Thus, proteins that comprise 1% of the total cell protein are made by mRNAs that usually comprise 1% of the total poly(A)+ mRNA, and the desired cDNA clones should comprise about 1% of the clones in the cDNA library.

Introduction

Screening a YAC library. In the typical genomic libraries maintained in E. coli (described in Chapter 5), the size of the insert is limited to 20 to 25 kb for lambda vectors or to 40 to 45 kb for cosmid vectors. Yeast artifical chromosome (YAC) vectors, by contrast, are designed to carry much larger genomic DNA fragments and thereby facilitate genomic analysis, with inserts ranging from 0.3 to ∼1 Mb in size. Conventional screening of YAC

6.0.4 Supplement 27


libraries by hybridization is difficult, both because of the unfavorable signal-to-noise ratio and the sheer numbers of replica films required to represent an entire library. For example, a standard YAC library representing 5 to 8 genome equivalents requires over 500 microtiter plates (and corresponding filters for screening by hybridization). Thus, most core laboratories screen YAC libraries using a locus-specific PCR assay whose primers define a particular sequence. The PCR screening is initially performed using pools (representing up to 4 microtiter plates or 384 YAC clones) or superpools (representing up to 20 microtiter plates or nearly 2000 clones), followed by subsequent rounds of screening to narrow down the possible candidates. Specialized screening strategies. For particular applications, there exist specialized approaches to screening. For example, cloned cDNAs encoding cell surface or intracellular proteins can be identified by expression screening, involving rounds of transient expression of a library and subsequent screening by immunoselection (UNIT 6.11). The technique of recombination-based screening provides a rapid and efficient approach for screening a complex genomic library in bacteriophage lamba (UNIT 6.12). The library is screened for homology against a plasmid carrying a particular cloned target sequence. If homology exists, a recombination event occurs, resulting in integration of the plasmid into the phage, and the recombinant is isolated by genetic selection. General considerations. When selecting the library it is critical that the base be larger than the number of clones to be screened. One problem with predicting the number of clones to screen is that most libraries are amplified and in the process of amplifying the library some clones are lost while others may grow more rapidly. Thus, if the desired clone is not found in a particular library, another independent library should be screened. Having selected the library, the investigator is ready to begin screening for the desired clone. The technologies used to screen libraries are mostly extensions of the techniques that have been described earlier in the manual. Libraries are plated out, transferred to nitrocellulose filters, and hybridized to 32P-labeled probes or bound to antibodies. The major problem associated with this technique is that “false” positives can be identified: the probe may hybridize to clones that do not encode the desired sequence. Approaches to minimize this problem are discussed in UNIT 6.7. A second source of undesired clones arises from the power of the screening procedures that are normally used to screen these libraries. The investigator will be screening as many as one million clones. If the library contains any contaminating recombinant DNA clones that have been previously grown in the laboratory, it will be identified in the screening procedure. Thus, extreme care must be exercised to prevent contamination of the library with previously isolated recombinant clones. Despite these problems the ability to screen large DNA libraries to isolate the desired clone provides a powerful tool for molecular biologists. J.G. Seidman



Supplement 27

PLATING LIBRARIES AND TRANSFER TO FILTER MEMBRANES

SECTION I

The basic principle of screening recombinant DNA libraries is that bacteriophage plaques, or bacterial colonies containing plasmids or cosmids, contain relatively large amounts of insert DNA that can be detected either directly by hybridization (see below) or indirectly by the protein that may be expressed from the cloned segment (UNIT 6.7). The first step in the nucleic acid hybridization screening procedure is to grow large numbers of colonies or plaques on agar plates. Replica copies of these colonies are transferred to nitrocellulose filters, where they can be screened. In this section the techniques for producing large numbers of colonies and plaques, and for transferring these to filter membranes, are discussed. Prerequisites to these procedures are that the library must already be chosen and the number of clones to be screened must be determined (see introduction to this chapter).

Plating and Transferring Bacteriophage Libraries Bacteriophage are plated onto agar plates at high density so that as many as 1 million different plaques can be screened. The bacteriophage plaques are then transferred to nitrocellulose filters, denatured, and baked. The library and the number of clones to be screened are predetermined. Principles for choosing the plaque density and the number of plates to be used are outlined in the commentary.

UNIT 6.1

BASIC PROTOCOL

Materials Host bacteria, selection strain if applicable (UNIT 1.10; Table 1.4.5; Table 5.10.1) Recombinant phage (UNIT 5.10) 0.7% top agarose (prewarmed; UNIT 1.1) 82-mm or 150-mm LB plates; or 245 × 245–mm Nunc bioassay LB plates (UNIT 1.1) 0.2 M NaOH/1.5 M NaCl 0.4 M Tris⋅Cl, pH 7.6/2× SSC 2× SSC (APPENDIX 2) Nitrocellulose membrane filters (or equivalent) 20-G needle 46 × 57–cm Whatman 3MM or equivalent filter paper 80°C vacuum oven or 42°C oven Plating bacteriophage 1. Determine the titer of the library by serial dilution as described in UNITS 1.11 & 5.7. For λ vectors that allow genetic selection against nonrecombinants, plating should be done on the appropriate bacterial strain (e.g., P2 lysogen for EMBL vectors). LB plates should be poured several days in advance to allow them to dry prior to plating. The large Nunc plates are particularly prone to condensation on the surface of the agar, but this can be alleviated by allowing them to sit on the benchtop with covers removed for a few minutes to several hours before use.

2. Mix recombinant phage and plating bacteria (prepared as described in UNIT 1.11) in a culture tube as outlined in Table 6.1.1 and incubate 20 min at 37°C. 3. Add 0.7% top agarose to culture tube and transfer mixture to LB plates. Disperse bacteria and agarose on plates by tilting the plates back and forth. Mix cells and agarose for the large Nunc plates by gently inverting several times in a capped 50-ml tube prior to plating. Contributed by Thomas Quertermous Current Protocols in Molecular Biology (1996) 6.1.1-6.1.4 Copyright © 2000 by John Wiley & Sons, Inc.

Screening Recombinant DNA Libraries

6.1.1 Supplement 34

Top agarose rather than top agar should be used as agar tends to lift off with the nitrocellulose filter. Melt the top agarose and cool to 45° to 50°C before use. If top agarose is too hot it will kill the bacteria, while if it is too cold the library will solidify in the tube.

4. Incubate plates at 37°C until plaques cover the plate but are not confluent. Incubation time varies between 6 and 12 hr and depends on type of phage and bacteria used. Store at 4°C. Do not incubate unattended overnight, but rather place at 4°C and allow to continue growth the next day. Allowing phage plaques to incubate for the correct amount of time is critical. The object is to optimize two parameters. First, the plaques must be large enough to contain sufficient DNA to give a good signal. Second, if the plaques are too large and become confluent they are difficult to purify in subsequent steps. Because most nucleic acid probes give a very strong signal, we tend to prefer having smaller plaques and weaker signals.

5. Incubate plates at 4°C for at least 1 hr before applying filters. Transferring to nitrocellulose filters 6. Label nitrocellulose filters with a ballpoint pen and apply face down (ink side up) on cold LB plates bearing bacteriophage plaques. This is best accomplished by touching first one edge of the filter to the agarose and progressively laying down more of the filter as it wets. Bubbles should be avoided. If difficulties are encountered the filter should not be adjusted on the plate, but rather removed and replaced with a new filter. Nitrocellulose filters should be handled only with forceps or gloved hands.

7. Leave filters on plates for 1 to 10 min to allow transfer of phage particles to the filter. During this transfer period the orientation of the filter to the plate is recorded by stabbing a 20-G needle through the filter into the agar at several asymmetric points around the edge of the plate. Up to five replicas can be made from each plate. Remove the filter slowly from the plate with blunt, flat forceps and place face up on paper towels or filter paper. Some investigators dip the needle used to orient the filter in India ink to more clearly mark the filter and agar. Other investigators mark the back of the agar plate with a black marker. Making two replicas from each filter, hybridizing both to the DNA probe, and comparing the autoradiographs of the replica filters eliminates many possible artifacts. This is particularly helpful when screening with an oligonucleotide probe.

8. Dry the filters on the benchtop for at least 10 min. This drying process binds the plaques to the filter.

Table 6.1.1

Plating and Transferring Bacteriophage Libraries

Recommended Mixtures for Plating Bacteriophage Libraries

Plate size

LB plate ingredient

82 mm

Bacteriab (ml) Phage, pfu Top agarose, ml

0.2 5,000 3

245 × 245 mma

150 mm 0.5 20,000-30,000 7

2 150,000 30

aNunc Bioassay plates distributed by Vangard International. bPlating bacteria are prepared as described in Chapter 1.

6.1.2 Supplement 34


Denaturation and baking 9. Place 46 × 57–mm Whatman 3MM paper on the benchtop and saturate with 0.2 M NaOH/1.5 M NaCl. Place filters on the paper face up for 1 to 2 min. The 3MM paper should be wet enough to allow immediate saturation of the filters, but not so wet that the solution pools on the surface.

10. Transfer filters (face up) to 3MM paper saturated with 0.4 M Tris⋅Cl, pH 7.6/2× SSC for 1 to 2 min and then to 3MM paper saturated with 2× SSC for 1 to 2 min. Some investigators immerse the filters in all three solutions. This procedure can make the plaques detected by hybridization appear diffuse.

11. Dry filters in a vacuum oven 90 to 120 min at 80°C or overnight in a regular oven at 42°C. Store at room temperature in folded paper towels or other absorbent paper until needed for hybridization (described in UNIT 6.3 or 6.4). COMMENTARY Background Information There are two parts to this protocol—plating the library and preparing filters. The number of bacteriophage per plate determines the number of plates that must be poured. This number is defined by the number of recombinants in the library (i.e., base of the library) and the frequency of the expected clone in the library. There is no advantage to screening more than 3 to 5 times the base of the library. The frequency of the clone in the library is determined as follows. cDNA libraries: the expected frequency of the desired RNA among the total RNA of the cell, ranging from 1⁄100 to 1⁄50,000. Genomic libraries: the size of the insert divided by the total genome size. Subgenomic libraries: the size of insert per total genome size times the fold purification of the DNA fragment (usually 10- to 50-fold). The usefulness of a recombinant phage library depends on the ability to screen a large number of phage and identify the clone that carries the DNA sequence of interest. This has been made possible by the technique of in situ plaque hybridization described by Benton and Davis (1977). The phage are allowed to multiply in host bacteria in a thin layer of agarose on regular bacterial plates. When nitrocellulose is applied to the agarose, phage particles and unpackaged DNA adsorb to the filter to produce a replica of the plate surface. If the agarose surface is not excessively wet, there will be little spreading of the phage on the filter. Subsequent treatment of the filter with sodium hydroxide destroys the phage particles and denatures the phage DNA which then binds to the nitrocellulose. Neutralization of the filters is required to maintain the integrity of the nitrocellulose. Hy-

bridization of these filters to a DNA or RNA probe will identify the location of the phage plaque of interest, which can then be recovered from the plate. A common variation of this technique is the substitution of one of the nylon-based membranes for nitrocellulose (see UNIT 2.9). The advantage of nylon membranes is their durability, which allows multiple hybridizations to the same filter and allows one to sequentially clone several genes from the same library using a single set of filters. However, nylon filters do not offer an improvement in sensitivity and are often more expensive than nitrocellulose filter paper.

Literature Review

The molecular basis of λ phage replication and the adaptation of the λ genome for molecular cloning has been reviewed by Arber et al. (1983) and Williams and Blattner (1980). Principles governing the plating of λ phage have been outlined by Arber (1983); see also UNIT 1.10. Thorough understanding of these principles has led to a universal approach to plating phage libraries.

Critical Parameters To prevent recombination between different phage, do not allow them to overgrow, and grow them in recombination-minus hosts where possible. Calculations of the amount of phage stock to be used per plate should be based on a recent titration, and plating cells should be fresh. Filters must not become brittle during this procedure; brittle filters will be destroyed during the hybridization process. This can be avoided by limiting the time in the hydroxide solution to less than 5 min, making certain that



Supplement 13

the 0.4 M Tris⋅Cl, pH 7.6/2× SSC brings the filters to neutral pH, and limiting the baking to 2 hr.

Troubleshooting Plaques should be visible on the plate before filters are made. If there appears to be poor bacterial growth, it is possible that the top agarose was too warm and many bacteria were killed, or that the phage titer was higher than expected and most host cells were lysed. Lower than expected phage titer could be due to an inaccurate titration of the phage stock, poor host-cell preparation, or too little time for adsorption. The preparation of the nitrocellulose filters will only be tested after hybridization is complete. Occasionally, hybridization to a plaque will produce a streak instead of a discrete circle on the autoradiograph, making location of the correct plaque difficult. Steps that will often correct this problem include: (1) drying plates with the cover removed for 1 to 2 hr before applying the filter, (2) drying the filters well before the hydroxide treatment, and (3) making certain that the face (phage side) of the filters is not directly in contact with the solutions.

Anticipated Results This plating procedure characteristically produces plates with an even distribution of dense phage particles. It is sensitive enough to allow identification of a phage by hybridization even when the phage are plated at high density (>5000 plaques per 82-mm plate). A signal is easily visible after 18 to 24 hr, when filters are hybridized to a nick-translated DNA probe with activity of >107 counts/µg DNA.

Time Considerations Usually plaques will become visible within 6 to 10 hr after plating. Bacteriophage should generally not be allowed to grow longer than necessary to visualize the plaques. Using the procedure outlined, even a large number of filters can be processed in a single day.

Literature Cited Arber, W. 1983. A beginner’s guide to lambda biology. In Lambda II (R.W. Hendrix, J.W. Roberts, F.W. Stahl, and R.A. Weisberg, eds.) pp. 381395. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Arber, W., Enquist, L., Hohn, B., Murray, N., and Murray, K. 1983. Experimental methods for use with lambda. In Lambda II (R.W., Hendrix, J.W. Roberts, F.W. Stahl, and R.A. Weisberg, eds.) pp. 433-466. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Williams, B.G. and Blattner, F.R. 1980. Bacteriophage lambda vectors for DNA cloning. In Genetic Engineering, Vol. 2 (J.K. Setlow and A. Mullander, eds.) p. 201. Plenum, NY.

Key References

Benton, W.D. and Davis, R.W. 1977. Screening λgt recombinant clones by hybridization to single plaques in situ. Science 196:180-182. Describes the method of plaque hybridization developed by the authors to allow isolation of phage possessing specific cloned DNA sequences.

Contributed by Thomas Quertermous Massachusetts General Hospital Boston, Massachusetts

Plating and Transferring Bacteriophage Libraries

6.1.4 Supplement 13


Plating and Transferring Cosmid and Plasmid Libraries A bacterial suspension is suctioned through a porous membrane, leaving the bacteria bound to the membrane surface. The membrane is transferred, bacteria up, to an agar plate upon which the bacteria will receive enough nutrients to grow into colonies. These filters can then be used for replica platings and for hybridization with specific DNA probes.

UNIT 6.2

BASIC PROTOCOL

Materials LB plates containing antibiotic (UNIT 1.1) LB medium (UNIT 1.1) LB plates containing 50 µg/ml chloramphenicol (UNIT 1.1) 0.5 M NaOH 1 M Tris⋅Cl, pH 7.5 0.5 M Tris⋅Cl, pH 7.5/1.25 M NaCl 10- or 15-cm Whatman 3MM or equivalent filter paper discs Sintered glass filter with vacuum Nitrocellulose membrane filters (10- or 15-cm, Millipore HATF) 20 × 20–cm Whatman 3MM or equivalent filter paper 20 × 20–cm glass plate 20-G needle 46 × 57–cm Whatman 3MM or equivalent filter paper 80°C vacuum oven NOTE: All materials coming into contact with E. coli must be sterile. Plating cosmids 1. Start with plasmid or cosmid library produced after transformation, transfection, or amplification (UNIT 5.7). 2. Determine titer of the library by serial dilutions using plates containing antibiotics (see UNIT 1.3). Remaining library suspension can be held at 4°C overnight with only minimal loss of viable bacteria. A 10-cm nitrocellulose filter can accommodate 10,000 to 20,000 colonies, while a 15-cm filter can hold up to 50,000.

3. Calculate the appropriate amount of the bacterial suspension for plating and dilute the suspension in LB medium such that there is the desired amount of bacteria in 5 ml (10-cm filter) or 10 ml (15-cm filter) of solution. 4. Meanwhile, prepare a layer of 10- or 15-cm Whatman 3MM paper discs on either the bottom part of a sintered glass Buchner funnel or on a porcelain filter funnel. Pour 10 to 20 ml LB medium over two or three layers of 3MM paper discs to make a level bed. The same pad of discs can be used for many filters. Sterilize filter apparatus and filter paper before use. The 3MM and nitrocellulose filters can be sterilized by autoclaving them while wrapped in aluminum foil. The purpose of this step is to spread the bacteria uniformly across the surface of a nitrocellulose filter. The filtering apparatus must be level, it must create a uniform suction to all the surface of the filter, and it should be easy to move the filters to and from the apparatus.

Contributed by Thomas Quertermous Current Protocols in Molecular Biology (1987) 6.1.1-6.1.4 Copyright © 2000 by John Wiley & Sons, Inc.


6.2.1

5. Label a nitrocellulose filter with a ballpoint pen on the side opposite that where the bacteria will be plated. Place the filter on the surface of the LB/antibiotic plate to wet it. The antibiotic plate must be permissive for cosmid- or plasmid-bearing bacterial cells and usually is ampicillin or tetracycline. Most ballpoint pen inks do not smudge during the hybridization reaction. If the one you choose runs, try another type.

6. Remove the wet filter from an antibiotic plate to the filtration apparatus. The suction should be off.

Carefully pipet the 5 to 10 ml of bacterial suspension onto the surface of the nitrocellulose filter, leaving the outer 4 to 5 mm of the filter free of solution. This outside bacteria-free ring leaves enough surface area to work with the filter without smearing or losing the colonies.

7. Slowly suction the solution down through the filter, taking care not to create any preferential suction pockets that would concentrate the bacteria. After suctioning all of the solution through the filter, transfer the filter back to the antibiotic plate on which it was wetted. In laying the filter down on the agar surface, take care to avoid trapping any air bubbles between the surface of the plate and the filter.

8. Plate the entire library in this way and incubate the plates upside down (agar side up) at 37°C until the colonies are ∼1 mm in diameter. Do not overgrow the filters, as smaller colonies can be lost beneath larger, faster-growing recombinant bacteria.

Preparing replica filters 9. Label and wet another set of nitrocellulose filters, as described in step 5. 10. Remove the initial library filter from its plate and place on several sheets of 20 × 20 cm 3MM paper, bacteria side up. While wearing gloves, carefully position the wetted replica filter above the bacterial lawn. Lay the second filter upon the first, leaving the two filters offset by 2 to 3 mm. This overlap will help in the separation of the two filters after the replica transfer. Do not allow air bubbles to form between the two filters. These are excluded by touching the second filter to the first in the middle and then allowing the edges to fall.

11. Lay three sheets of 20 × 20–cm 3MM paper on the two filters, followed by a 20 × 20 cm glass plate. Using the palms of your hands, press with all your weight down on the glass plate, thus transferring the bacterial colonies from the library filter to the replica filter. 12. Remove the glass plate and the filter paper and, using a 20-G needle, punch holes 2 to 4 cm apart through both of the filters. These holes will allow the orientation of the film produced from the replica filter down on the library filter for the isolation of the correct clones.

Plating and Transferring Cosmid and Plasmid Libraries

13. Carefully peel the two filters apart, placing them both bacteria up, on their respective agar plates. Grow the replica colonies at 37°C overnight, leaving the library filters at 25°C overnight. After overnight growth, store the library filters on the agar plates at 4°C, while screening the replica filters. Multiple replica filters can be made from the same library filter. Incubate library filters 2 to 4 hr at 37°C or overnight at 25°C to allow regrowth of the colonies.


Then repeat steps 9 to 13. Normally, two copies of the cosmid are hybridized to each probe.

14. After the bacterial colonies have grown, the cosmids or plasmids on the replica filter are amplified by transferring them to an LB plate containing 50 µg/ml chloramphenicol and incubating at 37°C for 4 to 10 hr. This step will increase the signal produced by hybridization. Preparing filters for hybridization 15. Remove the replica filters from the LB/chloramphenicol plates, place filters bacteria side up on a sheet of 46 × 57–cm 3MM paper soaked with 0.5 M NaOH, and leave them for 5 min. 16. Carefully transfer to a sheet of 46 × 57–cm 3MM paper soaked with 1 M Tris⋅Cl, pH 7.5. Allow neutralization to occur for 5 min. 17. Transfer to a third 46 × 57–cm filter soaked in 0.5 M Tris⋅Cl, pH 7.5/1.25 M NaCl. Neutralize 5 min. 18. Transfer filter to a dry sheet of 3MM paper to allow filter to dry. After filters are completely dry, stack them on paper towels or other adsorbent paper. Each nitrocellulose filter should be separated by paper towels from other filters.

19. Transfer the stacked filters to a vacuum oven at 80°C for 90 min. Remove filters and hybridize with a nick-translated probe, as described in UNITS 6.3 and 6.4. COMMENTARY Background Information There are two commonly used protocols for the screening of recombinant bacteria with hybridization probes. The first method involves the spreading of bacteria on the surface of agar using a sterile spreader (UNIT 1.3). A nitrocellulose membrane filter is then placed on top of the colonies and most of each colony is transferred to the filter. The filter is then treated as described in steps 15 to 19. This method works well when relatively small numbers of positive colonies are being selected (up to several thousand). The second method employs a matrix of some type (here nitrocellulose filters are used) upon which bacteria can be plated and grown when the filter is placed on top of a nutrient agar surface. Once the plated bacteria have grown into visible colonies, the filters can be used for replica plating and in situ hybridization analysis.

Critical Parameters In order to provide a uniform lawn of recombinant bacteria for screening, it is critical to ensure that the suction applied to the filters is uniform and not spotty. The best way to accomplish this is to suction the suspension through

the filter slowly and to avoid any preferential suction sites in the filter. Make sure that the apparatus is level and that adequate layers of LB-soaked chromatography paper are used. Air bubbles will prevent bacterial growth, so be certain that air is not trapped between the filter and the agar surface.

Time Considerations

Once the apparatus is set up, it takes ∼5 min per filter to wet the filter, suction the bacteria, and transfer to an LB plate. The colonies take ∼15 hr to grow at 37°C, after which they can be transferred to 4°C until ready for the replica platings. Replica plating also requires 5 min per filter, and resulting filters will be ready for denaturation and hybridization after 15 hr at 37°C.

Key Reference Hanahan, D. and Meselson, M. 1983. Plasmid screening at high density. Meth. Enzymol. 100:333-342.

Contributed by John H. Weis Harvard Medical School Boston, Massachusetts Screening Recombinant DNA Libraries


Supplement 24

SECTION II

HYBRIDIZATION WITH RADIOACTIVE PROBES After plaques or colonies have been transferred to nitrocellulose filters, the desired clone can be detected by its ability to hybridize to a DNA probe. This is a rapid, effective screening procedure that allows the identification of a single clone within a population of millions of other clones. The filters are hybridized with a 32P-labeled nucleic acid probe, the excess and incorrectly matched probe is washed off the filter, and the filter is autoradiographed. Two features of the nucleic acid probe used for these experiments are critical to the successful screening of recombinant DNA libraries. First, the probe must hybridize only to the desired clones and not to any other clones. Thus, the nucleic acid sequence used for a probe must not contain any reiterated sequences or sequences that will hybridize to the vector. Second, the specific activity of the probe must be at least 107 cpm/µg. Most of the procedures for labeling DNA or copy RNA molecules are described in Chapter 3, and a support protocol is presented here that allows the 5′ end-labeling of a mixture of oligonucleotides. The two basic protocols presented in this section describe steps required to hybridize labeled probes to recombinant DNA clones on filters. Two protocols are presented because conditions for hybridizing short oligonucleotide probes and longer nucleic acid probes to filters are different.


Using DNA Fragments as Probes HYBRIDIZATION IN FORMAMIDE Bacteriophage plaques or bacterial colonies bound to a filter membrane are detected by hybridization with a radioactive probe. Hybridization proceeds on prewet filters placed in a sealable plastic bag. After hybridization the filters are removed from the sealed bag, excess probe is washed off, and the filters are autoradiographed to identify the clones that have hybridized with the probe. Materials Nitrocellulose membrane filters bearing plaques, colonies, or DNA (UNITS 6.1 & 6.2) Hybridization solution I Radiolabeled probe, 1 to 15 ng/ml (UNIT 3.5) 2 mg/ml sonicated herring sperm DNA High-stringency wash buffer I Low-stringency wash buffer I Sealable bags 42°C incubator Water bath adjusted to washing temperature (see commentary) Glass baking dish Additional reagents and equipment for autoradiography (APPENDIX 3) Incubate filters with probe 1. Wet filters with hybridization solution I. Lay a filter membrane bearing plaques on top of 5 to 20 ml of hybridization solution I and allow solution to seep through filter. It is important to wet only one surface at a time to prevent trapping air in filter. Wet each filter in turn, producing a stack of wet filters.

Using DNA Fragments as Probes

6.3.1 Supplement 24

When multiple filters are to be hybridized to the same probe, no more than twenty 8.2-cm discs or ten 20 × 20 cm square filters should be placed in one stack. Contributed by William M. Strauss Current Protocols in Molecular Biology (1993) 6.3.1-6.3.6 Copyright © 2000 by John Wiley & Sons, Inc.

Estimate the volume of hybridization solution used to wet the filters; this is a significant fraction of the volume of the hybridization reaction.

2. Transfer the stack of wetted filters to an appropriately sized sealable bag. Add enough hybridization solution to generously cover filters and seal. Note the volume of hybridization solution used to cover the filters.

3. Prehybridize filters by placing the bag in a 42°C incubator for at least 1 hr. Some investigators omit this step.

4. While filters are prehybridizing, pipet the radioactive probe into a screw-cap tube, add 2 mg (1 ml) sonicated herring sperm DNA, and boil 10 min. Place boiled probes directly into ice to cool. The amount of probe used is important, and should be in the range of 1 to 15 ng/ml of hybridization reaction. The volume of the hybridization reaction can be assumed to be the amount of hybridization solution added to the filters.

5. Add 2 ml hybridization solution I to the boiled probe. 6. Remove bag containing filters from the 42°C incubator. Open bag, add probe mixture, exclude as many bubbles as possible, and reseal. A good way to add the radioactive probe is to take it up in a syringe with an 18-G needle and then inject it into the bag. Reseal the bag after adding probe.

7. Mix probe in the bag so that filter is evenly covered. Replace bag in the 42°C incubator and let hybridize overnight. Wash filters to remove nonhybridized probe 8. Warm 1 liter high-stringency wash buffer I to the “washing temperature” in a water bath. The stability of washing temperature and salt concentrations are critical features of this experiment. See discussion in commentary.

9. Remove bag containing hybridizing filters from the 42°C incubator. Cut bag open and squeeze hybridization solution out of the bag. CAUTION: Handle material carefully as it is extremely radioactive. This should be done on disposable paper bench covers.

10. Quickly immerse the filters in 500 ml low-stringency wash buffer I at room temperature in a glass baking dish. Separate all the filters, as they may stick together during hybridization. The volume of the low-stringency wash buffer is not important as long as the filters are completely covered. The filters must not be allowed to dry as the radioactive probe will irreversibly bind the filters if the filters dry in contact with probe. (The type of container used to hold the filters is not important as long as it transfers heat well. Thus glass, metal, or enamel containers are better than plastic.) The low-stringency wash only removes nonhybridized probe formamide and hybridization solution; it does not determine the stringency of the hybridization.

11. Rinse the filters three times with 500 ml low-stringency wash buffer. Let the filters sit 10 to 15 min at room temperature in low-stringency wash buffer with each rinse. 12. Pour off the low-stringency wash buffer and pour in 500 ml high-stringency wash buffer (prewarmed to washing temperature).



13. Replace the high-stringency wash buffer with another 500 ml of high-stringency wash buffer, then place the glass dish containing the filters in incubator at wash temperature. Make sure that the temperature in the glass dish reaches the desired washing temperature by placing a thermometer directly into the bath and measuring the temperature. Usually 15 to 20 min at the desired wash temperature is sufficient to remove most of the background radioactivity. Of course, if the glass dish is placed in a water bath, be careful that the water from the water bath does not get into the filters.

Autoradiographing filters 14. Remove filters and mount them either wet or dry on a plastic backing. If the filter(s) is to be exposed wet, then isolate it from the film by covering it with plastic wrap. Used X-ray film provides a good form of plastic backing for filters.

15. Mark the filters with radioactive ink to assist in alignment and autoradiograph. An easy way to apply radioactive ink is to mark adhesive-backed paper labels with radioactive ink and then attach the stickers to the plastic wrap cover. X-ray intensifying screens greatly decrease the amount of exposure time required. ALTERNATE PROTOCOL

HYBRIDIZATION IN AQUEOUS SOLUTION This method differs mainly in that formamide is not used in the hybridization solution. Follow the basic protocol except use the reagents and alternate parameters listed below. Additional Materials Hybridization solution II Low-stringency wash buffer II High-stringency wash buffer II 65°C incubator 1. Prehybridize as in basic protocol except that the filters are prehybridized at 65°C using hybridization solution II. Hybridization solution II may have to be prewarmed to solubilize the SDS.

2. Prepare probe as in step 4 of basic protocol and dilute with 2 ml of hybridization solution II. 3. Hybridize overnight as in steps 6 and 7 of basic protocol except use a hybridization temperature of 65°C. 4. Remove bag containing hybridization from the 65°C incubator. Squeeze out the hybridization solution, taking care to avoid contamination with the excess radioactive hybridization solution. 5. Immediately rinse filters twice with low-stringency wash buffer II. It is unnecessary to maintain a given temperature for this wash; just let the filters sit in wash buffer at room temperature until ready to proceed.

Using DNA Fragments as Probes

6. At 65°C, proceed to wash filters with high-stringency wash buffer II. Employ multiple quick washes (5 to 8) and immerse filter in a final wash for ∼20 min. Check the radioactivity of the filters with a Geiger counter and be certain that they produce a signal only a fewfold above background levels.


REAGENTS AND SOLUTIONS High-stringency wash buffer I 0.2× SSC (APPENDIX 2) 0.1% sodium dodecyl sulfate (SDS) High-stringency wash buffer II 1 mM Na2EDTA 40 mM NaHPO4, pH 7.2 1% SDS Hybridization solution I Mix following ingredients for range of volumes indicated (in milliliters): Formamide 20× SSC 2 M Tris⋅Cl, pH 7.6 100× Denhardts solution Deionized H2O 50% dextran sulfate 10% SDSa Total volume aIn place

24 12 0.5 0.5

48 24 1.0 1.0

72 36 1.5 1.5

120 60 2.5 2.5

240 120 5.0 5.0

2.5 10 0.5 50

5.0 20 1 100

7.5 30 1.5 150

12.5 50 2.5 250

25 100 5 500

480 240 10 10 50 200 10 1000

of SDS, N-lauroylsarcosine (Sarkosyl) may be used.

Add the SDS last. The solution may be stored for prolonged periods at room temperature. The dextran sulfate should be of high quality. Pharmacia produces acceptable-grade dextran sulfate. Recipes for SSC and Denhardt’s solution are in APPENDIX 2.

Hybridization solution II 1% crystalline BSA (fraction V) 1 mM EDTA 0.5 M NaHPO4, pH 7.2 (134 g Na2HPO4⋅7H2O plus 4 ml 85% H3PO4/liter = 1 M NaHPO4) 7% SDS Low-stringency wash buffer I 2× SSC (APPENDIX 2) 0.1% SDS Low-stringency wash buffer II 0.5% BSA (fraction V) 1 mM Na2EDTA 40 mM NaHPO4, pH 7.2 5% SDS Sonicated herring sperm DNA, 2 mg/ml Resuspend 1 g herring sperm DNA (Boehringer Mannheim #223636) in a convenient volume (about 50 ml of water) by sonicating briefly. The DNA is now ready to be sheared into short molecules by sonication. Place the tube containing the herring sperm DNA solution in an ice bath (the tube must be stable even if the ice begins to melt). The sonicator probe is placed in the DNA solution (without touching the bottom of the vessel). The sonicator is turned on to 50% power 20 min, or until there is a uniform and obvious decrease in viscosity. At no time should the tube containing the DNA become hot to the touch. After sonication, the DNA is diluted to a final concentration of 2 mg/ml, frozen in 50-ml aliquots, and thawed as needed.



Supplement 13

COMMENTARY Background Information All hybridization methods depend upon the ability of denatured DNA to reanneal when complementary strands are present in an environment near but below their Tm (melting temperature). In a hybridization reaction involving double-stranded DNA on a filter and a singlestranded DNA probe there are three different annealing reactions occurring. First, there are the desired probe-DNA interactions which result in signal. Second, there are mismatch interactions that occur between related but nonhomologous sequences; these mismatch hybrids are the ones that must be eliminated during the washing of the filters. Non-sequence-specific interactions also occur and these result in noise. The ability to extract information from a particular hybridization experiment is a function of the signal-to-noise ratio. High background or poor specific signal can both result in uninterpretable results. Washing nitrocellulose filters is required to remove excess radioactive probe, as well as radioactive probe that has bound to the DNA on the filter as mismatch hybrids. Temperature and salt concentration dramatically affect the maintenance of specific hybrids. Detergents and other charged species can have a profound effect upon the nonspecific binding of species that contribute to background. In this protocol, hybridization is achieved in a solution containing 50% formamide. Excess probe is rinsed away under low-stringency conditions so that further hybridization will not occur. Once the hybridization solution is rinsed away, it is possible to proceed to a high-stringency wash without fear of further hybridization. When washing is complete, the filters should produce very little “noise” when monitored with a Geiger counter. Although single-copy sequence probe normally does not produce a signal that is detectable with a Geiger counter, a probe corresponding to more abundant sequences will produce a signal that can be “heard” with a Geiger counter.

Literature Review Hybridization to filter membranes forms a basis of recombinant DNA technology and is described in detail earlier in the manual (UNIT 2.9). The protocols described here vary from those used for Southern blot filter hybridization in that the volume of the hybridization is usually larger and the washing conditions are different. Dextran sulfate is an important component of the hybridiUsing DNA Fragments as Probes

zation solution as it increases the rate of reassociation of the nucleic acids. The protocols in this unit describe methods for hybridizing radioactive probes to membranebound plaques or colonies. These procedures for screening recombinant clones were first suggested by Grunstein and Hognes (1975) and by Benton and Davis (1977). The conditions of hybridization proposed in the basic protocol involving hybridization in formamide was originally described by Denhardt (1966) and Gillespie and Spiegelman (1965) while the alternate protocol using aqueous hybridization solution was introduced by Church and Gilbert (1984). The method of washing filters under stringent conditions to remove background was first proposed by Southern (1975). Botchan et al. (1976) described the benefit of adding SDS to the wash solution. Jeffreys and Flavell (1977) first employed the wash conditions described in the protocols presented here.

Critical Parameters Hybridization. Kinetically, the hybridization of DNA (or RNA) probes to filter-bound DNA is not significantly different from hybridization in solution. For single-stranded probes, the rate of hybridization follows first-order kinetics, since probe is available in excess. Under conditions of excess probe, the time for hybridization is inversely proportional to the probe concentration. For double-stranded probes the rate of hybridization displays a more complex relationship to the initial probe concentration. However, to a first approximation the initial probe concentration is inversely proportional to the rate of hybridization. To determine the actual time required for the successful hybridization of a given probe, either empirical data must be available or the following formula can be used to determine the length of time (in hours) required to achieve 50% hybridization (T50): 1⁄

x

× y⁄5 × z⁄10 × 2 = T50

where x is the weight of probe in micrograms; y is the complexity of probe in kilobases; and z is the volume of hybridization solution in milliliters. The length of time T is given in hours. Maximum hybridization signal will be obtained if the reaction is allowed to proceed to 5 × T50, although 1 to 2 × T50 is often used. It is also clear that nonspecific interactions

6.3.5 Supplement 13


occur and that in any hybridization, sources of noise will be present. Therefore, from a practical standpoint one conventionally utilizes concentrations of nick-translated probe on the order of 1 to 15 ng/ml of hybridization, where the specific activity of the probe is from 5 × 107 cpm/µg to >108 cpm/µg. Too much probe in a hybridization is as bad as too little. One important source of background hybridization to filters is due to the hybridization of the probe to vector sequences or to E. coli DNA. Be certain that there is no vector or E.coli DNA sequences in the probe. This can best be ensured by isolating the probe from one type of vector (e.g., plasmid) and screening a library made with a different type of vector (e.g., bacteriophage). Washing temperature. Washing at low stringency is a straightforward proposition. Buffer is added at room temperature and washing proceeds at room temperature. High-stringency wash is determined empirically. The relative homology between the probe and target sequence is a determining parameter. If the homology is 100%, a high temperature (65° to 75°C) can be used. As the homology drops, lower washing temperatures must be used. In general one starts at 37° to 40°C, raising the temperature by 3° to 5°C intervals until background is low enough not to be a major factor in the autoradiography. The length of the probe is also important. Very short probes (5 × 107 cpm/µg and an overnight hybridization reaction with a 1-kb unique sequence probe, hybridizing bacterial colonies or bacteriophage plaques can be visualized after a 1 to 18 hr exposure.

Time Considerations Generally hybridizations are carried on overnight for 12 to 16 hr. This is sufficient for most probes and blots. However, with probes of increasing complexity longer hybridization times are required. This is preferable to increasing the probe concentration from experiment to experiment. Autoradiography requires 1 to 18 hr.

Literature Cited

Benton, W.D. and Davis, R.W. 1977. Screening λgt recombinant clones by hybridization to single plaques in situ. Science 196:180. Botchan, M., Topp, W., and Sambrook, J. 1976. The arrangement of simian virus 40 sequences in the DNA of transformed cells. Cell 9:269-287. Church, G. and Gilbert, W. 1984. Genomic sequencing. Proc. Natl. Acad. Sci. U.S.A. 81:1991-1995. Denhardt, D. 1966. A membrane filter technique for the detection of complementary DNA. Biochem. Biophys. Res. Commun. 23:641-646. Gillespie, D. and Spiegelman, S. 1965. A quantitative assay for DNA–RNA hybrids with DNA im mobiliz ed on a m em bra ne. J. M ol. Biol.12:829-842. Grunstein, M. and Hogness, D. 1975. Colony Hybridization: A method for the isolating of cloned DNA’s that contain a specific gene. Proc. Natl. Acad. Sci. U.S.A. 72:3961. Jeffreys, A.J. and Flavell, R.J. 1977. A physical map of the DNA region flanking the rabbit β globin gene. Cell 12:429-439. Southern, E.M. 1975. Detection of specific sequence among DNA fragments separated by gel electrophoresis. J. Mol. Biol. 98:503-517.

Contributed by William M. Strauss Harvard Medical School Boston, Massachusetts



Supplement 2

UNIT 6.4

Using Synthetic Oligonucleotides as Probes The protocols in this unit describe procedures for using mixtures of 32P-labeled oligonucleotides to screen recombinant DNA clones bound to nitrocellulose filters. A partial amino acid sequence of a protein is used to predict the nucleotide sequence of the gene that would encode it. A mixture of oligonucleotides is chosen that includes all possible nucleotide sequences encoding that amino acid sequence. This mixture of oligonucleotides is then used to screen a recombinant DNA library for the corresponding clones. In some cases however, the exact nucleotide sequence of a desired clone is known and it is possible to use a unique oligonucleotide as a probe.

BASIC PROTOCOL

HYBRIDIZATION IN SODIUM CHLORIDE/SODIUM CITRATE (SSC) This procedure outlines the steps necessary to screen nitrocellulose filters bearing DNA from bacteriophage or plasmids with mixtures of synthetic oligonucleotide probes. Hybridization and washing steps are carried out in solutions containing SSC. The washing temperature that produces the lowest background is determined empirically. Materials Membrane filters bearing plasmid, bacteriophage, or cosmid libraries (UNITS 6.1 & 6.2) 3× SSC/0.1% SDS Prehybridization solution SSC hybridization solution 6× SSC/0.05% sodium pyrophosphate, prewarmed to wash temperature Filter forceps (e.g., American Scientific Products #2568-1) Sealable bags (or equivalent) Additional reagents and equipment for autoradiography (APPENDIX 3) Prehybridize the filters 1. Prepare duplicate nitrocellulose filters of bacterial colonies or bacteriophage plaques. These should be processed and baked as described in UNITS 6.1 and 6.2. Although some authors recommend wiping the wet filters prior to baking to remove bacterial debris, we do not advise this procedure because the hybridization signal may be reduced. Filter forceps (i.e., without serrated tips) should be used to handle membrane filters to prevent marring the surface.

2. Wash the filters 3 to 5 times in 3× SSC/0.1% SDS at room temperature; about 50 82-mm filters can be washed in 500 ml. Then wash them once in the same solution at 65°C for at least 1.5 hr or overnight. This step removes much of the bacterial debris from the filters.

3. Remove filters from 3× SSC/0.1% SDS and prehybridize them 1 hr at 37°C in prehybridization solution. Herring sperm DNA in the prehybridization solution blocks nonspecific binding of probe to the filters and thus decreases the background level of radioactivity.

Using Synthetic Oligonucleotides as Probes

6.4.1 Supplement 2

Hybridize oligonucleotides to the filters 4. Remove filters from the prehybridization solution and put them into sealable bags containing SSC hybridization solution. Place up to 20 filters and ≥20 ml SSC hybridization solution into each bag. Add 0.125 to 1.0 ng of each 32P-labeled oligonucleotide per ml of hybridization solution to each bag. The mixed oligonucleotide probe is end-labeled with 32P as described in the support protocol. For example, to 20 ml of hybridization solution that will contain a mixture of 128 Contributed by Allan Duby, Kenneth A. Jacobs, and Anthony Celeste Current Protocols in Molecular Biology (1993) 6.4.1-6.4.10 Copyright © 2000 by John Wiley & Sons, Inc.

17-base oligonucleotides, add 320 ng (0.125 ng/ml × 128 oligonucleotides × 20 ml) of labeled probe. Hybridize filters 14 to 48 hr at the temperature indicated below: 14-base oligonucleotide 17-base oligonucleotide 20-base oligonucleotide 23-base oligonucleotide

room temperature 37°C 42°C 48°C

For bacterial colonies, adding much more than 0.125 ng of each oligonucleotide probe per ml of hybridization solution significantly increases the background on the autoradiogram. For bacteriophage plaques, there is less DNA per plaque than in a bacterial colony; as high backgrounds are not a problem with filters bearing bacteriophage plaques, more probe should be added to the hybridization mixture.

5. Remove filters from the hybridization bag and wash filters for 5 to 15 min, 3 to 5 times, in 6× SSC/0.05% pyrophosphate at room temperature. It is important that the filters are well separated from each other and that the solution is occasionally or continuously gently agitated.

Wash the filters 6. Wash filters for 30 min in prewarmed 6× SSC/0.05% sodium pyrophosphate at the temperature indicated below: 14-base oligonucleotide 17-base oligonucleotide 20-base oligonucleotide 23-base oligonucleotide

37°C 48°C 55°C 60°C

Adjust the temperature of 6× SSC/0.05% pyrophosphate and filters. Measure the temperature of the filters and surrounding solution by putting the thermometer into the solution, not into the water bath. Make sure the filters are separated and are occasionally or continuously gently agitated.

7. Examine the filters with a Geiger counter; they should not exhibit above-background radioactivity. If the filters still show a significant degree of radioactivity above background, increase the temperature by 2° to 3°C for 15 to 30 min and reexamine the filters with the Geiger counter. Do not exceed the following temperatures: 14-base oligonucleotide 17-base oligonucleotide 20-base oligonucleotide 23-base oligonucleotide

41°C 53°C 63°C 70°C

The background level of bound radioactivity depends upon the amount of bacterial debris left on the filters, the amount of labeled oligonucleotides added to the hybridization mixture, and the guanosine-cytosine (G-C) content of the oligonucleotide mixture.

Perform autoradiography 8. When the filters exhibit a low level of radioactivity or the maximum temperatures referred to in step 7 have been reached, the filters should be removed from the wash solution and mounted wet on a solid support before exposure at −70°C to X-ray film, using an intensifying screen. Cover filters with plastic wrap. Do not allow the filters to dry out. Allow films to expose for 14 to 72 hr. Autoradiograms made from filters with a high background may still yield interpretable results.



Supplement 13

9. Develop the films; if a high background prevents proper interpretation of the films, rewash the filters at a higher temperature. 10. Number and mark the orientation of the films as described in UNIT 6.3. Spots that appear in precisely the same place on duplicate filters are “positives” (winners) and should be processed as described in UNIT 6.5. It is impossible to identify the characteristics of a true positive spot. Only colonies or plaques that produce evidence of hybridization on both filter copies should be processed as described below. Note that the intensity of the spot can vary dramatically between the duplicate filters. If a clear-cut spot appears on one filter and only a darkening of the background appears on the other, this should be considered positive and the plate should be processed as described in UNIT 6.5. Note that if two different oligonucleotide mixtures representing two different parts of the protein are available, either the positives obtained with one probe can then be hybridized with the other probe or four filter copies of the library can be made and hybridized to the two probes. Of course, depending on how far apart the sequences that hybridize to the two probes are, it is possible that neither will be present on a less than full-length cDNA clone. BASIC PROTOCOL

HYBRIDIZATION IN TETRAMETHYLAMMONIUM CHLORIDE (TMAC) This procedure is similar to the SSC protocol except that hybridization and washing are performed in solutions containing TMAC. In TMAC, the melting temperature of an oligonucleotide is a function of length and is independent of base composition; thus, spurious hybridization due to high G-C content of some of the oligonucleotides is reduced. Conditions are described for using 17-base oligonucleotides, but information is provided for determining the conditions when oligonucleotides of various lengths are employed. Materials Nitrocellulose or nylon membrane filters bearing plasmid, bacteriophage, or cosmid libraries (UNITS 6.1 and 6.2) 150-mm LB agarose plates (UNIT 1.1), prewarmed to 37°C 2× SSC/0.5% SDS/50 mM EDTA, pH 8.0, prewarmed to 50°C TMAC hybridization solution, prewarmed to hybridization temperature TMAC wash solution 2× SSC/0.1% SDS 15-cm glass crystallizing dishes Filter forceps (e.g., American Scientific Products #2568-1) Additional reagents and equipment for autoradiography (APPENDIX 3) Process and prehybridize the filters 1. Process filters bearing bacterial colonies as described in bearing amplified bacteriophage plaques as follows:

UNIT 6.2.

Produce filters

a. Plate the bacteriophage from the library on LB agarose plates and transfer to nitrocellulose filters as described in UNIT 6.1, steps 1 to 7. To obtain maximum sensitivity with oligonucleotide probes when the amplification procedure is used, plating density should be reduced to 8,000 to 10,000 plaques per 150-mm plate.

Using Synthetic Oligonucleotides as Probes

Either nitrocellulose or nylon (Colony/Plaque Screen Filters by New England Nuclear) filters can be used in this procedure. Nitrocellulose filters become fragile when hybridized in TMAC and must be handled very carefully. If this becomes a problem and nylon filters are substituted, the phage plaques must be amplified overnight. The rest of the protocol is unchanged.

6.4.3 Supplement 13


b. Amplify the bacteriophage by transferring the wet filter to a prewarmed (37°C) LB agarose plate so that the surface bearing the bacteriophage is faceup. Refrigerate the master plates upon which the recombinant phage library were plated to prevent any further plaque expansion.

c. Incubate the plates at 37°C until the bacterial lawn re-forms on the surface of the nitrocellulose and plaques are evident. Plaque size will be somewhat larger than those on the original plate. This usually requires a 5- to 12-hr incubation period. Longer periods of growth will produce a dense bacterial lawn without significantly increasing plaque size or affecting hybridization signal. Bacteriophage that produce small plaques (e.g., EMBL) are usually plated in the evening and allowed to grow overnight. The plaques are transferred to nitrocellulose filters the following morning and the phage are amplified on the filters by incubation for 5 to 7 hr during the day. Phage that produce large plaques (e.g., λgt10) are plated early in the morning, allowed to grow 5 to 7 hr, transferred to nitrocellulose filters (steps 6 and 7 of UNIT 6.1), transferred to fresh plates, and then incubated for amplification overnight.

d. Denature and bind the bacteriophage DNA to nitrocellulose filters as described in steps 8 to 11 of UNIT 6.1. 2. Wash filters bearing bacterial colonies as described in step 1 of the SSC protocol. Wet bacteriophage-bearing filters in a prewarmed (50°C) solution of 2× SSC/0.5% SDS/50 mM EDTA (pH 8.0). Float the filters on top of the solution (with the surface containing the dried bacteria and plaques faceup) to allow the nitrocellulose to wet completely. Submerge the filters and, with a gloved hand, gently rub the surface of the filters to remove the dried bacterial debris. Transfer the filters to a container of fresh solution of 2× SSC/0.5% SDS/50 mM EDTA to remove bacterial debris. Alternatively, the filters can be incubated in this solution at 65°C for one to several hours and then scrubbed. Inadequate scrubbing of the filters results in an increase of nonspecific background hybridization, obscuring positive hybridization signals in the subsequent screening procedure.

3. Transfer the filters to a 15-cm glass crystallizing dish containing 5 to 10 ml TMAC hybridization solution (per filter), which has been prewarmed to the appropriate hybridization temperature (48°C for 17-mer oligonucleotides; see Fig. 6.4.1 and commentary for other oligonucleotides) and seal the dishes with plastic wrap and rubberbands. Prehybridize 1 to 2 hr at the hybridization temperature, which is 5° to 10°C below the melting temperature. Prehybridization and hybridization can be performed in glass crystallizing dishes that are slightly larger in diameter than the nitrocellulose filters. Gentle agitation on an orbital platform shaker will allow the solution to pass freely between the stacked filters and prevent the filters from sticking together. Place no more than 25 to 30 filters in each dish. Alternatively, prehybridization and hybridization can be performed in a sealable bag (see SSC protocol) with 7000 Ci/mmol) 25 to 50 U T4 polynucleotide kinase (UNIT 3.10) and 10× kinase buffer (UNIT 3.4) Ice-cold 10% trichloroacetic acid (TCA) 1. Set up reaction mixture on ice in microcentrifuge tube as follows: 2.5 to 250 pmol mixed oligonucleotides 7.5 µl 10× T4 polynucleotide kinase buffer 66 pmol [γ-32P]ATP (200 µCi) 25 to 50 U T4 polynucleotide kinase H2O to 75 µl Incubate 30 min at 37°C. The reaction mixture should have either equimolar amounts of label and oligonucleotide ends, or the label should be in molar excess. 1 mol deoxyribonucleotide ≅ 330 g 1 OD260 ≅ 40 ìg/ml oligonucleotide 1 ìg 14-base oligonucleotide ≅ 0.24 nmol 1 ìg 17-base oligonucleotide ≅ 0.18 nmol 1 ìg 20-base oligonucleotide ≅ 0.15 nmol

2. At the end of the reaction, check for incorporation of label by precipitating 1 µl of a diluted aliquot with ice-cold 10% TCA (acid precipitation, UNIT 3.4) and counting the incorporated radioactivity. Using equimolar amounts of oligonucleotide and label, ∼30% to 90% of the counts are incorporated. The labeled oligonucleotide can be further purified by a combination of phenol extraction and/or ethanol precipitation (UNIT 2.1). To remove unincorporated label, oligonucleotides of 17 bases or longer can be quantitatively precipitated from a solution of 2.5 M ammonium acetate containing 25 ìg carrier DNA plus 9 vol of 100% ethanol. The resulting pellets are washed with 70% ethanol, followed by 95% ethanol, air dried, and resuspended in 100 ìl TE buffer.

3. Store mixture in appropriate container at −20°C. REAGENTS AND SOLUTIONS Prehybridization solution 6× SSC (APPENDIX 2) 5× Denhardts solution (APPENDIX 2) 0.05% sodium pyrophosphate 100 µg/ml boiled herring sperm DNA continued



Supplement 9

0.5% sodium dodecyl sulfate (SDS) SSC hybridization solution 6× SSC (APPENDIX 2) 1× Denhardt’s solution (APPENDIX 2) 100 µg/ml yeast tRNA 0.05% sodium pyrophosphate TMAC hybridization solution 3 M tetramethylammonium chloride (see recipe below for stock solution) 0.1 M NaPO4, pH 6.8 1 mM EDTA, pH 8.0 5× Denhardt’s solution (APPENDIX 2) 0.6% SDS 100 µg/ml denatured salmon sperm DNA TMAC wash solution 3 M tetramethylammonium chloride (see recipe below for stock solution) 50 mM Tris⋅Cl, pH 8.0 0.2% SDS Tetramethylammonium chloride (TMAC), 6 M stock solution Dissolve 657.6 g TMAC (mol wt = 109.6) in H2O and bring to 1 liter. Filter the solution through Whatman No. 1 filter paper and determine the precise concentration of the solution by measuring the refractive index (n) of a 3-fold diluted solution. The molarity (M) of the diluted solution = 55.6(n − 1.331) and the molarity of the stock solution = 3 × M. TMAC can be stored at room temperature in brown bottles. CAUTION: TMAC can irritate eyes, skin, and mucous membranes. It should be used with adequate ventilation in a fume hood. Used TMAC solutions should be collected and discarded as hazardous and/or radioactive waste. Small amounts (1 Mb have been produced and these are routinely propagated with apparent stability, suggesting that the major limitation to the size of YAC inserts is the quality of the starting genomic DNA. Large “core” laboratories that generate human YAC libraries— such as the Center for Genetics in Medicine, Washington University School of Medicine, St. Louis; the Centre d’Etude du Polymorphisme Humain (CEPH), Paris; and the Genome Analysis Laboratory, Imperial Cancer Research Fund, London—prepare human YACs with average insert sizes ranging from 0.3 to 1.2 Mb. Additional high-quality YAC libraries have been constructed using inserts from Drosophila melanogaster, Caenorhabditis elegans, Schizosaccharomyces pombe, and mouse (Burke et al., 1991; Rossi et al., 1992). Anecdotal reports indicate YAC libraries may support the propagation of certain insert sequences that are poorly represented in Escherichia coli–based libraries. The YAC cloning system also offers the advantage that large genomic YAC inserts can be easily manipulated in yeast by homologous recombination. Thus, it is relatively simple to truncate a YAC insert or to introduce specific deletions, insertions, or point mutations with high efficiency using methods such as those described in UNIT 13.10. This unit provides an introduction to the use of yeast artificial chromosome–bearing yeast clones (hereafter referred to as YAC clones) in genome analysis. It describes criteria for de-

signing a polymerase chain reaction (PCR) assay to be used in screening a YAC core library and discusses the rationale for verification and characterization of YAC clones obtained from these core laboratories. Protocols for maintaining YAC clones, analyzing YAC insert structure, preparing YAC DNA, and subcloning YAC inserts into other vectors are presented in UNIT 6.10. These protocols are outlined in the flow chart in Figure 6.9.1.

GENERATING YAC LIBRARIES Although YAC cloning is the method of choice when insert sizes >100 kb are required, a number of features of the system have interfered with its rapid assimilation for routine cloning. Because the S. cerevisiae genome is at least an order of magnitude more complex than the E. coli genome and existing YACs are carried as only a single copy within yeast cells, the signal-to-noise ratio is less favorable for identifying a cognate clone in a YAC library than in a λ or cosmid library. Moreover, efforts to develop high-density screening methods for YACs have enjoyed only limited success. Most laboratories that maintain YAC libraries organize them as collections of individual clones in 96-well microtiter plates, which can be replicated faithfully and kept frozen for storage; in this form, a standard library representing 5 to 8 genome-equivalents comprises more than 500 microtiter plates. As a result, the effort and resources required to construct YAC libraries and prepare them for screening are enormous. Consequently, it is generally most practical for investigators wishing to obtain YACs carrying a specific DNA sequence to arrange for screening of a preexisting library maintained by a core laboratory. Initially, YAC libraries were constructed with total genomic DNA (Burke et al., 1987). More recently, there has been interest in generating libraries from targeted DNA using somatic cell hybrids carrying a specific chromosome or portion of a chromosome. The feasibility of this approach has been demonstrated with the construction of a library carrying a portion of the human X chromosome (Abidi et

Contributed by David D. Chaplin and Bernard H. Brownstein Current Protocols in Molecular Biology (1992) 6.9.1-6.9.7 Copyright © 2000 by John Wiley & Sons, Inc.


6.9.1 Supplement 20

al., 1990). Additional targeted libraries are in the late stages of development and should reduce the cost and effort of screening for loci whose chromosomal location has been established.

YAC LIBRARY SCREENING BY A CORE LABORATORY Methods used by YAC core laboratories for library screening evolve rapidly. It is possible to screen a library by hybridizing a single-copy probe to nylon filters stamped with a replica of one or more microtiter arrays. However, because of the low signal-to-noise ratio for hybridization and the substantial cost required to produce all of the nylon filter replicas, most laboratories perform library screening using PCR (Green and Olson, 1990). At the time of this writing, most core facilities first extract DNA from pools of clones, usually representing 1 to 4 microtiter plates (96 to 384 YACs) per pool, and then combine this pooled DNA into more superpools of 1500 to 2000 YACs. The pools and superpools are screened by PCR to identify candidate microtiter plates containing at least one amplifying YAC clone. Final identification of the clone is most commonly performed either by colony hybridization using the PCR product as the probe or by screening pools of rows and col-

umns from the same microtiter plate using PCR. The time required for a YAC core laboratory to verify the specificity and parameters of the PCR assay and screen complex clone pools and subpools is usually 3 to 8 weeks. As an example, the screening strategy used by one major core laboratory is described in the accompanying box. It should be noted that this procedure may change as technology advances; for instance, the recent advent of techniques providing reliable DNA extraction from small quantities of thousands of individual clones has made it feasible to screen individual wells on a plate and eliminate the laborious filter-hybridization step.

DESIGNING A LOCUS-SPECIFIC PCR ASSAY FOR SCREENING An investigator arranging with a core laboratory for library screening is required to design a strategy for detecting the inserted genomic DNA and to provide the appropriate probe(s). It is worth investing considerable effort to create a convenient and reliable assay because the assay’s success depends on its ability to detect the target sequence with high sensitivity while being insensitive to the presence of large excesses of yeast and plasmid sequences. Because a core laboratory must adopt PCR assays that have been imported

EXAMPLE: SCREENING OF HUMAN-GENOME YAC LIBRARY AT THE WASHINGTON UNIVERSITY SCHOOL OF MEDICINE At this core facility, screening of the human-genome YAC library proceeds in three stages: (1) initial evaluation of the PCR assay; (2) screening of pools of YACs; and (3) identification of individual YACs from subpools to the single well by filter hybridization. To permit pretesting of assays before they are sent to a screening core, new PCR assays are evaluated using four control DNA samples as templates: (1) CGM-1 human genomic DNA (33 ng/µl) from a lymphoblastoid cell line established from the donor whose DNA was used in preparing the YAC library; (2) YY212 DNA from a yeast strain carrying a YAC whose insert is yeast chromosomal DNA; (3) “single-membrane-pool” DNA (33 ng/µl) prepared from a pool of 396 YAC isolates; and “spiked-pool” DNA, which is single-membrane-pool DNA augmented with 5 ng/µl of CGM-1 DNA.

Yeast Artificial Chromosome Libraries

CGM-1 DNA serves as a positive control to demonstrate that the sensitivity of the PCR assay is adequate. YY212 DNA serves as a negative control, demonstrating that no product is amplified from either yeast host genomic DNA or the YAC vector. DNA from the singlemembrane pool and the spiked pool provide additional negative/positive controls that more closely mimic library screening conditions. A negative signal from single-membrane-pool DNA demonstrates lack of cross-reactivity of the probe with the YAC vector, yeast genomic DNA, or common human repetitive sequences. A positive signal obtained from the spikedpool DNA (containing only 5 ng/µl of CGM-1 DNA) is a strong indication that the assay possesses sufficient sensitivity against a yeast DNA background for successful library screening.

6.9.2 Supplement 20


design a PCR assay for identifying YAC clone of interest from genomic YAC library (UNIT 6.9 )

obtain isolated YAC clone from core facility (UNIT 6.9 )

validate identify of YAC using PCR (UNITS 6.9 & 15.2 )

grow and store YAC clone (UNIT 6.10, first basic protocol)

prepare DNA from isolated YAC clone and analyze by Southern blotting (UNIT 6.10, second basic protocol, and UNIT 2.9 )

analyze isolated YAC clone for chimerism using PCR (UNIT 6.10, fourth basic protocol)

prepare DNA from isolated YAC clone using agarose plugs and analyze by PFGE (UNIT 6.10, third basic protocol, and UNIT 2.5B )

analyze isolated YAC clone for chimerism by subcloning in bacterial vector (UNIT 6.10, alternate and support protocols)

for high-resolution analysis (optional) prepare high-molecular-weight YAC-containing DNA (UNIT 6.10, fifth basic protocol)

subclone high-molecular-weight YAC-containing DNA into cosmid or λ vector (UNIT 6.10, sixth basic protocol)

Figure 6.9.1 Flow chart showing protocols used to obtain and analyze YAC clones.



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from outside laboratories, it is a good idea to inquire in advance about the protocols preferred by the specific core facility that will be performing the screening. In general, any highly specific, sensitive, and robust PCR assay is suitable for screening a YAC library (see Chapter 15). Typically, two 18- to 30-mer oligonucleotide primers for use in amplifying a single-copy 75- to 750-bp product are satisfactory. Such primers define a landmark for genome mapping called an STS (sequence-tagged site; see introduction to Chapter 7 and Olson et al., 1989). When designing a PCR assay from scratch, it is useful to consider the following:

Fragment size The STS should be 75 to 750 bp in length. Fragments in this range are most efficiently amplified by PCR and are easily detected by either polyacrylamide (UNIT 2.7) or standard agarose (UNIT 2.5A) gel electrophoresis.

assay, it does help eliminate some of the most trivial causes of assay failure.

ANALYZING INDIVIDUAL YAC CLONES Once library screening has been successfully completed and the isolated YAC clone has been furnished to the investigator, attention should be directed to analyzing its structure. Initial studies should focus on determining whether the genomic insert is chimeric, checking for evidence of rearrangement within the insert, and verifying that the YAC is propagated in stable fashion in the yeast cell (see below). Simply analyzing several isolates of the same YAC in parallel may provide a means of recognizing instability, as each isolate serves as a control for the others. The following sections give an overview of strategies for analyzing YAC clones; specific protocols are given in UNIT 6.10.

Chimerism of the YAC Insert Primer length Each primer should ideally be 18 to 30 nucleotides long, be composed of 50% to 55% G + C, and be contained within a single-copy human-genomic-DNA segment. This ensures efficient priming and decreases the probability of false priming, enhancing the sensitivity and specificity of the assay. This also permits the amplified fragment to be used as a hybridization probe in the final hybridization-dependent steps of library screening (see below). If it is not possible to amplify a single-copy fragment, then some other single-copy probe (e.g., a synthetic oligonucleotide 30 nucleotides long) should also be prepared. Oligonucleotide design strategies are discussed further in UNITS 2.11 & 15.1.

Primer affinity


Primers should show little affinity for selfannealing or for annealing with each other. This prevents the production of small, template-independent PCR products that compete for primers in the reaction. A number of academic and commercial DOS-based and Macintosh software programs permit rapid selection of non-self-annealing primers from within a known DNA sequence (e.g., Oligo 4.0, National Biosciences; Primer, S. Lincoln and M. Daly, Whitehead Institute for Biomedical Research, Cambridge, Mass., and OSP, Hillier and Green, 1991; see UNIT 7.7). Although the use of these programs cannot remove all the uncertainty associated with designing a new PCR

A consistent problem in YAC cloning is chimerism of the YAC insert—i.e., the insert is composed of two or more separate genomic fragments joined in a single YAC. The mechanism(s) giving rise to chimeric YAC clones are currently not fully understood (Green et al., 1991). In most existing total genomic YAC libraries, chimeric clones represent from 5% to 50% of the total clones. Preliminary data suggest that targeted, chromosome-specific libraries may contain only 5% to 15% chimeric clones. Although future generations of YAC libraries are likely to contain lower frequencies of chimeric clones, chimerism will probably remain a significant problem requiring assessment for every new YAC clone being analyzed. The most reliable way to determine if a YAC insert is chimeric is to isolate a small fragment from each end of the insert and determine its chromosome of origin and whether it shares sequences with overlapping YACs derived from the same chromosomal region. Many approaches have been suggested for isolating such YAC genomic insert end fragments, all of them relying upon the fact that end fragments are marked by their adjacent YAC vector sequences. Thus, it is possible to determine whether a YAC insert is chimeric by preparing probes from the two YAC vector arms and using these to demonstrate that both ends of the YAC map to the same general chromosomal region. This is generally done using hybridization or PCR analysis of a somatic hybrid cell line containing the appropriate human chromo-

6.9.4 Supplement 20


some (or preferably a fragment thereof) as its sole human DNA. The appropriate end-fragment probes may be produced in several ways, two of which are presented in UNIT 6.10. The most rapid and versatile approaches to producing end-fragment probes use PCR amplification (Riley et al., 1990; Green, 1992). The template is a restriction fragment produced by cutting the YAC DNA at sites near the two ends of the genomic insert. The YAC DNA is digested with a frequently cutting restriction endonuclease to produce a collection of small restriction fragments. One of the fragments contains the distal portion of the YAC insert still associated with a portion of the left vector arm, while another contains the other end of the insert associated with a portion of the right vector arm (the arm of the YAC vector containing the yeast centromere is arbitrarily designated the left vector arm and the arm containing the ura3 selection marker is arbitrarily designated the right vector arm). These fragments are prepared for PCR amplification by ligation of a synthetic double-stranded DNA tag to both ends. This tag contains a 29-nucleotide “bubble” of noncomplementary sequence flanked by two 12-nucleotide complementary sequences (Fig. 6.10.2). The two YAC-insert end fragments can then be selectively amplified using one PCR primer derived from the YAC vector and one primer with the sequence of the noncomplementary portion of the bubble. These methods are generally favored because of their speed, but they depend on the fortuitous placement of restriction sites close enough to the ends of the genomic insert that a fragment suitably sized for PCR amplification can be generated. Moreover, if highly repetitive sequences are present at the distal portions of the insert, the PCR method may fail to generate useful information. A reliable but more time-consuming method of generating probes for end-fragment analysis is conventional subcloning of larger YAC-derived restriction fragments into plasmid or λ vectors (Bronson et al., 1991). Subcloning an end fragment several kilobases in size is timeconsuming, but reliably assures identification of nonrepeated sequences for use as probes. The subcloning protocol given in UNIT 6.10 involves double-digesting the YAC DNA to enrich for end fragments in the course of subcloning the insert into a pUC19-based vector. One of two specific enzymes that cut rarely in yeast and human genomic DNA, ClaI or SalI, is included in the double digestion mixture. A ClaI recognition sequence lies in the left arm

of the YAC vector, while a SalI recognition site lies in the right arm (Fig. 6.10.1). When one of these rarely cutting restriction enzymes is used together with a frequently cutting enzyme, doubly-digested fragments constitute only a small fraction of the total digested product. Ligation to a doubly-digested plasmid vector eliminates all of the single-digested fragments, resulting in a substantial enrichment for the YAC end fragment.

Internal Rearrangement or Instability of the YAC Insert Internal rearrangement of a YAC insert is more difficult to identify than chimerism, and may become apparent only after high-resolution analysis of the clone. Existing reports of internal rearrangement of YAC inserts are anecdotal, infrequent, and usually identify only rather large-scale changes. It is likely, however, that subtle rearrangements will be recognized as more clones are analyzed. Nevertheless, the data suggest that important rearrangements will remain relatively infrequent and will not impede most YAC cloning efforts. Although YACs are usually stable in culture, deletion or other rearrangements of the insert may occur months after the initial isolation of a clone. Thus, it is wise to verify the size of a YAC following prolonged passage in culture or after it has been thawed from a frozen stock. Several different colonies of the same YAC strain should be analyzed in parallel, using the protocols in UNIT 6.10, to confirm that the artificial chromosome is the same size in each of the isolates. Because cytosine methylation, which is quite frequent in the DNA of higher eukaryotic species, does not occur in yeast (Proffitt et al., 1984), it is not possible to perform direct structural comparisons of the YAC inserts and the corresponding genomic DNA isolated from higher eukaryotic cells using infrequently cutting restriction enzymes to create large-scale restriction maps. Consequently, direct structural comparisons must be carried out using methylation-insensitive restriction enzymes and frequently spaced probes. Evidence of internal rearrangement within a YAC clone can be obtained by preparing chromosomes from the clone (UNIT 6.10) and analyzing them by pulsed-field gel electrophoresis (PFGE; UNIT 2.5B). The CHEF gel system (Vollrath and Davis, 1987) is particularly useful in that it permits excellent resolution in the size range most common for individual YAC clones. Following electrophoresis, the artificial chro-



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mosome can be visualized using ethidium bromide staining as an extra chromosome not present in the host yeast strain. Occasionally, the YAC is not immediately recognizable because it comigrates with one of the endogenous yeast chromosomes. As described in the third basic protocol in UNIT 6.10, the PFGE gel should be Southern blotted and analyzed by successive hybridization with probes specific for the locus used in the library screening and for one or both of the YAC vector arms [e.g., for pYAC4, the 351-bp ClaI/BamHI fragment of pBR322 (left YAC arm) and the 276-bp BamHI/SalI fragment of pBR322 (right YAC arm)]. These blots should show hybridization to a single chromosome of the same size in all isolates from the same YAC strain as well as no hybridization to the AB1380 host strain.

CONSTRUCTION AND ANALYSIS OF A YAC-INSERT SUBLIBRARY


Although the large genomic DNA fragments provided by the YAC cloning system are easy to manipulate, it is often convenient to reduce a YAC to smaller fragments by subcloning it into a cosmid or λ vector. In particular, such smaller fragments are more amenable to highresolution analysis; this is important because information concerning the specific content of the YAC insert is typically limited, and often the only internal probe that is available is the one used for YAC library screening. Protocols for preparing YAC insert DNA and constructing a cosmid sublibrary are provided in UNIT 6.10. Two general strategies are available for preparing YAC insert DNA in order to create a saturating collection of subclones. The more elegant strategy is to purify the artificial chromosome itself by preparative CHEF gel electrophoresis (UNIT 2.5B). This permits isolation and analysis of the resulting recombinant clones without further selection, assuming that only a small amount of contaminating yeast DNA is present in the purified YAC, and that essentially all subclones isolated are derived from the human YAC insert. In practice, however, it is difficult to recover sufficient quantities of purified YAC DNA to permit construction of a cosmid or λ library. An alternate approach is to prepare a library from the total DNA of the YAC-carrying yeast strain. YACspecific subclones must then be selected by hybridization. An initial round of screening is usually performed with total human genomic DNA (rich in repetitive sequences) as the probe.

This detects subclones that contain human repetitive elements and eliminates subclones consisting of yeast DNA. Additional analysis is performed to identify overlapping sequences and thereby establish an approximate map of the original YAC insert. Ultimately, one or more rounds of chromosome walking may be required to fill in gaps between contiguous groups of subclones.

Literature Cited Abidi, F.E., Wada, M., Little, R.D., and Schlessinger, D. 1990. Yeast artificial chromosomes containing human Xq24-Xq28 DNA: Library construction and representation of probe sequences. Genomics 7:363-376. Bronson, S.K., Pei, J., Taillon-Miller, P., Chorney, M.J., Geraghty, D.E., and Chaplin, D.D. 1991. Isolation and characterization of yeast artificial chromosome clones linking the HLA-B and HLA-C loci. Proc. Natl. Acad. Sci. U.S.A. 88:1671-1675. Burke, D.T., Carle, G.F., and Olson, M.V. 1987. Cloning of large segments of exogenous DNA into yeast by means of artificial chromosome vectors. Science 236:806-812. Burke, D.T., Rossi, J.M., Leung, J., Koos, D.S., and Tilghman, S.M. 1991. A mouse genomic library of yeast artificial chromosome clones. Mamm. Genome 1:65-69. Green, E.D. and Olson, M.V. 1990. Systematic screening of yeast artificial chromosome libraries by use of the polymerase chain reaction. Proc. Natl. Acad. Sci. U.S.A. 87:1213-1217. Green, E.D., Riethman, H.C., Dutchik, J.E., and Olson, M.V. 1991. Detection and characterization of chimeric yeast artificial-chromosome clones. Genomics 11:658-669. Green, E.D. 1992. Physical mapping of human chromosomes: Generation of chromosome-specific sequence-tagged sites (STS). Methods Mol. Genet. In press. Hillier, L. and Green, P. 1991. A computer program for choosing PCR and DNA sequencing primers. PCR Meth. Appl. 1:124-128. Olson, M., Hood, L., Cantor, C., and Botstein, D. 1989. A common language for physical mapping of the human genome. Science 245:1434-1435. Proffitt, J.H., Davie, J.R., Swinton, D., and Hattman, S. 1984. 5-Methylcytosine is not detectable in Saccharomyces cerevisiae DNA. Mol. Cell Biol. 4:985-988. Riley, J., Butler, R., Ogilvie, D., Finniear, R., Jenner, D., Powell, S., Anand, R., Smith, J.C., and Markham, A.F. 1990. A novel, rapid method for the isolation of terminal sequences from yeast artificial chromosome (YAC) clones. Nucl. Acids Res. 18:2887-2890.

6.9.6 Supplement 20


Rossi, J.M., Burke, D.T., Leung, J.C., Koos, D.S., Chen, H., and Tilghman, S.M. 1992. Genome analysis using a yeast artificial chromosome library with mouse DNA inserts. Proc. Natl. Acad. Sci. U.S.A. 89:2456-2460. Vollrath, D. and Davis, R.W. 1987. Resolution of DNA molecules greater than 5 megabases by contour-clamped homogeneous electric fields. Nucl. Acids Res. 15:7865-7876.

Contributed by David D. Chaplin and Bernard H. Brownstein Howard Hughes Medical Institute and Washington University School of Medicine St. Louis, Missouri

Key Reference Burke, et al., 1987. See above. Initial description of the YAC cloning system, covering general features of library construction.



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UNIT 6.10

Analysis of Isolated YAC Clones The preceding unit gives an overview of methods involved in screening a YAC library to isolate a particular clone of interest (UNIT 6.9), with the sequence of methods illustrated in a flow chart (Fig. 6.9.1). This unit provides a series of protocols describing the analysis and manipulation of an isolated YAC clone. The procedures are based upon the use of the YAC vector pYAC4. Once an isolated YAC clone has been obtained from a core laboratory (UNIT 6.9), the clone can be analyzed as described herein. As depicted in Figure 6.9.1, methods for analysis involve growing and storing YAC-containing yeast strains and purifying YAC DNA in a form suitable for assessing the size of the artificial chromosome and for conventional Southern blotting. Preparation of yeast chromosomes in agarose plugs for subsequent analysis by pulsed-field gel electrophoresis is also described. Additional protocols are provided for recovering DNA fragments from the ends of a YAC genomic insert to be used as probes for detecting chimerism and for chromosome walking. Finally, preparation of high-molecular-weight YAC DNA is described and a general method for subcloning YAC inserts into cosmid or λ vectors for higher-resolution analysis is provided. NOTE: All solutions, media, glassware, and plasticware coming into contact with yeast or bacterial cells must be sterile, and sterile techniques should be followed throughout.

BASIC PROTOCOL

PROPAGATION AND STORAGE OF YAC-CONTAINING YEAST STRAINS YACs prepared using the pYAC4 vector (Figs. 6.10.1 and 13.4.6; pYAC4 contains an EcoRI site within the SUP4 gene in addition to the SnaBl site found in pYAC3, but is otherwise identical to pYAC3, carrying selectable markers TRP1 and URA3) and the S. cerevisiae host strain AB1380 (trp1−, ura3−, ade2-1) are grown on AHC plates. They can be stored short-term on AHC plates or stored long-term (after growth in YPD medium) in YPD containing glycerol at −80°C.

EcoRI

Hinfl Clal

CEN4

SUP4

EcoRl Hin fl

genomic insert

Sal l

URA3

SUP4

1 2

3 4

Analysis of Isolated YAC Clones


1

HYAC-C

2

LS-2

3

RA-2

4

HYAC-D

Figure 6.10.1 Structure of a representative pYAC4 clone at the vector/insert junction. Open boxes represent portions of the pYAC4 vector derived from yeast sequences (CEN4, the two halves of the SUP4 element, and URA3). Thin lines represent sequences derived from pBR322 and the bold line represents the YAC genomic insert fragment. Sites of annealing of the HYAC-C, LS-2, RA-2 and HYAC-D oligonucleotides are indicated by arrows 1, 2, 3, and 4, respectively. The EcoRI cloning site, the ClaI and Sal I sites (used for the end-fragment subcloning alternate protocol), and the HinfI sites that are utilized in the bubble linker end-fragment isolation protocol are indicated. Contributed by David D. Chaplin and Bernard H. Brownstein Current Protocols in Molecular Biology (1992) 6.10.1-6.10.19 Copyright © 2000 by John Wiley & Sons, Inc.

Materials S. cerevisiae strain AB1380 containing pYAC4 with insert (from core facility; UNIT 6.9) AHC plates (ura−, trp−) YPD medium (UNIT 13.1) 80% (v/v) glycerol in YPD medium 30°C orbital shaking incubator (e.g., New Brunswick Scientific #G-24) Cryovials Additional reagents for preparation of yeast media (UNIT 13.1) and growth and manipulation of yeast (UNIT 13.2) 1. Streak strain AB1380 containing pYAC4 with insert onto AHC plates. AHC medium selects for the presence of both arms of the YAC vector and thereby favors high stability of the YAC through successive passages.

2. Invert plate and incubate at 30°C until colonies are 1 to 3 mm in size. The AB1380 strain carries the ade2-1 ochre mutation, a block in the purine biosynthetic pathway that leads to accumulation of red-hued intermediates. Because a genomic insert in the YAC interrupts the SUP4 (ochre) gene in the YAC vector, colonies will have a red pigmentation.

3a. For short-term storage: Seal plates with Parafilm and store at 4°C for 4 to 6 weeks. 3b. For long-term storage: Inoculate an individual colony into 3.2 ml YPD medium and shake overnight at 30°C. Add 1 ml of 80% glycerol in YPD medium, mix thoroughly, and transfer in 0.2- to 1.0-ml aliquots to cryovials. Store at −80°C. YPD is a nonselective medium used to favor rapid growth and high cell viability. Strains stored in this fashion are stable for ≥5 years. Before strains are used in an experiment, they should first be grown on selective medium (e.g., AHC plates) to avoid recovery of a contaminant clone or one that has lost its YAC.

PREPARATION OF YAC-CONTAINING DNA FROM YEAST CLONES FOR ANALYSIS BY SOUTHERN BLOTTING

BASIC PROTOCOL

Procedures used by core laboratories for isolating an individual clone from a YAC library ensure that the purified YAC supports amplification of an appropriately sized PCR product using the screening primer pair. However, it is best to confirm the identity of the clone by hybridization analysis. Various methods can be used to prepare DNA suitable for Southern blot analysis using frequently cutting restriction enzymes. This protocol yields substantial quantities of DNA in the size range of 50 to 200 kb; it involves growing and lysing a single red colony containing pYAC4 with the insert DNA, then obtaining the DNA from the supernatant after centrifugation and analyzing by Southern blotting. Yeast chromosomes prepared in agarose plugs or very-high-molecular-weight DNA prepared in solution (third and fifth basic protocols) may also be used. Materials Single colony of S. cerevisiae AB1380 containing pYAC4 with insert (first basic protocol) AHC medium (ura−, trp−) SCE buffer SCEM buffer 50 mM Tris⋅Cl (pH 7.6)/20 mM EDTA (Tris/EDTA lysis buffer)



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10% (w/v) sodium dodecyl sulfate (SDS) 5 M potassium acetate, pH 4.8, ice-cold (UNIT 1.6) 95% ethanol, room temperature TE buffer, pH 8.0 (APPENDIX 2) 1 mg/ml DNase-free RNase A (UNIT 3.13) Isopropanol, room temperature 5 M NaCl Total genomic DNA of the species or individual from which the library was made (e.g., UNITS 2.2, 2.3 & 5.3) Appropriate single-copy probe designed to hybridize with the YAC insert (see UNITS 2.9 & 6.9) Orbital shaker (e.g., New Brunswick Scientific #G-24) 50-ml conical plastic centrifuge tubes Beckman JS-4.2 rotor or equivalent Additional reagents and equipment for digestion of DNA with restriction endonucleases (UNIT 3.1), Southern blotting and hybridization (UNIT 2.9), and pulsed-field gel electrophoresis (UNIT 2.5B) Culture and lyse cells from YAC clone 1. Inoculate a single red colony of a YAC-containing clone into 20 ml AHC medium in a 250-ml Erlenmeyer flask. Shake 24 hr at 250 rpm, 30°C, on an orbital shaker. The culture should begin to turn pink. If not, continue incubation an additional 24 hr. If culture is still not pink, discard and start over with a new red colony. Orbital shakers are preferred because they give much better aeration.

2. Inoculate 1 ml of culture from step 1 into 100 ml AHC medium in a 1-liter Erlenmeyer flask. Shake 24 hr at 250 rpm, 30°C. 3. Transfer culture to 50-ml plastic conical centrifuge tubes. Centrifuge 5 min at 2000 × g (2800 rpm in Beckman JS-4.2 rotor), 4°C. 4. Discard supernatants and resuspend cell pellets in a total of 5 ml SCE buffer. Pool into a single tube. 5. Add 1 ml SCEM buffer. Mix gently 1 to 2 hr at 100 rpm, 37°C, on an orbital shaker. SCEM buffer contains lyticase, which will digest the cell wall.

6. Centrifuge 5 min at 2000 × g, 4°C. Discard supernatant and resuspend cell pellet in 5 ml Tris/EDTA lysis buffer. 7. Add 0.5 ml of 10% SDS and invert several times to mix. Incubate 20 min at 65°C. Isolate nucleic acids 8. Add 2 ml of ice-cold 5 M potassium acetate, pH 4.8, and invert to mix. Keep 60 min on ice. 9. Centrifuge 10 min at 2000 × g, room temperature. Carefully pour nucleic acid–containing supernatant into a new tube. Add 2 vol room-temperature 95% ethanol and invert to mix.


10. Centrifuge 5 min at 2000 × g, room temperature. Discard supernatant and air-dry nucleic acid pellet 10 to 15 min. Add 3 ml TE buffer, pH 8.0, and dissolve overnight at 37°C.



Recover and analyze DNA 11. Add 0.1 ml of 1 mg/ml DNase-free RNase A and incubate 1 hr at 37°C. 12. Add 6 ml room-temperature isopropanol with swirling, then invert to mix. 13. Spool DNA using a capillary pipet and dissolve in 0.5 ml TE buffer, pH 8.0. Add 50 µl of 5 M NaCl and 2 ml of room-temperature 95% ethanol. Mix by inverting. 14. Spool DNA again and dissolve in 0.5 ml TE buffer. Store at 4°C. A yield of 1 to 1.5 ìg DNA/108 yeast cells can be expected.

15. Analyze 2-µg aliquots of YAC DNA and 15-µg aliquots of total genomic DNA from the species or individual from which the YAC library was made by digesting with several frequently cutting restriction enzymes. Proceed with Southern blotting and hybridization using a single-copy probe. The product amplified by PCR screening (see UNIT 6.9) may be used as probe. Because the YAC donor may exhibit a restriction-fragment-length polymorphism for this probe, two restriction fragments may be observed in the donor DNA. One of these fragments should be identified in the isolated YAC DNA.

16. Once the YAC clone has been verified by Southern blotting, determine its size and obtain a preliminary assessment of its stability by preparing chromosomes in agarose plugs (third basic protocol) and analyzing by pulsed-field gel electrophoresis. PREPARATION OF YEAST CHROMOSOMES IN AGAROSE PLUGS FOR PULSED-FIELD GEL ELECTROPHORESIS

BASIC PROTOCOL

In order to assess size, stability, and possible rearrangements within YACs, and to identify overlapping YACs, it is useful to isolate the YACs by embedding them in agarose plugs for subsequent analysis by pulsed-field gel electrophoresis (PFGE). Most methods of pulsed-field gel electrophoresis can be used (UNIT 2.5B); the CHEF (contour-clamped homogeneous electric-field electrophoresis) gel system is particularly suitable in that it reliably permits excellent resolution in the size range most common for YACs. Materials AHC medium (ura−, trp−) Single colony of S. cerevisiae containing pYAC4 with insert (first basic protocol) 0.05 M EDTA, pH 8.0 (APPENDIX 2) SEM buffer 10 mg/ml Lyticase (Sigma #L-8137 or ICN Biomedicals #190123) 2% InCert or SeaPlaque agarose (FMC Bioproducts), dissolved in SEM buffer and equilibrated to 37°C SEMT buffer Lithium lysis solution 20% (v/v) NDS solution 0.5× TBE (APPENDIX 2) or GTBE buffer (UNIT 2.5B) 30°C rotary platform shaking incubator Beckman JS-4.2 rotor or equivalent Gel sample molds (e.g., CHEF gel molds, Bio-Rad #1703622) 60-mm tissue culture plate Additional reagents and equipment for pulsed-field gel electrophoresis (UNIT 2.5B) Screening of Recombinant DNA Libraries


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Prepare and lyse YAC clone 1. Inoculate 25 ml AHC medium with a single red colony of a YAC-containing clone. Shake 48 to 60 hr at 250 rpm, 30°C. The culture should be pink. If it is not, discard and start over with a new red colony. To assess the stability of an individual YAC and to facilitate distinguishing of the artificial chromosome from the native yeast chromosomes, it is useful to analyze 4 or 5 individual colonies from the same YAC strain as well as a colony of the untransformed yeast host.

2. Centrifuge 10 min at 2000 × g (2800 rpm in a Beckman JS-4.2 rotor), 4°C. Discard supernatant and resuspend cell pellet in 10 ml of 0.05 M EDTA, pH 8.0. 3. Centrifuge 10 min at 2000 × g, 4°C. Remove all liquid from pellet and resuspend in 150 µl SEM buffer. Prepare agarose molds 4. Warm YAC sample to 37°C and add 25 µl Lyticase. Add 250 µl of 2% InCert or SeaPlaque agarose that has been melted in SEM buffer and equilibrated to 37°C. 5. Mix quickly and pour into CHEF gel sample molds. Chill 10 min at 4°C. Transfer solidified plugs to a 60-mm tissue culture plate. 6. Cover each plug with 4 ml SEMT buffer. Incubate 2 hr with gentle shaking at 37°C. 7. With a pipet, remove SEMT buffer and replace with 4 ml lithium lysis solution. Incubate 1 hr with gentle shaking at 37°C. 8. Remove and replace lithium lysis solution two or three times, shaking ≥1 hr each time. Shake the last change overnight. 9. Remove lithium lysis solution, replace with 4 ml of 20% NDS solution, and shake 2 hr at room temperature. Repeat once. Electrophorese samples in individual agarose plugs 10. Cut into plugs of suitable size to fit into wells of a pulsed-field gel. Store plugs individually in 20% NDS solution at 4°C. Plugs prepared and stored in this manner are usually stable for 4 to 8 weeks.

11. Soak each plug 30 min in 1 ml of 0.5× TBE or GTBE buffer. Change three times. 12. Analyze by pulsed-field gel electrophoresis. Following electrophoresis, the artificial chromosome can be visualized in an ethidium bromide–stained gel as an extra chromosome not present in the host yeast strain. If desired, Southern blot hybridization (UNIT 2.9) with appropriate probes can be carried out.




END-FRAGMENT ANALYSIS USING PCR AMPLIFICATION This protocol provides a means for recovering end fragments from the YAC insert using PCR amplification of end fragments. Digestion of YAC-containing DNA with a frequently cutting restriction enzyme produces a collection of small fragments: among these, one contains the distal portion of the YAC insert associated with part of the left vector arm, and another contains the other end of the insert associated with part of the right vector arm. Fragments encoding these vector sequences are prepared for PCR amplification by ligation of a double-stranded DNA tag containing a “bubble” of noncomplementary sequence flanked by short complementary sequences (Fig. 6.10.2). Selective amplification of these two end-fragment sequences is achieved using one PCR primer derived from the YAC vector (HYAC-C or LS-2 for the left arm or HYAC-D or RY-2 for the right arm) and one primer containing the sequence of the noncomplementary portion of the bubble (the 224 primer template, created by extension from the YAC-vector-specific primer; Fig. 6.10.3). Occasionally, nonspecific DNA fragments are amplified from the bubble PCR reaction. If this occurs, specificity may be restored using a hemi-nesting strategy. A small aliquot of the product of the initial PCR reaction (containing a mixture of the specific and nonspecific amplified fragments) is reamplified in a second round of PCR using an internal sequence from the vector arm as one of the primers. Because this sequence is not present in the nonspecific fragments, only the specific fragment will be amplified.

BASIC PROTOCOL

Materials “Bubble-top” and “bubble-bottom” oligonucleotide primers (Fig. 6.10.2) YAC-containing DNA (second basic protocol) RsaI and HinfI restriction endonucleases and appropriate buffers (UNIT 3.1) 10× T4 DNA ligase buffer and 1 U/µl T4 DNA ligase (UNITS 3.4 & 3.14) PCR reaction mix PCR amplification primers HYAC-C, HYAC-D, 224, and RA-2, 4 µM each (Fig. 6.10.2) Thermal cycling apparatus 65° and 68°C water baths Additional reagents and equipment for phosphorylating synthetic oligonucleotides (UNIT 3.10), restriction endonuclease digestion (UNIT 3.1), PCR (UNIT 15.1), nondenaturing PAGE (UNIT 2.7), preparing radiolabeled oligonucleotide probes (UNITS 3.10, 4.6 & 15.2), and blunt-end ligation (UNIT 3.16) Prepare bubble oligonucleotide tags 1. Phosphorylate the bubble-top oligonucleotide. This step can usually be eliminated, but may modestly increase efficiency.

2. Adjust bubble-top and bubble-bottom oligonucleotide concentrations to 4 nmol/ml with water. Mix together 1 nmol of each, then anneal by heating 15 min at 68°C in a water bath, followed by slow cooling to room temperature over 30 to 60 min. Digest YAC DNA and ligate to bubble oligonucleotides 3. Digest 2.5-µg aliquots of purified YAC-containing DNA to completion with RsaI or HinfI in 20 µl final volume, 37°C. Digestion of separate samples with RsaI and HinfI increases the chance of obtaining an end fragment of a size suitable for PCR amplification (75 bp).

4. Heat samples 15 min at 65°C to inactivate the restriction enzymes.



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A GCTGTCTGTCGAAGGTAAGGAACGGACGA

5 ′ - GAAGGAGAGGAC

Rsal bubble-top

GAGAAGGGAGAG - 3 ′

3 ′- CTTC C T C T CC TG CTCT T C C C TC TC - 5 ′ TCGCTAAGAGCATGCTTGCCAATGCTAAG

Universal bubble-bottom

* * * * * * * * * * * * * * * * * * * * * * * *

224 primer

3 ′- TCGCTAAGAGCATGCTTGCCAATGCTAAGC- 5 ′

B GCTGTCTGTCGAAGGTAAGGAACGGACGA

GAGAAGGGAGAG - 3 ′ Hin f l bubble-top

5 ′- A (N)TGAAGGAGAGGAC

3 ′ - C T T C C T C T CC T G

CTCT T C C C TC TC - 5 ′ Universal bubble-bottom TCGCTAAGAGCATGCTTGCCAATGCTAAG

C HYAC - C primer HYAC - D primer RA - 2 primer LS - 2 primer Bubble sequencing primer

5 ′- GCTACTTGGAGCCACTATCGACTACGCGAT- 3 ′ 5 ′- GGTGATGTCGGCGATATAGGCGCCAGCAAC- 3 ′ 5 ′- TCGAACGCCCGATCTCAAGATTAC- 3 ′ 5 ′- TCTCGGTAGCCAAGTTGGTTTAAGG- 3 ′ 5 ′- CGCTGTCCTCTCCTTC - 3 ′

Figure 6.10.2 Oligonucleotides for amplification and sequencing of YAC insert end-fragments. (A) Annealing of the 53-mer universal “bubble-bottom” oligonucleotide to the 53-mer RsaI bubble oligonucleotide yields a blunt-ended DNA duplex in which 12-bp complementary sequences flank a 29-nucleotide “bubble” of noncomplementary sequence. This bubble linker can be ligated to any blunt-ended fragment (e.g., one generated by digestion with RsaI). The 224 primer does not anneal to either strand of the bubble, but is fully complementary to any DNA strand that is generated during PCR using the universal bubble-bottom strand as a template (see Fig. 6.10.3). (B) Annealing of the 53-mer universal bubble-bottom oligonucleotide to the 56-mer HinfI bubble-top oligonucleotide yields a DNA duplex with one blunt end and one cohesive end with the degenerate HinfI site. A mixture of all four nucleotides at a specific position is indicated by (N). (C) The HYAC-C, HYAC-D, RA-2, and LS-2 primers anneal to sequences in the pYAC4 vector (see Fig. 6.10.1). The bubble sequencing primer anneals to the RsaI and HinfI bubble-top sequences near their 5′ ends, permitting DNA sequencing from the bubble linker back into the YAC insert end-fragment.

5. Prepare the following ligation mix (50 µl total): 2 µl (250 ng) digested DNA 1 µl (2 pmol) annealed bubble oligonucleotides (from step 2) 5 µl 10× ligase buffer 2 µl (2 U) T4 DNA ligase 40 µl H2O. Incubate 2 hr at 37°C or overnight at room temperature.


The blunt-ended bubble composed of the universal bubble-bottom oligonucleotide and the RsaI bubble-top oligonucleotide should be used with RsaI-digested YAC DNA. Likewise, the HinfI cohesive bubble composed of the universal bubble-bottom oligo-



A random insert fragment

5′ 3′

B

3′ 5′

vector primer site insert end-fragment

5′ 3′

3′ 5′

YAC vector

1st cycle of PCR

3′

5′

insert end-fragment

3′ 5′

224 primer site

Figure 6.10.3 Selective PCR amplification from the YAC insert end-fragment. (A) Result of ligation of the bubble linker to a random fragment from the internal portion of the YAC insert. Because this fragment is not derived from the end of the YAC genomic insert, it contains no sequences from the YAC vector and has no site for annealing any of the HYAC-C, HYAC-D, RA-2, or LS-2 primers or for annealing of the 224 primer. Consequently, no fragment is amplified by PCR. (B) Result of ligation of the bubble linker to a fragment derived from the end of the YAC genomic insert and containing its associated YAC vector sequences. During the first cycle of PCR, extension from the YAC vector priming site produces sequences complementary to the universal bubble-bottom primer. This extended fragment provides a template for annealing of the 224 primer, thus permitting successful amplification of the insert end-fragment.

nucleotide and the HinfI bubble-top oligonucleotide should be used with HinfI-digested YAC DNA.

6. Add 200 µl water to bring the DNA concentration to 1 ng/µl final. Amplify fragments containing YAC-insert end sequences 7. Prepare the following PCR on ice (10 µl total): 8 µl PCR reaction mix 1 µl (2 µM each) PCR primer pair mix 1 µl (1 ng) digested, bubble-ligated YAC DNA. Carry out 35 cycles of amplification as follows: 1 min at 92°C, 2 min at 65°C, and 2 min at 72°C. To amplify left end of YAC insert, use primer pair mix made from equal amounts of primers 224 and HYAC-C (Fig. 6.10.1). To amplify right end of YAC insert, use primer pair mix made from equal amounts of primers 224 and RA-2 (specific for the SUP4 region of pYAC4; Fig. 6.10.1).



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These parameters were optimized using the Perkin-Elmer TC1 thermal cycler. If another instrument is used, some adjustment of parameters may be required.

8. Analyze a 1-µl aliquot of PCR product on a 5% polyacrylamide gel. A single, clearly visible amplified fragment should be observed after staining the gel with ethidium bromide. Produce end fragments using hemi-nested amplification To amplify the left end (either RsaI- or HinfI-digested DNA): 9a. Amplify 1 µl digested, bubble-ligated YAC DNA (from step 6) with primers 224 and HYAC-C, using 20 cycles of 1 min at 92°C, 2 min at 62°C, and 2 min at 72°C. 10a. Dilute the amplification product 1:100 with water and add 1 µl to a new PCR reaction containing primers 224 and LS-2 (specific for the SUP4 region of pYAC4; see Figs. 6.10.1 and 13.4.6). Carry out 30 cycles of 1 min at 92°C, 2 min at 65°C, and 2 min at 72°C. To amplify the right end using RsaI-digested DNA: 9b. Amplify 1 µl digested, bubble-ligated YAC DNA (from step 6) with primers 224 and HYAC-D, using 20 cycles of 1 min at 92°C, 2 min at 62°C, and 2 min at 72°C. 10b. Dilute amplification product 1:100 with water and add 1 µl to a new PCR reaction (see step 7) containing primers 224 and RA-2. Carry out 30 cycles of 1 min at 92°C, 2 min at 65°C, and 2 min at 72°C. Hemi-nesting of the right end cannot be performed with HinfI-digested DNA, because there is a HinfI site only 24 bp from the EcoRI YAC vector cloning site.

11. Analyze a 1-µl aliquot of each final PCR reaction on a 5% polyacrylamide gel. 12. End label amplified fragments with 32P and use as hybridization probes or for nucleotide sequencing to produce an end-specific STS. Alternatively, subclone by blunt-end ligation to a plasmid vector prior to further manipulation. Blunt-end subcloning of DNA fragments that have been amplified by PCR must be preceded by “polishing” of the ragged PCR ends with S1 nuclease or T4 DNA polymerase (see UNIT 15.7). ALTERNATE PROTOCOL

END-FRAGMENT ANALYSIS BY SUBCLONING INTO A BACTERIAL PLASMID VECTOR This method (an alternative to the previous protocol for recovering end fragments from the YAC insert) uses the strategy of double digesting the YAC-containing DNA to enhance the efficiency of subcloning into a pUC19-based vector; the SUP4 element containing the YAC EcoRI cloning site is located within a portion of the YAC vector derived from pBR322 (Figs. 1.5.2 and 6.10.5). The cloning strategy enriches for end-fragment-containing subclones because the restriction endonuclease used for digestion of YAC-containing DNA is either ClaI (for the left arm) or SalI (for the right arm). Both enzymes cut rarely in human or yeast genomic DNA; therefore, when one is combined with a more frequently cutting enzyme, the resulting doubly digested fragments will represent a minor portion of the total DNA pool. Size fractionation on an agarose gel prior to subcloning affords a still further enrichment for end fragments. Note that after the first step of the protocol, all steps are performed in duplicate to identify both the right and left end fragments of the YAC insert.




Additional Materials ClaI, SalI, and other appropriate restriction endonucleases and digestion buffers (UNIT 3.1) Left- and right-vector-arm probes (Fig. 6.10.4) pUC19-ES and pUC19-HS plasmid vectors (support protocol and Fig. 6.10.5) Transformation-competent Rec− strain of E. coli (e.g., DH5; Table 1.4.5) 2× TY or LB agar plates (UNIT 1.1) containing 50 to 100 µg/ml ampicillin Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5A), subcloning of DNA fragments (UNIT 3.16), transformation of E. coli (UNIT 1.8), Southern blotting and hybridization (UNIT 2.9), labeling by random-primed synthesis (UNIT 3.5), isolation and purification of DNA fragments from agarose gels (UNIT 2.6), replica plating (UNIT 1.3), and purification of plasmid DNA (UNITS 1.6 & 1.7) Perform an appropriate double digest and analyze by hybridization 1a. For left arm: Digest 5-µg aliquots of YAC-containing DNA with ClaI and then with each possible second cloning enzyme—SacI, KpnI, SmaI, BamHI, XbaI, and SphI. In addition to SmaI, other blunt-cutters not represented within the ClaI-EcoRI interval of the YAC vector may be tested.

1b. For right arm: Digest 5-µg aliquots of YAC-containing DNA with SalI and then with each of the following possible second cloning enzymes: SacI, KpnI, SmaI, BamHI, XbaI, SphI, or HindIII. All of the above restriction endonucleases have compatible cleavage sites within the polylinker of the modified pUC19; AccI provides a cohesive site for ClaI.

Carry out all remaining steps in parallel for the left- and right-arm probes: 2. Electrophorese doubly digested DNA on an agarose gel and transfer to a filter for Southern hybridization. 3. Prepare left- and right-vector-arm probes by PCR as described in Fig. 6.10.4. Probes can also be obtained by digestion and fractionation of pBR322 DNA with subsequent labeling.

4. Hybridize each probe to the appropriate filter from step 2.

Left arm:

5′ ATCGATAAGCTTTAATGCGGTAGT 3′ (pBR322 bases 23-46) 5′ GATCCACAGGACGGGTGTGGTCGC 3′ (pBR322 bases 379-356)

Right arm: 5′ GATCCTCTACGCCGGACGCATCGT 3′ (pBR322 bases 375-399) 5′ GTCGACGCTCTCCCTTATGCGACT 3′ (pBR322 bases 656-632)

Figure 6.10.4 Generation of left- and right-vector-arm probes. The 351-bp ClaI-BamHI and 276-bp BamHI-SalI fragments of pBR322, which hybridize to sequences immediately flanking the sup4 sequences of the YAC vector, are appropriate probes for the YAC left and right vector arms. These probes can be obtained by restriction digestion and gel fractionation of pBR322 plasmid DNA or generated by PCR using 10 ng pBR322 as template for the primers illustrated here. Perform PCR using 25 cycles of 1 min at 92°C, 1 min at 50°C, and 2 min at 72°C. Extract the amplified material once with phenol and once with chloroform, then precipitate with ethanol (UNIT 2.1). Label directly by random priming (UNIT 3.5) without further purification.



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5. Examine autoradiogram and choose an enzyme combination that yields a hybridizing DNA fragment in the 2- to 7-kb size range. Digest a 50-µg aliquot of YAC-containing DNA with these two enzymes. This should yield ∼5 times more size-fractionated DNA than needed.

Isolate the DNA 6. Electrophorese doubly digested DNA on an agarose gel. Using a scalpel or razor blade, cut out the segment of gel that should contain the doubly digested DNA fragment. To avoid missing the critical portion of the gel, it may be useful to excise adjacent gel slices containing fragments larger than and smaller than the expected size, and to process them in parallel.

7. Purify size-fractionated DNA from gel slice and resuspend in a final volume of 20 µl TE buffer, pH 8.0. For purifying the DNA, the best results have been obtained by using the Geneclean II kit (BIO 101, La Jolla, CA).

Subclone the end fragments 8. Ligate 20% of the purified YAC-derived insert DNA with 0.2 µg of gel-purified, compatibly digested pUC19-HS or -ES vector DNA overnight in a total volume of 20 µl. Because the pUC19 plasmid from which they are derived has no homology with the portion of pBR322 detected by the ClaI/BamHI and BamHI/SalI probes (Fig. 6.10.4), these probes can be used to detect YAC-insert end-fragment-containing subclones in pUC19, and will not cross-hybridize to the pUC vector.

9. Transform the ligated DNA into a transformation-competent Rec− host strain of E. coli. Plate sufficient transformation mix on 2× TY/ampicillin or LB/ampicillin plates to obtain ∼200 colonies, a sufficiently low density that individual colonies may be recovered following hybridization. Invert plates and incubate overnight at 37°C. 10. Prepare colony-lift filters and hybridize overnight with ∼1–2 × 107 cpm of appropriate 32 P-labeled left- or right-arm probes. Wash and autoradiograph. Because of the enrichment afforded by double digestion, 1% to 4% of colonies will contain the end fragment.

11. Purify plasmid DNA from hybridizing colonies. 12. Verify the structure of the plasmid by comparing its restriction map to the data obtained during the initial analytical double digests of the YAC (steps 1 to 3). SUPPORT PROTOCOL

DESIGN AND PREPARATION OF pUC19-ES and pUC19-HS SUBCLONING VECTOR


This protocol describes the construction of two vectors for subcloning YACs (previous basic protocol). pUC19 is modified by insertion of a “stuffer” fragment in both possible orientations (see UNIT 3.16; Fig. 6.10.5). If a double digest is performed on the resulting construct (UNIT 3.1), using AccI (cohesive with ClaI) or SalI and any of the other enzymes in the pUC polylinker, the presence of the stuffer makes it possible to visualize whether the vector has been fully cut. Complete double digestion is critical to the success of the end-fragment subcloning described in the previous protocol. For example, digestion with AccI or SalI will linearize the pUC19-ES vector. Subsequent digestion with EcoRI, SacI, KpnI, SmaI, BamHI, or XbaI will result in a shift in vector size from 3161 bp to 2686 bp



EcoRI Sacl Kpnl Smal BamHI Xbal

ampr

pUC19-ES 3161 bp

475-bp stuffer

SaII / AccI / HincII Ps tI SphI Hin dIII EcoRI Sacl Kpnl Smal BamHI Xbal SaI I / AccI / Hin cII

ori

ampr

pUC19-HS 3161 bp

ori

475-bp stuffer

Ps tI SphI Hin dIII

Figure 6.10.5 Structure of the pUC19-ES and pUC19-HS plasmids.

and free stuffer fragment will be generated. The doubly digested vector can then be isolated by fractionation in an agarose gel (UNIT 2.5A) and purified (UNIT 2.6). pUC19-ES: Modify the pUC19 (see Fig. 1.5.2) vector by inserting a stuffer consisting of 475-bp TaqI fragment of pBR322 (positions 653-1128) into the pUC19 polylinker AccI (HincII) site. In the resulting plasmid, the AccI (and SalI and HincII) site adjacent to the polylinker PstI site is preserved, but the AccI site previously found next to the polylinker XbaI site (which would now be at the other end of the stuffer) is lost (Fig. 6.10.5). pUC19-HS: Insert the 475-bp TaqI fragment stuffer described above into the same pUC19 AccI site but in the opposite orientation. In the resulting plasmid, the polylinker AccI site adjacent to the XbaI site is preserved, but the AccI site adjacent to the PstI site is lost (Fig. 6.10.5).



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BASIC PROTOCOL

PREPARATION OF HIGH-MOLECULAR-WEIGHT YAC-CONTAINING YEAST DNA IN SOLUTION This protocol describes the purification of YAC-containing DNA of sufficiently high molecular weight to provide a source of YAC insert material for subcloning in λ or cosmid vectors. This DNA is also suitable for restriction mapping or other genetic manipulations. A cell lysate is fractionated on a sucrose gradient; the DNA-containing fraction is subsequently dialyzed, concentrated, and examined by electrophoresis through a pulsedfield gel. Materials Single colony of S. cerevisiae containing pYAC4 with insert (first basic protocol) AHC medium (ura−, trp−) SCEM buffer Lysis buffer Step-gradient solutions: 50%, 20%, and 15% (w/v) sucrose TE buffer, pH 8.0 (APPENDIX 2) Dry granular sucrose 30°C orbital shaking incubator (e.g., New Brunswick Scientific #G-24) 250-ml conical centrifuge bottles (e.g., Corning #25350) 65°C water bath 25 × 89–mm tube (e.g., Beckman #344058) Beckman JS-4.2 and SW-27 rotors (or equivalents) Dialysis tubing (APPENDIX 3) Pyrex baking dish CHEF pulsed-field gel apparatus or equivalent (UNIT 2.5B) Additional reagents and equipment for size fractionation using a sucrose gradient (UNIT 5.3) and estimating DNA concentration (UNIT 2.6) Grow and prepare the cells 1. Inoculate a single red colony of a YAC-containing clone into 25 ml AHC medium in a 250-ml flask. Shake at 250 rpm, 30°C, until culture reaches saturation (∼3 days). 2. Transfer 1 ml of saturated culture to 100 ml AHC medium in a 1-liter flask. Shake 16 to 18 hr at 250 rpm, 30°C. 3. Harvest yeast cells by centrifuging 10 min at 2000 × g (2800 rpm in Beckman JS-4.2 rotor), room temperature, using a 250-ml conical centrifuge bottle. Discard supernatant. 4. Resuspend cells in 50 ml water. Centrifuge 5 min at 2000 × g, room temperature. Discard supernatant. A cell pellet of ∼4 g should be obtained.

5. Resuspend cells in 3.5 ml SCEM buffer. Lyse the cells 6. Incubate 2 hr at 37°C with occasional gentle mixing. The mixture will become highly viscous. 7. Gradually add cell mixture to 7 ml lysis buffer in a 250-ml Erlenmeyer flask by allowing viscous cell suspension to slide down side of flask. Analysis of Isolated YAC Clones

8. Gently mix by swirling flask until mixture is homogeneous and relatively clear. 9. Incubate 15 min at 65°C, then cool rapidly to room temperature in a water bath.



Fractionate cell contents 10. Fractionate on a sucrose step gradient. In a 25 × 89–mm tube, prepare a step gradient consisting of: 3 ml 50% sucrose 12 ml 20% sucrose 12 ml 15% sucrose 11 ml lysed sample. Centrifuge 3 hr at 125,000 × g (26,000 rpm in a Beckman SW-27 rotor), room temperature. 11. Discard ∼25 ml from top of gradient using a 10-ml pipet. Dialyze and analyze DNA 12. Collect viscous DNA-containing solution at the 20% to 50% sucrose interface (∼5 ml total volume) and place in dialysis tubing, leaving room for volume to increase ≥2- to 3-fold. Dialyze overnight against 2 liters TE buffer, pH 8.0, at 4°C. 13. Reconcentrate dialyzed DNA by placing dialysis tubing in an autoclaved Pyrex baking dish and covering with granular sucrose. Recover dialysis tubing when volume of contents has been reduced to ∼2 ml. 14. Squeeze DNA solution to one end of dialysis tubing and tie an additional knot to keep DNA in a small volume. Dialyze overnight against 1 liter of TE buffer, pH 8.0, at 4°C. 15. Recover dialyzed DNA and check a small aliquot by electrophoresing in a CHEF pulsed-field gel. Stain with ethidium bromide and estimate DNA content by comparison to a known amount of λ DNA. The DNA sample will contain a substantial amount of yeast RNA but should also contain a population of YAC DNA fragments migrating at a size of >100 kb. The presence of the RNA may make it difficult to determine the DNA concentration accurately; the concentration may be estimated by comparison to known DNA standards in an ethidium bromide– stained gel. The RNA will not affect restriction digestion of the DNA.

PREPARATION AND ANALYSIS OF A YAC-INSERT SUBLIBRARY Construction of a sublibrary of fragments of the YAC insert facilitates high-resolution analysis of the insert sequence. This protocol details the steps required to produce a cosmid library, followed by a series of screenings to identify regions of interest and “walking” to establish a contiguous map of the insert.

BASIC PROTOCOL

Materials High-molecular-weight YAC-containing DNA (fifth basic protocol) Vector DNA (e.g., SuperCos 1, Stratagene #251301) 32 P-labeled (UNIT 3.10) probes: total genomic DNA of the individual or species from which the library was made (e.g., UNITS 2.2, 2.3 & 5.3), end-specific DNA (UNIT 3.10) or RNA (UNIT 3.8), and end fragment from YAC (fourth basic protocol or alternate protocol) Additional reagents and equipment for restriction endonuclease digestion (UNIT 3.1), genomic DNA library production (UNIT 5.7), plating and transferrin a cosmid library (UNIT 6.2), and hybridization with radioactive probes (UNITS 6.3 & 6.4)



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Construct the library 1. Partially digest 1 to 2 µg of YAC-containing DNA with restriction endonuclease(s) appropriate for cosmid vector to be used. For example, to clone into the BamHI site of SuperCos 1, digest the YAC DNA with either MboI or Sau3A. The quantity of restriction endonuclease should be adjusted to produce digested fragments with an average size of ∼40 kb. Although only a small fraction of the YAC-containing DNA used as starting material is actual YAC DNA (the rest being yeast genomic DNA), because of the low complexity of the yeast genome (e.g., compared to the human genome), only 3000 to 5000 cosmid clones are required to yield 3 yeast genome equivalents. Thus, only 1 to 2 ìg of yeast DNA are required to make an adequate library.

2. Perform a series of test ligations as described in UNIT 5.7. Using optimal conditions, ligate insert DNA to vector DNA. 3. Package cosmid recombinants; dilute packaged extract and determine the titer. 4. Plate and transfer the sublibrary as appropriate for the vector, and prepare resulting filters for hybridization. Screen the sublibrary 5. Perform a preliminary screen of the library using a 32P-labeled probe of total genomic DNA of the individual or species from which the library was made. This probe is a source of repetitive sequences. Because these repetitive sequences are spaced frequently throughout the source genome, and are absent from yeast, this probe will identify most of the source-DNA insert cosmids from the excess of yeast insert cosmids.

6. Organize this first set of cosmid clones into contigs by analyzing shared restriction fragments and by hybridizing with probes contained in the YAC insert or prepared from the ends of individual cosmid inserts. Cosmid end-fragment-specific probes can be generated by digesting cosmid DNA and end-labeling the purified restriction fragment(s) that contain(s) the cloning site. If the cloning vector is SuperCos 1 (or a comparable vector), end-specific RNA probes may also be transcribed from the ends of the cosmid clones using T3 and T7 polymerase (see critical parameters).

7. Establish a complete contiguous collection of cosmid clones of the original YAC insert by screening the library with specific YAC-derived probes and cosmid end-specific probes. Note that nitrocellulose filters may be reused for hybridization in subsequent steps without further washing or removal of probe. Repeated hybridization with sequential “walking probes” should reveal new hybridizing colonies at each step.




REAGENTS AND SOLUTIONS AHC medium and plates (ura−, trp−) 1.7 g yeast nitrogen base without amino acids and without ammonium sulfate (Difco) 5 g ammonium sulfate 10 g casein hydrolysate-acid, salt-free and vitamin-free (U.S. Biochemical #12852) 50 ml (for medium) or 10 ml (for plates) of 2 mg/ml adenine hemisulfate (Sigma #A-9126) Dissolve in a final volume of 900 ml H2O Adjust pH to 5.8 Autoclave 30 min, then add 100 ml sterile 20% (w/v) glucose. For AHC plates, add 20 g agar prior to autoclaving. Store at 4°C for ≤6 weeks. Lithium lysis solution 1% lithium dodecyl sulfate (Sigma # L-4632) 100 mM EDTA 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2) Filter sterilize and store indefinitely at room temperature Lysis buffer 0.5 M Tris⋅Cl, pH 8.0 (APPENDIX 2) 3% (v/v) N-lauroylsarcosine (Sarkosyl) 0.2 M EDTA, pH 8.0 (APPENDIX 2) Store indefinitely at room temperature. Add 1 mg/ml proteinase K just before use. 100% NDS solution Mix 350 ml H2O, 93 g EDTA, and 0.6 g Tris base. Adjust pH to ∼8.0 with 100 to 200 pellets of solid NaOH. Add 5 g N-lauroylsarcosine (predissolved in 50 ml water) and adjust to pH 9.0 with concentrated NaOH. Bring volume to 500 ml with water. Filter sterilize and store indefinitely at 4°C. Dilute 1:5 with H2O (20% final) just before use. PCR reaction mix 1.5 mM MgCl2 50 mM KCl 10 mM Tris⋅Cl, pH 8.3 (APPENDIX 2) 0.2 mM each dATP, dCTP, dGTP, and dTTP 0.05 U AmpliTaq polymerase (Perkin-Elmer/Cetus)/µl reaction mixture 0.03 µl Perfect Match Enhancer (Stratagene)/µl reaction mixture Store all components at −20°C and mix just before use SCE buffer 0.9 M sorbitol (Fisher, molecular biology grade) 0.1 M sodium citrate 0.06 M EDTA, pH 8.0 Adjust pH to 7.0 Store at room temperature ≤3 months SCEM buffer 4.9 ml SCE buffer (see above) 0.1 ml 2-mercaptoethanol (2-ME) Add 1 to 2 mg Lyticase (Sigma #L-8137 or ICN Biomedicals #190123) just before use.



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SEM buffer 1 M sorbitol 20 mM EDTA, pH 8.0 14 mM 2-ME Filter sterilize Store at 4°C for ≤6 weeks SEMT buffer 1 M sorbitol 20 mM EDTA 14 mM 2-ME 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2) Filter sterilize Add 1 mg/ml Lyticase (Sigma #L-8137 or ICN Biomedicals #190123) just before use. COMMENTARY Background Information An overview of strategies for screening YAC libraries and analyzing YAC clones is presented in UNIT 6.9.

Critical Parameters and Troubleshooting


The protocols provided in this unit are intended to describe the analysis and characterization of particular YAC clones of interest. It is initially desirable to assure the integrity of the clone; that is, to ensure that the YAC indeed carries the proper insert, that the insert is not chimeric or rearranged, and that it is stably maintained and propagated in the yeast host. Growth of YAC-containing strains. In YAC clones, genomic DNA is inserted into a cloning site carried within the SUP4 gene of the vector (see Figs. 6.10.1 and 13.4.6). In the parent vector, the SUP4 product complements the ade2-1 ochre mutation carried in the host AB1380. This mutation causes a block in purine biosynthesis, resulting in accumulation of red pigment in the culture. Thus, disruption of SUP4 by insertional inactivation prevents complementation of the ochre mutation in the host. Before a strain is used, it is important to check that upon growth the colonies or cultures exhibit a red pigmentation. If not, another isolate should be used. Additionally, growth on selective AHC medium requires the presence of both arms of the YAC vector and favors stability of the clone through passage. Analysis of YAC DNA. Restriction analysis of purified YAC DNA (second basic protocol) can be used to assess YAC structure. If fragments of unanticipated sizes are detected in the YAC, then it is likely that the YAC contains

sequences homologous to, but different from, the desired clone, or that the YAC insert has undergone some sort of rearrangement during cloning. Alternatively, lack of methylation of the YAC DNA at the restriction enzyme recognition site may give a digestion pattern not seen in uncloned genomic DNA. Determination of size and stability of the YAC clone is made by preparing chromosomes in agarose plugs and subsequent pulsed-field gel electrophoresis. The PFGE gel can be blotted and analyzed by hybridization with sequence-specific probes and end-fragment probes. The results should reveal hybridization to the same size chromosome in all isolates of a given YAC clone. Variation in the size of the artificial chromosome between yeast isolates derived from one clone indicates YAC instability. Hybridization of a YAC vector arm probe to more than one artificial chromosome may indicate multiple transformation of the strain at the time of library construction. Alternatively, it may represent strain instability, with the smaller chromosome(s) representing deletion products of the original YAC. When two or more artificial chromosomes are identified with a single-copy genomic insert-specific probe, instability of the YAC insert is the most likely cause. Assessing chimerism. In most existing YAC libraries, chimeric clones are observed to represent from 5% to 50% of the total clones. One of the most reliable ways to identify a chimeric YAC insert is to isolate a small fragment from each end of the YAC insert and define its chromosome of origin and whether it is contained in overlapping YACs from the same chromosomal region. Two procedures are given for analyzing the end fragments of



the YAC clone, which can be used to identify chimerism in the YAC insert—one based upon subcloning (which for large-scale mapping projects can require prohibitive amounts of time and effort) and one based upon PCR. With the PCR strategy, the resulting amplified product should migrate as a single fragment in a polyacrylamide gel. If multiple bands are present, it may be possible to demonstrate that one is an appropriately amplified fragment because it should be digested by EcoRI to yield the vector-linker fragment plus the insert end fragment (see Fig. 6.10.3). If digestion by EcoRI cannot be confirmed or if no amplified band is observed, it is useful to try a “hemi-nested” PCR amplification in which the initially amplified product is reamplified using another primer that should be contained only in the properly amplified fragment. Each end-fragment probe should be shown to be single-copy by hybridization to a Southern blot of total DNA from the species used to prepare the YAC library. If a smear of hybridization is obtained, repetitive sequences are present within the probe. In the case of fragments obtained by subcloning, it is usually possible to identify a single-copy probe by digesting the fragment into several smaller pieces using selected restriction enzyme. The end fragments recovered by PCR are usually small, so that it is generally impossible to salvage a single-copy probe. It may be possible to suppress the repetitive DNA hybridization by including in the hybridization reaction an excess of unlabeled denatured repetitive DNA fragments from the species used to prepare the YAC library. If this is unsuccessful, the alternate protocol (subcloning into a bacterial vector) is usually necessary. Construction of a cosmid sublibrary. For further high-resolution analysis and mapping of the YAC insert, it is desirable to construct a sublibrary (final basic protocol) from the YACcontaining DNA (fifth basic protocol). A number of cosmid and phage vectors are available that are suitable for subcloning YACs into bacterial vectors (UNITS 3.16 & 5.7). One excellent candidate is the SuperCos-1 cosmid vector (Stratagene), which can accommodate inserts in the 35- to 42-kb range. It contains a neomycin-resistance cassette that permits selection of transfected clones in mammalian cells. It also contains T3 and T7 phage promoters flanking the genomic insert, which facilitate generation of RNA probes from the ends of the genomic inserts. This feature is useful for verifying overlaps of clones or to permit chromo-

some walking if a complete cosmid contig is not established in the first round of screening. Once a sublibrary has been constructed, it should be screened with a probe consisting of 32P-labeled total genomic DNA from the individual or species that was originally used to construct the library (a source of repetitive sequences). This will identify most of the cosmids containing DNA inserts from the source genome. Individual clones can be analyzed by Southern blotting with probes from the YAC insert or with genomic repetitive sequences. This data, together with results of Southern blots using probes derived from the ends of the cosmid clones using T3 or T7 polymerase, can be used to organize the cosmids into contigs. If a complete contig is not established, the cosmid library can be screened again with probes representing the ends of the YAC inserts, or derived from the ends of cosmid clones. Empirically, different portions of the YAC insert have been found to be nonrandomly represented in the cosmid library. Consequently, it is common for one or more rounds of chromosome walking to be required to fill in gaps between cosmid contigs.

Anticipated Results The basic protocol for preparation of DNA from YAC clones can be expected to yield ∼1 to 1.5 µg of DNA in the size range of 50 to 200 kb from 108 yeast cells. The yield of DNA obtained by purification using preparative CHEF gel electrophoresis is ≥10-fold lower. The basic protocol for preparation of yeast chromosomes in agarose plugs for PFGE should yield sufficient material for ∼40 lanes of a pulsed-field gel from a 25-ml yeast culture. The basic and alternate protocols for analysis of the YAC insert end fragments (by PCR and by subcloning into a plasmid vector) should each yield DNA fragments that identify single hybridizing bands in genomic DNA when used as probes for Southern blots. The basic protocol for preparation of high-molecular-weight YACcontaining yeast DNA in solution should yield ∼25 to 50 µg of DNA ≥100 kb in size from a 100-ml culture. Basic protocol for preparation of a YAC insert cosmid sublibrary should yield ∼1500 colonies/µg of starting yeast DNA. From these, ∼30 to 50 genomic DNA–containing cosmids should be recovered.

Time Considerations Purifying YAC DNA using the first basic protocol requires ∼3 days to grow the culture, 3.5 hr to isolate the DNA, and after an overnight



Supplement 20

resuspension, ∼2 hr to remove RNA and reprecipitate the DNA. Preparation of yeast chromosomes in agarose plugs takes ∼4 days to grow the yeast culture, ∼11⁄2 hr to form the yeast-containing plugs, ∼6 hr to lyse the yeast, an overnight incubation, and ∼6 hr to wash and prepare the plugs for electrophoresis. Preparation of YAC insert end fragments by PCR takes from 6 hr to overnight to anneal the bubble primers, digest the YAC DNA, and ligate it to the bubbles. Another 3 to 5 hr are needed to amplify the end fragment by PCR. Isolation of YAC end fragments by subcloning requires 2 to 3 days to perform the preliminary analytical Southern blot to identify the enzyme combination of choice for subcloning. Once this is identified, 1 day is required for preparative isolation of the doubly digested DNA fragments, followed by an overnight ligation to the modified pUC 19 vector. An additional day is needed to transform bacteria and grow colonies, and 1 to 2 days are required to identify the specific end-fragment

subclones by hybridization. Finally, 2 days are needed to purify the subcloned DNA and to verify its structure by restriction enzyme analysis. Preparation of high-molecular-weight YACcontaining DNA requires ∼4 days to grow the culture, 7 hr to lyse the cells and perform the sucrose-gradient fractionation, an overnight dialysis, and 2 to 3 hr to concentrate the DNA. Following an additional overnight dialysis, the DNA is ready for use. YAC insert cosmid sublibrary preparation and analysis takes 2 days to perform test digestions and ligations. Another 2 days are required to perform the preparative digestion, ligation, packaging, and plating of the library. Two more days are required for preparation of filters, hybridization, washing, and autoradiography. Contributed by David D. Chaplin and Bernard H. Brownstein Howard Hughes Medical Institute and Washington University School of Medicine St. Louis, Missouri




SPECIALIZED STRATEGIES FOR SCREENING LIBRARIES

SECTION VI

Use of Monoclonal Antibodies for Expression Cloning

UNIT 6.11

This unit details the use of transient expression in mammalian cells to screen cDNA libraries with monoclonal antibodies (MAb) to isolate cDNA clones encoding cell-surface and intracellular proteins. The first basic protocol describes the cloning of cDNAs encoding cell-surface antigens. Several steps in this protocol involve transfection procedures that are described in greater detail in UNIT 16.12. The second basic protocol is a modification that facilitates isolation of cDNAs encoding antigens that are expressed intracellularly. Both protocols are designed for use with the expression vector CDM8, which contains a polylinker for subcloning double-stranded cDNA (Fig. 16.12.1). ISOLATION OF cDNA CLONES ENCODING CELL-SURFACE ANTIGENS This protocol is designed to isolate cDNAs encoding cell-surface proteins by screening cDNA libraries transiently expressed in mammalian cells. The procedure requires multiple rounds of transfection and immunoselection and is divided into four sections: (1) COS cell transfection by the DEAE-dextran method, (2) immunoselection by panning, (3) plasmid recovery and E. coli transformation, and (4) COS cell transfection by the spheroplast fusion method. A total of four rounds of transfection and immunoselection (one using DEAE-dextran, three using spheroplast fusions; Fig. 6.11.1) are used. After the final round of immunoselection, plasmid DNA is prepared from individual bacterial colonies. COS cells are then transfected with this DNA by the DEAE-dextran method and examined for their ability to express the foreign protein of interest by immunofluorescence microscopy (UNIT 14.6) or flow cytometry analysis (Holmes and Fowlkes, 1991).

BASIC PROTOCOL

DEAE-dextran transfection is a highly efficient means of introducing the cDNA library into COS cells to ensure that the transfected cells receive as complete a library representation as possible. Typically, ten 100-mm tissue culture plates of COS cells are transfected (Fig. 6.11.1). The subsequent panning steps allow rapid and efficient culling of cells expressing the protein of interest from the bulk of the transfected cells (each 60-mm, antibody-coated plate can be used to pan 1–2 × 107 transfectants). Plasmid DNA can be rescued from the panned cells by obtaining a Hirt supernatant (Hirt, 1967) and following amplification in E. coli, the plasmid DNA can be reintroduced into COS cells using spheroplast fusion. This transfection procedure ensures that a single plasmid type is delivered into each transfectant, allowing greater enrichment in subsequent rounds of transfection and immunoselection. Each round of screening usually requires a set of six fusions and each set of six fusions requires 100 ml of cells in broth. NOTE: All incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise noted. See Chapter 9 introduction for critical parameters concerning media components and preparation. Materials Complete Dulbeccos minimum essential medium containing 10% (v/v) NuSerum or 10% (v/v) calf serum (complete DMEM-10 NS or complete DMEM-10 CS; APPENDIX 3F) 100-mm tissue culture plates seeded with COS cells (∼50% confluent) Contributed by Diane Hollenbaugh, Alejandro Aruffo, Bryan Jones, and Peter Linsley Current Protocols in Molecular Biology (1998) 6.11.1-6.11.16 Copyright © 2003 by John Wiley & Sons, Inc.



cDNA library: plasmid expression vector DNA containing >106 of cDNA clones (UNIT 5.8; see background information), CsCl-purified (UNITS 1.7 & 9.1) Phosphate-buffered saline (PBS; APPENDIX 2) DEAE-dextran/chloroquine solution: PBS containing 10 mg/ml DEAE-dextran (Sigma) and 2.5 mM chloroquine (Sigma) 10% (v/v) DMSO in PBS Trypsin/EDTA solution: PBS containing 0.5 mg/ml trypsin + 0.2 mg/ml EDTA 0.5 mM EDTA/0.02% (v/v) azide in PBS 0.5 mM EDTA/0.02% (v/v) azide/5% (v/v) calf serum in PBS 1 µg/ml purified monoclonal antibody (MAb) or 1:100 dilution of ascites fluid (UNIT 11.1) 0.5 mM EDTA/0.02% (v/v) azide/2% (w/v) Ficoll 60-mm antibody-coated plates (first support protocol) 5% (v/v) calf serum in PBS 0.6% (w/v) SDS/10 mM EDTA 5 M NaCl (APPENDIX 2) Phenol (extracted twice with 1 M Tris⋅Cl, pH 7.5) 2 µg/µl linear polyacrylamide TE buffer, pH 7.5 (APPENDIX 2) Electroporation-competent E. coli cells (UNIT 1.8) LB medium (UNIT 1.1) 100 mg/ml spectinomycin or 35 mg/ml chloramphenicol in ethanol 20% (w/v) sucrose/50 mM Tris⋅Cl, pH 8.0, ice cold 5 mg/ml lysozyme (Sigma #L6876), freshly prepared in 250 mM Tris⋅Cl, pH 8.0 250 mM EDTA, ice cold (APPENDIX 2) 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2)

cDNA library

DEAE-dextran transfection (steps 1- 4)

panning (steps 6-12)

plasmid recovery (steps 13-18)

repeat 3 times

protoplast fusion (steps 20- 36)

plasmid amplification (step 19) isolation of DNA from single colonies

DEAE-dextran transfection


immunofluorescence analysis

Figure 6.11.1 Isolation of a cDNA clone encoding a cell-surface antigen by transient expression in mammalian cells.



10% (w/v) sucrose/10 mM MgCl2 in DMEM (GIBCO/BRL #320-1960AJ) without serum, filter sterilized 60-mm tissue culture plates seeded with COS cells (∼50% confluent) 50% (w/w) PEG 1000 or 1450 in DMEM (no serum), adjusted to pH 7 with 7.5% (w/v) sodium bicarbonate (Baker or Kodak) DMEM without serum Complete DMEM-10 CS (APPENDIX 3F) containing 15 µg/ml gentamycin sulfate Nylon mesh, 100-µm pore size (Tetco) Sorvall GSA rotor or equivalent Swinging-bucket centrifuge (e.g., Sorvall RT-6000B) Additional reagents and equipment for transformation of E. coli by electroporation (UNIT 1.8), phenol extraction and ethanol precipitation (UNIT 2.1), alkaline lysis miniprep (UNIT 1.7), and immunofluorescence (UNIT 14.6) Transfect COS cells using DEAE-dextran 1. Add 5 ml complete DMEM-10 NS to each 100-mm plate of COS cells to be transfected. Each 100-mm plate should be ∼50% confluent the day of transfection (∼5 × 106 cells). This protocol is designed to be used with COS cells and is too harsh for WOP or MOP cells (see background information). If these murine lines must be used, it is important to reduce both the concentration of DEAE-dextran used to 200 ìg/ml final and the time that the cells are exposed to the transfection medium to 2 hr, and to use IMDM (Iscoves modified Dulbeccos medium; GIBCO/BRL #430-2200) in place of DMEM (prepare complete IMDM media as for complete DMEM media, but omit amino acids).

2. To each dish, add 5 µg cDNA library and mix, then add 0.2 ml DEAE-dextran/chloroquine solution and mix. Incubate 4 hr. Typically, libraries of >106 clones are used to obtain plasmid DNA. It is important that the DNA and the DEAE-dextran form a fine, invisible precipitate. If the DNA is not diluted prior to addition of DEAE-dextran, a large DNA/DEAE-dextran precipitate forms (it is easily seen), which is not readily taken up by the cells. Check the cells after ∼3 hr exposure to the DEAE transfection mix, as their health can decline rapidly. This is particularly true of chloroquine transfections, and it is usually better to shorten the transfection than to allow too many cells to die.

3. Aspirate the medium and add 2 ml of 10% DMSO. Incubate ≥2 min at room temperature. The time that the cells are exposed to the DMSO is not critical.

4. Remove DMSO and replace with 10 ml complete DMEM-10 CS. Incubate overnight. 5. Aspirate the medium, add PBS, then aspirate the PBS. Add 2 ml trypsin/EDTA to each plate and incubate 5 to 15 min until cells have lifted from the plate. Replate the cells on two new 100-mm plates and incubate overnight. Replating the cells allows them to recover more effectively from the transfection. In addition, the DEAE-dextran transfection makes the cells sticky and replating allows them to be lifted from the plates with EDTA to initiate the panning step.

Immunoselect the cells by panning 6. Aspirate the medium, add 2 ml EDTA/azide solution, and incubate 10 to 20 min to detach cells from plates. 7. Pipet vigorously with a short Pasteur pipet to dislodge the cells, then transfer cells from each plate into a 15-ml centrifuge tube.



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8. Centrifuge 4 min at 200 × g (e.g., 1000 rpm in a Sorvall RT-6000B with GSA rotor or in a tabletop centrifuge) and discard supernatant. 9. Resuspend cells in 0.5 to 1.0 ml EDTA/azide/calf serum solution and add purified MAb to 1 µg/ml final or ascites at a 1:100 dilution final. Incubate 30 to 60 min on ice. 10. Add an equal volume of EDTA/azide solution and carefully layer on 3 ml EDTA/azide/Ficoll solution. Centrifuge 4 min at 200 × g. Aspirate supernatant in one smooth movement. 11. Add 3 ml EDTA/azide/calf serum solution to each antibody-coated plate. Resuspend cells in 0.5 ml EDTA/azide solution, then add aliquots of the cells to these plates by pipetting them through a nylon mesh. Leave 1 to 3 hr at room temperature. Four 6-mm antibody-coated plates are used in each round of panning. It is important to pass the cells through the nylon mesh to break up large clumps of cells which might contain both positive and negative cells. This ensures that individual antibody-coated cells bind to the panning plate.

12. Remove excess cells not adhering to the plate by gently washing two to three times with 3 ml of 5% calf serum (or complete DMEM-10 NS or complete DMEM-10 CS). Washing gently means swirling the plate with a smooth motion for ∼30 sec. The plate obtained after these washes is known as a panned plate.

Recover plasmid DNA and transform E. coli 13. Add 0.4 ml SDS/EDTA solution to the panned plate and leave 20 min at room temperature (to lyse the cells). This incubation period can be as little as 1 min if there are only a few cells on the plate.

14. Pipet the viscous mixture into a microcentrifuge tube. Add 0.1 ml of 5 M NaCl, mix, and place ≥3 hr on ice or leave overnight at 4°C. The viscosity is primarily due to the genomic DNA. It is important to avoid shearing the genomic DNA so that it will not contaminate the plasmid DNA. Keeping the mixture as cold as possible seems to improve the quality of the Hirt supernatant.

15. Microcentrifuge 4 min at top speed, 4°C, and remove supernatant carefully. 16. Extract with phenol (twice if the first interface is not clean) and add 5 µl (10 µg) of 2 µg/µl linear polyacrylamide (or other carrier). 17. Fill the tube to the top with 100% ethanol and precipitate. Resuspend the pellet in 0.1 ml TE buffer, pH 7.5. 18. Add 10 µl of 3 M sodium acetate and 300 µl of 100% ethanol, and repeat precipitation. Resuspend the pellet in 0.1 ml TE buffer, pH 7.5. 19. Transform electroporation-competent E. coli cells by electroporation using DNA obtained from step 18. Incubate overnight at 37°C. Approximately 105 bacterial colonies should be obtained. It is advisable to transform E. coli with an aliquot of DNA to determine the amount necessary to obtain 105 colonies. Generally 1⁄10 to 1⁄4 of the recovered DNA will be used. Use of Monoclonal Antibodies for Expression Cloning



Prepare the spheroplasts 20. Rinse the plate from step 19 several times with LB medium while scraping with a spreader to dislodge the bacteria. Use 1⁄10 to 1⁄5 of the pooled scrapings to inoculate 200 ml of LB medium. Grow to OD600 = 0.5 at 37°C with shaking. 21. Add 100 mg/ml spectinomycin to 100 µg/ml or 35 mg/ml chloramphenicol to 150 µg/ml. Incubate with shaking 10 to 16 hr at 37°C. Do not let the cells grow >16 hr or they will begin to lyse. If the cells lyse, do not proceed.

22. Centrifuge 100 ml of the culture in a 250-ml bottle, 5 min at 4000 × g (e.g., 5000 rpm in a Sorvall with GSA rotor), room temperature or 4°C. 23. Drain well and resuspend pellet in 5 ml of ice-cold sucrose/Tris⋅Cl, pH 8.0. 24. Add 1 ml of 5 mg/ml lysozyme solution. Incubate 5 min on ice. 25. Add 2 ml of ice-cold 250 mM EDTA, pH 8.0, and incubate 5 min on ice. 26. Add 2 ml of 50 mM Tris⋅Cl, pH 8.0, and incubate 5 min in a 37°C water bath. 27. Place on ice. Check percent conversion to spheroplasts by microscopy. A good spheroplast preparation gives about 80% to 90% conversion; anything 106 cDNA clones (UNIT 5.8; see background information), CsCl purified (UNITS 1.7 & 9.11) Trypsin/EDTA solution: PBS containing 0.5 mg/ml trypsin + 0.2 mg/ml EDTA Phosphate-buffered saline (PBS; APPENDIX 2) Methanol 1% (w/v) nonfat dry milk in PBS with and without monoclonal antibody (MAb) 1% (w/v) nonfat dry milk in PBS containing 0.25 µCi/ml of 125I-labeled protein A 0.6% (w/v) SDS/10 mM EDTA buffer LB medium (UNIT 1.1) Polyvinylidene-wrapped plates (second support protocol) X-ray film Polyvinylidene wrap (e.g., Saran Wrap) Rubber cement Luminescent stickers Additional reagents and equipment for alkaline lysis miniprep (UNIT 1.7) and autoradiography (APPENDIX 3) Transfect cells and fix with methanol 1. Transfect ten 100-mm plates of COS cells with the cDNA library using the DEAEdextran method as described in steps 1 to 4 of the first basic protocol. 2. Trypsinize each plate of transfected COS cells using trypsin/EDTA as described in step 5 of the first basic protocol and replate onto polyvinylidene-wrapped plates. Incubate 1 to 2 days. Cells should be split at a ratio that will provide 50% to 75% confluency the following day. Complete DMEM containing penicillin and streptomycin should be used to avoid minor contamination that may occur because the plates are not sterile.

3. Remove medium from transfected COS cells. Wash plates by adding 5 ml PBS and aspirating at room temperature. Repeat wash one time. Generally, 15 to 20 plates are used.

4. Add ∼6 ml methanol and incubate 5 min at room temperature. 5. Wash plates three times with PBS, leaving the first addition of PBS on the plate for 2 to 3 min before aspirating. 6. Add 4 ml of 1% dry milk containing MAb to each plate. Incubate 30 to 60 min at room temperature. 7. Wash plates twice by adding 5 ml of 1% dry milk (without MAb), swirling gently, and removing solution. Radiolabel and locate positive cells 8. Add 4 ml of 1% dry milk in PBS containing Incubate 30 min at room temperature.

125

I-labeled protein A to each plate.

9. Rinse plates four times with 1% dry milk in PBS and one time with PBS at room temperature. Remove all excess liquid. Use of Monoclonal Antibodies for Expression Cloning

It is helpful to prop the plates on edge to remove the excess liquid.

10. Completely cover a piece of X-ray film with polyvinylidene wrap and tape the edges



of the wrap to the film. Paint the wrap with a thin layer of rubber cement. Allow rubber cement to dry briefly. The X-ray film acts as a support and may be a used piece that would otherwise be discarded. The presence of rubber cement does not appear to affect DNA recovery. Five plates will fit on an 8 × 10–inch (20.3 × 25.4–cm) film or 15 plates will fit on a 14 × 17–inch (35 × 43–cm) film.

11. Paint the bottom surface of the polyvinylidene-wrapped plate with a thin layer of rubber cement and allow the rubber cement to dry briefly. 12. Place the plates on the support. Lift the plate slightly and cut the wrap away from the edge of the plate. The wrap with the cells will be left on the X-ray film. By slightly lifting the plate, a scalpel held nearly horizontal along the inside of the plate can be used to cut the wrap from the plate without cutting the wrap on the support. While this technique is not difficult, it is advisable to practice it on nonradioactive samples.

13. Affix luminescent stickers to the support. Cover the support and samples with polyvinylidene wrap. There are now three layers of wrap on the support. Without the luminescent stickers, the film cannot be aligned over the samples to recover positive cells. Careful alignment is necessary to recover positive cells (there will be insufficient background to use the location of the samples for alignment).

14. Autoradiograph with an intensifying screen for 1 to 2 days at −70°C. Develop the film. 15. Align the film over the support using the luminescent stickers. Mark the location of positive cells by piercing the film alongside the spot with a needle (this leaves a mark on the polyvinylidene wrap). A light box is helpful for locating the positive cells on the film.

16. Remove the film. Cut small squares of ∼3 mm in the polyvinylidene wrap at the places it is marked. Recover plasmid DNA and isolate the positive clone 17. Add 400 µl of SDS/EDTA buffer to a microcentrifuge tube. Place squares in tube (10 to 25 squares per tube) and incubate 30 min at room temperature. All three layers of wrap are placed in the tube.

18. Recover plasmid DNA and transform E. coli as in steps 14 to 19 of the first basic protocol, plating transformed bacteria on separate LB plates to form pools of appropriate numbers of clones. Incubate overnight at 37°C. 19. Collect the bacteria from the LB plates by rinsing several times with LB medium while scraping with a spreader to dislodge the bacteria. 20. Prepare plasmid DNA from the scraped colonies using an alkaline lysis miniprep. 21. Transfect COS cells with the pools of plasmid DNA by the DEAE-dextran method as described in steps 1 to 4 of the first basic protocol. In general, 1⁄10 to 1⁄5 of the DNA obtained from 1000 colonies is used to transfect one 100-mm plate of COS cells. Screening of Recombinant DNA Libraries


Supplement 23

22. Repeat steps 2 to 14 above to identify pools enriched with the gene of interest. Store the appropriate plasmid DNA in TE buffer at −20°C. When screening pools, it is possible to use 60-mm plates (reduce all volumes to 1⁄3).

23. Prepare DNA from a single bacterial colony, transfect COS cells, and analyze as in steps 38 and 39 of the first basic protocol. SUPPORT PROTOCOL

PREPARATION OF POLYVINYLIDENE-WRAPPED PLATES Plates are prepared in which the growth surface is a polyvinylidene wrap (to be used in the second basic protocol). The plates are quite sturdy and can be used in the same manner as standard tissue culture plates. Plates may be prepared a day or two in advance but the wrap will stretch and become floppy on prolonged storage. Additional Materials Chloroform 70% ethanol 0.1 mg/ml poly-L-lysine HCl (Sigma) in 50 mM Tris⋅Cl, pH 8.0, freshly prepared 100-mm or 60-mm tissue culture plates 1. Break the bottoms out of a 100- or 60-mm tissue culture plate with a blunt object. Safety glasses are advisable. Strike the plate near the sides of the plate rather than in the center. If too much force is used, the sides of the plate will break as well. Structural stability is increased if the outer edges of the bottoms are not removed.

2. In a fume hood at room temperature, dip the top rim of the plate in chloroform to a depth of ∼3 mm. 3. Shake off excess chloroform and place plate on a piece of polyvinylidene wrap laid flat. Place the plate and attached film into the lid of the plate to force the wrap into contact with the edges of the plate. 4. Remove the lid and gently but firmly pull the film tight to form a smooth surface. Adhesion of the film to the outside of the plate helps maintain the strength of the seal.

5. Cut excess wrap from the plate with a razor blade. The plate is now essentially inverted. The lid is placed over the opening that had been the bottom of the plate. When using 100-mm plates, a second lid is used to support the new wrap bottom.

6. Wash the plate two times with 70% ethanol. Allow the wrap to soak in the ethanol ∼30 min. 7. Wash the plate with water. 8. Add 0.1 mg/ml poly-L-lysine to the plate—11 ml for a 100-mm plate and 4 ml for a 60-mm plate. Incubate 2 hr to overnight at room temperature. 9. In a tissue culture hood, rinse the plates twice with PBS. Dishes should not be stored more than a few days because the wrap will stretch and become loose. Use of Monoclonal Antibodies for Expression Cloning



COMMENTARY Background Information Transient expression in mammalian cells has emerged as a powerful method for isolating cDNA clones that encode secreted cell-surface and intracellular proteins. It was first used to isolate cDNA clones encoding the lymphokine granulocyte/macrophage colony stimulating factor (GM-CSF; Lee et al., 1985; Wong et al., 1985). This cloning strategy is well suited for isolating any secreted proteins for which a rapid and sensitive bioassay exists, and has since been applied to isolate cDNA clones encoding a number of different lymphokines. Transient expression cloning was combined with the simple but powerful immunoselection technique of panning (Wysocki and Sato, 1978) to isolate a cDNA encoding the T cell–surface proteins CD2 and CD28 (Seed and Aruffo, 1987; Aruffo and Seed, 1987). This procedure has since been used to isolate cDNA clones encoding a number of different cell-surface proteins when antibodies against them were available. When antibodies against a cell-surface receptor of interest are not available, but its ligand is, transient expression in mammalian cells has been combined with ligand-binding assays to isolate cDNA clones encoding the receptor. This strategy was first used to isolate a cDNA clone encoding the receptor for the lymphokine interleukin 1 (Sims et al., 1988). Modifications to this protocol (Gearing et al., 1989) have since allowed the use of this strategy to isolate a number of receptors. Recent improvements have allowed the use of transient expression in mammalian cells to isolate cDNA clones encoding intracellular proteins, including the major DNA-binding protein of the erythroid lineage (Tsai et al., 1989), the lysosomal membrane glycoprotein CD63 (Metzelaar, 1991), and fucosyltransferase, which adds fucose to N-acetylglucosamine with α(1,3) linkages, allowing the expression of the sialyl CD15 antigen (Goelz et al., 1990). Transient expression in mammalian cells. Mammalian cells are ideal hosts for screening cDNA libraries prepared using mRNA isolated from higher eukaryotes. These cells are able to synthesize transcripts correctly from the cDNA clones in the library and are likely to process the proteins that they encode appropriately, thus maximizing the likelihood that the foreign proteins will be present in their native state and will be detectable using functional or immunological assays.

Mammalian cells were initially used as the host for isolation of genomic DNA fragments encoding oncogenes by stably introducing genomic DNA fragments derived from human tumors into murine cells (Goldfarb et al., 1982; Shih and Weinberg, 1982). The gene encoding the oncogene was then rescued from the transfected cells that had acquired the transformed phenotype. Stable transfections of mammalian cells were subsequently combined with immunoselection procedures to isolate genomic DNA fragments encoding the human HLA and β2 microglobulin genes (Kavathas and Herzenberg, 1983) and cDNA clones encoding the T cell–surface proteins CD8 (Kavathas et al., 1984; Littman et al., 1985) and CD4 (Maddon et al., 1985). The time involved in obtaining stable transfectants expressing the gene of interest and the difficulties associated with recovering the transfected DNA from the host’s chromosomal DNA has limited the number of genes isolated using this cloning strategy. These difficulties prompted the development of transient expression systems for use in cloning with mammalian cells as the screening host. Unlike stable transfectants, these methods permit rapid preparation and detection of transfectants expressing the protein of interest and efficient recovery of the DNA encoding it. Many technical advances have permitted efficient and routine screening of cDNA libraries in transiently expressing mammalian cells. These include the following developments: shuttle vectors that contain the appropriate eukaryotic transcription elements for highlevel protein expression in transfected mammalian cells, mammalian cell lines that can act as effective heterologous expression hosts, and transfection protocols that allow efficient introduction of plasmid DNA into mammalian cells. Expression vectors. A number of mammalian expression vectors permit screening of cDNA libraries by transient expression in mammalian cells (Chapter 16; Kaufman, 1990). These plasmids contain at least four basic elements: an efficient eukaryotic transcription unit, a viral-derived origin of replication, a prokaryotic origin of replication, and a prokaryotic selectable marker. One particular expression vector, CDM8 (Seed, 1987; Fig. 16.12.1; available from Invitrogen #V308-20) is especially engineered for these purposes, and is described in detail in UNIT 16.12. CDM8 con-



Supplement 62


tains origins of replications derived from polyoma and SV40 viruses that allow for plasmid replication in cell lines expressing either the polyoma or SV40 large T antigens, respectively; usually, WOP (Dailey and Basilico, 1985) and COS (Gluzman, 1981) cells, respectively. Bacterial and M13 origins of replication (ori) are also present, allowing for plasmid amplification in bacteria and production of single-stranded DNA, respectively. The plasmid contains a supF gene as a prokaryotic selectable marker and a T7 RNA polymerase promoter for the preparation of mRNA in vitro from the subcloned cDNAs. Mammalian cell lines. A number of cell lines have been developed that are excellent hosts for screening cDNA libraries prepared in the vector described above. Perhaps the most popular is the COS cell line (Gluzman, 1981), which was derived from the African green monkey kidney cell line CV-1 by transformation with an origin-defective SV40 virus. This cell line produces wild-type SV40 large T antigen but no viral particles. When plasmids containing an SV40 virus–derived ori are transfected into COS cells, the plasmid is replicated to a high copy number 48 hr posttransfection (10,000 to 100,000 copies/cell). This high-level replication has two important consequences. First, it allows for amplification of all DNA templates available for transcription. Second, it allows for recovery of the plasmid encoding the protein of interest from the immunoselected cells. This last point is of importance in the two basic cloning protocols because each cycle of transfection and immunoselection is followed by a plasmid-rescue step. Other cell lines that have been used for expression cloning include the murine cell lines WOP (Dailey and Basilico, 1985) and MOP (Muller et al., 1984). These two cell lines express the polyoma large T antigen allowing the replication of plasmids containing a polyoma origin of replication in the transfected cells (1,000 to 10,000 copies/cell, 48 hr posttransfection). Another cell line, CV-1/EBNA, has been developed for screening cDNA libraries in conjunction with the expression vector pDC406, which contains an Epstein-Barr virus ori (McMahan et al., 1991). The ability of COS cells to endure the transfection protocols and their ability to replicate the transfected plasmid to a very high copy number make them the cells of choice when screening a cDNA library with the methods described here. However, when the antibodies to be used in the immunoselection step cross-

react with proteins expressed by COS cells, another cell line must be used. In these cases, WOP or MOP cells can be used successfully, in spite of their more delicate nature and lower copy numbers of transfected plasmid. Mammalian cell transfection. A number of transfection protocols have been developed for efficient introduction of foreign DNA into mammalian cells, including calcium phosphate, DEAE-dextran, spheroplast fusion, lipofection, and electroporation (UNITS 9.1-9.5). Two factors determine which transfection procedure should be used when screening a cDNA library by transient expression in mammalian cells: first, the efficiency of transfection, and second, the number of different plasmids that are introduced into each cell during transfection. Two transfection protocols, DEAE-dextran (McCutch and Pagano, 1968) and spheroplast fusion (Sandri-Goldin et al., 1981), are discussed below. The mechanism by which DEAE-dextran transfections allow for introduction of foreign DNA into cells is poorly understood. It is believed that the positive charge of the DEAEdextran polymer neutralizes the negative charge of the DNA polymer, forming a fine precipitate that can come into contact with the plasma membrane of the host cell. The DEAEdextran/DNA complex is then internalized by pinocytosis. Some of this DNA makes its way to the host-cell nucleus, where it is replicated and transcribed. Because the foreign DNA enters the cell via endosomes, DNA integrity is enhanced by the addition of chloroquine to the transfection medium to prevent endosome acidification. DEAE-dextran transfections are very efficient, allowing for transfection of up to 70% of the host cells and delivery of up to 200 different plasmids into each transfected cell. Introducing foreign DNA into mammalian cells by spheroplast fusion is very inefficient, allowing for transfection of only 1% to 2% of the host cells. Bacteria containing the foreign DNA are treated with lysozyme to remove their cell walls. The resulting spheroplasts are then fused with the host mammalian cell using polyethylene glycol (PEG), allowing introduction of the foreign DNA directly into the host cell cytoplasm. The DNA is then replicated and transcribed in the nucleus. Because of the inefficiency of the procedure, each host cell fuses with only one spheroplast, on average, introducing only a single plasmid type into each transfected cell. The immunoselection screening method de-



scribed in the first basic protocol involves multiple rounds of mammalian cell transfection, immunoselection, and plasmid rescue steps. The rescued plasmids are amplified in E. coli and reintroduced into mammalian cells to initiate additional rounds of enrichment. To take full advantage of immunoselection, the two methods of transfection are used. When the cDNA library is first introduced into mammalian cells, it is important to obtain a complete representation of the library in the transfected host cells, ensuring that the protein of interest is expressed by the transfectants. For this reason, the first round of enrichment is initiated using DEAE-dextran transfection. In subsequent cycles of enrichment, it is important that a single plasmid type be delivered into each of the transfected cells to maximize the level of enrichment obtained in the subsequent immunoselection steps. This is accomplished using the spheroplast fusion transfection. Immunoselection procedures. Two immunoselection techniques designed to rapidly select and enrich for plasmids encoding proteins of interest from a cDNA library transfected into mammalian cells are described. The first strategy is designed to isolate cDNA clones encoding surface proteins. A cDNA library prepared in a mammalian expression vector is transfected into COS cells using DEAE-dextran transfection. Forty-eight hours posttransfection, the cells are lifted from the plate and incubated with antibodies directed against the protein of interest. Cells expressing the foreign protein on their cell surface are easily culled from the bulk of the transfected cells by panning on plastic plates coated with anti-antibody antibodies (Wysocki and Sato, 1978). Plasmid DNA is then recovered from the transfected cells by the method of Hirt (Hirt, 1967), amplified in E. coli, and reintroduced into COS cells by spheroplast fusion. Two additional rounds of spheroplast fusion and panning are usually required to enrich for the plasmid encoding the protein of interest. Panning has many advantages over other immunoselection procedures. It is fast, efficient (107 cells can easily be panned on two 60-mm plastic plates in 30 min), and very inexpensive. Other immunoselection techniques, such as sorting of fluorescence-labeled cells (Holmes and Fowlkes, 1991), may be used to screen cDNA libraries in transiently expressing mammalian cells (Yamasaki et al., 1988); but the greater demands on time, equipment, and technical expertise make these methods much less attractive.

The second strategy is designed to isolate cDNA clones encoding intracellular antigens. This method is a combination of the techniques described by Munro and Maniatis (1989) and Metzelaar et al. (1991). COS cells are transfected with a cDNA library by DEAE-dextran transfection. The day after transfection, they are replated onto poly-L-lysine-coated polyvinylidene wrap and allowed to grow for 1 to 2 additional days. They are then washed and fixed with methanol. The permeabilized cells are incubated with antibodies directed against the protein of interest, washed, and incubated with radiolabeled protein A (125I). After washing, they are exposed to film to identify radiolabeled cells, which are then recovered by cutting the polyvinylidene wrap. The plasmid DNA is recovered from these cells by the method of Hirt, amplified in E. coli, and subjected to additional rounds of transfection and immunoselection.

Critical Parameters The cloning strategy described in the first basic protocol is well suited for isolating cDNA clones encoding cell-surface proteins (see discussion above). If a cDNA library is thought to contain cDNA clones encoding a number of proteins of interest, it is possible to isolate all of them simultaneously by simply using a mixture of antibodies against all of the proteins of interest in the panning steps of the first three rounds of enrichment. The last cycle of immunoselection is carried out independently with antibodies against each of the different proteins. Many times antibodies directed against the protein(s) of interest are of multiple isotypes. It is important, in this case, to use panning plates that have been coated with anti-Ig antibodies that bind to each of the isotypes present in the initial antibody pool. Alternatively, individual panning plates can be prepared for each of the antibody isotypes present in the initial antibody pool, but no significant advantage is achieved. Although this cloning strategy has allowed isolation of a large number of cDNA clones encoding cell-surface proteins, it has some serious limitations. As with any expression cloning system, the gene of interest must initially be present in the library, the target protein must be functional or immunoreactive as a single chain, and the host system must posttranslationally modify the protein appropriately when these modifications are required for function or immunoreactivity. Compared with earlier bacterial expression cloning systems, the mam-



Supplement 23


malian expression systems presented here are more likely to appropriately modify the gene products of higher eukaryotes. However, these methods require the host mammalian cell to express the target protein on its surface as a single molecule. This may not occur in cases where the target protein is part of a heterocomplex that requires more than one member for surface expression. Several factors contribute to the successful application of this cloning strategy. The most critical parameter when screening a cDNA library by expression in mammalian cells is the quality of the cDNA library (for a more complete discussion, see UNIT 5.8). In addition, the quality of the COS cells and transfectability is of utmost importance. COS cells maintained in culture for prolonged periods tend to become refractory to transfection. For this reason, it is important to check the cells periodically for transfectability and replace them with cells from frozen stocks when necessary. The competency level of the bacterial cells used to amplify the DNA rescued from the immunoselection step is very important and should be determined prior to the start of the experiment. If only a few positive cells are immunoselected by panning, it is of utmost importance that this plasmid DNA work its way into the bacteria so that it can potentially be amplified and thus be available for subsequent rounds of transfection and immunoselection. Ideally, only cells whose competency level is ≥109 cfu/µg DNA should be used. For most bacterial strains, this can be achieved using electroporation. When using panning to immunoselect the transfected cells, it is important to check that the antibody directed against the protein of interest does not bind to COS cells. If it does, another cell line should be used for screening. The researcher must also be mindful that DEAE-dextran treatment of cells changes their phenotype and thus the antibodies to be used in the panning step should be tested for cross-reactivity with mock-transfected cells. The most difficult step, technically, is the spheroplast-fusion step. Careful timing of cell exposure to PEG is necessary to promote fusion while minimizing cell death. Although lowermolecular-weight PEG (PEG 1000) results in more efficient fusion, it is more toxic to the cells. The cloning strategy described in the second basic protocol will not be effective if the antibodies used recognize epitopes that are sensitive to methanol treatment. Testing of the target

cells may reveal this limitation. If methanol sensitivity cannot be assayed, multiple antibodies against a given protein should be used when available.

Anticipated Results In the first basic protocol, the authors typically use DNA prepared from a cDNA library of ≥106 clones for screening. After four rounds of transfection, immunoselection, and plasmid rescue, 12 individual colonies are picked and plasmid DNA prepared from them. If the cloning procedure has been successful, at least one DNA preparation directs the expression of the protein of interest. In general, if the screening is unsuccessful, this indicates that the clone of interest may not be present in the library. In this case, screening of a new cDNA library may be successful. Unsuccessful screening may indicate that the target gene is refractory to cloning using this strategy. Possible alternative methods are described in Chapter 5. In the second basic protocol, three to five spots are generally obtained on each 100-mm plate. It is possible to enrich a mixture that is 1:10,000 in the desired clone to 1:100 with a single immunoselection step.

Time Considerations In the first basic protocol, the cloning strategy involves multiple steps. Each cycle of transfection, immunoselection, plasmid rescue, and amplification can be comfortably accommodated in 1 week. On this schedule, it is possible to screen a cDNA library in 1 month. However, the more ambitious can screen a library in 3 weeks. In the second basic protocol, successful application allows identification of positive pools in 11⁄2 weeks. The screening of subsets of the pool can be accelerated by using 60-mm plates. When screening individual clones, transfectants may be assayed by immunofluorescence. The length of time required to obtain a single isolated positive clone will depend on the pool sizes used.

Literature Cited Aruffo, A. and Seed, B. 1987. Molecular cloning of a CD28 cDNA by a high-efficiency COS cell expression system. Proc. Natl. Acad. Sci. U.S.A. 84:8753-8577. Dailey, L. and Basilico, C. 1985. Sequence in the polyomavirus DNA regulatory region involved in viral DNA replication and early gene expression. J. Virol. 54:739-749.



Gearing, D.P., King, J.A., Gough, N.M., and Nicola, N.A. 1989. Expression cloning of a receptor for human granulocyte-macrophage colony-stimulating factor. EMBO J. 8:3667-3676. Gluzman, Y. 1981. SV40-Transformed simian cells support the replication of early SV40 mutants. Cell 23:175-182. Goelz, S.E., Hession, C., Goof, D., Griffiths, B., Tizard, R., Newman, B., Chi-Rosso, G., and Lobb, R. 1990. ELFT: A gene that directs the expression of an ELAM-1 ligand. Cell 63:13491356. Goldfarb, M., Schimizu, K., Perucho, M. and Wigler, M. 1982. Isolation and preliminary characterization of a human transforming gene from T24 bladder carcinoma cells. Nature (Lond.) 296:404-409. Hirt, B. 1967. Selective extraction of polyoma DNA from infected mouse cell cultures. J. Mol. Biol. 26:365-369.

McMahan, C.J., Slack, J.L., Mosley, B., Cosman, D., Lupton, S.D., Brunton, L.L., Grubin, C.E., Wignall, J.M., Jenkins, N.A., Brannan, C.I., Copeland, N.G., Huebner, L., Croce, C.M., Cannizzarro, L.A., Benjamin, D., Dower, S.K., Spriggs, M.K., and Sims, J.E. 1991. A novel IL-1 receptor, cloned from B cell by mammalian expression, is expressed in many cell types. EMBO J. 10:2821-2832. Metzelaar, M.J., Wijngaard, P.L.J., Peters, P.J., Sixma, J.J., Nieuwenhuis, H.K., and Clevers, H.C. 1991. CD63 antigen. J. Biol. Chem. 266:3239-3245. Muller, W.J., Naujokas, M.A., and Hassell, J.A. 1984. Isolation of large T antigen-producing mouse cell lines capable of supporting replication of polyomavirus-plasmid recombinants. Mol. Cell. Biol. 4:2406-2412. Munro, S. and Maniatis, T. 1989. Expression cloning of the murine interferon γ receptor cDNA. Proc. Natl. Acad. Sci. U.S.A. 86:9248-9252.

Holmes, K. and Fowlkes, B.J. 1991. Preparation of cells and reagents for flow cytometry. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 5.3.1-5.3.11. Greene Publishing and John Wiley & Sons, New York.

Sandri-Goldin, R.M., Goldin, A.L., Glorioso, J.C., and Levine, M. 1981. High-frequency transfer of cloned herpes simplex virus type I sequences to mammalian cells by protoplast fusion. Mol. Cell. Biol. 1:743-752.

Kaufman, R.J. 1990. Overview of vectors used for expression in mammalian cells. Methods Enzymol. 185:487-511.

Seed, B. 1987. An LFA-3 cDNA encodes a phospholipid-linked membrane protein homologous to its receptor CD2. Nature (Lond.) 329:840842.

Kavathas, P. and Herzenberg, L.A. 1983. Stable transformation of mouse L cells for human membrane T-cell differentiation antigens, HLA and 2-microglobulin: Selection by fluorescence-activated cell sorting. Proc. Natl. Acad. Sci. U.S.A. 80:524-528. Kavathas P., Sukhatme, V.P., Herzenberg, L.A., and Parnes, J.R. 1984. Isolation of the gene encoding the human T-lymphocyte differentiation antigen Leu-2 (T8) by gene transfer and cDNA subtraction. Proc. Natl. Acad. Sci. U.S.A. 81:7688-7692. Lee, F., Yokota, T., Otsuka, T., Gemmell, L., Larson, N., Luh, J., Arai, K.-I., and Rennick, D. 1985. Isolation of cDNA for a human granulocytemacrophage colony-stimulating factor by functional expression in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 82:4360-4364. Littman, D.R., Thomas Y., Maddon, P.J., Chess, L., and Axel, R. 1985. The isolation and sequence of the gene encoding T8: A molecule defining functional classes of T lymphocytes. Cell 40:237-246. Maddon, P.J. Littman, D.R., Godfrey, M., Maddon, D.E., Chess, L., and Axel, R. 1985. The isolation and nucleotide sequence of a cDNA encoding the T cell surface protein T4: A new member of the immunoglobulin gene family. Cell 42:93-104. McCutchan, J.H. and Pagano, J.S. 1968. Enhancement of the infectivity of simian virus 40 deoxyribonucleic acid with diethylaminoethyl-dextran. J. Natl. Cancer Inst. 40:351-357.

Seed, B. and Aruffo, A. 1987. Molecular cloning of the CD2 antigen, the T-cell erythrocyte receptor, by a rapid immunoselection procedure. Proc. Natl. Acad. Sci. U.S.A. 84:3365-3369. Shih, C. and Weinberg, R.A. 1982. Isolation of a transforming sequence from a human bladder carcinoma cell line. Cell 29:161-169. Sims, J.E., March, C.J., Cosman, D., Widmer, M.B., MacDonald, H.R., McMahan, C.J., Grubin, C.E., Wignall, J.M., Jackson, J.L., Call, S.M., Friend, D., Alpert, A.R., Gillis, S., Urdal, D.L., and Dower, S.K. 1988. cDNA expression cloning of the IL-1 receptor, a member of the immunoglobulin superfamily. Science 241:585-589. Tsai, S.F., Martin, D.I., Zon, L.I., D’Andrea, A.D., Wong, G.G., and Orkin, S.H. 1989. Cloning of cDNA for the major DNA-binding protein of the erythroid lineage through expression cloning. Nature (Lond.) 339:446-451. Wong, G.G., Witek, J.S., Tempel, P.A., Wilkens, K.M., Leary, A.C., Luxenberg, D.P., Jones, S.S., Brown, E.L., Kay, R.M., Orr, E.C., Shoemaker, C., Golde, D.W., Kaufman, R.J., Hewick, R.M., Wang, E.A., and Clark, S.C. 1985. Human GMCSF: Molecular cloning of the complementary DNA and purification of the natural and recombinant proteins. Science 228:810-815. Wysocki, L.J. and Sato, V.L. 1978. Panning for lymphocytes: A method for cell selection. Proc. Natl. Acad. Sci. U.S.A. 75:2844-2848. Screening of Recombinant DNA Libraries


Supplement 23

Yamasaki, K., Taga, T., Hirata, Y., Yawata, H., KaWanishi, Y., Seed, B., Taniguchi, T., Hirano, T., and Kishimoto, T. 1988. Cloning and expression of the human interleukin-6 (BSF-2/ IFNβ 2) receptor. Science 241:825-828.

Metzelaar et al., 1991; Munro and Maniatis, 1989. See above.

Key References

Contributed by Diane Hollenbaugh and Alejandro Aruffo (cell-surface and intracellular antigens)

Aruffo and Seed, 1987; Seed and Aruffo, 1987. See above. Contain original descriptions of cDNA library construction in CDM8 and isolation of cDNA clones encoding cell-surface antigens by expression cloning.

Contain descriptions of growth of COS cells on wrap and screening for extracellular ligands.

Bryan Jones and Peter Linsley (intracellular antigens) Bristol-Myers Squibb Seattle, Washington




Recombination-Based Assay (RBA) for Screening Bacteriophage Lambda Libraries The recombination-based assay represents a convenient way to screen a complex library constructed in bacteriophage λ for homology to a given sequence cloned into a specially designed plasmid. The technique serves to screen a bacteriophage library rapidly and efficiently with a sequence cloned into a plasmid; counterselection then yields the gene product of interest with its plasmid carrier deleted. Because 106 to 107 plaque-forming units (pfu) may be screened using several petri dishes, and the homology for crossing-over need only be >25 bp, the RBA represents an efficient way to screen complex λ libraries rapidly for homology to a given sequence.

UNIT 6.12

BASIC PROTOCOL

In this procedure (outlined in Fig. 6.12.1), a λ library is screened using a specially designed R6K supF plasmid, pAD1 (Fig. 6.12.2), carrying the desired target sequence. Recombinants arising from cross-over events between the plasmid and a bacteriophage carrying a corresponding region of homology are selected by their ability to grow on strain DM21 (Fig. 6.12.3). Growth of λ on DM21 requires the presence of the supF allele encoded on the plasmid to suppress an amber mutation in the host strain that prevents λ propagation. Recovery of the original phage carrying the target sequence requires a reversal of the homologous recombination event. This reversal occurs spontaneously, and is detected by PCR amplification using primers that flank the cloning site in the λ vector (Fig. 6.12.4).

supF

supF

Kmr

Kmr

ori ori DM21 blue plaque lacZam P1 ban dnaBam co

s

cos

λ-plasmid chimera colorless plaque plasmid cos cos

λ phage

human probe

cos cos

phage

Figure 6.12.1 The recombination-based assay (RBA). Homology between sequences in a plasmid and a bacteriophage >25 bp long (Watt et al., 1985; Shen and Huang, 1986, 1989; King and Richardson, 1986) mediates a recombination event between the two vectors. As a result supF is integrated into the bacteriophage, allowing it to plate on the dnaBam host DM21 (see Table 6.12.1). The cointegrate yields a blue plaque in the presence of IPTG and Xgal on the lacZam host DM21, as supF suppresses the amber mutations in both the dnaB and lacZ genes. Different shadings indicate origins of DNA regions. Contributed by David M. Kurnit Current Protocols in Molecular Biology (1994) 6.12.1-6.12.12 Copyright © 2000 by John Wiley & Sons, Inc.



Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

DNA fragment encoding sequence of interest Plasmid pAD1 (Fig. 6.12.2; available from Dr. D. Kurnit) recA+ E. coli strain (Table 1.4.5 or commercial suppliers) L broth (see recipe) with 50 µg/ml kanamycin (Table 1.4.1) Bacteriophage λ library (UNIT 5.8) Lambda top agar (see recipe) Lambda plates (see recipe), some with 50 µg/ml kanamycin and some with 100 µg/ml streptomycin (Table 1.4.1) Suspension medium (SM; see recipe) Chloroform E. coli DM21, DM75, DM392, and DM1061 (Fig. 6.12.3 and Table 6.12.1), saturated overnight cultures freshly grown in LB medium (UNIT 1.1) with 100 µg/ml streptomycin 100 mM IPTG (isopropyl thiogalactoside; Table 1.4.2) 2% Xgal in DMF (see recipe) Additional reagents and equipment for subcloning DNA into plasmids (UNIT 3.16), culturing (UNIT 1.1) and transformation (UNIT 1.8) of bacteria, plating and titering λ phage (UNIT 1.11), β-galactosidase assay (UNIT 1.4), and PCR amplification (UNIT 15.1) NOTE: All incubations are at 37°C unless otherwise specified. Screen library and select recombinants 1. Clone the sequence of interest into a pAD1 plasmid and transform into recA+ E. coli strain yielding a kanamycin-resistant recA+ strain. Prepare a saturated overnight culture grown with aeration in L broth containing 50 µ/ml kanamycin.

EcoRI Sfi l

Notl Sacll

polylinker

Pstl Pvull supF

BamHl

pAD1 4 kb Kmr R6K ori RBA for Screening Bacteriophage Lambda Libraries

Figure 6.12.2 Structure of pAD1. This plasmid incorporates the R6K replicon, Kmr, supF, and a polylinker. It is not homologous to ColE1 plasmids.



2. Mix 3 ml lambda top agar, 200 µl of overnight culture, and 106 to 107 pfu of a bacteriophage λ library. Mix well and pour mixture onto a lambda/kanamycin plate. Incubate 7 hr to overnight until total lysis occurs. If more convenient, incubation overnight is perfectly acceptable, because there is no need to harvest the plates just as lysis occurs.

3. Add 3 ml SM and 0.5 ml chloroform to each plate. Swirl lightly. Incubate 2 hr to overnight at room temperature to allow the plates to elute. SM and chloroform are immiscible; swirling them together ensures that the SM is saturated with chloroform, killing any eluted bacteria and minimizing phage adsorption to bacterial debris. The easiest method is to rotate a stack of plates slowly by hand after adding the liquid. Care should be taken not to get chloroform on the petri dish cover, as this can cause fusion of the cover and the plate bottom. If fusion occurs, the cover can be pried from the bottom (e.g., with a screwdriver).

4. Using a nonsterile disposable transfer pipet, harvest the eluate from each plate into a 1.5-ml polypropylene microcentrifuge tube. Although the transfer pipets are polyethylene, they hold chloroform-saturated SM for too short a time-span to be damaged by the solvent. At this stage harvested eluates can be stored ≤1 week at 4°C before continuing the procedure.

5. Add 50 µl of eluate (5 × 108 to 1 × 109 pfu) to 200 µl DM21 culture. Add 3 ml lambda top agar and pour mixture onto a lambda/streptomycin plate. Incubate 7 hr to overnight until plaques form. DM21 is selective (dnaBam lacZam) and resistant to streptomycin. DM75, DM1061, and DM392 (used in later steps) are also streptomycin-resistant, with growth and plating conditions identical to those for DM21.

supF dnaBam

Kmr

ori

lacZam λ-plasmid chimera

tonA co

s

cos

bacterial chromosome

λ imm21 P1 ban

plasmid human probe

cos cos

phage

Figure 6.12.3 Bacterial strain DM21 (outer rectangle) containing λ plasmid chimera with supF integrated (inner circle). DM21 has the genotype lacZYA536(am), dnaB266(am), Smr, hsdR+, hsdM+, tonA− (λ imm21 b515 b519 nin5 att+P1 ban), supO lacZ(am) dnaB(am). The dnaB amber allele selects for λ phage that have supF integrated as shown. SupF also suppress the lacZ amber mutation, yielding blue plaques. Different shadings indicate origins of DNA regions.



Supplement 27

Titer eluates on permissive strain 6. Add 10 µl of each eluate to be titered to 990 µl SM to obtain a 1/100 dilution. Prepare a 100-fold dilution series (to 10−8) in SM. Several random eluates should be titered on the permissive (supF-bearing) strain DM392 to ensure that an appropriate number of phage have been added to the DM21 lawn.

7. Pour a lawn of DM392 (200 µl culture in 3 ml top agar) on a lambda/streptomycin plate. Drop 10-µl aliquots of each eluate dilution onto lawn. Dry 15 min in a forced-air hood (or for longer on bench or in incubator). Incubate 7 hr to overnight until total lysis occurs. This drop-titer procedure is the most convenient method of titering the eluates.

8. Count plaques in the lowest dilution that yields plaques. Convert the result to pfu/ml by multiplying it by the appropriate dilution factor and by a factor of 102. Titration ensures that sufficient phage have been added to the DM21 lawn. Should too many be added (more pfu than cells), the lawn will not materialize due to lysis from without. This phenomenon occurs because every cell that is infected with a bacteriophage will die, even though only cells infected by a phage carrying supF will yield a productive burst that then goes on to infect other cells. In rare cases of lysis from

supF

supF Kmr

ori

Kmr plasmid ori

blue plaque A

co

s

B

cos

A plasmid or recombined phage

B

plasmid human probe

cos A

RBA for Screening Bacteriophage Lambda Libraries

cos

phage

B

PCR primers

cos cos

library phage

Figure 6.12.4 Counterselection. Reversal of the recombination event (which is an equilibrium event) occurs spontaneously. PCR using primers abutting the cloning site of the bacteriophage is employed preparatively to obtain the cDNA without the genomic sequence in pAD1 that was used to retrieve it. The cDNA insert + pAD1 + genomic insert is too large to be amplified by PCR; in contrast, the cDNA insert alone can be amplified. Because there is an equilibrium between the selected and the counterselected phage, the counterselected insert can be amplified directly from the selected blue plaque, which contains a mixture of the two phages. Different shadings indicate origins of DNA regions.



without, the plating should be repeated; either the eluate should be titered or less eluate used. Plaques on DM21 are very small, because suppression of the dnaBam mutation (which is not fully efficient) is required for growth. This makes it difficult to confirm that supF is present via simultaneous suppression of the lacZam mutation by supF; therefore, phage must be transferred to another strain as described in the following steps.

Confirm phages have integrated supF 9. Elute plaques on DM21 (from step 5) into 100 µl SM. Mix: 10 µl eluate 200 µl DM75 culture 3 ml lambda top agar 10 µl 100 mM IPTG 100 µl 2% Xgal in DMF. Plate on lambda/streptomycin plates. Incubate 7 hr to overnight until total lysis occurs. To mix water and DMF, the tubes of top agar must be inverted and righted several times, taking care not to create bubbles. It is best not to prepare more than several tubes at once, because cells do not tolerate the heating block for very long. Light blue plaques are the desired phage containing supF. A larger number of colorless plaques that have not integrated supF will also plate on this strain; these correspond to phage that were not adsorbed originally on DM21 and therefore remain viable. In addition, for a phage such as λgt11, in which interruption of an intact lacZ gene serves as evidence of successful cloning, blue color can result from an intact lacZ gene in the phage. To differentiate between the two, note that the desired supF suppression of the single-copy chromosomal lacZ locus results in a light blue color that extends only to the plaque margins, whereas the high-copy-number lacZ gene on λgt11 yields a dark blue halo that extends past the plaque margins.

10. Elute each plaque thought to contain an integrated supF (from step 9) into 100 µl SM. Pour lawns of DM75 and DM1061 (200 µl culture in 3 ml top agar/IPTG/Xgal, as in previous step) onto separate lambda/streptomycin plates. Drop 10-µl aliquots of each phage eluate onto a lawn of each strain. Incubate 7 hr to overnight until total lysis occurs. This serves to confirm that plaques result from phage with supF rather than lacZ. Phage with supF will be blue on DM75 (lacZam) but colorless on DM1061 (which contains a lacZ deletion), whereas phage carrying an intact lacZ gene will be blue on both strains.

Counterselect with PCR 11. Pour a lawn of 200 µl DM75 in 3 ml top agar onto lambda/streptomycin plate. Drop 10-µl aliquots of phage eluate onto lawn. Incubate 7 hr to overnight, until a single large plaque (“macroplaque”) appears. 12. PCR amplify the cloned product from the macroplaque using primers that abut the EcoRI cloning site of the λ phage vector used to construct the library. This reverses the selection process and accomplishes counterselection (see Fig. 6.12.4). Using the large macroplaque ensures that sufficient template is present. Because the recombination reaction is an equilibrium reaction, a small fraction of phage within a blue macroplaque represent colorless revertants that have excised the pAD1 plasmid and its insert. In contrast, the major product in the macroplaque carries the phage insert, the plasmid, and the insert. Because this is too large to be amplified efficiently by PCR, the technique preferentially yields the desired genic insert from the phage without the unwanted plasmid and its insert.



Supplement 27

13. If desired, sequence the isolated genic clone (UNITS 7.1-7.5) and compare it to a database of known expressed sequences (UNIT 7.7) to obtain information about its possible significance, if available. Repeatedly performing this protocol with different cDNA libraries allows determination of the timing of development and the tissue(s) in which the gene of interest is expressed. The latter can also be determined by using PCR primers specified by the sequence to see if amplification of different cDNA libraries occurs; given the sensitivity of this method, only cDNA library eluates, rather than DNA preparations, need be screened.


Lambda plates 10 g tryptone 5 g NaCl 13 g agar 3 ml 1 M MgCl2 H2O to 1 liter Sterilize by autoclaving. Allow to cool until comfortable to touch. Add antibiotics as needed, mix gently to avoid bubbles, and pour plates. Store up to several months at 4°C. Lambda top agar 10 g tryptone 5 g NaCl 8 g agar 3 ml 1 M MgCl2 H2O to 1 liter Sterilize by autoclaving. Maintain ≤1 month molten at 60°C. L broth 10 g tryptone 5 g NaCl 5 g yeast extract 5 g MgSO4⋅7H2O 1 g glucose 160 ml 12.5 M NaOH (to pH 7.2) H2O to 1 liter Sterilize by autoclaving. Allow to cool until comfortable to touch. Add antibiotics as needed and mix. Store up to several months at 4°C.




Suspension medium (SM) 5.8 g NaCl 2 g MgSO4⋅7H2O 50 ml 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2) 5 ml 2% (w/v) gelatin H2O to 1 liter Sterilize by autoclaving. Store up to several months at 4°C. Gelatin is prepared by adding 2 g gelatin to 100 ml H2O, then autoclaving to dissolve when needed.

Xgal, 2% (v/v) in DMF Dissolve 2% Xgal (5-bromo-4-chloro-3-indolyl-β-D-galactoside; see Table 1.4.2) in dimethylformamide (DMF). Place in polypropylene tube (not polystyrene, which will be dissolved by DMF), wrapped in aluminum foil. Store indefinitely at −20°C (solution will not freeze). COMMENTARY Background Information The recombination-based assay (RBA) permits screening of a complex library or group of libraries with a given probe using only two petri dishes. As a result, the RBA is unparalleled in its efficiency and speed. The crux of the RBA is the insertion of a DNA fragment into a plasmid containing supF, followed by screening of a complex λ library (106 to 107 recombinants) for homology to the fragment. If such homology exists, a recombination event ensues between the inserts in the plasmid and homologous phage at a frequency of 10−2 to 10−3 (see Fig. 6.12.1). As a result of this homology-mediated recombination event, the plasmid with supF is integrated into λ. Genetic selection for λ phage carrying the plasmid with supF results in the isolation of λ phage carrying an insert homologous to the insert in the plasmid. Given the high frequency of homologous recombination (10−2 to 10−3), and the fact that 5 × 108 to 109 pfu can be plated onto a single petri dish, it is feasible to screen rapidly a λ library with a complexity of 106 to 107. Bacterial host characteristics This assay employs a bacterial strain, DM21 (see Figs. 6.12.1 and 6.12.3), that has been constructed to require the presence of supF in λ for phage propagation. As a result, sequences from a λ library that are homologous to a sequence cloned into the supF-bearing plasmid can be isolated on this strain. By screening a λ library carrying human genomic DNA sequences (Lawn et al., 1978), the copy number of a given sequence can be determined analytically. Plasmids carrying repetitive sequences rescue more phage clones from a human

genomic library than do plasmids carrying nonrepetitive sequences (Neve and Kurnit, 1983). By screening a λ library corresponding to the genes encoded by a given tissue with singlecopy sequences, the tissue and time in which a single-copy sequence is transcribed can be determined analytically. Selection for the desired supF-bearing phage is done using the dnaB/P1 ban balanced lethal system. In constructing the host, the dnaB unwinding protein that is normally essential for λ phage growth was replaced by the related, but not identical, P1 ban gene for E. coli growth. The resulting streptomycin-resistant dnaBam P1 ban lacZam host, DK21 (Kurnit and Seed, 1990), was then protected against a contaminating large (?T1) phage infection by a ?tonA mutation to yield the strain DM21 that is used in the protocol (the question mark notes characteristics that are likely but not definite). Analogously, strains LE392, LG75, and MC1061 have each been altered to carry a ?tonA mutation and resistance to streptomycin for use in the protocol, and have been renamed DM392, DM75, and DM1061, respectively (Table 6.12.1). DM21 selects for the plasmid-borne supF by requiring the suppression of an amber mutation in the dnaB gene to permit λ propagation. Furthermore, supF also suppresses the amber mutation in the lacZ gene of DM21, yielding a blue plaque upon addition of the chromogenic substrate Xgal in the presence of IPTG. This makes it possible to discard rare (100 copies; Neve and Kurnit, 1983). 2. Determining tissue- and time-specific transcriptional activity of single-copy fragments and isolating genes. Gene libraries containing >106 independent recombinants are constructed: each corresponds to the totality of genes made in a given tissue at a given time in development. Screening a pool of 106 recombinants from a cDNA library requires only two petri dishes. The phage are first plated on a bacterial lawn carrying the sequence to be tested cloned in a supF-bearing plasmid. Following confluent lysis, 5 × 108 to 5 × 109 pfu are eluted and plated on DM21 to select for phage that have integrated supF. If no phage plaques are observed on DM21, this indicates that the sequence is not transcribed in the tissue

at the developmental stage present when the cDNA libraries were made. If plaques are observed on DM21, this indicates that the sequence is transcribed at that stage. The transcribed sequence is isolated free of the genomic sequence initially used to screen for it by reversing the recombination event (Fig. 6.12.4). In all the libraries used to date—λgt10 (Huynh et al., 1985), λgt11 (Young and Davis, 1983), and Sumo 15A (Kurachi et al., 1989)—the desired sequence is liberated as an EcoRI fragment that can be subcloned. As well as liberating the sequence, the reversal also makes it possible to discard rare nonhomologous (or imperfect) recombination events, which are identified by the fact that they reverse at the same low 10−9 frequency that they occur (for nonhomologous events) and at an intermediate frequency (for partially homologous events). In contrast, homologous recombination events, which can occur in a forward direction at a similar 10−8 frequency (assuming a worst case where a sequence is present only once per genome equivalent in a phage library of 106 recombinants, which is multiplied by a 10−2 chance of recombining if there is homology), reverse at a much higher 10−2 frequency. Thus, reversal of the recombination reaction will yield the cDNA free of the genomic sequence and will simultaneously allow rarer nonhomologous or partially homologous exchange events to be identified and discarded. The RBA can be employed to determine the tissue and time of transcription of candidate genes discovered by other technologies as well as to obtain the gene of interest (in the form of the larger gene sequence that is transcribed). The technique is useful either alone or in combination with other methods for defining single-copy transcribed sequences. If DNA sequencing (as part of the genome initiative) or techniques to define transcribed sequences are used to identify genes, the RBA is still useful for determining the tissue and developmental timing of transcription, as well as for isolating a larger gene of interest. Technologies for defining transcribed sequences include exon trapping/amplification (Nisson and Watkins, 1994; Duyk et al., 1990; Buckler et al., 1991), use of somatic cell hybrids (Liu et al., 1989), and the use of hybridization-based schemes (Hochgeschwender and Brennan, 1994; Hochgeschwender et al., 1989; Kao and Yu, 1991), including hybrid selection (Lovett, 1994; Lovett et al., 1991; Parimoo et al., 1991). The RBA will proceed cooperatively, rather than competitively, with these other methods be-



cause it efficiently accomplishes two necessary tasks: identifying the timing and tissue of gene transcription and isolating a large transcribed sequence.

Critical Parameters Plaque size is a major issue in this assay because plaques on the dnaB am strain DM21 are so small. Fresh λ plates should be used to maximize plaque size, because plaques will be smaller on older (drier) plates; likewise, it is important to plate cells on lambda plates, because plaques will be smaller on richer (e.g., LB) plates. It is essential that there be no homology between the screening plasmid and sequences in the λ libraries (see Background Information). Therefore, screening should be performed solely with R6K supF plasmids, not with ColE1 supF plasmids. Although titering all eluates would be too time-consuming, a few eluates should be titered to ensure that lysis and elution are occurring as expected. This is especially important because a lysed plate may vary from clear to grainy, rendering it difficult to determine visually whether complete lysis has occurred. Eluates should be saved until the DM21 plates have been scored as a precaution in case too many phage have been added, resulting in lysis from without. If this happens, the eluate may be titered or a lesser amount plated on DM21.

Anticipated Results

The abundance of sequences in screened λ libraries should be reflected in the number of phage that plate on DM21. Assuming a recombination rate of 1/500 (the exact number that will depend on the extent of homology), a sequence abundance of 1/106 should yield one plaque on DM21 per 5 × 108 phage plated. A higher abundance should yield a correspondingly greater number of plaques on DM21. If mismatching occurs in an interspersed “saltand-pepper” manner (as for Alu sequences), recombination will be depressed (e.g., ∼1000fold for Alu sequences; Neve et al., 1983).

Time Considerations The major advantage of the RBA is its rapidity: selection can be completed in four days using the following schedule. Day 1, grow bacterial cultures; day 2, add λ library and perform lysis; day 3, elute and plate on DM21; and day 4, identify plaques on DM21. Counterselection takes an additional four days. One day is necessary for elution of

plaques from DM21 that are plated on DM75 with IPTG and Xgal in top agar. A second day is required for elution of putative light blue plaques and confirmatory macroplaque plating on DM75 and DM1061 with IPTG and Xgal. PCR counterselection of macroplaques that are blue on DM75 and colorless on DM1061 takes one day and a final day is necessary to isolate the counterselected PCR band from the gel.

Literature Cited Bolivar, F., Rodriguez, R., Green, P.J., Betlach, M., Heyneker, H.L., Boyer, H.W., Crosa, J., and Falkow, S. 1977. Construction and characterization of new cloning vehicles. Gene 2:95113. Buckler, A.J., Chang, D.D., Graw, S.L., Brook, J.D., Haber, D.A., Sharp, P.A., and Housman, D.E. 1991. Exon amplificaton: A strategy to isolate mammalian genes based on RNA splicing. Proc. Natl. Acad. Sci. U.S.A. 88:4005-4009. Casadaban, M.J. and Cohen, S.N. 1980. Analysis of gene control signals by DNA fusion and cloning in Escherichia coli. J. Mol. Biol. 138:179-207. Duyk, G.M., Kim, S., Myers, R.M., and Cox, D.R. 1990. Exon trapping: A genetic screen to identify candidate transcribed sequences in cloned mammalian genomic DNA. Proc. Natl. Acad. Sci. U.S.A. 87:8995-8999. Guarente, L., Lauer, G., Roberts, T.M., and Ptashne, M. 1980. Improved methods for maximizing expression of a cloned gene: A bacterium that synthesizes rabbit β-globin. Cell 20:543-553. Hanzlik, A.J., Hauser, M.A., Osemlak-Hanzlik, M.M., and Kurnit, D.M. 1993. The recombination-based assay demonstrates that the fragile X sequence is transcribed widely during development. Nature Genet. 3:44-48. Hochgeschwender, U. 1994. Identifying transcribed sequences in arrayed bacteriophage or cosmid libraries. In Current Protocols in Human Genetics (Dracopoli, N., Haines, J.L., Korf, B., Moir, D.T., Morton, C.M., Seidman, C.E., Seidman, J.G., and Smith, D.R., eds.) pp. 6.2.1-6.2.15. John Wiley & Sons, New York. Hochgeschwender, U., Sutcliffe, J.G., and Brennan, M.D. 1989. Construction and screening of a genomic library specific for mouse chromosome 16. Proc. Natl. Acad. Sci. U.S.A. 86:8482-8486. Huynh, T., Young, R.A., and Davis, R.W. 1985. Constructing and screening cDNA libraries in λgt10 and λgt11. In DNA cloning, Vol. II (D. Glover, ed.). IRL Press, Eynsham, U.K. Ikeda, H., Aoki, K., and Naito, A. 1982. Illegitimate recombination mediated in vitro by DNA gyrase of Escherichia coli: Structure of recombinant DNA molecules. Proc. Natl. Acad. Sci. U.S.A. 79:3724-3728. Jankowski, S., Stewart, G.D., Buraczynska, M., Galt, J., Van Keuren, M., and Kurnit, D.M. 1990. Molecular approaches to trisomy 21. Prog. Clin. Biol. Res. 360:79-88.



Supplement 27

Kao, F.-T. and Yu, J.-W. 1991. Chromsome microdissection and cloning in human genome and genetic disease analysis. Proc. Natl. Acad. Sci. U.S.A. 88:1844-1848. King, S.R. and Richardson, J.P. 1986. Role of homology and pathway specificity for recombination between plasmids and bacteriophage λ. Mol. Gen. Genet. 204:141-147. Kurachi, S., Baldori, N., and Kurnit, D.M. 1989. Sumo 15A: A lambda plasmid that permits easy selection for and against cloned inserts. Gene 85:35-43. Kurnit, D.M. and Seed, B. 1990. Improved genetic selection for screening bacteriophage libraries by homologous recombination in vivo. Proc. Natl. Acad. Sci. U.S.A. 87:3166-3169.

Parimoo, S., Patanjali, S.R., Shukla, H., Chaplin, D.D., and Weissman, S.M. 1991. cDNA selection: Efficient PCR approach for the selection of cDNAs encoded in large chromosomal DNA fragments. Proc. Natl. Acad. Sci. U.S.A. 88:9623-9627. Poustka, A., Rackwitz, H.-R., Frischauf, A., Hohn, B., and Lehrach, H. 1984. Selective isolation of cosmid clones by homologous recombination in Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 81:4129-4133. Rubin, C.M., Houck, C.M., Deininger, P.L., and Schmid, C.W. 1980. Partial nucleotide sequence of the 300 nucleotide interspersed repeated human DNA sequences. Nature 284:372-374.

Lawn, R.M., Fritsch, E.H., Parker, R.C., Blake, G., and Maniatis, T. 1978. The isolation and characterization of linked δ- and β-globin genes from a cloned library of human DNA. Cell 15:11571174.

Saiki, R.K., Scharf, S., Faloona, F., Mullis, K.B., Horn, G., Erlich, H.A., and Arnheim, N. 1985. Enzymatic amplification of β-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230:13501354.

Liu, P., Legerski, R., and Siciliano, M.J. 1989. Isolation of human transcribed sequences from human-rodent somatic cell hybrids. Science 246:813-815.

Seed, B. 1983. Purification of genomic sequences from bacteriophage libraries by recombination and selection in vivo. Nucl. Acids Res. 11:24272445.

Lovett, M. 1994. Direct selection of cDNAs using genomic contigs. In Current Protocols in Human Genetics (Dracopoli, N., Haines, J.L., Korf, B., Moir, D.T., Morton, C.M., Seidman, C.E., Seidman, J.G., and Smith, D.R., eds.) pp. 6.3.16.3.15. John Wiley & Sons, New York.

Shen, P. and Huang, H.V. 1986. Homologous recombination in Escherichia coli: Dependence on substrate length and homology. Genetics 112:441-457.

Lovett, M., Kere, J., and Hinton, L.M. 1991. Direct selection: A method for the isolation of cDNAs encoded by large genomic regions. Proc. Natl. Acad. Sci. U.S.A. 88:9628-9632. Lutz, C.T., Hollifield, W.C., Seed, B., Davie, J.M., and Huang, H.V. 1987. Syrinx 2A: An improved λ phage vector designed for screening DNA libraries by recombination in vivo. Proc. Natl. Acad. Sci. U.S.A. 84:4379-4383.

Shen, P. and Huang, H.V. 1989. Effect of base pair mismatches on recombination via the recBCD pathway. Mol. Gen. Genet. 218:358-360. Short, J.M., Fernandez, J.M., Sorge, J.A., and Huse, W.D. 1988. λ ZAP: A bacteriophage λ expression vector with in vivo excision properties. Nucl. Acids Res. 16:7583-7599. Stewart, G.D., Hauser, M.A., Kang, H., McCann, D.P., Osemlak, M.M., Kurnit, D.M., and Hanzlik, A.J. 1991. Plasmids for recombination-based screening. Gene 106:97-101.

Marvo, S.L., King, S.R., and Jaskunas, S.R. 1983. Role of short regions of homology in intermolecular illegitimate recombination events. Proc. Natl. Acad. Sci. U.S.A. 80:2452-2456.

Watt, V.M., Ingles, C.J., Urdea, M.S., and Rutter, W.J. 1985. Homology requirements for recombination in Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 82:4768-4772.

Neve, R.L. and Kurnit, D.M. 1983. Comparison of sequence repetitiveness of human cDNA and genomic DNA using the miniplasmid vector piVX. Gene 23:355-367.

Yanisch-Perron, C., Vieira, J., and Messing, J. 1985. Improved M13 phage cloning vectors and host strains: Nucleotide sequences of the M13mp18 and pUC19 vectors. Gene 33:103-119.

Neve, R.L., Bruns, G.A.P., Dryja, T.P., and Kurnit, D.M. 1983. Retrieval of human DNA from rodent-human genomic libraries by a recombination process. Gene 23:343-354.

Young, R.A. and Davis, R.W. 1983. Efficient isolation of genes by using antibody probes. Proc. Natl. Acad. Sci. U.S.A. 80:1194-1198.

Nisson, P.E. and Watkins, P.C. 1994. Isolation of exons from cloned DNA by exon trapping. In Current Protocols in Human Genetics (Dracopoli, N., Haines, J.L., Korf, B., Moir, D.T., Morton, C.M., Seidman, C.E., Seidman, J.G., and Smith, D.R., eds.) pp. 6.1.1-6.1.14. John Wiley & Sons, New York.

Contributed by David M. Kurnit University of Michigan Medical Center Ann Arbor, Michigan




CHAPTER 7 DNA Sequencing INTRODUCTION For many recombinant DNA experiments, knowledge of a DNA sequence is a prerequisite for its further manipulation. DNA sequencing followed by computer-assisted searching for restriction endonuclease cleavage sites is often the fastest method of obtaining a detailed restriction map (UNITS 3.1-3.3). This information is particularly useful when vectors designed to overexpress proteins or to generate protein fusions are used to subclone a gene of interest (Chapter 16). Computer-assisted identification of protein-coding regions (open reading frames, or ORF) within the DNA sequence, followed by computer-assisted similarity searches of DNA and protein data bases, can lead to important insights about the function and structure of a cloned gene and its product (UNIT 7.7). In addition, the DNA sequence is a prerequisite for a detailed analysis of the 5′ and 3′ noncoding regulatory regions of a gene. DNA sequence information is essential for sitedirected mutagenesis (UNIT 8.1). Small amounts of DNA sequence information (sequence tagged sites, or STS; or expressed sequence tags, or ESTs) are the basis of methods for mapping and ordering large DNA segments cloned into yeast or bacterial artificial chromosomes (CYACs; BACs) or cosmids (Olson et al., 1989; Stephens et al., 1990; Green and Olson, 1990; Adams et al., 1991; Shizuya et al., 1992). EST databases are extremely valuable in gene discovery. DNA sequencing techniques are primarily based on electrophoretic procedures using high-resolution denaturing polyacrylamide gels. These so-called sequencing gels are capable of resolving single-stranded oligonucleotides up to 800 bases in length that differ in size by a single deoxynucleotide. In practice, for a given region to be sequenced, a set of labeled, single-stranded oligonucleotides is generated whose members have one fixed end, and which differ at the other end by each successive deoxynucleotide in the sequence. The key to determining the sequence of deoxynucleotides is to generate, in four separate enzymatic or chemical reactions, all oligonucleotides that terminate at the variable end in

A, T, G, or C. The oligonucleotide products of the four reactions are then resolved on adjacent lanes of a sequencing gel. Because all possible oligodeoxynucleotides are represented among the four lanes, the DNA sequence can be read directly from the four “ladders” of oligonucleotides as shown in Figure 7.0.1. In automated four-color fluorescent DNA sequencers, the oligonucleotide products terminated from each of the four bases (A, C, G, T) are run in a single lane and resolved on the basis that the DNA fragments ending with each of the four bases are labeled with four different fluorescent tags. The practical limit on the amount of information that can be obtained from a set of sequencing reactions is the resolution of the sequencing gel (see UNIT 7.6 for protocols on setting up and running sequencing gels). Current technology allows ∼500 nucleotides of sequence information to be reliably obtained in one set of sequencing reactions, although more information (up to 800 nucleotides) is often obtained using an automated sequencer. Thus, if the region of DNA to be sequenced is 30 kb), such as BAC clones, is very important for closing the gaps in large genomic sequencing and mapping projects. Such large templates are very difficult to

488 nm

sequence due to the smaller number of available priming sites (fewer molecules) compared with plasmid templates of the same mass. The higher sensitivity provided by primers/terminators tagged with ET dyes make it possible to sequence these large-template DNAs directly, which is a significant advantage for large-scale sequencing and mapping projects (Marra et al., 1996; Heiner et al., 1998). The sequencing protocol for large DNA templates are provided in UNIT 7.4

Solid-Phase Sequencing Another recent innovation that is applicable to both manual and automated DNA sequencing is the use of solid-phase capture strategy to generate single-stranded DNA templates (Hultman et al., 1989, 1991; Jones et al., 1991; Kaneoka et al., 1991; Zimmerman et al., 1992).

energy transfer

605 nm hv

donor

spacer

CAGGAAACAGCTATGACC

ET cassette

desired primer sequence

Relative fluorescence intensity

5′−

1.0

acceptor

ETF10F

ETF10G

ETF10T

ETF10R

0.8 0.6 FAM

0.4 JOE 0.2 TAM 0

Introduction

–3′

525

555 580 Wavelength (nm)

ROX 605

Figure 7.0.5 In ET primers, a common donor with a high absorbance at the excitation wavelength harvests energy and transmits it efficiently to acceptor fluorophores that emit in distinct wavelength regions. To avoid fluorescence quenching, the donor and acceptor are separated by a spacer that can be an oligonucleotide or other chemical functionality. The fluorescence emission intensity of current ET primers is 2- to 24-fold higher than that of conventional single dye-labeled primers, leading to high-quality DNA sequencing data.



In this approach, one strand of a doublestranded DNA molecule is biotinylated (e.g., by amplification using PCR in which one of the two primers is biotinylated; Chapter 15). The hemibiotinylated DNA molecule is then bound to streptavidin-ferromagnetic beads. The strands are denatured by treating the beads with alkali and the biotinylated strands are separated from the nonbiotinylated strands using a magnet that traps the bead complex to which the biotinylated strands are bound. Sequencing reactions can be performed using either the biotinylated strand-bead complex or the nonbiotinylated strand preparation as the template. Fluorescent sequencing procedures (both dye primer and dye terminator methods) have disadvantages, most notably false termination and background noise. In the dye primer method, all the extended DNA fragments—including false-terminated fragments—from the primer carry a fluorescent dye, and thus all are detected by the fluorescent sequencer. This causes background noise and results in inaccurate sequencing data. In the dye terminator method, the excess dye-labeled dideoxynucleotides need to be cleaned up completely. Furthermore, if RNAs and nicked DNAs are present in the DNA templates, they will act as primers to generate false termination or high background noise. Thus, a DNA sequencing method that overcomes these disadvantages is desirable. A sequencing chemistry using solid-phasecapturable dideoxynucleotides was recently developed that produces much cleaner sequencing data on both slab gel and capillary array sequencers, eliminating the disadvantages of current dye primer and dye terminator chemistries (Ju et al., 1997; Ju, 1999). The procedure involves coupling fluorescent ET primers that produce high fluorescent signals with solid phase–capturable terminators such as biotinylated dideoxynucleotides. After the sequencing reaction, the extension DNA fragments are captured with magnetic beads coated with streptavidin, while the other components in the sequencing reaction are washed away. Only the pure dideoxynucleotide terminated extension products are released from the magnetic beads and loaded on the sequencing gel, producing high-quality data.

Sequencing with Mass Spectrometry Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDITOF MS; UNITS 10.21 & 10.22) has recently been explored for DNA sequencing (Koster et al.,

1996; Monforte et al., 1997; Fu et al., 1998). The Sanger dideoxy procedure is used to generate the DNA sequencing fragments and no labels are required. The fragments are mixed with a matrix compound, 3-hydroxypicolinic acid, forming microcrystals on a flat plate. The sample plate is then placed in the vacuum chamber of the mass spectrometer. Upon irradiation by a laser light, a microscopic portion of the matrix molecules desorbs off the plate, entraining the DNA fragments. In an electric field, the charged DNA molecules (ions) are accelerated toward the detector, which measures the mass of the fragments based on their time of flight, which is inversely proportional to their mass. Since the mass of each nucleotide (A, C, G, T) is different, the mass difference between adjacent peaks in the mass spectrum is used to establish the sequence of the DNA templates. For example, if the mass difference between the two adjacent peaks in a mass spectrum of an unknown DNA is the mass of “A” base, then the peak with the higher mass is identified as an “A”. Thus simple computation software is all that is needed to assemble the sequence of the DNA from its mass spectrum. Compared to gel-based sequencing systems, MS produces very high resolution of the sequencing fragments (sometimes as good as 1 Da) and extremely fast separation, on a time scale of microseconds. The high resolution allows accurate detection of mutations and heterozygosity. Another advantage of sequencing with mass spectrometry is that the compressions associated with gel-based systems are completely eliminated. However, in order for accurate measurements of the masses of the sequencing DNA fragments to be obtained, the sample must be free from alkaline and alkaline earth salts. The samples must therefore be desalted before the MS analysis. Solid-phase procedures using either biotinylated primer or immobilized templates are generally used for desalting (Koster et al., 1996; Monforte and Becker, 1997). Both approaches introduce false-terminated DNA fragments into the mass detector. An elegant method to obtain pure sequencing fragments is to use solid phase–capturable dideoxynucleotides—such as biotinylated terminators—in the Sanger reactions to generate sequencing fragments. In this procedure, only the correctly terminated DNA fragments are isolated by streptavidin-coated beads, which are subsequently released and loaded on the mass spectrometer, resulting in accurate sequencing data (Ju et al., 1997; Ju, 1999). The current limit of mass spectrometry for sequenc-

DNA Sequencing


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ing is in the 100-bp range. Optimized solidphase sequencing chemistry plus improvement in detector sensitivity for large DNA fragments will further improve mass spectrometry for DNA sequencing.

Sequencing by Hybridization Sequencing by hybridization (SBH) makes use of an array of all possible short oligonucleotides to identify a segment of sequences present in an unknown DNA (Drmanac et al., 1989, 1993). This can be clearly explained by the following example. A pentanucleotide 5′ CAGTA-3′, with a complementary sequence of 5′-TACTG-3′ is the sequence that need to be determined from a pool of all possible trinucleotides (43 = 64). This pentanucleotide will specifically hybridize only with the complementary trinucleotides TAC, ACT, and CTG, revealing the presence of these blocs in the complementary sequence. From this the sequence 5′-TACTG-3′ can be reconstructed. Thus with a library of 8- to 10-mer oligonucleotides, much larger segments of DNA sequences can be established. Computational approaches are then used to assemble the complete sequence. In the current state of the art of this technology, SBH has been used for detecting mutations and for resequencing a genome as well as for detecting polymorphisms (Chee et al., 1996; Drmanac et al., 1998). Robust de novo sequencing has not yet been demonstrated. Potential applications of SBH include physical mapping (ordering) of overlapping DNA clones, sequence checking, DNA fingerprinting comparisons of normal and diseasecausing genes, and identification of DNA fragments with particular sequence motifs in complementary DNA and genomic libraries.

COMPUTER ANALYSIS

Introduction

Once the gels are run and autoradiograms are obtained, computer software is practically indispensable for analysis of the sequence information. Computer software can assist at three stages. First, DNA sequence data can be entered into a computer data base either by “reading” the sequencing gels manually with a digitizer system or by using an automated gel scanner. Second, several software packages are available for detecting overlaps in sequence data and then assembling contiguous DNA sequences (contigs) from individual templates. Third, computer assistance is indispensable for analyzing final sequence data, e.g., in finding open reading frames or finding homologies to other sequences present in the nucleotide (UNIT

19.2) or protein databases (UNIT 19.3). UNIT 7.7 provides an overview of software and technology currently available.

LITERATURE CITED Adams, M.D., Keller, J.M., Gocayne, J.D., Dubnick, M., Polymeropoulos, M.H., Xaio, H., Merril, C.R., Wu, A., Olde, B., Moreno, R.F., Kerlavage, A.R., McCombie, W.R., and Venter, J.C. 1991. Complementary DNA sequencing: Expressed sequence tags and human genome project. Science 252:1651-1656. Adams, S. and Blakesley, R. 1991. Linear amplification sequencing. Focus (BRL) 13:56-57. Ambrose, B. and Pless, R. 1987. DNA sequencing: Chemical methods. Methods Enzymol. 52:522538. Ansorge, W., Sproat, B., Stegemann, J., Schwager, C., and Zenke, M. 1987. Automated DNA sequencing: Ultrasensitive detection of fluorescent bands during electrophoresis. Nucl. Acids Res. 15:4593-4602. Applied Biosystems. 1989 to present. Model 370 and 370A Automated Sequencer User Bulletin. Applied Biosystems, Foster City, Calif. Arnold, W. and Puhler, A. 1988. A family of high copy number plasmid vectors with single end-label sites in rapid nucleotide sequencing. Gene 70:171-179. Beck, S., O’Keefe, T.O., Coull, J.M., and Koster, H. 1989. Chemiluminescent detection of DNA: Application for DNA sequencing and hybridization. Nucl. Acids Res. 17:5115-5123. Brumbaugh, J., Middendorf, L., Grone, D., and Ruth, J. 1988. Continuous on-line DNA sequencing using oligonucleotide primers with multiple fluorophores. Proc. Natl. Acad. Sci. U.S.A. 85:5610-5614. Carothers, A.M., Urlab, G., Mucha, J., Grunburger, D., and Chasin, L.A. 1989. Point mutation analysis in a mammalian gene: Rapid preparation of total RNA, PCR amplification of cDNA and Taq sequencing by a novel method. BioTechniques 7:494-499. Chee, M., Yang, R., Hubbell, E., Berno, A., Huang, X.C., Stern, D., Winkler, J., Lockhart, D.J., Morris, M.S., and Fodor, S.P. 1996. Accessing genetic information with high-density DNA arrays. Science 274:610-614. Chen, E.Y. and Seeburg, P.H. 1985. Supercoil sequencing: A fast and simple method for sequencing plasmid DNA. DNA (N.Y.) 4:165-170. Church, G. and Gilbert, W. 1984. Genomic sequencing. Proc. Natl. Acad. Sci. U.S.A. 81:1991-1995. Church, G. and Kiefer-Higgins, S. 1988. Multiplex DNA sequencing. Science 240:185-188. Connel, C., Fung, S., Heiner, C., Bridgeham, J., Chakerian, V., Heron, E., Jones, B., Menchen, S., Mordan, W., Raff, M., Racknor, M., Smith, L., Springer, J., Woo, S., and Hunkapiller, M. 1987. Automated DNA sequence analysis. BioTechniques 5:342-348.



Craxton, M. 1991. Linear amplification sequencing: A powerful method for sequencing DNA. Methods, a companion to Methods Enzymol. 3:20-26. Creasey, A., D’Angio, L.M., Dunne, T., Kissinger, C., O’Keefe, T., Perry-O’Keefe, H., Moran, L., Roskey, M., Shildkraut, I., Sears, L., and Slatko, B. 1991. Application of a novel chemiluminescent-based DNA detection method to single-vector and multiplex DNA sequencing. BioTechniques 11:102-109. D’Cunha, J., Berson, B.J., Brumly, R.L., Wagner, P.R., and Smith, L.M. 1990. An automated instrument for the performance of enzymatic DNA sequencing reactions. BioTechniques 9:80-90. Drmanac, R., Labat, I., Brukner, I., and Crkvenjakov, R. 1989. Sequencing of megabase-plus DNA by hybridization: Theory of the method. Genomics 4:114-128. Drmanac, R., Drmanac, S., Strezoska, Z., Paunesku, T., Labat, I., Zeremski, M., Snoddy, J., Funkhouser, W.K., Koop, B., Hood, L., et al. 1993. DNA sequence determination by hybridization: A strategy for efficient large-scale sequencing. Science 260:1649-1652. Drmanac, S., Kita, D., Labat, I., Hauser, B., Schmidt, C., Burczak, J.D., and Drmanac, R. 1998. Accurate sequencing by hybridization for DNA diagnostics and individual genomics. Nature Biotechnol. 16:54-58. Eckert, R. 1987. New vectors for rapid sequencing of DNA fragments by chemical degradation. Gene 51:245-252. Evans, S. 1991. Millipore’s system speeds up DNA sequencing and eliminates radioactivity. Genet. Eng. News 14:29-41. Frank, R., Bosserhoff, A., Boulin, C. Epstein, A. Gausepohl, H., and Ashman, K. 1988. Automation of DNA sequencing reactions and related techniques: A workstation for micromanipulation of liquids. Bio/Technology 6:1211-1213. Fu, D.J., Tang, K., Braun, A., Reuter, D., DarnhoferDemar, B., Little, D.P., O’Donnell, M.J., Cantor, C.R., and Koster, H. 1998. Sequencing exons 5 to 8 of the p53 gene by MALDI-TOF mass spectrometry. Nature Biotechnol. 16:381-384. Fujita, M., Usui, S., Kiyama, M., Kambara, H., Murakawa, K., Suzuki, S., Sambe, H., and Takachi, K. 1990. Chemical robot for enzymatic reactions and extraction processes of DNA in DNA sequence analysis. BioTechniques 9:584591. Green, E.D. and Olson, M.V. 1990. Chromosomal region of the cystic fibrosis gene in yeast artificial chromosomes: A model for human genome mapping. Science 250:92-98. Haltiner, M., Kempe, T., and Tjian, R. 1985. A novel strategy for constructing clustered point mutations. Nucl. Acids Res. 13:1015-1025. Hattori, M. and Sakaki, Y. 1986. Dideoxy DNA sequencing method using denatured plasmid templates. Anal. Biochem. 152:232-238.

Hawkins, T.L., McKernan, K.J., Jacotot, L.B., MacKenzie, J.B., Richardson, P.M., and Lander, E.S. 1997. A magnetic attraction to highthroughput genomics. Science 276:1887-1889 Heiner, C.R., Hunkapiller, K.L., Chen, S.M., Glass, J.I., and Chen, E.Y. 1998. Sequencing multimegabase-template DNA with BigDye terminator chemistry. Genome Res. 8:557-561. Huang, X.C., Quesada, M.A., and Mathies, R.A. 1992. DNA sequencing using capillary array electrophoresis. Anal. Chem. 64:2149-2154. Hultman, T., Stahl, S., Hornes, E., and Uhlen, M. 1989. Direct solid phase sequencing of genomic and plasmid DNA using magnetic beads as solid support. Nucl. Acids Res. 17:4937-4946. Hultman, T., Bergh, S., Moks, T., and Uhlen, M. 1991. Bidirectional solid phase sequencing of in vitro amplified plasmid DNA. BioTechniques 10:84-93. Johnston-Dow, E., Mardis, E., Heiner, C., and Roe, B.A. 1987. Optimized methods for fluorescent and radiolabeled DNA sequencing. BioTechniques 5:754-765. Jones, D. S., Schofield, J.P., and Vaudin, M. 1991. Fluorescent and radioactive solid phase dideoxy sequencing of PCR products in microtitre plates. J. DNA Seq. Map. 1:279-283. Ju, J. 1999. Nucleic Acid Sequencing with Solid Phase Capturable Terminators. U.S. Patent no. 5,876,936. Ju, J., Ruan, C., Fuller, C.W., Glazer, A.N., and Mathies, R.A. 1995. Energy transfer fluorescent dye-labeled primers for DNA sequencing and analysis. Proc. Natl. Acad. Sci. USA. 92:43474351. Ju, J., Glazer, A.N., and Mathies, R.A. 1996. Energy transfer primers: A new fluorescence labeling paradigm for DNA sequencing and analysis. Nature Med. 2:246-249. Ju, J., Yan, H., Zaro, M., Doctolero, M., Goralski, T., Konrad, K., Lachenmeier, E., and Cathcart., R. 1997. DNA sequencing with solid phase capturable terminators Microb. Comp. Genomics 2:223. Kambara, H., Nishikawa, T., Katayama, K., and Yamaguchi, T. 1988. Optimization of parameters in a DNA sequenator using fluorescence detection. Bio/Technology 6:816-821. Kaneoka, H., Lee, D.R., Hsu, K.-C., Sharp, G.C., and Hoffman, R.W. 1991. Solid phase DNA sequencing of allele specific polymerase chain reaction amplified HLA-DR genes. BioTechniques 10:30-40. Kheterpal, I. and Mathies, R.A. 1999. Capillary array electrophoresis DNA sequencing. Anal. Chem. 71:31A-37A. Koster, H., Tang. K., Fu, D.J., Braun, A., van den Boom, D., Smith, C.L., Cotter, R.J., and Cantor, C.R. 1996. A strategy for rapid and efficient DNA sequencing by mass spectrometry. Nature Biotechnol. 14:1123-1128. DNA Sequencing


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Krishnan, B.R., Blakesley, R.W., and Berg, D.E. 1991. Linear amplification DNA sequencing directly from single phage plaques and bacterial colonies. Nucl. Acids. Res. 19:1153. Lee, L.G., Spurgeon, S.L., Heiner, C.R., Benson, S.C., Rosenblum, B.B., Menchen, S.M., Graham, R.J., Constantinescu, A., Upadhya, K.G., and Cassel, J.M. 1997. New energy transfer dyes for DNA sequencing. Nucl. Acids Res. 25:28162822. Mardis, E.R. and Roe, B.A. 1989. Automated methods for single-stranded DNA isolation and dideoxynucleotide DNA sequencing reactions on a robotic workstation. BioTechniques 7:840-850. Marra, M., Weinstock, L.A., and Mardis, E.R. 1996. End sequence determination from large insert clones using energy transfer fluorescent primers. Genome Res. 6:1118-1122. Martin, W., Warmington, J., Galinski, B., Gallager, M., Davies, R., Beck, M., and Oliver, S. 1985. A system to perform the Sanger dideoxy sequencing reactions. Bio/Technology 3:911-915. Martin, C., Bresnick, L., Juo, R.-R., Voyta, J.C., and Bronstein, I. 1991. Improved chemiluminescence DNA sequencing. BioTechniques 11:110113. Maxam, A.M. and Gilbert, W. 1977. A new method for sequencing DNA. Proc. Natl. Acad. Sci. U.S.A. 74:560-564. Maxam, A.M. and Gilbert, W. 1980. Sequencing end-labeled DNA with base-specific chemical cleavages. Methods Enzymol. 65:499-559. Messing, J. 1983. New M13 vectors for cloning. Methods Enzymol. 101:20-78. Messing, J. 1988. M13, the universal primer and the polylinker. Focus (BRL) 10:21-26. Middendorf, L., Brumbaugh, J., Grone, D., Morgan, C., and Ruth, J. 1988. Large scale DNA sequencing. Am. Biol. Lab. (August) 14-22. Monforte, J.A. and Becker, C.H. 1997. Highthroughput DNA analysis by time-of-flight mass spectrometry. Nature Med. 3:360-362. Murray, V. 1989. Improved double-stranded DNA sequencing using the linear polymerase chain reaction. Nucl. Acids Res. 17:8889. Olson, M., Hood, L., Cantor, C., and Botstein, D. 1989. A common language for physical mapping of the human genome. Science 245:1434-1435. Prober, J., Trainor, G., Dam, R., Hobbs, F., Robertson, C., Zagursky, R., Cocuzza, R., Jensen, M., and Baumeister, K. 1987. A system for rapid DNA sequencing with fluorescent chain-terminating dideoxynucleotides. Science 238:336341. Rosenthal, A., Sproat, B., Voss, H., Stegemann, J., Schwager, C., Erfle, H., Zimmerman, J., Courelle, C., and Ansorge, W. 1990. Automated sequencing of fluorescently labeled DNA by chemical degradation. J. DNA Seq. Map. 1:6371.

Rubin, J. and Schmid, C. 1980. Pyrimidine-specific chemical reactions useful for DNA sequencing. Nucl. Acids Res. 8:4613-4619. Salas-Solano, O., Carrilho, E., Kotler, L., Miller, A.W., Goetzinger, W., Sosic, Z., and Karger, B.L. 1998. Routine DNA sequencing of 1000 bases in less than one hour by capillary electrophoresis with replaceable linear polyacrylamide solutions. Anal. Chem. 70:3996-4003. Sanger, F., Nicklen, S., and Coulson, A.R. 1977. DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. U.S.A. 74:5463-5467. Sanger, F., Coulson, A.R., Barrell, B.G., Smith, A.J.M., and Roe, B.A. 1980. Cloning in singlestranded bacteriophage as an aid to rapid DNA sequencing. J. Mol. Biol. 143:161-178. Sears, L., Moran, L., Kissinger, C., Creasey, T., Perry-O’Keefe, H., Roskey, M., Sutherland, E., and Slatko, B. 1992. CircumVent thermal cycle sequencing and alternative manual and automated DNA sequencing protocols using the highly thermostable VentR (exo−) DNA polymerase. BioTechniques. 13:626-633. Shizuya, H., Birren, B., Kim, U.-J., Mancino, V., Slepak, T., Tachiiri, Y., and Simon, M. 1992. Cloning and stable maintenance of 300-kilobase-pair fragments of human DNA in Escherichia coli using an F-factor-based vector. Proc. Nat. Acad. Sci. U.S.A. 89:8794-8797. Smith, L., Fung, S., Hunkapiller, M., Hunkapiller, T., and Hood, L. 1985. The synthesis of oligonucleotides containing an aliphatic amino group at the 5′ terminus: Synthesis of fluorescent DNA primers for use in DNA sequencing analysis. Nucl. Acids Res. 13:2399-2412. Smith, L., Sanders, J., Kaiser, R., Hughes, P., Dodd, C., Heiner, C., Kent, S., and Hood, L. 1986. Fluorescence detection in automated DNA sequence analysis. Nature 321:674-679. Smith, V., Brown, C.M., Banker, A.T., and Barrell, B.G. 1990. Semiautomated preparation of DNA template for large scale sequencing projects. J. DNA Seq. Map. 1:73-78. Stephens, J.C., Cavanaugh, M.L., Gradie, M.I., Mador, M.L., and Kidd, K.K. 1990. Mapping the human genome: Current status. Science 250:237-244. Tabor, S. and Richardson, C.C. 1987a. Selective oxidation of the exonuclease domain of bacteriophage T7 DNA polymerase. J. Biol. Chem. 262:15330-15333. Tabor, S. and Richardson, C.C. 1987b. DNA sequence analysis with a modified bacteriophage T7 DNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 84:4767-4771. Tabor, S. and Richardson, C.C. 1989a. Selective inactivation of the exonuclease activity of bacteriophage T7 DNA polymerase by in vitro mutagenesis. J. Biol. Chem. 264:6447-6458.

Introduction



Tabor, S. and Richardson, C.C. 1989b. Effect of manganese ions on the incorporation of dideoxynucleotides by bacteriophage T7 DNA polymerase and Escherichia coli DNA polymerase I. Proc. Natl. Acad. Sci. U.S.A. 86:4076-4080. Tabor, S. and Richardson, C.C. 1990. DNA sequence analysis with a modified bacteriophage T7 DNA polymerase: Effect of pyrophosphorolysis and metal ions. J. Biol. Chem. 265:83228328. Tabor, S., Huber, H., and Richardson, C.C. 1987. Escherichia coli thioredoxin confers processivity of the DNA polymerase activity of the gene 5 protein of bacteriophage T7. J. Biol. Chem. 262:16212-16223. Tizard, R., Cate, R.L., Ramachandran, K.L., Wysek, M., Voyta, J.C., Murphy, O.J., and Bronstein, I. 1990. Imaging of DNA sequences with chemiluminescence. Proc. Natl. Acad. Sci. U.S.A. 87:4514-4518. Volckaert, G., de Vleeschouwer, E., Blocker, H., and Frank, R. 1984. A novel type of cloning vector for ultrarapid chemical degradation sequencing of DNA. Gene Anal. Tech. 1:52-59. Wilson, R., Yuen, A., Clark, S., Spence, C., Arakelian, P., and Hood, L. 1988. Automation of dideoxynucleotide DNA sequencing reactions using a robotic workstation. BioTechniques 6:776-787.

Zagursky, R.J., Conway, P.S., and Kashdan, M.A. 1991. Use of 33P for Sanger DNA sequencing. BioTechniques 11:36-38. Zimmerman, J., Voss, H., Schwager, C., Stegemann, J. Erfle, H., and Ansorge, W. 1989. Automated preparation and purification of M13 templates for DNA sequencing. Methods Mol. Cell Biol. 1:29-34. Zimmerman, J., Dietrich, T., Voss, H., Erfle, H., Schwager, C., Stegemann, J., Hewitt, N., and Ansorge, W. 1992. Fully automated Sanger sequencing protocol for double-stranded DNA. Methods Mol. Cell Biol. 3:39-42.

KEY REFERENCES Maxam and Gilbert, 1977. See above. Describes the chemical cleavage method. Sanger et al., 1977. See above. Describes the traditional Sanger procedure. Tabor and Richardson, 1987b. See above. Describes the labeling/termination method. Sears et al., 1992. See above. Describes the thermal cycle sequencing procedure.

Young, A. and Blakesley, R. 1991. Sequencing plasmids from single colonies with the dsDNA cycle sequencing system. Focus (BRL) 13:137.

Frederick M. Ausubel, Lisa M. Albright, and Jingyue Ju

Zagursky, R. and McCormick, R. 1990. DNA sequencing separations in capillary gels on a modified commercial DNA sequencing instrument. BioTechniques 9:74-79.

The chapter editors wish to acknowledge the substantial assistance of Richard L. Eckert of Case Western Reserve and Barton E. Slatko of New England Biolabs.

Zagursky, R., Baumeister, K., Lomax, N., and Berman, M. 1985. Rapid and easy sequencing of large double-stranded DNA and supercoiled plasmid DNA. Gene Anal.Tech. 2:89-94.

DNA Sequencing


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DNA Sequencing Strategies This unit contains a general discussion of factors that should be considered before embarking on a DNA sequencing project. In general, any sequencing strategy should include plans for sequencing both strands of the DNA fragment. Complementary strand confirmation leads to higher accuracy, especially when sequencing regions where artifacts such as “compressions” are a problem. Sequencing the opposite strand is often required to obtain accurate data for such regions. The most commonly used methods for generating appropriately sized DNA fragments for dideoxy and chemical sequencing are discussed below. The biochemistry underlying these procedures, as well as how to choose between these and alternative sequencing methods, are discussed in the introduction to this chapter.

DIDEOXY SEQUENCING Planning for Dideoxy Sequencing Sequencing determination of a fragment of 100 µg), the lipid can be toxic. For each particular liposome mixture tested, it is important to vary the amount as indicated in Table 9.1.3. Concentration of DNA. In many of the cell types tested, relatively small amounts of DNA are effectively taken up and expressed. In fact, higher levels of DNA can be inhibitory in some cell types with certain liposome preparations. In the optimization protocol outlined in Table 9.1.3, the standard reporter vector pSV2CAT is used; however, any plasmid DNA whose expression can be easily monitored would be suitable.

Introduction of DNA into Mammalian Cells


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Table 9.1.3

Dish (35-mm) 1 2 3 4 5 6 7 8 9 10

Optimization of Liposome-Mediated Transfection

pSV2CAT (µg) 0.1 0.1 0.1 0.1 0.1 0.5 0.5 0.5 0.5 0.5

Liposomes (µl) 1 2 4 8 12 1 2 4 8 12

Dish (35-mm)

pSV2CAT (µg)

11 12 13 14 15 16 17 18 19 20

5 5 5 5 5 10 10 10 10 10

Liposomes (µl) 5 10 15 20 30 5 10 15 20 30

Time of incubation. When the optimal amounts of lipid and DNA have been established, it is desirable to determine the length of time required for exposure of the liposome-DNA complex to the cells. In general, transfection efficiency increases with time of exposure to the liposome-DNA complex, although after 8 hr, toxic conditions can develop. HeLa or BHK-21 cells typically require ∼3 hr incubation with the liposome-DNA complex for optimal tranfection, while CV-1 and COS-7 cells require 5 hr of exposure.

UNIT 9.1

Calcium Phosphate Transfection This unit presents two methods of calcium phosphate–based eukaryotic cell transfection that can be used for both transient and stable (UNIT 9.5) transfections. In these protocols, plasmid DNA is introduced to monolayer cell cultures via a precipitate that adheres to the cell surface. A HEPES-buffered solution is used to form a calcium phosphate precipitate that is directly layered onto the cells (see Basic Protocol). For some cells, shocking the cells with glycerol or DMSO (see Support Protocol) improves transfection efficiency. In the alternate high-efficiency method, a BES-buffered system is used that allows the precipitate to form gradually in the medium and then drop onto the cells (see Alternate Protocol). The alternate method is particularly efficient for stable transformation of cells with circular plasmid DNA. For transformation with linear plasmid or genomic DNA, or for transient expression, however, the Alternate Protocol is comparable to the Basic Protocol. Both methods of transfection require very high-quality plasmid DNA. Factors that can be optimized for calcium phosphate transfections are presented in the introduction to Section I, and protein expression strategies are discussed in Chapter 16. Additional details of mammalian cell culture are given in APPENDIX 3F.

BASIC PROTOCOL

Calcium Phosphate Transfection

TRANSFECTION USING CALCIUM PHOSPHATE–DNA PRECIPITATE FORMED IN HEPES A precipitate containing calcium phosphate and DNA is formed by slowly mixing a HEPES-buffered saline solution with a solution containing calcium chloride and DNA. This precipitate adheres to the surface of cells and should be visible in the phase contrast microscope the day after transfection. Depending on the cell type, up to 10% of the cells on a dish will take up the DNA precipitate through an as yet undetermined mechanism. Glycerol or dimethyl sulfoxide shock increases the amount of DNA absorbed in some cell types (see Support Protocol).

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Materials Exponentially growing eukaryotic cells (e.g., HeLa, BALB/c 3T3, NIH 3T3, CHO, or rat embryo fibroblasts) Complete medium (depending on cell line used) CsCl-purified plasmid DNA (10 to 50 µg per transfection) 2.5 M CaCl2 (see recipe) 2× HEPES-buffered saline (HeBS; see recipe) PBS (APPENDIX 2) 10-cm tissue culture dishes 15-ml conical tube Additional reagents and equipment for ethanol precipitation (UNIT 2.1A) and mammalian cell tissue culture (APPENDIX 3F) NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Split exponentially growing eukaryotic cells into 10-cm tissue culture dishes the day before transfection. Feed cells with 9.0 ml complete medium 2 to 4 hr prior to precipitation. When transfecting adherent cells that double every 18 to 24 hr, a 1:15 split from a confluent dish generally works well. On the day of the transfection, it is important that cells are thoroughly separated on the dish, as the ability to take up DNA is related to the surface area of the cell exposed to the medium. Cells should be split in a manner that accomplishes this. The desired density of cells on dishes to be transfected will vary with cell type and the reason for doing the transfection. The optimal density is that which produces a near confluent dish when the cells are harvested or split into selective medium.

2. Ethanol precipitate the DNA to be transfected and air dry the pellet by inverting the microcentrifuge tube on a fresh Kimwipe inside a tissue culture hood. Resuspend the pellet in 450 µl sterile water and add 50 µl of 2.5 M CaCl2. The amount of DNA that is optimal for transfection varies from 10 to 50 ìg per 10-cm plate, depending on the cell line to be transfected. DNA to be transfected should be purified twice by CsCl gradient centrifugation (UNIT 1.7). DNA can also be prepared using column procedures (UNIT 2.1B). Some column procedures produce DNA that does not transfect well, so column-purified DNA should be tested and compared to CsCl-purified DNA for transfection efficiency. Supercoiled DNA works well in transfections. Impurities in the DNA preparation can be deleterious to transfection efficiency. A description of how to optimize the amount of DNA to transfect and other parameters of calcium phosphate–mediated transfection is provided in the discussion of Optimization of Transfection. Ethanol precipitation sterilizes the DNA to be transfected. For transfections that will be harvested within 3 to 4 days (transient analysis), this is not necessary. For transient experiments, many researchers make a 450-ìl aqueous solution containing the DNA directly, without ethanol precipitation. If this is done, care should be taken to keep the amount of Tris in the solution to a minimum, as Tris may alter the pH of the precipitate and therefore reduce transfection efficiency.

3. Place 500 µl of 2× HeBS in a sterile 15-ml conical tube. Use a mechanical pipettor attached to a plugged 1- or 2-ml pipet to bubble the 2× HeBS and add the DNA/CaCl2



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Figure 9.1.1 Formation of calcium phosphate precipitate.

solution dropwise with a Pasteur pipet (see Fig. 9.1.1). Immediately vortex the solution for 5 sec. If no mechanical pipettor is available, the solution can be bubbled by blowing through rubber tubing that is attached to a pipet via a filter. The filter is necessary to maintain sterility. This does not give as reproducible results as the mechanical pipettor.

4. Allow precipitate to sit 20 min at room temperature. 5. Use a Pasteur pipet to distribute the precipitate evenly over a 10-cm plate of cells and gently agitate to mix precipitate and medium. 6. Incubate the cells 4 to 16 hr under standard growth conditions. Remove the medium. Wash cells twice with 5 ml PBS and feed cells with 10 ml complete medium. The amount of time that the precipitate should be left on the cells will vary with cell type. For hardy cells such as HeLa, NIH 3T3, and BALB/c 3T3, the precipitate can be left on for 16 hr. Other cell types will not survive this amount of exposure to the precipitate. See discussion of Optimization of Transfection for optimization of this step as well as for a discussion of how to determine whether glycerol shock is useful.

7. For transient analysis, harvest the cells at the desired time point (UNITS 9.6-9.8 & 14.6). For stable transformation, allow the cells to double twice before plating in selective medium (UNIT 9.5). Calcium Phosphate Transfection

9.1.6 Supplement 63


GLYCEROL/DMSO SHOCK OF MAMMALIAN CELLS The Basic Protocol works well for cell lines such as HeLa, BALB/c 3T3, NIH 3T3, and rat embryo fibroblasts. Transfection efficiency in some cell lines, such as CHO DUKX, is dramatically increased by “shocking” the cells with either glycerol or DMSO. Precipitates are left on the cell for only 4 to 6 hr, and the cells are shocked immediately after removal of the precipitate.

SUPPORT PROTOCOL

Additional Materials (also see Basic Protocol) 10% (v/v) glycerol solution or DMSO in complete medium, sterile PBS (APPENDIX 2), sterile Replace step 6 of the Basic Protocol with the following: 6a. Incubate the cells 4 to 6 hr and remove the medium. Add 2.0 ml of a sterile 10% glycerol solution. Let the cells sit 3 min at room temperature. Alternatively, 10% or 20% DMSO can be used. DMSO tends to be somewhat less harmful to the cells, but also may not work as well.

6b. Add 5 ml of PBS to the glycerol solution on the cells, agitate to mix, and remove the solution. Wash twice with 5 ml of PBS. Feed the cells with complete medium. It is important to dilute the glycerol solution on the cells with PBS before removing the glycerol solution so the cells do not stay in glycerol too long. Excessive exposure to glycerol will kill cells.

HIGH-EFFICIENCY TRANSFECTION USING CALCIUM PHOSPHATE–DNA PRECIPITATE FORMED IN BES

ALTERNATE PROTOCOL

A solution of calcium chloride, plasmid DNA, and N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES) buffer, pH 6.95, is added to a plate of cells containing culture medium. The plates are incubated overnight while a calcium phosphate–DNA complex forms gradually in the medium under an atmosphere of 3% CO2. With this method, 10% to 50% of the cells on a plate stably integrate and express the transfected DNA. Transient expression under these conditions is comparable to that obtained with the Basic Protocol. Glycerol or DMSO shock does not increase the number of cells transformed. Materials Exponentially growing mammalian cells (see Critical Parameters) Complete medium: Dulbecco modified Eagle medium containing 10% (v/v) fetal bovine serum (FBS) CsCl-purified plasmid DNA TE buffer, pH 7.4 (APPENDIX 2) 2.5 M CaCl2 (see recipe) 2× BES-buffered solution (BBS; see recipe) PBS (APPENDIX 2) Selection medium (UNIT 9.5; optional) 10-cm tissue culture dishes 35°C, 3% CO2 humidified incubator 35° to 37°C, 5% CO2 humidified incubator Fyrite gas analyzer (optional; Fisher Scientific or Curtin Matheson) NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly.



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1. Seed exponentially growing mammalian cells at 5 × 105 cells/10-cm tissue culture dish in 10 ml complete medium the day prior to transfection. There should be 10- to 100-fold higher when the BESbased protocol is used, with 10% to 50% of the cells on a plate stably transformed (Chen and Okayama, 1987). Transient expression is comparable in the Basic and Alternate Protocols (Chen and Okayama, 1988).

Time Considerations For the Basic Protocol, preparation of twelve DNA precipitates and addition of the precipitates to the cells takes 1 to 2 hr. Without the ethanol precipitation step, the procedure can be done in 1 hr. With practice, the actual mixing of the CaCl2 and 2× HeBS solutions will take ∼1 min. This means that up to eighteen precipitates can be made before the first precipitate is ready to apply to the cells.

For the Alternate Protocol, no ethanol precipitation step has been necessary for either transient or stable transfections. It takes slightly less time than the Basic Protocol because the 2× BBS does not need to be added dropwise to the calcium chloride–DNA solution.

Literature Cited Chen, C. and Okayama, H. 1987. High efficiency transformation of mammalian cells by plasmid DNA. Mol. Cell. Biol. 7:2745-2752. Chen, C. and Okayama, H. 1988. Calcium phosphate–mediated gene transfer: A highly efficient system for stably transforming cells with plasmid DNA. BioTechniques 6:632-638. Graham, F.L. and van der Eb, A.J. 1973. A new technique for the assay of infectivity of human adenovirus 5 DNA. Virology 52:456. Ishiura, M., Hirose, S., Uchida, T., Hamada, Y., Suzuki, Y., and Okada, Y. 1982. Phage particle– mediated gene transfer to cultured mammalian cells. Mol. Cell. Biol. 2:607-616. Wigler, M., Pellicer, A., Silverstein, S., and Axel, R. 1978. Biochemical transfer of single-copy eucaryotic genes using total cellular DNA as donor. Cell 14:725.

Key References Chen and Okayama, 1987. See above. Ishiura et al., 1982. See above. Provides the basis for BES-mediated transfection.

Contributed by Robert E. Kingston (HEPES method) Massachusetts General Hospital and Harvard Medical School Boston, Massachusetts Claudia A. Chen (BES method) National Institute of Mental Health Bethesda, Maryland Hiroto Okayama (BES method) Osaka University Osaka, Japan John K. Rose (optimization) Yale University School of Medicine New Haven, Connecticut



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Transfection Using DEAE-Dextran

UNIT 9.2

Transfection of cultured mammalian cells using diethylaminoethyl (DEAE)-dextran/DNA can be an attractive alternative to other transfection methods in many circumstances. The major advantages of the technique are its relative simplicity and speed, limited expense, and remarkably reproducible interexperimental and intraexperimental transfection efficiency. Disadvantages include inhibition of cell growth and induction of heterogeneous morphological changes in cells. Furthermore, the concentration of serum in the culture medium must be transiently reduced during the transfection. Any of these factors may adversely affect or be incompatible with some bioassays or experimental goals. In addition, for nonstandard cell types there may be a requirement for extensive preliminary investigation of optimal transfection conditions. Together, these factors influence the suitability of this technique to specific purposes. In general, DEAE-dextran DNA transfection is ideal for transient transfections with promoter/reporter plasmids in analyses of promoter and enhancer functions, and is suitable for overexpression of recombinant protein in transient transfections or for generation of stable cell lines using vectors designed to exist in the cell as episomes. The procedure may also be used for expression cloning (Aruffo and Seed, 1987; Kluxen and Lubbert, 1993; Levesque et al., 1991), although electroporation is usually preferred for this purpose (Puchalski and Fahl, 1992). This unit presents a general description of DEAE-dextran transfection (see Basic Protocol) as well as two more specific protocols for typical experimental applications (see Alternate Protocols 1 and 2). The Basic Protocol is suitable for transfection of anchorage-dependent (attached) cells. For cells that grow in suspension, electroporation (UNIT 9.3) or lipofection (UNIT 9.4) is usually preferred, although DEAE-dextran-mediated transfection can be used (Fregeau and Bleackley, 1991). For suspension cells, the transfection step should be performed on collected cells that have been resuspended at 107 cells/ml in transfection medium, using reagents and conditions that are otherwise similar to those of the Basic Protocol. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and proper sterile technique should be followed accordingly. NOTE: All culture incubations are performed in a 37°C, 5% CO2 incubator unless otherwise specified. GENERAL PROCEDURE FOR DEAE-DEXTRAN TRANSFECTION Cultured cells are incubated in medium containing plasmid DNA and DEAE-dextran, which form complexes that are taken up by cells via endocytosis. Chloroquine can be included to inhibit degradation of plasmid DNA. Cells are exposed transiently to DMSO or another permeabilizing agent to increase DNA uptake (DMSO “shock”). Important variables include the concentration of DEAE-dextran, the ratio of DNA to DEAE-dextran, the duration of transfection, and the presence and timing of chloroquine exposure (see Critical Parameters). This protocol is suitable for transfection of COS and CV1 cells; Alternate Protocols 1 and 2 describe two examples of transfection experiments. Materials Cells to be transfected and appropriate culture medium (e.g., complete DMEM; APPENDIX 3F) with and without 10% FBS 100 mM (1000×) chloroquine diphosphate in PBS, filter-sterilized (store at 4°C) Plasmid DNA(s), prepared by CsCl density-gradient centrifugation or affinity chromatography (UNIT 1.7) Contributed by Tod Gulick Current Protocols in Molecular Biology (1997) 9.2.1-9.2.10 Copyright © 1997 by John Wiley & Sons, Inc.

BASIC PROTOCOL


9.2.1 Supplement 40

TE buffer (APPENDIX 2) 10 mg/ml DEAE-dextran stock solution (see recipe) 10% (v/v) dimethyl sulfoxide (DMSO) in PBS, filter-sterilized (store up to 1 month at room temperature) Phosphate-buffered saline (PBS; APPENDIX 2) Appropriate-sized tissue culture vessels (Table 9.2.1) Inverted microscope Additional reagents and equipment for mammalian cell culture (APPENDIX 3F) 1. Plate cells at a density to achieve 50% to 75% confluence on the target day for transfection. For COS or CV1 cells, perform a 1:10 split 2 days prior to transfection. The surface area of various cell culture vessels given in Table 9.2.1 can be used to determine how to split cells to the desired density. Some cell types including many primary cells show particular sensitivity to the toxicity of DEAE-dextran. These cells should be plated at higher density or transfected after reaching near-confluence.

2. Determine the total volume of medium to be used in the transfection based on the number of culture vessels containing cells to be transfected and the volume per vessel shown in Table 9.2.1. Make up this amount of medium (plus some excess) to contain 2.5% FBS by combining 1 part medium containing 10% FBS with 3 parts serum-free medium. DEAE-dextran can precipitate in the presence of high medium protein, necessitating use of a low FCS concentration. Alternatively, NuSerum (Collaborative Research), which contains only ∼30% serum, can be used at a final concentration of 10%.

3. Add 100 mM (1000×) chloroquine diphosphate stock solution to the 2.5%-FBS-containing transfection medium prepared in step 2 to achieve a final concentration of 100 µM. Warm transfection medium to 37°C.

Table 9.2.1 Surface Areas of Commonly Used Tissue Culture Vessels and Corresponding Appropriate DEAE-Dextran Transfection Medium Volumes

Vessel T175 flask T150 flask T75 flask T25 flask 150-mm dish 100-mm dish 60-mm dish 35-mm dish 6-well plate (35-mm wells) 12-well plate (22-mm wells) 24-well plate (15.5-mm wells) Transfection Using DEAE-Dextran

Area (cm2) 175 150 75 25 148b 55b 21b 8b 9.4b 3.8b 1.9b

Appropriate vol. DEAE-dextran mediuma (ml)

10 4 2 1 1 0.5 0.25

aThese volumes are roughly a linear function of vessel surface area. To ensure that cells are completely covered by medium during the transfection, small wells require proportionately larger volumes due to annular sequestration of medium because of surface tension at the periphery. bCostar; other manufacturer products may deviate slightly.

9.2.2 Supplement 40


Chloroquine is toxic to all cells, so exposure time should be limited to 15 min (up to an hour) work just as well.

3. Dilute cationic lipid reagent into dilution medium in a second tube and mix. Transfection of Cultured Eukaryotic Cells Using Cationic Lipid Reagents

4. Combine precomplexed DNA and diluted cationic lipid reagent, mix, and incubate for 15 min at room temperature. Incubation times >15 min (up to an hour) work just as well when LipofectAmine is the cationic lipid and Plus reagent is the enhancer.

9.4.6 Supplement 48


5. While complexes are forming, replace medium on the cells with the appropriate volume of fresh transfection medium. The medium can be the same as the dilution medium. It is possible to use serum in the transfection medium at this step. In some cells (e.g., HeLa and NIH 3T3), transfection in medium containing serum is as efficient or more efficient than in medium without serum.

6. Add DNA-enhancer-lipid complexes to each well containing cells. Mix complexes into the medium gently, holding the plate at a slant. Incubate at 37°C in 5% CO2 for several hours. The exposure time with LipofectAmine Plus may be as short as 3 hr or up to overnight. Be sure there is sufficient medium to prevent the cells from drying out (it is not necessary to increase other components if this is done).

7. After incubation, add cell culture medium to reach normal volume and add serum to bring the final concentration to that of normal cell culture medium. 8. Perform transient or stable expression analysis (see Basic Protocol 1, steps 8a and 8b). CATIONIC LIPID–MEDIATED TRANSFECTION OF SUSPENSION CELLS WITH DNA

BASIC PROTOCOL 2

This protocol is essentially the same as for adherent cells (see Basic Protocol 1) in that lipid and DNA are diluted separately into dilution medium, mixed, and allowed to form complexes before exposing to cells. However, complexes are formed in the wells of multiwell culture plates, and cells are then distributed into the wells containing complexes and allowed to transfect. Materials Dilution medium: cell culture medium without serum or specialized medium for transfection (e.g., Opti-MEM I, Life Technologies) Cationic liposome reagent (e.g., DMRIE-C or LipofectAmine 2000, Life Technologies; also see Table 9.4.1) Plasmid DNA, purified by anion-exchange chromatography (UNIT 2.14 or Goldsborough et al., 1998), cesium chloride density gradient (UNIT 1.7), or alkaline lysis (UNIT 1.6) Cell suspension: 1 × 107 cells/ml in normal cell culture medium without serum or antibiotics Cell culture medium (e.g., complete DMEM; APPENDIX 3F) Serum 6-well tissue culture plates 1. To each well of a 6-well tissue culture plate add 0.5 ml dilution medium. Commercial medium that is specialized for lipid-mediated transfection (e.g., Opti-MEM I), without serum or antibiotics, gives the best results. However, other serum-free media may also be used. When transfecting in different-sized culture plates, change the amounts of DNA, cationic lipid reagent, and medium in proportion to the difference in surface area (see Table 9.4.3).

2. Add 0, 2, 4, 6, 8, or 12 µl cationic lipid reagent to each well and mix gently by swirling the plate. DMRIE-C was found to give high efficiency transfection of DNA in Jurkat (human T cell lymphoma), K562 and KG-1 (human myeologenous leukemia), and MOLT-4 (human lymphoblastic leukemia) cell lines. It is a lipid suspension that may settle with time. To ensure that a homogenous sample is taken, mix thoroughly by inverting the tube 5 to 10 times before removing a sample for transfection.



Supplement 48

3. Add 0.5 ml dilution medium containing 4 µg plasmid DNA to each well. Mix by swirling plate. The amount of DNA should be optimized for each cell line.

4. Incubate at room temperature for 15 to 45 min to allow formation of lipid-DNA complexes. 5. Add 0.2 ml cell suspension (2 × 106 cells) to each well and mix gently. The single most important factor in reproducible, high-efficiency transfection is a consistent number of healthy, proliferating cells. Transfection is most efficient when the cells are maintained in mid-log growth.

6. Incubate several hours at 37°C in a 5% CO2 incubator. A 4-hr incubation is adequate for DMRIE-C transfections.

7. To each well add 2 ml cell culture medium containing 1.5× the usual amount of serum. For Jurkat and MOLT-4 cells, addition of 1 ìg/ml phytohemagglutinin (PHA) and 50 ng/ml phorbol myristate acetate (PMA) enhances promoter activity and gene expression. For K562 and KG-1 cells, PMA alone enhances promoter activity.

8. Assay the cells at 24 or 48 hr posttransfection for transient or stable expression (see Basic Protocol 1, step 8a or 8b). BASIC PROTOCOL 3

CATIONIC LIPID–MEDIATED TRANSFECTION OF ADHERENT CELLS WITH RNA In this protocol, lipid is first diluted into dilution medium and mixed. RNA is then mixed directly into the diluted lipid and immediately added to cells (which have been rinsed with serum-free medium), and cells are incubated for transfection. Materials Adherent cells Cell culture medium with serum (e.g., complete DMEM; APPENDIX 3F) Dilution medium: serum-free cell culture medium or specialized medium for transfection (e.g., Opti-MEM I, Life Technologies) Cationic lipid reagent (e.g., DMRIE-C, Life Technologies; also see Table 9.4.1) mRNA (UNIT 4.5) 6-well or 35-mm tissue culture plate 12 × 75–mm polystyrene tubes Additional reagents and equipment for trypsinizing, counting, and plating cells (APPENDIX 3F) 1. The day before transfection, trypsinize and count adherent cells (APPENDIX 3F). In each well of a 6-well tissue culture plate, or in six 35-mm tissue culture plates, seed ∼2–3 × 105 cells in 2 ml of the appropriate cell culture medium supplemented with serum. Transfection is most efficient when the cells are growing rapidly. Cultures should be maintained carefully and passaged frequently. As transfection efficiency may be sensitive to culture confluency, it is important to maintain a standard seeding protocol from experiment to experiment.

Transfection of Cultured Eukaryotic Cells Using Cationic Lipid Reagents

2. Incubate at 37°C in a 5% CO2 incubator until the cells are ∼80% confluent. This will usually take 18 to 24 hr, but the time will vary among cell types.

9.4.8 Supplement 48


3. On the day of transfection, wash the cells in each well with 2 ml dilution medium at room temperature. Commercial medium that is specialized for lipid-mediated transfection (e.g., Opti-MEM I), without serum or antibiotics, gives the best results. However, other serum-free media may also be used.

4. Add 1.0 ml dilution medium to each of six 12 × 75–mm polystyrene tubes. 5. Add 0, 2, 4, 6, 8, or 12 µl cationic lipid reagent to each tube and mix or vortex briefly. DMRIE-C was found to give high-efficiency transfection of RNA in adherent cell lines (Ciccarone et al., 1995). It is a lipid suspension that may settle with time. To ensure that a homogenous sample is taken, mix thoroughly by inverting the tube 5 to 10 times before removing a sample for transfection.

6. Add 2.5 to 5.0 µg RNA to each tube and vortex briefly. mRNA that is capped and polyadenylated is translated more efficiently and is more stable within the cell.

7. Immediately add lipid-RNA complexes to washed cells and incubate 4 hr at 37°C in a 5% CO2 incubator. The time of exposure of cells to lipid-RNA complexes, as well as the amount of RNA added to the cells, should be adjusted for each cell type.

8. Replace transfection medium with cell culture medium containing serum. 9. Allow cells to express the RNA for 16 to 24 hr and analyze them for expression of the transfected RNA as appropriate for the transgene used. CATIONIC LIPID–MEDIATED TRANSFECTION OF ADHERENT Sf9 AND Sf21 INSECT CELLS WITH BACULOVIRUS DNA

BASIC PROTOCOL 4

As for transfecting mammalian cells (see Basic Protocol 1), cationic lipid reagent and nucleic acid are diluted separately into serum-free medium and then mixed and allowed to form complexes. Complexes are then diluted with fresh transfection medium and added to the cells for transfection. After the cells are fed and incubated, budded virus can be isolated from the medium. Materials Insect cells: Sf 9 or Sf 21 cells (UNIT 16.9) Insect medium (UNIT 16.9; e.g., Sf-900 II SFM, Life Technologies) with and without serum and antibiotics Baculovirus DNA: purified DNA or bacmid DNA miniprep (UNITS 16.9 & 16.10; Anderson et al., 1995) Cationic lipid reagent (Table 9.4.1) 6-well tissue culture plate 27°C incubator 12 × 75–mm polystyrene tubes, sterile Additional reagents and equipment for culturing insect cells and harvesting baculovirus from cell supernatants (UNIT 16.10) 1. In each well of a 6-well tissue culture plate, seed ∼9 × 105 insect cells in 2 ml insect medium without serum or antibiotics (UNIT 16.10). Insect cells must be plated when they are in mid-log growth phase. Cells that have reached stationary phase transfect and infect at very low efficiency. Therefore, it is advisable to



Supplement 48

maintain a standard cell passage protocol that keeps the cells in log growth. For Sf9 or Sf21 cells adapted in Sf-900 II SFM, cells are passaged twice weekly to a density of 3 × 105 cells/ml in suspension, and plated for transfection on the third day postseeding, when they are in mid-log phase. For other cell culture media and growth conditions, adjust conditions to maintain similar growth characteristics. For culture of insect cells, use 50 units/ml penicillin and 50 ìg/ml streptomycin (half the usual final concentration). For transfection, it is preferable to omit antibiotics from the medium to avoid toxicity.

2. Allow cells to attach at 27°C for ≥1 hr. 3. For each transfection, dilute 1 to 2 µg baculovirus DNA into 100 µl insect medium without serum or antibiotics in a 12 ×75–mm polystyrene tube. 4. For each transfection, dilute 1.5 to 9 µl cationic lipid reagent into 100 µl insect medium without serum or antibiotics in a separate 12 ×75–mm polystyrene tube. The suggested amount is 6 ìl, but this should be optimized for each system. CellFectin gives high-efficiency transfection of DNA in insect cell lines (Anderson et al., 1995). It is a lipid suspension that may settle with time. To ensure that a homogenous sample is taken, mix thoroughly by inverting the tube 5 to 10 times before removing a sample for transfection.

5. Combine the two solutions, mix gently, and incubate at room temperature for 15 to 45 min to form lipid-DNA complexes. 6. For each transfection, add 0.8 ml insect medium without serum or antibiotics to each tube containing lipid-DNA complexes and mix gently. 7. Aspirate medium from cells and overlay diluted lipid-DNA complexes onto the washed cells. Alternatively, the medium on the cells can be replaced with 0.8 ml fresh insect medium and the undiluted complexes can be added directly to the fresh medium on the cells.

8. Incubate cells for 5 hr in a 27°C incubator. Protect plates from evaporation by putting them in a humidified container. 9. Remove transfection mixture and add 2 ml insect medium containing antibiotics and serum, if desired. Incubate cells in a 27°C incubator for 72 hr. 10. Harvest baculovirus from cell supernatants (UNIT 16.10). Gene expression may also be evaluated in the cells after removal of virus-containing medium. SUPPORT PROTOCOL

FINE TUNING OR OPTIMIZING CONDITIONS FOR CATIONIC LIPID REAGENT TRANSFECTIONS This protocol provides an example of a simple one-step procedure for determining conditions conducive to high-efficiency transfections using cationic lipid reagents in any target cell type. A matrix of DNA and lipid reagent concentrations is used on transfections performed in a multiwell plate (Fig. 9.4.3). Once the best conditions have been determined, the transfections may be scaled up to larger vessels using the relative surface area (see Table 9.4.3) to increase the amounts of all reagents proportionately. This protocol can be modified for use with any transfection protocol.




Additional Materials (also see Basic Protocol 1 and Alternate Protocol) 24-well tissue culture plates 96-well round-bottom plates (sterile, with lid) 1. The day before transfection, trypsinize and count cells (APPENDIX 3F). Plate cells in each well of a 24-well tissue culture plate using normal cell culture medium with serum, so that they are 50% to 95% confluent on the day of transfection. Avoid antibiotics at the time of plating and during transfection. The single most important factor in high-efficiency transfection is healthy, proliferating cell cultures. Antibiotics may cause some toxicity if present during transfection. In a 24-well plate, seeding 4 × 104 to 2 × 105 cells per well will usually give good plating density. Any type of plate may be used by scaling the reagent and cell amounts in proportion to the relative surface area (see Table 9.4.3).

2. Dilute DNA into dilution medium (appropriate for the lipid being optimized) without serum or antibiotics in four microcentrifuge tubes. Use a range of DNA concentrations, and use a volume of dilution medium that is 7× the appropriate protocol volume (see Basic Protocol 1, step 2, and Table 9.4.1). Mix gently after each addition. This makes enough DNA per tube for seven wells on a 24-well plate. Good ranges include 0.2 to 1.6 ìg DNA per well. If the Plus enhancer is being used, include it in the diluted DNA tubes, using 10 ìl Plus reagent per ìg DNA. Add the Plus reagent to the diluted DNA after mixing well. If the Plus reagent is added first, precipitation may occur.

3. Dilute cationic lipid reagent into dilution medium without serum or antibiotics in six microcentrifuge tubes. Use a range of DNA concentrations, and use a volume of dilution medium that is 5× the appropriate protocol volume (see Basic Protocol 1, step 3, and Table 9.4.1). Mix gently after each addition. Be sure to observe timing that works best for the cationic lipid reagent being used. This makes enough diluted lipid per tube for five wells on a 24-well plate. Good ranges include 0.5 to 5 ìl lipid reagent per well.

Cationic lipid reagent (µl)

1.0

1.5

2.0

2.5

3.0

3.5

DNA (µg)

0.2

0.4

0.8

1.2

Figure 9.4.3 A sample matrix for fine tuning (optimizing) transfection reagent efficiencies using cationic lipid reagents.



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4. Pipet equal per-well volumes of diluted DNA and diluted cationic lipid reagent into the wells of a 96-well plate in a matrix corresponding to the wells on the 24-well plate (Fig. 9.4.3). Mix the complexes with the pipet tip by triturating. Cover the plate and incubate for 15 min at room temperature. Incubation times >15 min (up to an hour) work just as well, but be sure the complexes do not dry by covering them well.

5. While complexes are forming, replace medium on the cells with fresh transfection medium. See Basic Protocol 1, steps 5 and 6, for alternate procedures for combining complexes, medium, and cells. The medium can be the same as the serum-free dilution medium. It is possible to use serum in the transfection medium at this step. It is also possible to omit this step when using LipofectAmine 2000.

6. Add aliquots of DNA-lipid complexes (total volume from wells in step 4) to each well containing cells with fresh transfection medium. Mix complexes into the medium gently, holding the plate at a slant. Incubate at 37°C in 5% CO2 for 5 hr. The exposure time may be >5 hr (up to overnight). Be sure there is sufficient medium to prevent the cells from drying out (it is not necessary to increase other components if this is done). If using the Plus enhancer, a 3-hr exposure is sufficient.

7. After 5 hr incubation, add cell culture medium to reach normal volume and add serum to bring the final concentration to that of normal cell culture medium. If necessary to maximize cell growth, replace the medium containing the complexes with fresh complete medium after 5 hr incubation. This step may be omitted entirely for some protocols.

8. Check expression as described (see Basic Protocol 1, steps 8a and 8b). If peak activity is found to occur on the edge of the matrix of concentrations tested, adjust the concentrations to include the observed peak at the center of a new matrix and repeat the transfection.



There are currently at least eight companies that market cationic lipid–based transfection reagents. A partial listing of companies and products may be seen in Table 9.4.1. Many companies offer more than one type of reagent. Among the more popular ones are LipofectAmine 2000 and LipofectAmine Plus (Life Technologies), DOTAP and FuGENE 6 (Roche), and Effectene (Qiagen). Some of the structures are proprietary. The structures that are published can be classified into two general categories based on the number of positive charges in the lipid headgroup. The first cationic lipid (DOTMA) has a single positive charge per molecule and is used in Lipofectin (Life Technologies; Felgner et al., 1987). Several other cationic lipid–based transfection reagents such as DOTAP liposomal transfection reagent (Roche) and DMRIE-C (Life Technologies) also make use of singly charged cationic lipid mole-

cules. Increasing the number of positive charges per cationic lipid molecule to as many as five improved transfection efficiency dramatically in most cell types. This can be seen in the examples of DOGS, the cationic lipid in Transfectam (Promega; Behr et al., 1989); DOSPA in LipofectAmine (Life Technologies; Hawley-Nelson et al., 1993); and TMTPS in CellFectin (Life Technologies; Anderson et al., 1995). Further increase in transfection efficiency can sometimes be achieved by precomplexing DNA with a proprietary enhancer. Two commercially available transfection kits with enhancers are LipofectAmine Plus (Life Technologies; Shih et al., 1997) and Effectene (Qiagen). Life Technologies has designed cationic lipid reagents with specialized applications such as high-efficiency transfection of insect cells (see Basic Protocol 4) or delivery of RNA (see Basic Protocol 3). Lipofectin has high



activity for endothelial cell transfection (Tilkins et al., 1994). Basic Protocol 1 and the Alternate Protocol described in this unit are the procedures with the highest potential for efficient DNA transfection of adherent mammalian cells (Shih et al., 1997; Ciccarone et al., 1999). LipofectAmine 2000 has a simple protocol that yields the highest transfection efficiencies in many cell types. Using the enhancer reagent results in more reproducible transfections without extensive optimization because of the overall high activity. Prior to the availability of enhanced cationic liposome transfections (e.g., using LipofectAmine 2000 and Effectene), the most effective procedure for transfection of adherent mammalian cells with DNA was with other polycationic reagents (e.g., LipofectAmine; Hawley-Nelson et al., 1993) following Basic Protocol 1. In order to achieve high-efficiency transfections with Basic Protocol 1, it is necessary to optimize lipid and DNA concentrations with the target cells at the desired plating density using a procedure similar to that described in the Support Protocol. Many cationic lipid reagents, as well as transfection reagents based on other chemistries, are available that can be used in Basic Protocol 1 for adherent mammalian cell DNA transfection, but they may yield lower efficiencies than the Alternate Protocol with the enhancer. Optimization using the Support Protocol is highly recommended when not using the enhancer, and the protocol can be modified for use with any cationic lipid reagent.

Critical Parameters The most critical parameter for successful transfection is cell health. Cells should be proliferating as rapidly as possible at the time they are plated for transfection. On the day of transfection, mitoses should be abundant in healthy cultures. Fresh cultures with a finite life span should be used at the earliest possible passage. For reproducible transfection results, it is critical to plate the same number of healthy cells for each transfection. Cells should always be counted, preferably using a hemacytometer and trypan blue (APPENDIX 3F). Although optimization is not required for high-efficiency transfection when using an enhancer (see Alternate Protocol), it is essential for success without the enhancer, and may improve efficiency even with the enhanced method. The medium used to dilute and form complexes between the cationic liposomes and the DNA must not contain serum. Serum contains

sulfated proteoglycans and other proteins, which compete with the DNA for binding to the cationic lipids. The medium should also not contain antibiotics. There is toxicity to the cells when cationic lipid reagents are used in the presence of antibiotics. The dilution medium/plating medium for the cells may have some influence on transfection efficiency. Some proprietary serum-free media contain components that inhibit transfection and should be replaced with Opti-MEM I, DMEM, or other media without serum during transfection (Hawley-Nelson and Ciccarone, 1996). Serum may be present in the medium on the cells during transfection. For most cationic lipid reagents, on most cell types, transfection activity is not inhibited in the presence of serum provided the complexes were formed in serumfree medium (Brunette et al., 1992; Ciccarone et al., 1993, 1999; Shih et al., 1997). The specific serum-free medium used to dilute the lipid and DNA can have a slight effect on the efficiency of transfection. For the enhanced protocol (Alternate Protocol), normal culture medium such as DMEM is recommended. For the standard procedure (Basic Protocol 1, 2, and 3), Opti-MEM I medium works best. The improvements resulting from using the recommended media are less than two fold. When using Lipofectin, dilution in OptiMEM I followed by a 30- to 45-min incubation is recommended (Ciccarone and Hawley-Nelson, 1995). With LipofectAmine 2000, the reverse is true: extended incubation (>30 min) of LipofectAmine 2000 in Opti-MEM prior to addition of DNA results in lower transfection activity (Ciccarone et al., 1999). High-purity DNA will increase transfection efficiency. Miniprep DNA does work, however, when efficiency is not critical. A wide range of sizes of polynucleotides may be transfected, from 18-mer single-stranded oligonucleotides (Chiang et al., 1991; Bennett et al., 1992; Yeoman et al., 1992; Wagner et al., 1993) to 400-kb YAC DNAs (Lamb et al., 1993). Excess vortexing of complexes or DNA solutions may result in some shearing, especially with larger molecules. The concentration of EDTA in the diluted DNA should not exceed 0.3 mM. Transgene expression may be increased in some cell types by inducing the promoter. This is observed in Jurkat cells when phytohemagglutinin and phorbol myristate acetate are added following transfection to activate the cytomegalovirus promoter (Schifferli and Ciccarone, 1996).



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thing to try is to work with a freshly thawed culture or isolate (Hawley-Nelson and Shih, 1995). Be sure the same number of cells is plated in each experiment, since plating density affects efficiency and peak position (HawleyNelson et al., 1993). Low cell yield often results from the use of too much DNA or cationic lipid reagent. Use lower concentrations of these two components and examine the results for transfection efficiency as well as cell yield. Acceptable efficiency can usually be obtained with higher cell yield by using lower concentrations of lipid and DNA (Hawley-Nelson et al., 1993; Life Technologies, 1999).

Troubleshooting The most common complaints surrounding transfections include decreased transfection efficiency and low cell yield. Decreases in efficiency often result from changes in the target cell line. Cultured cell lines are usually aneuploid and often consist of a mixture of genotypes and phenotypes that can be subject to selection in the laboratory environment. Primary cultures, although usually genotypically uniform, often consist of a mixture of phenotypes from different tissues and can change their population characteristics in response to their environment. Whenever a decrease in transfection efficiency is observed, the first

DNA (µg/well)

A

Cationic lipid reagent (µl)

0.4

0

0.5

1

1.5

2

2.5

0.8

1

1.5

2

2.5

3

3.5

1.2

1

1.5

2

2.5

3

3.5

1.6

1

1.5

2

2.5

3

3.5

B


Figure 9.4.4 Results of fine-tuning or optimizing conditions for transfection. Before transfection, 293 H cells were plated at 2 × 105/well in a 24-well plate precoated with poly-D-lysine. The following day, cells were transfected with pCMV⋅SPORTβgal DNA using LipofectAmine 2000 as described (see Support Protocol). One day posttransfection, cells were fixed and stained with Xgal. (A) Amounts of DNA and lipid reagent used. (B) Results of Xgal staining.



Table 9.4.5

Plate 24-well 12-well 6-well 60-mm 100-mm

Activity for a Scaled-up Transfection Using LipofectAmine Plus in BHK-21 Cells

Surface area ratio to 24-well plate 1 2 5 10 28

Cells/well (× 104) 4 8 20 40 112

DNA/well Plus reagent (µg) (µl) 0.4 0.8 2 4 11.2

LipofectAmine reagent (µl)

2 4 10 20 560

2 4 10 20 56

β-gal (ng/cm2)a 188 ± 5 193 ± 12 179 ± 27 171 ± 16 157

aResults are the mean of three transfections ± the standard deviation.

Cell yield can also be improved in several other ways. (1) Increasing the plating density. This usually requires adjustment of lipid and DNA amounts, but often the transfection efficiency as well as the cell yield increases with higher plating input (Life Technologies, 1999). (2) Decreasing time of exposure of the cells to cationic lipid–DNA complexes. This can be done by increasing volume and adding back serum at earlier times or by removing the complexes from the cells at the end of transfection. (3) Performing the transfection in the presence of serum. Most cationic lipid reagents work well in transfection medium containing serum (Brunette et al., 1992; Ciccarone et al., 1993, 1999; Shih et al., 1997). One exception is LipofectAmine without the Plus enhancer. Some cationic lipid solutions are naturally cloudy. Sometimes cloudiness is observed when complexes are made with DNA. Usually this does not interfere with transfection efficiency. Most cationic liposome solutions (especially DMRIE-C and CellFectin) should be mixed gently by inversion just before use to produce a uniform suspension. With Plus reagent, it is possible to precipitate the DNA when the Plus reagent is diluted first into the DMEM and DNA is added second. Always dilute the DNA into DMEM and mix well before adding Plus reagent.

al., 1999). Efficiencies also vary for suspension cells. The authors note that while LipofectAmine Plus is relatively inefficient for transfecting Jurkat cells, DMRIE-C can yield up to 85% blue cells following pCMV⋅SPORTβgal plasmid DNA transfection, gene activation with phytohemagglutinin and phorbol myristate acetate, and Xgal staining (Ciccarone et al., 1995, Schifferli and Ciccarone, 1996). The result of a typical fine-tuning/optimization protocol is shown in Figure 9.4.4. The transfection reagent was LipofectAmine 2000, the DNA was pCMV⋅SPORTβgal, the cells were 293 H. Cells were stained the day following transfection and were allowed to stain overnight at 37°C. A selection of transfection conditions can be made. Conditions found to be advantageous for transfection in small wells may be scaled up, the results of a typical scale-up are given in Table 9.4.5. The results of a stable transfection of NIH 3T3 cells with pSV2neo DNA using an enhancer reagent (LipofectAmine Plus) are shown in Figure 9.4.5. The transfection was done on 24-well plates, the cells were passaged onto 6-well plates the following day, and selection with geneticin was done for 10 days. The figure shows optimization of conditions and the generally high efficiency that can be achieved using this method.

Anticipated Results Transfection should be observed for most adherent mammalian cells transfected with DNA using Basic Protocol 1. Efficiencies vary with cell type. For example, 293, COS-7, and CHO-K1 cells yield 95% or more blue cells following Xgal staining of cells transfected with pCMVSPORTβgal plasmid DNA using LipofectAmine 2000. The authors have noted efficiencies of other cell types as high as 49% for SK-BR3 breast cancer cell lines, 77% for BE(2)C human neuroblastoma cells, and 43% for MDCK canine kidney cells (Ciccarone et

Time Considerations Counting and plating the cells should be done the day before transfection and will usually require 12 hr (see Critical Parameters).

3. Prepare a transfection cocktail by adding 6 to 10 µg retroviral vector plasmid DNA to water and diluting to 438 µl final volume. Add 62 µl of 2 M CaCl2 to the DNA solution. Plasmid DNA prepared by cesium chloride gradients or by using a commercial kit appears to work equally well. It is unnecessary to perform additional phenol extraction or precipitation steps prior to using the DNA, and the DNA may be stored in either water or TE buffer. When cotransfecting retroviral vector with retroviral structural genes on multiple plasmids, the total amount of transfected DNA should be 10 to 15 ìg and the plasmids should be introduced in approximately equimolar ratios. These amounts and ratios may have to be altered to optimize retroviral titers (see Critical Parameters).

4. Add 500 µl of 2× HeBS and mix by shaking the tube or pipetting. Within 1 to 2 min, add this solution to the cells and gently swirl the plate to ensure uniform mixing. Incubate until transfected cells are close to 100% confluent (24 to 48 hr post transfection). Remove the medium and add fresh medium at ∼24 hr post transfection (see step 5). HeBS may also be added to the DNA/CaCl2 solution by bubbling. Transfection efficiency is highest if DNA/CaCl2/HeBS mix is added immediately to cells rather than waiting for a visible precipitate to form (30 to 60 min). Although a precipitate is usually visible on the plate within 1 to 2 hr after transfection, the quantity of precipitate appears to be related to the amount of input DNA (W. Pear, unpub. observ.), and the amount of precipitate is usually an unreliable indicator of transfection efficiency. If chloroquine is included in the medium, remove medium ∼10 hr after transfection and gently replace with fresh 293 cell growth medium. Addition of 10 mM sodium butyrate to the cells may enhance retroviral titers by several-fold (Soneoka et al., 1995). It should be added to the 293 growth medium 12 to 24 hr after transfection at a final concentration of 10 mM. Cells should be treated for 12 to 14 hr with sodium butyrate and washed with PBS prior to proceeding to step 5.

5. Approximately 16 to 24 hr prior to harvesting the retroviral supernatant, remove the medium and gently replace with 4 ml of 293 cell growth medium. The transfected cells should be close to 100% confluent at this time. The relative retroviral titer may be increased by adding 3 ml rather than 4 ml of medium. If the cells that will be infected require special medium (e.g., RPMI medium, growth factors), the special medium,



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rather than 293 cell growth medium, should be added to the cells. Except for the presence of fetal bovine serum, 293 cells appear to tolerate a wide variety of culture conditions.

6. Harvest retroviral supernatant ∼48 hr following transfection by gently removing the supernatant from the cells. Either filter the supernatant through a 0.45-µm filter or centrifuge it 5 min at 500 × g (1000 rpm in Sorvall RT-3000B rotor), 4°C, to remove live cells. At harvest, the supernatant will be yellow. Acidity does not appear to affect retroviral titer. Once retroviral supernatant is harvested, the transfected cells may be assayed for transfection efficiency by staining for a histochemical marker (such as lacZ, UNIT 9.10) or cell surface marker, by preparing DNA from the cells and assaying by Southern hybridization (UNIT 2.9A), or by lysing cells for protein immunoblot analysis (UNIT 10.8). Optimal transfection efficiencies should range from 30% to 60%. If the supernatant is to be tested for the presence of replication-competent (helper) virus, see UNIT 9.13. It is necessary to wait at least 36 hr post transfection to obtain high-titer retroviral supernatants. At this point, it may be possible to serially, harvest and replenish supernatant every 12 hr up to 72 hr post transfection without a significant drop in retroviral titer. If the cells are sparse at 24 hr post infection, it may be necessary to wait until 72 hr to obtain high-titer supernatant.

7. If the retroviral supernatant is to be used within 1 to 2 hr, store it on ice. For longer intervals, freeze the supernatant on dry ice and transfer it to −80°C. Retroviral half-life is 3 to 6 hr at 37°C (Sanes et al., 1986). To maintain high titers, supernatants should be left on ice or frozen following harvest. Freezing does not appear to cause >2-fold drop in titer, as long as the supernatant does not undergo more than one freeze/thaw cycle; exposure to more than one such cycle causes a marked drop in retroviral titer. For this reason, supernatant should be aliquoted in appropriate volumes for subsequent infection. To thaw frozen supernatants, warm for a minimal period at 37°C and use immediately. SUPPORT PROTOCOL

GROWTH AND STORAGE OF 293 CELLS The two most important variables in determining the success of retroviral production are the state of the packaging cells and the quality of the transfection reagents. This protocol discusses methods for optimizing growth of 293 cells, passaging the cells, and freezing stocks. Further details for the culture, freezing, and storage of mammalian cell lines can be found in APPENDIX 3F. Materials 293 cells or packaging cell lines (e.g., Bosc23; see Table 9.9.1) derived from 293 cells 293 cell growth medium (see recipe) Freezing medium: 90% (v/v) heat-inactivated FBS/10% (v/v) DMSO Trypsin solution: 0.05% (v/v) trypsin/0.53 mM EDTA PBS without Ca2+ or Mg2+ (APPENDIX 2) 10-cm tissue culture dishes Sorvall RT-3000B centrifuge and rotor (or equivalent) 2-ml cryogenic vials

Transient Production of Retroviral Supernatants



Culture and passage 293 cells 1. Passage and grow 293 cells or the packaging cell lines derived from them in 10 ml of 293 cell growth medium in 10-cm tissue culture dishes. Cells are typically grown in 10-cm dishes; however, other sizes can be used and the protocols scaled appropriately. It is recommended that cells be maintained at relatively high density and passaged at a split ratio ≤1:5 to maintain uniformity of the cells in culture. This is especially true for packaging cell lines. It is also recommended that when cells are first received they be expanded to prepare 50 to 100 vials of frozen cells to provide a uniform source of cells for virus production. Cells should not be allowed to become overconfluent, as this leads to formation of cell clumps that can cause uneven cell distribution after replating and result in less efficient transfection. Cells should be split when they reach 90% confluence and the medium is acidic. If clumping occurs, growing cells for 1 to 2 passages at high density (1:3 split) should remove the clumps. 293 cells and packaging lines derived from those cells appear to tolerate a wide variety of growth conditions and addition of factors without drop in retroviral titer. One variable that is important for maximal retroviral titers is to use heat-inactivated fetal bovine serum (FBS) in the tissue culture medium, rather than horse serum (HS) or newborn calf serum (NCS). To date, both DMEM and RPMI have given good results, and the addition of IL-2, IL-3, IL-6, and stem cell factor (SCF) has not caused the retroviral titers to decrease (W. Pear, unpub. observ.).

2. When the cells are confluent, remove the medium and gently rinse once with PBS without Ca2+ and Mg2+. 293 cells are much less adherent than murine fibroblasts. As a result, washes must be performed very gently to avoid removing cells.

3. Add 1 ml trypsin solution and incubate 2 to 3 min at 37°C. Although cells may be detached by tapping the plate after 30 sec, a more efficacious method to obtain a single-cell suspensions is to return the cells to the incubator for 2 to 3 min. The cells should then detach by themselves.

4. Quench trypsin with 4 ml of 293 growth medium and transfer the cell suspension to a conical centrifuge tube. Centrifuge 3 min at 500 × g (1000 rpm in Sorval RT-3000B rotor), 4°C or room temperature. To obtain single-cell suspensions, it is recommended that cells be vigorously pipetted in a conical tube rather than on the tissue culture dish.

5. Resupend pellet vigorously in 5 ml of 293 cell growth medium. Let the suspension stand 1 min to allow the largest clumps to settle. Aliquot the cells without using the last 200 µl of cell suspension (clumps). Freeze 293 cells 6. Incubate a 10-cm dish of 293 cells in 293 growth medium until the cells are 80% confluent. Maximal viability is achieved if cells are frozen prior to confluence. One 10-cm dish that is 80% confluent will provide enough cells for 2 to 3 vials of frozen cells.

7. Using trypsin solution, remove the cells from the dish (see steps 2 to 4). Centrifuge the cells 3 min at 500 × g, 4°C or room temperature. 8. Remove the medium and resuspend the cells in 2 to 3 ml freezing medium per 10-cm dish.



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9. Add 1 ml cell suspension to each 2-ml cryogenic vial and store overnight at −80°C. On the following day, transfer the vials to liquid nitrogen. Thaw 293 cells 10. Remove a vial of cells from liquid nitrogen and thaw rapidly, warming by hand or at 37°C. 11. Add 1 ml 293 growth medium to the cryogenic vial and add the cell suspension to a tube containing 10 ml growth medium. Centrifuge 3 min at 500 × g, 4°C or room temperature. 12. Remove the supernatant and resuspend the cell pellet in 10 ml growth medium. Transfer the cells to a 10-cm tissue culture dish. Incubate. BASIC PROTOCOL 2

PSEUDOTYPING A STABLE CELL LINE SEQUENTIALLY WITH VSV G PROTEIN Pseudotyping retroviruses with the vesicular stomatitis virus (VSV) G protein creates virus with a polytropic host range and makes it possible to concentrate the virus by centrifugation. Calcium phosphate–mediated transfection yields retroviral titers in the range of 105 to 106 CFU/ml; however, concentration of the viruses can increase the titer by several log factors. Because concentration requires larger initial volumes of retroviral supernatants, best results are obtained if a stable cell line that expresses gag-pol and the gene of interest is prepared first. A VSV G protein expression vector is then transiently expressed in this cell line for production of retroviral supernatants. This method is adapted from Burns et al. (1993) and Matsubara et al. (1996). An alternative method is to introduce retroviral vector and VSV G protein–expressing plasmids by cotransfection (see Alternate Protocol 1); it should be possible to introduce gag-pol construct at the same time. Although the latter approach may be laborious for obtaining large quantities of retroviral supernatants for concentration, it may yield higher titers for proteins that are detrimental to the long-term growth of 293 cells. Materials Retroviral packaging cell line with amphoteric host range (e.g., PA317, ΨCRIP, GP+Am12, or Bing; see Table 9.9.1) and appropriate culture medium Retroviral vector DNA gag-pol-expressing cell line (e.g., Anjou65, ATCC CRL 11268; see Table 9.9.1) and appropriate culture medium 293 cell growth medium (see recipe) VSV G protein expression plasmid: e.g., pHCMV-VSV-G (Matsubara et al., 1996) TNE buffer (see recipe) or 1% (v/v) Hanks basic salt solution (HBSS; APPENDIX 2) 60-mm tissue culture dishes Sorvall RT-3000B centrifuge and rotor (or equivalent) Beckman L3-50 centrifuge and SW 41 rotor (or equivalent) Additional reagents and equipment for culturing of mammalian cells (APPENDIX 3F), transfecting cells and harvesting virus (see Basic Protocol 1), infecting cells (see Basic Protocols 3 or 4 or Alternate Protocols 2, 3, or 4), selecting clones with drugs (UNIT 9.5), assaying clones for the gene of interest (e.g., UNITS 9.6-9.8 & 9.10), and freezing cells (see Support Protocol)


Create a stable cell line 1. Transfect a retroviral packaging cell line with an amphotropic host range with 6 to 20 µg retroviral vector DNA (see Basic Protocol 1). Incubate 48 hr.



2. Twenty-four hours before harvesting retroviral supernatant, plate 2.5 × 106 gag-polexpressing cells into the appropriate culture medium in 60-mm tissue culture dish. 3. Collect retroviral supernatant at 48 hr post transfection. Filter through a 0.45-µm filter or centrifuge 5 min at 500 × g (1000 rpm in Sorvall RT-3000B rotor), 4°C, to remove cells. 4. Use 1 ml retroviral supernatant to infect the gag-pol-expressing cell line. For production of stable producer cell lines, infection is preferred to transfection because there is a reduced risk of rearrangement.

5. Select individual clones of stable producer cell lines—i.e., by drug selection (UNIT 9.5), FACS sorting, or other method. 6. Identify clones that express the gene of interest (e.g., UNITS 9.6-9.8 & 9.10). These are the stable producer cell line(s) that will be used for subsequent experiments.

7. Aliquot and prepare frozen stocks of the stable producer cell line (see Support Protocol). Pseudotype the producer cell line 8. Twenty-four hours prior to transfection, plate 2.5 × 106 stable producer cells in appropriate culture medium in a 60-mm dish. If desired, the procedure can be scaled up for larger volumes. It is even possible to transfect spinner cultures (J.C. Burns, pers. comm.).

9. Transfect stable producer cells with 3 to 6 µg VSV G protein expression plasmid by modified calcium phosphate–mediated transfection (see Basic Protocol 1). Incubate 24 hr. Adjust the volumes proportionally if larger volumes are used in step 8.

10. Replace the medium with fresh 293 cell growth medium and continue incubation 24 to 48 hr. 11. Harvest retroviral supernatants between 48 and 72 hr by collecting the medium from the culture. Concentrate supernatant immediately (proceed to step 12) or store at −80°C. Syncytium formation is evident ∼36 hr following transfection. Syncytia are identified by the formation of multinucleated giant cells which are the result of membrane fusion caused by VSV G protein.

Concentrate VSV G protein–pseudotyped viral supernatant 12. Concentrate retroviral supernatant by centrifuging 90 min at 50,000 × g (17,000 rpm in an SW 41 rotor), 4°C. 13. Resuspend the pellet overnight at 4°C in 20 µl of TNE buffer or 0.1% HBSS. Pellet is resuspended in 1/200 of viral culture volume.

14. Use concentrated retroviruses immediately for infection, store them frozen at −80°C, or concentrate the stock by another round of centrifugation (steps 12 and 13).



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PSEUDOTYPING COTRANSFECTIONALLY WITH VSV G PROTEIN It is also possible to produce a retrovirus pseudotyped with vesicular stomatitis virus (VSV) G protein by transient cotransfection. This method is adapted from J. Burns (pers. comm.) and Pear et al. (1996). Materials 293 cells or 293-derived cells expressing retroviral gag-pol: e.g., Anjou65 (ATCC CRL 11269) 293 cell growth medium (see recipe) Retrovirus vector DNA VSV G protein expression plasmid: e.g., pHCMV-VSV-G (Matsubara et al., 1996) 60-mm tissue culture dishes Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F), modified calcium phosphate–mediated transient transfection of cells (see Basic Protocol 1), and concentrating pseudotyped viral supernatants (see Basic Protocol 2) 1. Twenty-four hours prior to transfection, plate 2.5 × 106 293 cells, or cells of a 293 derivative stably expressing the retroviral gag-pol genes, into 293 cell growth medium in a 60-mm dish. 2. Cotransfect 3 to 5 µg retroviral vector DNA and 3 to 5 µg of a VSV G protein expression plasmid by modified calcium phosphate–mediated transfection (see Basic Protocol 1). Incubate. If 293 cells were plated in step 1, also cotransfect a gag-pol expression plasmid.

3. On the following day, replace the medium with 4 ml fresh 293 growth medium. 4. Harvest retroviral supernatants between 48 and 72 hr by collecting the culture medium. Concentrate the virus immediately (see Basic Protocol 2) or store at −80°C. Syncytium formation is evident ∼36 hr following transfection. BASIC PROTOCOL 3

INFECTION OF ADHERENT CELLS WITH RETROVIRAL SUPERNATANT This protocol gives methods for infecting adherent cells with retroviral supernatant. This is the most common method for determining the retroviral titer (UNIT 9.10) and testing for the presence of helper virus (UNIT 9.13). Infectious titer can be quantitated based on marker staining—e.g., using lacZ (UNIT 9.10), or green fluorescent protein (GFP; UNIT 9.7C)—or drug resistance (UNIT 9.5). Recent reports suggest that viral transduction by centrifugation (spin infection; see Alternate Protocol 2) will increase infectious titers 2- to 4-fold (Kotani et al., 1994). These methods are written for cells of the commonly used murine fibroblast line NIH 3T3; however, they should be adaptable to other adherent cell lines with minor modifications.


Materials Retroviral supernatant, fresh or frozen (see Basic Protocols 1 or 2) Target cells: e.g., NIH 3T3 cells 4 mg/ml polybrene in PBS (APPENDIX 2), filtered and stored at 4° or −20°C Fibroblast growth medium Antibiotic for drug selection (UNIT 9.5; optional) 10-cm tissue culture dishes



Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F) and for assaying of reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10) or drug selection (UNIT 9.5). 1. Twelve to eighteen hours prior to transfection, plate 5 × 105 NIH 3T3 cells into appropriate tissue culture medium in a 10-cm tissue culture dish. Infection protocols can be scaled up or down. For a 60-mm plate, plate 2 × 105 cells and infect in a volume of 1 ml.

2. Prepare a 3-ml infection cocktail containing: Retroviral supernatant: ≤1.5 ml 4 µg/ml polybrene Fibroblast growth medium to a final volume of 3 ml. The retroviral supernatant should be ≤1⁄2 the final volume of the infection cocktail. 293 supernatants appear to contain a cytostatic factor that can be diluted out by changing the transfection medium 18 to 24 hr before retroviral harvest and by avoiding infectious stocks that contain viral supernatant at ratios >1:1 (Naviaux et. al., 1996).

3. Remove the medium from the dish of cells and add the infection cocktail. Incubate ≥3 hr. If infection continues >6 hr, a 5-ml infection cocktail is recommended to prevent dehydration.

4. After the infection incubation, add 7 ml fibroblast growth medium to the dish. Incubate 36 to 48 hr. Drug selection (e.g., G418, puromycin) can begin 24 hr post infection (UNIT 9.5). If the cells are not over-confluent (i.e., if they are 50% confluent, it is necessary to split the cells as described in UNIT 9.5.

5. Assay the cells for reporter gene expression (UNITS 9.6-9.8 & 9.10). INFECTION OF ADHERENT CELLS BY SPIN INFECTION Adherent cells can also be infected by centrifuging the cells in the presence of the infection cocktail (spin infection). This protocol is modified from the methods of Kotani et al. (1994).


Materials Target cells: e.g., NIH 3T3 cells Fibroblast growth medium Retroviral supernatant 4 mg/ml polybrene in PBS (APPENDIX 2), filtered and stored at 4° or −20°C 6-well tissue culture plates Beckman GS-6KR or Sorvall RT-3000B centrifuge with microplate carriers (or equivalent) Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F) and for assaying of reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10) or drug selection (UNIT 9.5)



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1. Twelve to eighteen hours prior to infection, plate 1 × 105 target cells per well into fibroblast growth medium in a 6-well tissue culture plate. 2. For each well to be transfected, prepare a 4-ml infection cocktail containing: Retroviral supernatant: ≤2 ml 4 µg/ml polybrene Fibroblast growth medium up to 4 ml. An infection cocktail of ≥4 ml is necessary to prevent dehydration of the cells. The retroviral supernatant should be ≤1⁄2 the final volume of the infection cocktail. 293 supernatants appear to contain a cytostatic factor that can be diluted out by changing the transfection medium 18 to 24 hr before retroviral harvest and by avoiding infectious stocks that contain viral supernatant at ratios >1:1 (Naviaux et. al., 1996).

3. Remove the medium from the cells and add the infection cocktail. 4. Centrifuge the cells 1.5 to 2 hr at 1000 × g (2500 rpm in Beckman GS-6KR or Sorvall RT-3000B rotor), room temperature. 5. Return the cells to the incubator. 6. Change medium the next day and incubate until 48 hr post infection. 7. Assay for reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10), add antibiotic for drug selection (UNIT 9.5), or test for effect of the gene of interest. BASIC PROTOCOL 4

INFECTION OF NONADHERENT CELLS BY RETROVIRAL SUPERNATANT Nonadherent cells can be infected by direct addition of retroviral supernatant or by cocultivation of nonadherent cells with retroviral producer cells (see Alternate Protocol 3). The advantage of cocultivation is that there is continuous retroviral production; however, this must be weighed against the disadvantage of harvesting producer cells together with target cells. This effect may be minimized by irradiating producer cells prior to cocultivation. Alternatively, nonadherent cells can be infected by spin infection (see Alternate Protocol 4). In general, infection by cocultivation is equivalent to spin infection. Materials Retroviral supernatant 4 mg/ml polybrene in PBS (APPENDIX 2), filtered and stored at 4° or −20°C Target cell growth medium Exponentially growing nonadherent target cells 15-ml centrifuge tube 60-mm tissue culture dishes Sorvall RT-3000B centrifuge and rotor (or equivalent) Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F) and for assaying of reporter gene expression (UNITS 9.6-9.8 & 9.10) or drug selection (UNIT 9.5) 1. Prepare a 3-ml infection cocktail containing:


Retroviral supernatant: ≤1.5 ml 2 µg/ml polybrene Target cell growth medium up to 3 ml. The retroviral supernatant should be ≤1⁄2 the final volume of the infection cocktail. 293 supernatants appear to contain a cytostatic factor that can be diluted out by changing



the transfection medium 18 to 24 hr before retroviral harvest and by avoiding infectious stocks that contain viral supernatant at ratios >1:1 (Naviaux et. al., 1996). Many nonadherent cells are more stringent in their growth requirements than fibroblasts. For this reason, it may be necessary to use a medium other than 293 cell growth medium during the 24 hr prior to viral harvest and/or add growth factors or cytokines to the medium. It is essential, however, that heat-inactivated fetal bovine serum (FBS) be used as the serum source.

2. Centrifuge 3 × 105 to 3 × 106 exponentially growing target cells 5 min at 500 × g (1000 rpm in Sorvall RT-3000B rotor), 4°C or room temperature. 3. Remove supernatant and resuspend the cells in the infection cocktail at a final density of 105 to 106 cells/ml. 4. Add the cell suspension to a 60-mm dish and incubate 24 hr. 5. Twenty-four hours post transfection, centrifuge the cells 5 min at 500 × g, 4°C or room temperature, and resuspend the pellet in target cell growth medium. Incubate cells another 24 hr. 6. Assay for reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10), add antibiotic for drug selection (UNIT 9.5), or determine infectious titer (UNIT 9.10). INFECTION OF NONADHERENT CELLS BY COCULTIVATION Nonadherent cells can also be infected by cocultivating them with retroviral producer cells for 48 hr.


Materials Transfected packaging cells (see Basic Protocol 1) in 60-mm tissue culture dishes Retroviral supernatant 4 mg/ml polybrene in PBS (APPENDIX 2), filtered and stored at 4° or −20°C 105 to 106 cells/ml nonadherent target cells Target cell growth medium 15-ml conical centrifuge tubes 60-mm tissue culture dishes Sorvall RT-3000B centrifuge (or equivalent) Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F) and for assaying of reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10) or drug selection (UNIT 9.5) 1. Twenty-four hours after packaging cells have been transfected, prepare a 3-ml infection cocktail containing: Retroviral supernatant: ≤1.5 ml 2 µg/ml polybrene 3 × 105 to 3 × 106 nonadherent target cells Target cell growth medium to 3 ml. Replication of transfected retroviral packaging cells without loss of retroviral production can be achieved by irradiating them with 1500 rads (C. Zent, Univ. of Chicago, unpub. observ.) prior to addition of infection cocktail.

2. Aspirate the medium from the transfected packaging cells and gently add the infection cocktail to the cells. Incubate 24 hr. To prevent detachment of adherent transfected packaging cells, add cocktail to the side of the dish rather than directly onto the cells.



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3. Twenty-four hours after the infection cocktail is added (48 hr post transfection), gently remove 2 ml medium (which will contain many nonadherent cells) and transfer to a conical tube. Centrifuge 5 min at 500 × g, 4°C or room temperature. It is not necessary to remove all of the nonadherent cells from the dish because the purpose of this step is to replenish the medium (which is acidic). Sufficient residual medium should remain on the dish to prevent dehydration during the short centrifugation step.

4. Prepare a fresh infection cocktail containing: Retroviral supernatant: ≤1.5 ml (optional) 2 µg/ml polybrene Target cell growth medium to 3 ml. Adding virus in this step is optional, but it may increase infection efficiency.

5. Remove the supernatant and gently resuspend the cell pellet in freshly prepared infection cocktail. 6. Using extreme care to avoid disruption of the adherent cell layer, add the fresh infection cocktail to the wall of the plate. Incubate 24 hr. 7. At 72 hr post transfection, collect nonadherent cells from the dish by gentle pipetting. It is necessary to sufficiently wash the plate with FBS or medium to remove most nonadherent cells while minimizing contamination with adherent cells. Typically, there is about 10% contamination of nonadherent cells. Because adherent cells detach in sheets, rather than as individual cells, many of the cells can be removed by allowing th large clumps of cells to settle out and removing the others (see Support Protocol, step 5). Alternatively, cells can be transferred to a new 60-mm dish and given sufficient time for the adherent cells to reattach (∼3 hr), after which the nonadherent cells can be removed.

8. Centrifuge the suspension of nonadherent cells 5 min at 500 × g, 4°C or room temperature. Resuspend the cells in target cell growth medium. Plate the target cells in 60-mm tissue culture dishes. Incubate an additional 24 to 48 hr. 9. Assay for reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10) or add antibiotics for drug selection (UNIT 9.5), or assay for the gene of interest. ALTERNATE PROTOCOL 4

INFECTION OF NONADHERENT CELLS BY SPIN INFECTION Alternatively, nonadherent target cells can be infected by centrifuging the cells in the presence of infection cocktail, a method modified from Kotani et al. (1994). Materials Retroviral supernatant 4 mg/ml polybrene in PBS (APPENDIX 2), filtered and stored at 4° or −20°C Target cell growth medium Exponentially growing nonadherent target cells 24-well tissue culture plates Sorvall RT-3000B centrifuge with microplate carrier (or equivalent) 10-cm tissue culture dishes Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F) and for assaying of reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10) or drug selection (UNIT 9.5)




1. For each well to be transfected, prepare a 2-ml infection cocktail containing: Retroviral supernatant: ≤1 ml 4 µg/ml polybrene Target cell growth medium to 2 ml. 2. Add 1–2 × 106 exponentially growing nonadherent target cells per well of a 24-well tissue culture plate. Centrifuge 5 min at 500 × g (1000 rpm in Sorvall RT-3000B rotor), 4°C or room temperature. The cell number should be adjusted so that cells completely cover the surface of the well and form a monolayer following centrifugation.

3. Remove the supernatant and add infection cocktail. Centrifuge the cells 1.5 to 2 hr at 1000 × g (2500 rpm in Sorvall RT-3000B rotor), room temperature. 4. Either immediately after centrifugation or after ≤6 hr of incubation, transfer the cells in the infection medium to a 10-cm dish. Dilute to 10 ml final volume with target cell growth medium. Incubate to 48 hr post infection. If the cells are left overnight in only 1 to 2 ml of infection cocktail, the medium will be very acidic by the following morning.

5. Assay cells for reporter gene expression (e.g., UNITS 9.6-9.8 & 9.10) or add antibiotic for drug selection (UNIT 9.5). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

293 cell growth medium Dulbecco’s modified Eagle medium (high glucose) containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 100 U/ml penicillin 100 U/ml streptomycin 2 mM L-glutamine Store at 4°C This medium should be stable for at least 3 months except that L-glutamine has a half-life of 1 month.

HEPES-buffered saline (HeBS), 2× 50 mM N-2-hydroxyethylpiperazine-N′-ethanesulfonic acid (HEPES) 10 mM KCl 12 mM dextrose 280 mM NaCl 1.5 mM Na2HPO4 Adjust pH to 7.05 ± 0.05 Filter through 0.2-µm filter Store in 50-µl aliquots at −20°C Avoid multiple freeze/thaw cycles. To thaw, warm to room temperature and invert or vortex the tube to achieve uniform mixing. Although it is unclear why, the ability of the 2× HeBS solution to produce working calcium phosphate precipitates deteriorates after 6 months to 1 year, even when the solution is stored at −20°C. Fresh 2× HeBS should therefore be prepared every 6 months.



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TNE buffer 50 mM Tris⋅Cl, pH 7.8 130 mM NaCl 1 mM EDTA Store at room temperature indefinately COMMENTARY Background Information


Creation of high-titer retroviral supernatants by transient transfection offers several advantages over the creation of stable cell lines, especially for experimental purposes. The transient methods described in this unit take advantage of the high transfectability and high protein expression in 293 cells. When retroviral vectors (e.g., see Figs. 9.9.5 & 9.9.6) are expressed with the retroviral packaging constructs in these cells, the vector RNA (containing the Ψ site) is efficiently packaged and budded from the surface of the packaging cells. The infectious retrovirus vector may be harvested by removing the packaging cell supernatant. Because the transient methods are not dependent on stable integration of the introduced plasmid(s), high-titer viral supernatant may be harvested from the packaging cells from 36 to 96 hr following transfection. Two different approaches, both utilizing 293 cells, have successfully produced helper-free high-titer retroviral supernatants expressing a wide variety of genes. In the method of Pear et al. (1993), three cell lines (Anjou, Bosc23, and Bing) were made by stably transfecting plasmids expressing gag-pol (Anjou), gag-pol and ecotropic envelope (Bosc23), or gag-pol and amphotropic envelope (Bing) into 293T cells. 293T cells are a derivative of 293 cells that express the SV40 large T antigen (DuBridge et al., 1987). These cell lines were selected for high levels of expression of reverse transcriptase and the appropriate envelope protein. All of the retroviral structural genes are expressed from the Moloney murine leukemia virus (MoMuLV) long terminal repeat (LTR). High-titer retroviral supernatants may be made from these cell lines by introducing either a retroviral vector alone (Bosc and Bing) or a retroviral vector and a plasmid-expressing envelope (Anjou). Because the retroviral structural genes are expressed from two different stably integrated plasmids that contain multiple mutations (Danos and Mulligan, 1988), more than two recombination events are necessary for the creation of replication-competent retrovirus. To date, generation of helper virus by these cell lines has not been reported.

A second generation retroviral packaging cell line, termed φNX, has recently been developed for high-titer, transient production (P. Achacoso and G.P. Nolan, pers. comm.). The major addition is that the gag-pol construct has an IRES-CD8 surface marker, so it is possible to follow gag-pol expression in living cells on a cell-by-cell basis. This facilitates checking the line for continued high-level gag-pol expression using CD8 as proxy. Additionally, non-MoMuLV-based promoters are used to express gag-pol (and env) genes, further minimizing the potential for recombination with vector LTRs. Otherwise, the construction of these cell lines is similar to Bosc23. Amphotropic and ecotropic versions of these have been prepared, and they stably express gag-pol and env proteins over several months. These lines are helper-free and produce retroviral supernatants that have titers similar to Bosc23 and Bing. The methods described in this unit work well with the φNX cell line. In the strategies of Finer et al. (1994), Soneoka et al. (1995) and Naviaux et al. (1996), both the retroviral vector and the retroviral packaging constructs are cotransfected into 293 cells and retroviral supernatants are harvested 48 hr later. These groups found that titers were highest when the packaging constructs were expressed from a cytomegalovirus (CMV) promoter and a retroviral vector in which the 5′ LTR was a hybrid between CMV and MoMuLV sequences. Although previous reports suggested that cotransfection of retroviral vector and packaging constructs could lead to replication-competent retrovirus (Danos and Mulligan, 1988; Pear et al., 1993), these groups have engineered their constructs to minimize regions of homology that might lead to helper virus formation. In particular, deletion of 3′ LTR sequences appears important for preventing the formation of replication-competent retrovirus (Soneoka et al., 1995).

Critical Parameters Several parameters are important to optimize for high-titer retroviral production. Cells should be at optimal density (usually ∼80% confluent) with minimal clumping at the time



of transfection. In addition to cell density, production of high-titer retroviral supernatants also requires that the cells continue to grow for ≥24 hr following transfection. A rough indicator is that the plate should be nearly 100% confluent ∼24 hr after transfection. Several factors that may account for failure to obtain this degree of confluence are the nature of the protein expressed by the retroviral vector, the quantity of input DNA, and the length of chloroquine and/or sodium butyrate treatment. Expression of some proteins (such as P210bcr/abl) appears detrimental to 293 cell growth. Hightiter retroviral supernatants can be obtained with vectors expressing such genes by increasing the number of cells plated prior to transfection and/or decreasing the quantity of transfected retroviral plasmid DNA. For expression of some genes, 293 cells may need to be nearly 100% confluent at the time of transfection. In addition to the protein product, it appears that the quantity of transfected DNA may have a toxic effect upon the cells. For up to 10 to 15 µg transfected DNA per 60-mm plate, increasing the amount of input DNA increases retroviral titer; however, as more DNA is added beyond that point, the quantity of DNA has a toxic effect on the cells. As discussed in the protocols, to prevent toxicity from chloroquine or sodium butyrate, cells should not be exposed to either of these reagents for ≥12 to 14 hr. When trying to optimize titers from a new retroviral construct, it is recommended that cell density, input DNA, and length of chloroquine and/or sodium butyrate treatment all be optimized. If the cells are not confluent at 24 hr post transfection, but reach confluence by 48 hr post transfection, it may be possible to obtain high-titer supernatants between 48 and 72 hr post transfection. Supernatants harvested >96 hr post transfection do not produce high retroviral titers. Another variable that must be optimized is the transfection reaction. Of particular importance is the pH of the 2× HEPES-buffered saline (HeBS), which should be 7.05 ± 0.05. For reasons that are not clear, 2× HeBS loses its effectiveness between 6 months and 1.5 years. Unfortunately, there is no way to detect this except for a bad experimental result. For this reason, it is recommended that fresh stocks of 2× HeBS be prepared every 6 months. Although the modified calcium phosphate– mediated transfection method is inexpensive and gives consistently good results, other methods of gene transfer, including lipid-mediated

transfection (UNIT 9.4), work equally well. A recent report suggests that receptor-mediated, adenovirus-augmented transfection is superior to calcium phosphate–mediated transfection for some cell types (von Ruden et al., 1995). It has not yet been determined if this method is superior for gene transfer into 293 cells. There are several techniques that may increase retroviral titer—adding factors to the transfection or cell growth medium, amplifying the plasmid DNA, decreasing the incubation temperature, and infecting by centrifugation. Addition of 25 µM chloroquine during transfection may increase titers 2-fold; presumably this is due to the lysosomal neutralizing activity of the chloroquine. Sodium butyrate has been reported to increase retroviral titers from several-fold to 1 log by acting as a transcriptional activator of the MoMuLV LTR, as well as other promoters including CMV (Soneoka et al., 1995). Increasing the extracellular concentration of dNTPs may increase retroviral titers up to 10-fold by enhancing endogenous reverse transcription (Zhang et al., 1995). 293T cells, a 293 cell derivative, contain a temperaturesensitive large T antigen, whose permissive temperature is 32°C (DuBridge et al., 1987). Thus, incubation at this temperature in conjunction with the use of constructs that contain the SV40 origin of replication may result in plasmid replication and higher retroviral titers. This may come at a cost, however, as large T–induced replication is associated with higher recombination rates (St.-Onge et al., 1993). Shifting the cells to lower incubation temperatures (32° to 34°C) following transfection has been reported to increase retroviral titers by 5to 15-fold (Kotani et al., 1994). This is hypothesized to result from both improved retroviral survival and increased production. Similarly, infecting by centrifugation may increase retroviral infection by 4- to 10-fold (Kotani et al., 1994). It is important to note that all of the above methods have given variable results in different investigators’ hands, and each should be optimized (see Troubleshooting). Unlike retroviruses expressing the amphotropic and ecotropic envelopes, VSV G protein–pseudotyped retroviruses can be efficiently concentrated 100- to 1000-fold because they can better withstand shearing forces during centrifugation. This attribute together with their polytropic host range makes them a very useful laboratory tool, especially for stable gene transfer into nonmammalian cells.



Supplement 36

Troubleshooting


To maximize retroviral production, it is recommended that conditions be optimized using a nontoxic reporter protein such as lacZ (UNIT 9.10), green fluorescent protein (GFP; UNIT 9.7C), placental alkaline phosphatase (PLAP), or a cell-surface marker. During optimization, both transfection frequency of the producer cells and infection rate of the target cells should be measured. A number of variables that can affect retroviral production have been previously discussed (see Critical Parameters). To avoid drift of the packaging cell lines or highly transfectable 293 cells, it is recommended that they be maintained for ∼20 passages (2 months) before establishing a fresh stock by thawing a frozen vial of cells. It may be possible to carry the cells for longer periods without loss of efficacy, and it is recommended that the cells be assayed at various times to check the ability to produce high-titer supernatants. Bosc and Bing cells were made using 293T cells that contain selectable markers for neomycin (1 mg/ml), hygromycin (400 µg/ml), and mycophenolic acid (50 µg/ml). There does not appear to be any advantage in growing the cells in selection medium. If retroviral titers drop when using these cells, their growth may be tested by growing in hygromycin- and xanthine-guanine phosphoribosyltransferase-containing selection medium. If a majority of the cells are not resistant to these antibiotics, a fresh vial of earlier-passage cells should be thawed rather than attempting to reselect the cells. Similarly, when cells are initially received, aliquots should be tested for resistance to these antibiotics. 293 cells do not contain a selectable marker. It should be noted that the amphotropic envelope–expressing Bing cells release ∼100 units of infectious retrovirus per ml of retroviral supernatant expressing hygromycin resistance which results from the introduction of the gene for hygromycin resistance into these cells by a retroviral vector (W. Pear, unpub. observ.). Replication-competent retrovirus has not been detected in this cell line. The φNX-A cells do not produce a hygromycin-resistant retrovirus (G.P. Nolan, pers. comm.). For efficient infection, addition of a polycation increases infection efficiency by >100-fold (reviewed in Burns et al., 1993). Polybrene gives the best results at concentrations between 2 and 8 µg/ml. Polybrene may, however, be toxic to some cell types. For example, NIH 3T3 cells tolerate 4 µg/ml polybrene for at least 18

hr; however, treatment of these cells with 8 µg/ml for this period is detrimental. If high polybrene concentrations are chosen, the medium should be changed after 3 to 5 hr. For nonadherent cells, including primary bone marrow cultures, 1 to 2 µg/ml polybrene is well tolerated for ≤48 hr. It is possible to use other cations, such as protamine sulfate or poly-L-lysine; however, infectious titers are diminished by 2- to 4-fold with these agents (Burns et al., 1993). Efficient infection of nonadherent cells has been cumbersome due to the need to cocultivate the retroviral producer cells with the target cells. It is difficult to separate the producer cells from target cells. An additional difficulty is imposed by 293 cells which, because of their high density, tend to rapidly acidify medium. For this reason, it is important to replenish medium daily when performing cocultivations. An alternative to cocultivation is spin infection. This technique appears to produce equivalent results as cocultivation for several nonadherent cell lines (W. Pear, unpub. observ.). Although the original protocols of Kotani et al. (1994) indicate that infections should be performed at 32°C, many centrifuges are unable to maintain that temperature; equivalent results are obtained by spin infecting at room temperature (W. Pear, unpub. observ.). One major problem with retroviral gene transfer is that, despite producing high-titer retroviruses, it is difficult to maintain gene expression following infection. Although there are a number of potential mechanisms at the transcriptional and posttranscriptional levels, a teleologic explanation is that the retrovirally expressed protein is detrimental to cell growth resulting in outgrowth of clones that express lower levels of the protein. This is a particular problem with vectors that contain an internal promoter expressing a selectable marker, because even though the marker is selected there is no selective pressure to express the gene of interest. One method to increase the likelihood that the gene of interest will be expressed (although there may still be posttranscriptional mechanisms that affect the level of expression) is to use a polycistronic retroviral vector (see UNIT 9.10). Other methods of overcoming gene toxicity are being developed. Particularly promising are the tetracycline-inducible promoters originally described by Gossen and Bujard (1992); these promotors have shown some degree of success when incorporated into retro-



viral vectors (Hoffman et al., 1996; Paulus et al., 1996).

Anticipated Results Transfection efficiencies of 293 cells should be in the range of 30% to 60%. Infectious titers of 106 to 107 infectious U/ml should be expected from most retroviral vectors when titered on murine NIH 3T3 cells. Titers may be slightly reduced with large cDNA inserts (>4 kb). Titers may be lower on other cell types, particularly cells expressing only the amphotropic envelope receptor. For VSV G protein pseudotyping followed by concentration by centrifugation, titers of 109 infectious U/ml should be expected.

Time Considerations Plating cells, transfections, infections, and medium changes may take from 200 ml), we recommend the longer spins and the use of a Beckman JA-14 rotor or its equivalent. With smaller volumes, we recommend the SW-27 or SW-41Ti swinging bucket rotors with shorter spins (2 hr at 20,000 rpm, 4°C). Swinging-bucket rotors make a firm pellet at the bottom of the tube, so that it is easy to pour off the supernatant without dislodging the pellet; in addition, it is easier to see the pellet in tubes spun in swinging-bucket rotors.

7. Discard supernatant while carefully avoiding discarding the pellet, which sometimes dislodges when using the large volumes.

Large-Scale Preparation and Concentration of Retrovirus Stocks

If using the JA-14 rotor, when removing the bottles from the rotor, mark the side of the bottle that should contain the pellet. The pellet will be smeared down the side of the bottle and will be difficult to see. Although normally the supernatant is discarded, a



small volume of the supernatant may be saved to titer to determine whether the virus was indeed pelleted.

8. Gently resuspend the pellet in DMEM-10 using 0.1% to 1% of the original volume. Resuspension may take 2 hr if the pellet is fairly sticky. It is convenient to leave the centrifuge bottle in an ice bucket in the tissue culture hood and pipet the suspension approximately every 15 min. A 1- or 2-ml pipet can be used if large volumes were concentrated. If so, leave the original pipet in the bucket and use this pipet for the complete resuspension. This is because the virus pellet often sticks to the first pipet that touches it. Sometimes the pellet can be seen as an almost translucent, gooey yellow pellet. If the JA-14 rotor was used, be sure to wash down the sides of the bottle, as this is where much of the stock will be located. To resuspend pellets made in the SW-27 or SW-41 rotors, use a pipettor suitable for small volumes. In either case, avoid making bubbles, as this can denature the viral proteins. Alternatively, an easier way to resuspend pellets made in the swinging-bucket rotors is to pour off almost all of the supernatant, e.g., leaving only 0.10 to 0.15 ml in an SW-27 large bucket; this may require keeping a Pasteur pipet handy to remove the last few drips of supernatant from the mouth of the tube as one pours it off. Return the tube to the bucket and replace the bucket cap. Place the buckets on a shaker at low speed in a 4°C cold room, or place them in an ice bucket and slowly shake 1 to 2 hr on the benchtop. After shaking, gently pipet up and down several times and transfer to a smaller tube.

Titer and freeze concentrated virus 9. Store the virus at −70° or −80°C in small aliquots (typically 5 to 50 µl). Systematic tests in our laboratory have indicated that, in general, stocks can be frozen and thawed several times with no loss of titer. However, we have occasionally observed loss of titer over time in the freezer, or upon freeze-thaw. Thus, the most conservative approach is to make small aliquots and to minimize the number of times they are thawed and refrozen. We have also found that small volumes can desiccate over long-term storage. To combat this, we place a drop of the oil used to cover PCR reactions onto each small aliquot before freezing.

10. Titer an aliquot of concentrated and unconcentrated virus as described in UNIT 9.10. The titer can also be determined using an aliquot before freezing the stock.

CONCENTRATION BY PEG PRECIPITATION AND CHROMATOGRAPHY A method that is rapid and easy is to precipitate virus with polyethylene glycol, with an optional step of chromatography on Sepharose (Aboud et al., 1982).


Additional Materials 5 M NaCl, filter sterilized Polyethylene glycol (PEG) 6000, filter sterilized NTE buffer (see recipe) Sepharose CL-4B or CL-2B (Pharmacia) Savant high-speed centrifuge or Beckman SW-41Ti rotor Additional equipment for preparation of Sepharose column (UNIT 5.6) Precipitate virus with PEG 1. Prepare a virus stock (see Basic Protocol, steps 1 to 4) 2. Add 5 M NaCl to the virus stock to 0.4 M final while stirring at 4°C. Introduction of DNA into Mammalian Cells


Supplement 37

3. Slowly add PEG 6000 to 8.5% final (w/v) and continue stirring 1 to 1.5 hr at 4°C. For reasons that are not clear, some batches of PEG do not work well as they lead to problems in resuspending the pellet.

4. Collect the precipitate by centrifuging 10 min at 7000 × g (e.g., at 7500 rpm in a SW-41Ti rotor or 10,000 rpm in a Savant high-speed centrifuge.) 5. Dissolve the pellet in NTE buffer in 1% of the original volume. If necessary, proceed to step 6 for column chromatography. The stock can be used directly after this step or stored at −70° or −80°C. It can also be further purified on a Sepharose CL-4B column to remove PEG.

Carry out Sepharose chromatography 6. Prepare a column of Sepharose CL-4B or CL-2B as described in Protocol), equilibrating in NTE buffer.

UNIT 5.6

(Support

Column size will depend upon the amount of virus pellet; 10-ml Econo-Columns from Bio-Rad work well for pellets prepared from 100 ml of virus stock.

7. Apply the virus to the top and chromatograph at 1 ml/min with NTE buffer. 8. Collect 0.3- to 0.5-ml fractions. 9. Assay fractions for virus by measuring absorbance at 280 or 260 nm (see Aboud et al., 1982) or by reverse transcriptase (RT) assay (UNIT 9.13). 10. Pool appropriate fractions and titer as in UNIT 9.10. For examples of virus recovery as determined by both Xgal CFU bioassay and RT assay, see Figure 9.12.1.

7000

100,000 kcpm/ml kCFU/ml

80,000

6000

60,000

4000 3000

40,000

kCFU/ml

kcpm/ml

5000

2000 20,000

1000 0

0 0

2

4

6

8

10

Elution volume (ml)

Figure 9.12.1 Ψ2 BAG supernatant (8 ml) was concentrated 100-fold by PEG precipitation. The resulting 80 µl was loaded onto 10 ml of Sepharose CL-2B in a Bio-Rad Econo-Column. Fractions of 0.5 ml were collected and tested for both reverse transcriptase activity and Xgal CFU. Large-Scale Preparation and Concentration of Retrovirus Stocks



CONCENTRATION USING MOLECULAR-WEIGHT-CUTOFF FILTERS This method is the simplest means of concentrating small to medium volumes of retrovirus stocks. Supernatants prepared as in steps 1 to 4 of the Basic Protocol are centrifuged through filters, either the Centricon-30 microconcentrator from Amicon or the CentriCell 60 from Polysciences, essentially following the manufacturers’ instructions. These filters allow passage of molecules that are much smaller than the virus (e.g., Centricon-30 filters allow passage of molecules of 500 Ci/mmol. Because volume will vary with different lots, the volume of distilled H2O to add should be varied to compensate.


Detection of Helper Virus in Retrovirus Stocks

The production of wild-type, replicationcompetent helper virus by packaging cell lines can be an issue of concern when using replication-incompetent vectors. If the vectors are being used for lineage analysis, the virus stock must be free of helper virus. Similarly, for most infections of young animals, where the wildtype viruses are leukemogenic, helper viruses must not be present. The genome that supplies the gag, pol, and env genes in the Ψ2, Ψam, PA12, PA317, and Q2bn packaging cell lines (UNIT 9.9) does not encode the Ψ sequence, but can still become packaged—although at a low frequency. If the helper genome is coencapsidated with the vector genome, recombination in the next cycle of reverse transcription can occur. If recombination allows the helper genome to acquire the Ψ sequence from the vector genome, a recombinant that is capable of autonomous replication will result. This recombinant can spread through an entire culture (although slowly due to envelope

interference). Once this occurs, it is best to discard the producer clone, as there is no convenient way to eliminate the wild-type virus. As would be expected, helper virus contamination happens with a greater frequency in stocks with high titer. The PA317 packaging line has a safer design than Ψ2, Ψam, PA12, and Q2bn, while the ΨCRE, GP + E-86, ΩE, ΨCRIP, and Isolde lines probably have the best design for not producing helper virus. Recombination leading to helper virus creation has not yet been observed in ΨCRE, GP + E-86, ΩE, ΨCRIP, or Isolde, but one must be cautious and assay stocks produced by these lines (for a review of lines, see Table 9.9.1). These protocols can also be used to determine the host range of the helper virus. This can be tested by infection of cells of different species [e.g., dog (D17) or human (HeLa) cells cannot be infected with ecotropic virus, only with amphotropic virus]. Alternative methods to detect helper virus include the XC plaque assay (Rowe et al.,



1970) and S+L− assay (Eckner and Hettrick, 1977; Miller and Buttimore, 1986). These assays require additional cell lines. The assay listed here for horizontal spread is quite sensitive but very time-consuming. The RT assay is sensitive, but does not always detect crippled helper (virus that can replicate, but is mutant and does not replicate as well as wild-type).

Time Considerations Virus supernatant generation (Basic Protocol) requires 10 days. To titer the stocks for presence of viruses carrying the selectable marker or lacZ, another 10 to 13 days are needed (see UNIT 9.11). To test supernatants for reverse transcriptase activity (Alternate Protocol 2), 50 samples can be processed in 2 to 3 hr.

Troubleshooting If supernatants are assayed by titration of selectable markers and many (e.g., greater than several thousand) marker-resistant colonies are observed, the original test stock contained wild-type helper virus. Once this occurs, it is best to discard the producer clone, as there is no convenient way to eliminate the helper virus. If a smaller number of colonies result than expected (see Anticipated Results), there are several possible explanations. There may be “crippled” helper in the original test stock generated by an imperfect recombination event. For many applications, crippled helper is not a confounding influence. Alternatively, some of the Ψ− genome (e.g., the genome carrying gag, pol, and env) can transfer to the first set of infected NIH 3T3 cells and lead to a low titer, without true helper virus contamination (e.g., see Danos and Mulligan, 1988).

Anticipated Results Virus stock that contains wild-type helper virus should produce more than several thousand colonies carrying the neo marker (G418resistant) colonies on the assay dishes generated by step 9 of the Basic Protocol or several thousand lacZ+ colonies generated by step 11 of Alternate Protocol 1. If the reverse transcriptase assay is used, the RT activity from the test supernatants should be compared to that in the control supernatants (cells infected with a known helper-free virus stock, and cells infected with a stock of known wild-type helper virus). Stocks contaminated with helper virus should show a >1000-fold enhancement of RT activity when compared to helper-free controls. A truly helper-free stock will be equal in RT activity to that generated by the helper-free control.

Literature Cited Danos, O. and Mulligan, R.C. 1988. Safe and efficient generation of recombinant retroviruses with amphotropic and ecotropic host ranges. Proc. Natl. Acad. Sci. U.S.A. 85:6460-6464. Eckner, R.J. and Hettrick, K.L. 1977. Defective Friend spleen focus-forming virus: Interfering properties and isolation free from standard leukemia-inducing helper virus. J. Virol. 24:383396. Goff, S., Trakman, P., and Baltimore, D. 1981. Isolation and properties of murine leukemia virus mutants: Use of a rapid assay for release of virion reverse transcriptase. J. Virol. 38:239-248. Miller, A.D. and Buttimore, C. 1986. Redesign of retrovirus packaging cell lines to avoid recombination leading to helper virus production. Mol. Cell. Biol. 6:2895-2902. Morgan, B.A. and Fekete, D.M. 1996. Manipulating gene expression with replication competent retroviruses. Methods Cell Biol. 51:185-218. Omer, C.A. and Faras, A.J., 1982. Mechanism of release of the avian retrovirus tRNATrp primer molecule from viral DNA by ribonuclease H during reverse transcription. Cell 30:797-805. Rowe, W.P., Pugh, W.E., and Hartley, J.W. 1970. Plaque assay techniques for murine leukemia viruses. Virology 42:1136-1139. Stoker, A.W. and Bissell, M.J. 1987. Quantitative immunocytochemical assay for infectious avian retroviruses. J. Gen. Virol. 68:2481-2485.

Contributed by Constance Cepko Harvard Medical School Boston, Massachusetts Warren Pear (proviral rescue) University of Pennsylvania Philadelphia, Pennsylvania



Supplement 36

Retrovirus Infection of Cells In Vitro and In Vivo There are many applications in which retrovirus vectors are used as transduction agents. In some cases, the vector carries a gene that one wishes to express in a target cell in order to study the function of that gene. In other cases, the virus is used to introduce a histochemical marker gene into cells in order to follow their fate. Retrovirus vectors can also be used in a variety of cells type to investigate regulatory sequences in which a reporter gene and regulatory sequences are carried by the vector and to immortalize or transform primary cells by transduction of oncogenes. For each application, the infection protocol may vary and must often be optimized. Guidelines for infection of cells in some typical in vivo and in vitro experiments are presented here. In addition to these guidelines, the cited literature contains protocols that have been used successfully to infect cells under specific circumstances and may provide a good starting point for similar cells. To optimize infection of a particular type of cell, it is often advantageous to use vectors that are easy to score. These include vectors carrying selectable or screenable markers (such as BAG; Figs. 9.10.1 and 9.10.2). CAUTION: When working with human blood, cells, or infecting agents, strict biosafety practices must be followed (see UNIT 16.15).

INFECTION OF CELLS IN VITRO Infection of target cells in vitro is accomplished by simply incubating the virus with the cells (e.g., UNIT 9.10, Basic Protocol 2). For most in vitro applications using a murine virus, a polycation such as polybrene is used to aid viral infection. These polycations apparently promote virus binding to the host cell surface by reducing electrostatic repulsion between the negatively charged surfaces of the cell and virion. When incubating target cells with virus in vitro, problems may occur if undiluted, hightiter virus is used directly on some cell types— i.e., fusion of target cells can occur with subsequent death of most fused cells within a few days. This is presumably due to the virus serving as a fusogen (i.e., binding to the surface of more than one cell and promoting membrane fusion). Alternatively, cells to be infected in vitro can be incubated with the packaging line that produces the desired vector (cocultivation Contributed by Constance Cepko and Warren Pear Current Protocols in Molecular Biology (1996) 9.14.1-9.14.6 Copyright © 1996 by John Wiley & Sons, Inc.

method). This method is used to infect hematopoetic cells and appears to greatly increase the infection efficiency (Williams et al., 1984; Lemischka et al., 1986). In some cases, it may be desirable to prevent cell division of the producer cells during or after cocultivation. This can be achieved by using protocols that allow virus production, but that prevent further cell division of the producer cells. For example, prior to cocultivation, confluent or nearly confluent producer cells can be killed by irradiation (2800 rad for NIH 3T3 cells, 1500 rad for 293 cells) or a 3-hr treatment with mitomycin C (10 µg/ml in medium) followed by several rinses with medium. After this treatment, the cells to be infected are plated onto producer cells. Target cells should continue to grow, but producer cells will die as they can no longer divide. If the target cells are nonadherent, they can be removed by simply washing the producer monolayer gently after ∼48 hr of cocultivation. An alternative to cocultivation is spin infection (UNIT 9.11; Kotani et al., 1994), which achieves similar infectious titers without the difficulties caused by the presence of producer cells. It is difficult to generalize about the efficiency of infection, although it can approach 100%. For the most part, the variables that influence infectability of a given cell are unknown. However, it is clear that for optimal results, the cells should be as mitotically active as possible. Although virus will enter cells and undergo partial reverse transcription in nonmitotic cells, there will be no integration or expression of the viral genes. If virus is applied to cells that are not dividing, it may remain competent to finish reverse transcription and integration for 1 to 2 days, although this has not been carefully determined for most cell types. One approach that may overcome the difficulties in transducing non-dividing cells is the use of retroviral packaging cell lines and vectors based on human immunodeficiency virus (HIV; Naldini et al., 1996). This approach has successfully produced high-titer, helper-free viruses that are able to infect cell lines arrested at various phases of the cell cycle at frequencies that far exceed those attainable using Moloneybased packaging systems. In addition, these vectors are able to infect adult rat neurons at low frequency. Expression of viral gene products is usually

UNIT 9.14



assayed two or three cell cycles after infection. When assaying lacZ expression in NIH 3T3 cells via Xgal staining (UNIT 9.10), we have found that the number of clones peaks at ∼48 hr postinfection (three cell cycles). However, Xgal staining is visible within 24 hr in a subset of the infected cells. Infections of tissue explants or cultured embryos can also be performed essentially as described above. A tissue explant can be bathed in as much virus stock as is desired for a few hours (when using murine viruses or avian viruses of any subgroup other than A, include polybrene at 8 µg/ml), or it can be cultured over a monolayer of producer cells (as above in the cocultivation method).

Creation of Retroviral cDNA Libraries

Retrovirus Infection of Cells In Vitro and In Vivo

Expression cloning has been extremely useful for isolating a wide variety of genes, including cell surface markers, cytokines, and multiple receptors (see UNIT 6.11). This methodology relies on high levels of gene expression in the transfected cell line and the ability to recover the appropriate cDNA by plasmid rescue. To accomplish the former, COS cells are frequently used for expression cloning, as the expression of SV40 large T antigen causes high levels of protein expression from vectors that contain the SV40 origin of replication. Although expression cloning in COS cells has been extremely useful for the expression cloning of a number of cDNAs, this approach is limited to cell lines that are efficiently transfected and to phenotypic assays with a short readout period. These limitations have made it difficult to use expression cloning strategies in primary hematopoietic cells due to their poor transfectability and the long-term culture required for phenotypic selection. In contrast to expression cloning in COS cells, retroviral cDNA libraries offer the possibility of introducing cDNAs into a wide variety of cell types, including primary cells. As a result, it is possible to devise novel assays to select cDNAs that are based on specific properties of the cells themselves, such as transformation, growth factor dependence, or in vitro differentiation. Retroviral expression cloning has been successfully used to clone cDNAs using NIH 3T3– based packaging lines. These strategies have been useful for identifying genes involved in factor-dependent growth (Rayner and Gonda, 1994; Wong et al., 1994) and transformation (Whitehead et al., 1995), as well as for the selection of peptides and antisense messages that induce etoposide resistance (Gudkov et al.,

1993, 1994). Although overcoming some of the limitations of COS-based expression cloning, use of NIH 3T3–based packaging lines has two major disadvantages for expression cloning. These are that (1) it is difficult to obtain high titers by transient transfection, limiting the complexity of the library, and (2) the frequency of each cDNA may change during selection of the transfected population and the resulting retroviral stock may not represent the original cDNA library. An advance in the ability to prepare cDNA expression libraries has been the methodology to prepare high titer retroviral supernatants by transient transfection (UNIT 9.11; reviewed in Onishi et al., 1996). These approaches overcome the major disadvantages of the NIH 3T3–based packaging cells—i.e., it is possible to obtain high transient transfection efficiencies and it is unnecessary to select the cells prior to harvest of the retroviral stock. An application of the use of transient retroviral production for expression cloning has been described for cloning cDNAs from factor-dependent cell lines (Kitamura et al., 1995). In this report, the authors were able to develop expression libraries with complexities of >106 and detect cDNAs at a frequency of 1 in 107 (Kitamura et al., 1995). The technology for retroviral expression cloning is presently evolving, and the reader should consult the most recent literature for the best choice of packaging cell line and retroviral vector. With the methods for transient retroviral production, the basic strategy is to clone a cDNA library into a retroviral vector, transfect the library into the packaging cells, harvest the retroviral stock between 48 and 72 hr posttransfection, and infect the target cells. These methods are described in UNIT 9.11. The target cells are subsequently assayed for the phenotype of interest. Some considerations in the choice of retroviral vectors are the presence of multiple cloning sites for efficient cloning of the cDNAs into the vector, the presence of the extended packaging signal for improved titer (Bender et al., 1987), the use of IRES (internal ribosome entry site) elements rather than internal promoters to increase the likelihood that the cloned cDNA will be expressed, and the presence of primers that surround the multiple cloning site so that the cDNA can be amplified by PCR following phenotypic selection. Another consideration is the infection efficiency of the target cells. For example, in the studies of Onishi et al. (1996), the authors were able to infect Ba/F3 and BW5147 cells with >50% efficiency but infection of LG3 myeloid cells



resulted in an infection efficiency of only 8.3%. To obtain adequate infection efficiencies, it m ay be necessar y to p seu do type the retroviruses (UNIT 9.11) or introduce a retroviral receptor into the cell line to be assayed (UNIT 9.9). One drawback to high infection efficiencies, however, is that with higher infection efficiency, the infected cells are more likely to contain multiple integrated proviruses (Onishi et al., 1996). This may increase the difficulty in identifying the cDNA that confers the phenotype of interest. Another consideration in the generation of retroviral cDNA libraries is whether the library should be normalized prior to screening. An advantage of normalization is that there will be an “enrichment” of rare messages; however, the cost is that the resulting cDNA library is no longer representative of initial population of cDNAs.

INFECTION OF RODENTS AND CHICKS IN VIVO Transduction of genes via retrovirus vectors can be performed in vivo. The following paragraphs describe in vivo infection of rodents, the laboratory animal most often used in experimental procedures; however, the techniques apply to other species (e.g., avian) as well. For more detailed information on infection of chick embryos, see Fekete and Cepko (1993a,b), Homburger and Fekete (1996), and Morgan and Fekete (1996). NOTE: Detailed protocols for the care and handling of laboratory animals are beyond the scope of this unit. The reader is referred to Current Protocols in Immunology, Chapter 1 (Donovan and Brown, 1995) for instructions on proper animal restraint, anesthesia, injections, and euthanasia techniques that are essential to the infection methods described.

Infection of Postnatal Animals In Situ Volume and titer of virus stock. The volume that can safely be delivered to a tissue in vivo is generally quite small—0.1 to 1 µl. It is therefore important to prepare high-titer (usually concentrated) virus stock (UNIT 9.12). In general, virus titer should range from 106 to 108 CFU/ml. Because of these limitations, it is quite important to deliver the virus directly to the area containing the highest percentage of mitotic target cells. It is unclear if there are factors in tissue fluids (e.g., ventricular fluids or cerebral spinal fluids) that inhibit or destroy viral infectivity. Delivery of virus. Virus delivery to postnatal animals is fairly straightforward. It is possible

to use a hand-held Hamilton syringe with a 33-G needle or use a drawn-out glass pipet. If glass pipets are used, the size of the needle tip should be determined empirically for the tissue under study. The skin, and even the skull, are soft enough on the first few days after birth for direct injection into the tissue. For tougher injection sites on older animals, it may be necessary to first make a hole using a stronger needle, such as one made of steel, prior to inserting a more delicate needle at a specific site. Coinjection with a dye such as 0.05% (w/v) trypan blue or 0.025% fast green aids in detecting the accuracy of injections and does not impair viral infectivity. Rodents ≤7 days old can be anesthetized by simply cooling on ice for a few minutes. Landmarks (e.g., sutures or blood vessels) near the area to be injected can be visualized using a fiber-optics light source. The injection is made directly into the desired area using a hand-held pipet. It is best to practice a series of injections with dye alone and then immediately dissect the animal for examination of the injection site. After the injection method is perfected, virus can be injected. Animals can then be examined for evidence of viral infection at any time, depending upon the experimental design. Setup of pilot experiments to optimize injection and expression efficiency. When performing infections of tissue for the first time, or when infection and expression efficiency of the target cells is unknown, it is useful to perform pilot experiments using a retrovirus vector carrying a histochemical marker gene such as lacZ. Infection with a lacZ-coding virus allows for examination of the injected animal a few days after infection to determine if the target cells have been infected and are capable of expressing the viral genes. For such experiments, it is necessary to optimize conditions of the Xgal staining reaction (UNIT 9.10) for the particular tissue under study. An excellent way to simultaneously determine the accuracy of injection, and whether the Xgal histochemistry is working, is to inject cells that contain the lacZ gene, such as the Ψ2 BAG producer cells (obtainable from ATCC, #CRL 9560; UNIT 9.10). The cells can be prelabeled with a fluorescent dye such as carboxyfluorescein diacetate succinimyl ester (CFSE, Molecular Probes; prepare 10 mM stock in DMSO and store in foil at 4°C). As detailed by Bronner-Fraser (1985), cells suspended in PBS are exposed to 0.3 mM CFSE solution at 37°C for 30 min just before trypsin treatment. After dissociation, the cells are pelleted and washed



Supplement 36

twice with medium or PBS, and resuspended to a concentration of 108 cells/ml. A few minutes after injection of the CFSE-labeled cells, the animal should be killed, the tissue fixed, then stained with Xgal (UNIT 9.10). The injected cells can then be monitored independently from the Xgal staining by viewing the CFSE fluorescence under UV illumination. If the Xgal histochemistry is working properly, no fluorescent cells should be visible because Xgal-stained cells are usually so full of indigo dye that all fluorescence is absorbed. Once the injection method and Xgal histochemistry have been optimized, BAG virus is injected into the site and the tissue is examined for viral infection several days later. If the Xgal histochemistry is not working well, fluorescent cells will be visible, indicating the site of the injection and the failure of the Xgal histochemistry. In this case, one should vary fixation and tissue preparation methods until good Xgal staining is achieved (Cepko, 1989). Alternatively, the DAP retrovirus vector (Fields-Berry et al., 1992) encoding human placental alkaline phosphatase can be used. A stable Ψ2 line that makes DAP at titers of 1–2 × 106 CFU/ml (more reliably than Ψ2 BAG) can be obtained from ATCC (CRL #1949). Histochemical detection of the alkaline phosphatase enzyme is as simple as Xgal histochemistry (Fields-Berry et al., 1992). For further discussion of both Xgal and alkaline phosphatase assays, see Cepko et al. (in press).

Infection of Prenatal Rodents In Utero


Injections made in utero (through uterine wall) are performed with drawn-out glass pipets (e.g., see Austin and Cepko, 1990). The actual diameter and shape of the tip should be determined empirically (e.g., for injections of the lateral ventricle through the uterine wall of rats at embryonic day 15—E15—the outside diameter of the tip is ∼50 µm and the inside diameter is ∼15 µm). Animals are anesthetized with a mixture of ketamine (20 to 40 mg/kg) and xylazine (3 to 5 mg/kg) and are opened via an incision along the midline. The orientation of an embryo can be determined by visualizing the head with fiber optics. Injections of 0.1 to 1.0 µl are made through the uterine wall into the area of interest. When the target of the injection is the lateral ventricle, the inoculum can be seen filling the lateral ventricle when properly delivered to this area. Other areas may not be as visible and practice injections followed by immediate dissection and examination are again recommended. The mother is

then closed with suture. The injected animals can be delivered prenatally, or allowed to finish gestation and be born normally, to be sacrificed at a postnatal age, depending upon the design of the experiment.

Infection of Prenatal Rodents Exo Utero It is difficult to make precisely directed injections into many of the mitotic zones of prenatal mammals through the uterine wall. The exo utero surgical procedure developed by Muneoka et al. (1986) at least partially circumvents this problem. In this procedure, mouse embryos at embryonic days 11 to 19 (E11 to E19) are released from the uterus by cutting the uterine wall, but remain attached to the uterus via the placenta. The abdominal cavity of the mother is filled with a buffered saline solution to protect the embryos. An incision can be made in the extra-embryonic membranes that surround the embryo so that embryo can be directly manipulated or injected. Subsequently, the extra-embryonic membranes are closed with fine sutures. The embryos can be brought to term in the abdominal cavity and delivered by Caesarean section. Injections can be made through an incision in the extra-embryonic membranes that are subsequently closed with 10-0 suture. However, in our hands (Turner et al., 1990), embryonic and neonatal survival was improved by injecting directly through the extra-embryonic membranes without an incision or suture. Although injections made by this approach were more difficult to target, 25% of the embryos injected at E13 survived to adulthood and all injected animals contained clones of retrovirally infected cells. Additional factors that influence the success of this method are choice of mouse strain and health of the mouse colony. Outbred mouse strains such as CD-1 or Swiss Webster appear to be best, but even these strains may have different embryo survival rates when obtained from different suppliers or colonies. Apparently healthy mouse colonies can harbor subclinical infections that do not affect unoperated embryos but that can stress operated embryos beyond their ability to survive.

Ex Vivo Infection of Murine Bone Marrow Cells and Bone Marrow Transplant Ex vivo retroviral infection of murine bone marrow has been used for in vitro experiments as well as for subsequent transfer and rescue of



lethally irradiated animals. The goal of many of these experiments is to transduce hematopoietic stem cells; however, the methods discussed below can be altered to optimize infection of a particular hematopoietic cell type. As discussed below, the conditions for infecting hematopoietic stem cells are still being developed, and the reader should consult the most recent literature to ascertain the best conditions. Optimal conditions for irradiating mice need to be established for each irradiator and are also dependent on weight, age, and mouse strain. Donor mice (age 5 to 8 weeks) are treated with 250 µg per gram (mouse weight) 5fluorouracil (5-FU) by tail vein injection in a volume of ≤200 µl. 5-FU treatment kills dividing cells, and in doing so, enriches for hematopoietic progenitor cells (van Zandt, 1984; Bodine et al., 1991). Five days later, donor mice are sacrificed and the tibia and fibias are removed. These bones are flushed with 2 to 3 ml per bone of PBS or DMEM using a 25- to 27-G needle attached to a syringe. The flushed marrow cells (including red blood cells) are pooled, centrifuged 10 min at 500 × g, 4°C, and resuspended in 106 cells/ml of a prestimulation cocktail consisting of DMEM, 15% fetal bovine serum, antibiotics (penicillin and streptomycin), 6 ng/ml IL-3, 10 ng/ml IL-6, and 100 ng/ml stem cell factor. The bone marrow cells are incubated in the prestimulation cocktail for 24 to 48 hr prior to infection. A number of other cytokines have been tried, including IL-1α, and the concentrations of the cytokines used vary widely from lab to lab (Bodine et al., 1989, 1994; Daley et al., 1990; Fraser et al., 1992; Pear et al., 1996). The cells are infected by either spin infection or cocultivation (UNIT 9.11) in the above infection cocktail with 4 µg/ml polybrene added. In addition, viral supernatant may replace one-third of the volume of DMEM. Following infection, nonadherent cells are removed and injected into the tail vein of lethally irradiated 4- to 8-weekold recipient mice in a volume of ≤200 µl/mouse. Approximately 2–5 × 105 unfractionated bone marrow cells per mouse are necessary to ensure long-term reconstitution of most mouse strains. The animals are subsequently monitored for bone marrow reconstitution.

LITERATURE CITED Austin, C.P. and Cepko, C.L. 1990. Cellular migration patterns in the developing mouse cerebral cortex. Development 110:713-732. Bender, M.A., Palmer, T.D., Gelinas, R.E., and Miller, A.D. 1987. Evidence that the packaging signal of Moloney murine leukemia virus extends into the gag region. J. Virol. 61:1639-1646. Bodine, D.M., Karlsson, S., and Nienhuis, A.W. 1989. Combination of interleukins 3 and 6 preserves stem cell function in culture and enhances retrovirus-mediated gene transfer into hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 86:8897-8901. Bodine, D.M., McDonagh, K.T., Seidel, N.E., and Nienhuis, A.W. 1991. Survival and retrovirus infection of murine hematopoietic stem cells in vitro: effects of 5-FU and method of infection. Exp. Hematol. 19:206-212. Bodine, D.M., Seidel, N.E., Gale, M.S., Nienhuis, A.W., and Orlic, D. 1994. Efficient retrovirus transduction of mouse pluripotent hematopoietic stem cells mobilized into the peripheral blood by treatment with granulocyte colony-stimulating factor and stem cell factor. Blood 84:1482-1491. Bronner-Fraser, M. 1985. Alterations in neural crest migration by a monoclonal antibody that affects cell adhesion. J. Cell Biol. 101:610-617. Cepko, C.L. 1989. Lineage analysis and immortalization of neural cells via retrovirus vectors. In Neuromethods, Vol. 16: Molecular Neurobiological Techniques (A.A. Boulton, G.B. Baker, and A.T. Campagnoni, eds.) pp. 367-392. Humana Press, Clifton, N.J. Cepko, C.L., Ryder, E., Fekete, D.M., and Bruhn, S. In press. Detection of β-galactosidase and alkaline phosphatase activities in tissue. In Methods in Cell Biology (D. Spector, L. Leinwand, and R. Goldman, eds.). Cold Spring Harbor Laboratory Press. Cold Spring Harbor, N.Y. Daley, G.Q., Van Etten, R.A., and Baltimore, D. 1990. Induction of chronic myelogenous leukemia in mice by the P210 bcr/abl gene of the Philadelphia chromosome. Science 247:824830. Donovan, J. and Brown, P. 1995. Care and handling of laboratory animals. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 1.0.1-1.8.4. John Wiley & Sons, New York. Fekete, D.M. and Cepko, C.L. 1993a. Retroviral infection coupled with tissue transplantation limits gene transfer in the chick embryo. Proc. Natl. Acad. Sci. U.S.A. 90:2350-2354. Fekete, D.M. and Cepko, C.L. 1993b. Replication competent retroviral vectors encoding alkaline phosphatase reveal spatial restriction of viral gene expression/transduction in the chick embryo. Mol. Cell. Biol. 13:2604-2613. Introduction of DNA into Mammalian Cells


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Fields-Berry, S.C., Halliday, A.L., and Cepko, C.L. 1992. Novel recombinant retrovirus encoding alkaline phosphatase confirms clonal boundary assignment in lineage analysis of murine retina. Proc. Natl. Acad. Sci. U.S.A. 89:693-697.

Naldini, L., Blomer, U., Gallay, P., Ory, D., Mulligan, R., Gage, F.H., Verma, I.M., and Trono, D. 1996. In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272:263-267.

Fraser, C.C., Szilvassy, S.J., Eaves, C.J., and Humphries, R.K. 1992. Proliferation of totipotent hematopoietic stem cells in vitro with retention of long-term competitive in vivo reconstituting ability. Proc. Natl. Acad. Sci. U.S.A. 89:19681972.

Onishi, M., Kinoshita, S., Morikawa, Y., Shibuya, A., Phillips, J., Lanier, L.L., Gorman, D.M., Nolan, G.P., Miyajima, A., and Kitamura, T. 1996. Applications of retrovirus-mediated expression cloning. Exp. Hematol. 24:324-329.

Gudkov, A.V., Zelnick, C.R., Kazarov, A.R., Thimmapaya, R., Suttle, D.P., Beck, W.T., and Roninson, I.B. 1993. Isolation of genetic suppressor elements, inducing resistance to topoisomerase II-interactive cytotoxic drugs, from human topoisomerase II cDNA. Proc. Natl. Acad. Sci. U.S.A. 90:3231-3235. Gudkov, A.V., Kazarov, A.R., Thimmapaya, R., Axenovich, S.A., Mazo, I.A., and Roninson, I.B. 1994. Cloning mammalian genes by expression selection of genetic suppressor elements: association of kinesin with drug resistance and cell immortalization. Proc. Natl. Acad. Sci. U.S.A. 91:3744-3748. Homburger, S.A. and Fekete, D.M. 1996. High efficiency gene transfer into the embryonic chick CNS using B-subgroup retroviruses. Dev. Dyn. 206:112-120. Kitamura, T., Onishi, M., Kinoshita, S., Shibuya, A., Miyajima, A., and Nolan, G.P. 1995. Efficient screening of retroviral cDNA expression libraries. Proc. Natl. Acad. Sci. U.S.A. 92:9146-9150. Kotani, H., Newton, P.B.R., Zhang, S., Chiang, Y.L., Otto, E., Weaver, L., Blaese, R.M., Anderson, W.F., and McGarrity, G.J. 1994. Improved methods of retroviral vector transduction and production for gene therapy. Hum. Gene Ther. 5:19-28. Lemischka, I.R., Raulet, D.H., and Mulligan, R.C. 1986. Developmental potential and dynamic behavior of hematopoietic stem cells. Cell 45:917927. Morgan, B.A. and Fekete, D.M. 1996. Manipulating gene expression with replication competent retroviruses. Methods Cell Biol. 51:185-218. Muneoka, K., Wanek, N., and Bryant, S.V. 1986. Mouse embryos develop normally exo utero. J. Exp. Zool. 239:289-293.

Pear, W.S., Aster, J.C., Scott, M.L., Hasserjian, R.P., Soffer, B., Sklar, J., and Baltimore, D. 1996. Exclusive development of T cell neoplasms in mice transplanted with bone marrow expressing activated Notch alleles. J. Exp. Med. 183:22832291. Rayner, J.R. and Gonda, T.J. 1994. A simple and efficient procedure for generating stable expression libraries by cDNA cloning in a retroviral vector. Mol. Cell Biol. 14:880-887. Turner, D.L., Snyder, E.Y., and Cepko, C.L. 1990. Lineage-independent determination of cell type in the embryonic mouse retina. Neuron 4:833845. van Zandt, G. 1984. Studies of hematopoietic stem cells spared by 5-fluorouracil. J. Exp. Med. 159:679-690. Whitehead, I., Kirk, H., and Kay, R. 1995. Expression cloning of oncogenes by retroviral transfer of cDNA libraries. Mol. Cell Biol. 15:704-710. Williams, D.A., Lemischka, I.R., Nathan, D.G., and Mulligan, R.C. 1984. Introduction of a new genetic material into pluripotent haematopoietic stem cells of the mouse. Nature 310:476-480. Wong, B.Y., Chen, H., Chung, S.W., and Wong, P.M. 1994. High-efficiency identification of genes by functional analysis from a retroviral cDNA expression library. J. Virol. 68:5523-5531.

Contributed by Constance Cepko Harvard Medical School Boston, Massachusetts Warren Pear University of Pennsylvania Philadelphia, Pennsylvania




INACTIVATION OF GENES IN MAMMALIAN CELLS

SECTION IV

Human Somatic Cell Gene Targeting

UNIT 9.15

Human somatic cell gene targeting provides a powerful tool to scientists studying gene function in cultured human cells. This technology allows scientists to knock out genes in human somatic cells in an fashion analogous to the creation of knockout mice. Human somatic cell gene targeting brings the power of genetics to the study of human genes in human cells by making it possible to compare cells or individuals that are genetically identical except for a single, well-defined mutation in an endogenous gene. A partial list of genes successfully targeted in human somatic cells is presented in Table 9.15.1. Once this “molecular scalpel” has been employed, a researcher has at his or her disposal a set of cultured human cells that are genetically identical except for a specific, targeted change—the presence (in the parental cells) or absence (in the knockout cells) of a functional human gene. Since the cells are otherwise genetically the same, any biological or biochemical difference between the cells sheds light on the function of the targeted gene. This type of classical genetic analysis—the study of gene function by comparing cells or organisms that are genetically identical except for a single, well-defined genetic change in an endogenous gene—has been a mainstay of genetics in model systems for decades and has yielded innumerable critical discoveries. Until relatively recently, however, this capability has been restricted to model organisms and was not applicable to human cells. The biological and biochemical characteristics of such isogenic sets of human cells can be analyzed in many different ways. For example, the growth properties of the cells can be studied in vitro to assess differences in their morphology and other growth characteristics (Shirasawa et al., 1993). The cells can be analyzed using flow cytometry to assess their cell-cycle profile during exponential growth or cell-cycle arrest, or after treatment with growth factors, small-molecule therapeutics, and lead compounds (Waldman et al., Table 9.15.1

Gene Targeting in Human Cells

Gene targeted

Reference

Smad4 p53

Zhou et al., 1998 Bunz et al., 1998

14-3-3σ DNMT1

Chan et al., 1999 Rhee et al., 2000

BAX PPARδ

Zhang et al., 2000a Park et al., 2001

Securin ORC2

Jallepalli et al., 2001 Dhar et al., 2001

Ferredoxin reductase ATR

Hwang et al., 2001 Cortez et al., 2001

DEC1 DNMT3b

Zawel et al., 2002 Rhee et al., 2002

β-Catenin

Kim et al., 2002a; Chan et al., 2002; Sekine et al., 2002

Contributed by Todd Waldman, Carolyn Lee, Tagvor G. Nishanian, and Jung-Sik Kim Current Protocols in Molecular Biology (2003) 9.15.1-9.15.20 Copyright © 2003 by John Wiley & Sons, Inc.



1995). The propensity of the cells to undergo apoptosis in vitro can be assessed using a variety of assays (Waldman et al., 1996). Microarrays can be employed to identify novel putative effectors of the targeted gene (Kim et al., 2002a). The cells can also be studied in vivo. Since the cell lines commonly used for gene targeting are derived from human cancers, they will form tumors when injected subcutaneously into immunodeficient mice. By creating and comparing so-called “isogenic tumors,” it is possible to examine the role of the targeted gene in a wide variety of processes—e.g., one can study the effects of oncogenes and tumor-suppressor genes in tumor formation and on the sensitivity of tumors to anticancer agents (Shirasawa et al., 1993; Waldman et al., 1997). This unit presents protocols for human somatic cell gene targeting. The Strategic Planning section presents a number of considerations that must be made when planning such an experiment. Basic Protocol 1 provides details for vector construction. Basic Protocol 2 describes the transfection procedures. Basic Protocol 3 is a method for excising the antibiotic-resistance gene from a heterozygous knockout to make it possible to target the remaining allele; the Alternate Protocol describes how to switch selectable markers for the same purpose. Basic Protocol 4 describes how to target the remaining allele, converting a heterozygous knockout cell line to a homozygous knockout cell line. STRATEGIC PLANNING Selection of a Parental Cell Line To date, six different human cell lines have been successfully used for human somatic cell gene targeting (listed in Table 9.15.2). The list is short because suitable cell lines must possess each of three distinct characteristics. The first of these, which is very important, is that suitable cell lines should be diploid (or near-diploid), such that there are two (and only two) copies of the gene to be targeted. Second, the cells should transfect with high efficiency, since the promoterless enrichment built into human gene targeting vectors leads to the formation of relatively few colonies (since randomly integrated promoterless targeting vectors generally fail to express neor and therefore do not form drug-resistant colonies). Third, the cells should grow quickly and clonally, since one will need to grow individual clones in 96-well plates, preferably in 2 to 3 weeks. Of the cell lines listed in Table 9.15.1, HCT116 cells have proven to be the most robust and therefore have been used in the majority of human somatic cell gene targeting projects. As such, the authors of this unit tend to focus initial targeting efforts on HCT116 cells, since it is preferable to test new targeting vectors in a cell line that has been well established as suitable for human somatic cell gene targeting. Once the efficacy of targeting in HCT116 cells has been demonstrated, only then is the attempt made to target the gene in other cell lines as well.

Table 9.15.2


Human Cell Lines for Gene Targeting

Cell line

Reference

HCT116 DLD1

Shirasawa et al., 1993 Shirasawa et al., 1993

HT1080 SW48

Porter and Itzhaki, 1993 Kim et al., 2002b

HaCaT Human fibroblasts

Zawel et al., 2002 Brown et al., 1997



In addition to meeting these three requirements, it is important for both experimental and technical reasons that the chosen cell line express the gene to be targeted. Experimentally, there is not much point in deleting a gene in somatic cells that is not expressed. Technically, homologous integration of a promoterless targeting vector into a transcriptionally silent locus will lead to the absence of neor expression and the failure to form a drug-resistant colony. Thus, the authors recommend first confirming that the gene of interest is expressed in candidate cell lines prior to initiating a targeting project (via immunoblotting, northern blotting, or RT-PCR). Architecture of a Promoterless Targeting Vector Promoterless targeting vectors require that the selectable marker gene be inserted in-frame into the gene being targeted. Homologous integration of such a targeting vector into the human genome then leads to the expression of a functionally active drug-resistance protein. There are several general factors to consider when planning to construct such a targeting vector. One early concern was that targeting vectors for human somatic cell gene targeting would need to be composed of isogenic DNA—i.e., DNA derived from the actual cell line in which the gene would be targeted. These concerns were based on early gene-targeting studies performed in mouse embryonic stem cells, which indicated that the polymorphisms present in genomic DNAs derived from different strains of mice could affect the targeting frequency in a dramatically adverse fashion (Deng and Capecchi, 1992; van Deursen and Wieringa, 1992). If this were true for gene targeting in human cells, it would be difficult to target a gene in multiple cell lines using the same targeting vector. Furthermore, since humans (unlike mice) are not inbred, the targeting vector for each allele of a gene would need to be composed of different genomic DNA. As such, the need for isogenic DNA would substantially increase the technical complexity of human somatic cell gene targeting. Fortunately, experience has shown that targeting vectors composed of nonisogenic genomic DNA are able to create knockouts in a wide variety of genetically unrelated human cell lines (Sedivy et al., 1999). In the authors’ experience, the neor protein is frequently unable to tolerate the fusion of unrelated amino acids onto its amino terminus (C. Lee., J.S. Kim., and T. Waldman, unpub. observ.). This being the case, it is inadvisable to design a targeting strategy which leads to the formation of a neor fusion protein. There are two effective strategies that circumvent this limitation. First, one can design the targeting vector such that the initiating methionine of the gene being targeted is replaced precisely with the initiating methionine of neor gene. Second, one can replace internal exon(s) with an IRES-neor gene. Finally, it is not possible to design a true promoterless targeting vector for deleting the first exon of a gene, since the left arm of such a targeting vector would contain promoter elements of the gene being targeted. If the initiating methionine of a gene is, in fact, in the first exon, the authors recommend designing the targeting vector to delete internal exon(s) using an IRES-neor strategy. Building a Targeting Vector Building promoterless targeting vectors is a nontrivial exercise in recombinant DNA technology. There are two factors that complicate construction of these vectors. First, there is virtually no sequence flexibility in the junction between the left arm of the targeting vector and the selectable marker cassette, making the use of standard restriction enzyme–based recombinant DNA strategies problematic. Second, it is frequently necessary to add several sequence features between the end of the selectable marker gene and



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the right homology arm of the targeting vector, such as restriction sites and a PCR priming site. The authors have found that the most straightforward way to circumvent these difficulties is to build the targeting vectors in two steps, exploiting the high efficiency of homologous recombination in S. cerevisiae (Storck et al., 1996; Oldenburg et al., 1997). First, a 5- to 10-kb fragment containing the genomic region of interest is cloned (generally from a bacterial artificial chromosome, or BAC; UNIT 5.9) into a yeast shuttle vector, which contains selectable markers and origins of replication for propagation in both E. coli and S. cerevisiae. Next, the selectable marker gene (generally either neor or IRES-neor) is PCR-amplified from a plasmid using long primers that add both the needed sequence features, along with 30 to 50 nucleotides of homology to the cloned genomic region for directing the desired recombination event. This PCR product is then cotransformed into S. cerevisiae together with linearized recombinant shuttle vector, and recombinants are identified and used as the final targeting vector. This strategy makes it possible to design any junction and to add desired sequence features all in one step, and does not rely on the serendipitous presence of needed restriction sites. It should be noted that it is also theoretically possible to exploit homologous recombination in E. coli to accomplish the same goals (Zhang et al., 2000b). However, unlike in yeast, in E. coli it is generally necessary to select for homologous recombination between the vector and the PCR product. Therefore, the PCR-amplified neor cassette would need to contain a selectable marker for growth in bacteria as well. Screening for Knockouts in Human Cells The simplest approach for identifying gene-targeted cell lines is to conduct an initial PCR-based screen, then confirm the putative knockouts with a Southern blot. There are two general factors to consider when planning the details of such a screening strategy. Historically, PCR screening for knockouts has relied on one primer in the selectable marker gene (generally neor), and another primer located in the genomic region just outside one of the homology arms of the targeting vector. Homologous integration of the targeting vector creates a template for PCR, whereas random integration does not. As such, diagnostic PCR products are produced only from gene-targeted clones. The fundamental difficulty with this classic PCR-based screening strategy is the lack of a positive control for PCR—i.e.,by definition there exists no template for PCR unless a knockout is created. As such, it is virtually impossible to optimize the PCR reaction prior to screening for knockouts, and therefore difficult to definitively interpret a negative result. If none of the clones are PCR-positive, the question remains—were there truly no knockouts, or did the PCR not work?


Recently, a strategy has been devised to overcome this limitation, providing an internal positive control for PCR in each reaction (Chan et al., 1999; for a schematic of this approach, see Fig. 9.15.1). This approach is very similar to the classic approach, except that the PCR primer generally located in the neor gene is substituted by a PCR primer located in the genomic region being deleted by the targeting vector. Importantly, this PCR priming site is built into the targeting vector such that homologous integration of the targeting vector simultaneously deletes the endogenous priming site, and reintroduces it in a slightly different location. As such, clones in which the targeting vector has integrated randomly create a PCR product of a certain size, and clones in which the PCR product has integrated via homologous recombination create a PCR product of a different size. Therefore, it is no longer the presence or absence of a PCR product that determines a knockout; instead, a PCR product is produced in all the PCR reactions and its size



priming site A

A neomycin

targeting vector

priming site A genomic DNA deleted region priming site B

B

+/+

+/–

+/–

+/–

+/– wild-type allele knockout allele

Figure 9.15.1 PCR screening for knockouts.

determines whether a knockout is present. This elegant modification of the classic PCR screening approach has significant implications for interpreting PCR screening data. The other practical implication is that the diagnostic PCR must be optimized prior to completion of the targeting vector, since the PCR priming site must be built into the targeting vector itself. In addition to altering the size of a PCR product, homologous integration of the targeting vector invariably modifies the restriction map of the local genomic region. By using a Southern blot–based approach, it is possible to discriminate between wild-type alleles and targeted alleles. The details of choosing restriction sites and probes are described in Basic Protocol 1. Targeting the Second Allele One particularly challenging aspect of human somatic cell gene targeting is the creation of homozygous knockouts by disruption of the second, remaining allele. Since heterozygous knockout cells are already drug-resistant, it is necessary to modify the targeting vector to change the selectable marker gene. While swapping selectable markers is not technically demanding (especially if homologous recombination in S. cerevisiae was employed to create the targeting vector), experience has demonstrated that selectable markers other than neor (e.g., hygr and puror, among others) are suboptimal for somatic cell gene targeting. In particular, targeting vectors relying on these selectable markers are frequently either unable to generate drug-resistant colonies at all or require such low concentrations of drug that it is difficult to kill the background cells (Hanson and Sedivy, 1995; J.S. Kim, C. Lee, and T. Waldman, unpub. observ.). While the reasons for these difficulties are unknown, some have speculated that endogenous promoters are unable to generate a high enough level of gene expression to confer resistance to these specific drugs. The most straightforward way to circumvent difficulties with selectable markers other than neor is to build a Cre-loxP recyclable neor targeting vector and use it to target each allele sequentially. In such a targeting vector, the neor gene is flanked by loxP sites. After



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creation of heterozygous knockout cells, Cre is added to the cells (generally via infection with a recombinant Cre-containing adenovirus), and drug-sensitive derivatives are identified. These cells are then retransfected with the same targeting vector for deletion of the remaining allele. The second inherent difficulty in targeting the second allele is that the frequency of homologous integration is exactly half that of the first allele. This is because half the second allele homologous integration events will disrupt the already targeted allele. BASIC PROTOCOL 1

BUILDING A TARGETING VECTOR The first step in the creation of a human promoterless targeting vector is to obtain human genomic DNA that will form the basis of the homology arms of the targeting vector. To do this, the authors generally identify BAC clone(s) that contain the needed genomic region (UNIT 5.9). Once BACs have been identified and prepped, the next step is to subclone a smaller fragment (generally 5 to 10 kb) into a yeast shuttle vector. This subclone will be used as raw material for final assembly of the targeting vector via homologous recombination in S. cerevisiae (Storck et al., 1996; Oldenburg et al., 1997). Alternatively, it is possible to consecutively subclone each homology arm into a vector. However, subcloning a single larger piece that will ultimately form both homology arms saves a subcloning step. The final step in construction of the targeting vector is to use homologous recombination in S. cerevisiae to replace critical exon(s) in the cloned genomic region with an in-frame neor or IRES-neor gene. The decision whether to use neor or IRES-neor is made according to the guidelines in step 6, below, and Critical Parameters and Troubleshooting. There are two feasible alternative approaches. First, it is possible to add the promoterless neor gene via conventional restriction enzyme–based cloning. However, the authors find that such an approach often requires complex multistep strategies because of the lack of convenient restriction sites. Second, it is possible to conduct similar homologous recombination reactions in strains of E. coli that overexpress recombinase proteins (Zhang et al., 2000b). In this case the disadvantage is that such an approach generally requires that the insert include an E. coli selectable marker gene. Before beginning the homologous recombination, it is necessary to identify a unique restriction site in the cloned genomic region for linearization of the recombinant shuttle vector prior to transformation of S. cerevisiae. The recombination junctions will need to flank this restriction site. Once such a restriction site is identified, the neor gene (or IRES-neor gene) is PCR-amplified with long primers designed to add needed sequence features to the ends (e.g., a priming site for PCR-based diagnosis of knockouts, restriction site or sites for Southern blot-based diagnosis of knockouts, and sequence homologies for recombination in S. cerevisiae). This PCR product is then cotransformed into S. cerevisiae with the linearized recombinant yeast shuttle vector. After plating the transformation mix on the appropriate selective medium, individual yeast colonies are screened via whole-cell PCR to identify those harboring the desired recombinant plasmid. PCR products derived from recombinant plasmids are then sequenced to exclude plasmids harboring mutations in the recombination junctions. These plasmids are then transferred into E. coli and prepped. Finally, the integrity of the finished targeting vector is confirmed by restriction analysis and sequencing of critical junctions.




Materials Genomic DNA (UNIT 2.2) derived from the parental cell line (e.g., HCT116; ATCC #CCL 247) BAC DNA purification kit: e.g., Nucleobond Plasmid Maxi kit (Clontech) or similar kit from Qiagen (also see UNIT 2.1B) Yeast shuttle vectors (e.g., YEp24, pRS423, pRS424, pRS425, or pRS426, available from ATCC) E. coli cells (DH10B) pMC1neopolyA (Stratagene) or an IRES-neor plasmid (available from John Sedivy at Brown University upon request) EXPAND High Fidelity PCR System (Roche) QIAX Gel Extraction Kit (Qiagen) or equivalent Appropriate selective yeast media (Qbiogene) Yeast PCR kit (Qbiogene) Yeast RPM kit (Qbiogene) Facility for BAC library screening (e.g., Research Genetics) 30°C incubator Additional reagents and equipment for primer design (UNIT 7.7), synthesis of oligonucleotides (UNIT 2.11), PCR (UNIT 15.1), gel purification of DNA fragments (UNIT 2.7), preparation and screening of a BAC library (http://www.chori.org/ bacpac/home.htm), plating bacteria (UNIT 1.3), single-cell PCR (UNIT 25A.3), restriction digestion (UNIT 3.1), agarose gel electrophoresis (UNIT 2.5A), restriction mapping (UNITS 3.2, 3.3, & 7.7), finding sequence information on the Web (UNITS 19.2 & 19.3), dephosphorylation of DNA by calf intestinal phosphatase (UNIT 3.10), DNA subcloning and ligation (UNIT 3.16), introduction of plasmid DNA into E. coli (UNIT 1.8), colony hybridizations (UNIT 6.3), preparation of plasmid DNA (UNIT 1.7), DNA sequencing (Chapter 7), phenol extraction and ethanol precipitation of DNA (UNIT 2.1A), gel purification of DNA fragments (UNITS 2.5A, 2.7, or 2.8), introduction of DNA into S. cerevisiae (UNIT 13.7), growth of S. cerevisiae (UNIT 13.2), and DNA preparation from S. cerevisiae (UNIT 13.11) Identify and isolate BAC clones 1. Design (UNIT 7.7) and synthesize (as described in UNIT 2.11; or order from synthesis facility) a PCR primer pair for amplification of the genomic region of interest from human genomic DNA derived from the parental cell line. Optimize and perform PCR amplification (UNIT 15.1; the PCR product should be 500 to 1000 nucleotides in length) to produce a probe for BAC library screening (step 2). Gel purify the DNA fragment obtained from PCR (UNIT 2.7). The primer pair will be used both to create a probe for library screening and to confirm that the BACs identified harbor the correct insert. Ideally, the probe will be derived from intronic sequence, since probes derived entirely from exons can hybridize to pseudogenes.

2. Perform a hybridization-based BAC library screening using the probe for the region of interest created in step 1. For protocol details, see the laboratory Web page of Pieter De Jong at the Children’s Hospital Oakland Research Institute (http://www.chori.org/bacpac/home.htm). Since the physical map and sequence of the human genome is by now largely completed, it is often possible to search GenBank (UNIT 19.2) and identify the needed BAC, then order it from a BAC repository. However, if such information is unavailable, or if the BAC or BACs ordered from the repository do not contain the correct insert, it is necessary to screen a library to identify the needed BACs. In fact, the authors often screen a library even if a single correct BAC is available, since it is generally advantageous to have several different BAC clones with the needed insert, each with different boundaries and junctions.



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Although the materials are available to screen BAC libraries in academic laboratories), the authors generally contract out the screening to Research Genetics, which performs this procedure routinely. Screening BAC libraries involves hybridization of radiolabeled probes to very large commercially available membranes that contain hundreds of thousands of gridded BAC clones. Performing the actual hybridizations is tricky because of the size of the membranes. Furthermore, identification of the correct clones on the huge filters takes practice. If sending the probe to a commercial facility (e.g., Research Genetics), it generally takes 2 to 6 weeks to receive the final product, which will be stabs inoculated with bacteria harboring the desired BACs. Alternatively, it is possible to skip this BAC step by generating the needed genomic DNA via PCR amplification of the needed homology arms from a genomic DNA template. However, the authors prefer to use BACs as the source of genomic DNA for building targeting vectors, since it can be difficult to create large homology arms using PCR. Also, PCR can introduce mutations into the homology arms and therefore reduce the targeting efficiency of the completed targeting vector.

3. Once the BACs are received, confirm that they contain the correct insert as follows. a. Streak out bacteria from the stabs onto the appropriate selective medium to generate single colonies (UNIT 1.3). b. Grid individual colonies (generally ten or so) to a grid plate (UNIT 1.3). c. Using the PCR primer pair optimized in step 1 above, test each of the grid positions with whole-cell PCR (UNIT 25A.3) to identify bacteria harboring the desired insert. Confirming the BAC clone is important since the bacterial stabs are created by directly inoculating bacteria stored as glycerol stocks in 384-well plates. It is possible (and in the authors’ experience not uncommon) for the bacteria in a particular well to be a mixed population caused by spillage from adjacent wells, or the wrong clone entirely.

4. Prepare BAC DNA from positive clones using the Nucleobond Plasmid Maxi or similar DNA purification kit. The authors generally use Nucleobond columns from Clontech, though kits available from other manufacturers (e.g., Qiagen; UNIT 2.1B) will undoubtedly also work well.

5. Digest the various BACs with several different restriction enzymes (UNIT 3.1) and compare the restriction digestion patterns by agarose gel electrophoresis (UNIT 2.5A). This step confirms the large size of the purified DNAs. Furthermore, it is generally expected that different BACs share some bands but differ in others, reflecting the fact that they are derived from the same genomic region but have different boundaries.


Subclone genomic DNA from the BAC into the yeast shuttle vector To choose a restriction fragment to subclone, one must first decide what exon(s) the completed vector will be designed to target. There are several factors to consider when choosing an exon to target. If possible, target the exon containing the initiating methionine, since it is then possible to build a promoterless targeting vector in which the initiating methionine of neor precisely replaces the initiating methionine of the targeted gene. However, avoid targeting the first exon, since the left arm of such a targeting vector would contain promoter elements of the gene itself, reducing the efficiency of the promoterless selection. If the initiating methionine is in the first exon, choose an internal exon and use an IRES-neor based strategy (Sedivy and Dutriaux, 1999). There are two main issues to consider when targeting an internal exon with an IRES-neor. First, it is desirable to target an early exon and therefore create a protein that is as dramatically altered as possible. Second, it is desirable to target an exon which, if it were skipped by the splicing machinery, would change the reading frame and result in a premature stop codon.



6. Using published or Web-based human genomic sequence information (UNITS 19.2 & 19.3), make a sequence-based restriction map of the desired genomic region (see UNITS 3.2, 3.3 & 7.7). Identify several suitable restriction fragments for subcloning into the yeast shuttle vector. When choosing potential restriction fragments, keep in mind that one arm of the final targeting vector will need to be short enough to PCR amplify across (i.e., less than ∼2 kb) when identifying gene-targeted cell lines.

7. Using the RepeatMasker Web site (http://repeatmasker.genome.washington.edu), analyze the sequence within and surrounding these restriction fragments to identify repeat elements such as Alu and LINE-1. It is important to avoid designing targeting vectors composed largely of repeat elements, since the arms of such targeting vectors will not provide the unique homologies required to guide homologous recombination in human cells.

8. To perform the actual subcloning, first digest both the BAC DNA and the yeast shuttle vector with the appropriate restriction enzyme or enzymes (see UNIT 3.1 and documentation for shuttle vector). There are numerous yeast shuttle vectors available for use in this subcloning step. The authors generally use one of five different yeast shuttle vectors available from ATCC: YEp24, pRS423, pRS424, pRS425, and pRS426 (Botstein et al., 1979; Christianson et al., 1992). YEp24 is a pBR322-based, low-copy-number vector, whereas the pRS series are pBluescript-based, high-copy-number vectors. All work well for creating targeting vectors via homologous recombination; however, in the authors’ experience it is slightly easier to clone larger fragments (>5 kb) into low-copy-number vectors such as YEp24.

9. Treat the linearized vector with calf intestinal alkaline phosphatase (CIP; UNIT 3.10) to limit recircularization of the vector during ligation. 10. Ligate the linearized, CIP-treated vector to the digested BAC DNA fragments (UNIT 3.16). Transform or electroporate appropriate E. coli cells with the ligated plasmid vector (UNIT 1.8). After plating on the appropriate selective medium, grid ∼500 colonies (UNIT 1.3). Also grid bacteria harboring various BACs to serve as positive and negative controls for the next step. 11. Identify bacteria harboring the desired recombinant plasmid by performing colony hybridizations with a radiolabeled oligonucleotide probe (UNIT 6.3). It is often possible to use one of the PCR primers from in step 1, above, as the probe. Note that the positive signals produced by bacteria harboring high-copy-number recombinant plasmids (e.g., pRS423-6) will be much stronger than the signals produced by bacteria harboring low-copy-number recombinant plasmids (e.g., BACs, YEp24).

12. Streak out the bacteria (UNIT 1.3) in positive grid positions to purify individual colonies. Prepare plasmid DNA using the Nucleobond Plasmid kit or equivalent (also see UNIT 1.7). 13. Confirm the integrity of the subclones by restriction analysis and sequencing of junctions (Chapter 7). It is important to identify and avoid subclones with multiple, concatamerized inserts.

Identify priming sites and restriction sites for PCR and Southern blot–based identification of gene-targeted cell lines It is important to identify these sequence features before completing the targeting vector, since the sequence comprising a PCR primer and possibly a restriction site will be built into the targeting vector during the final step of construction.



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14. To identify suitable PCR primers, design (UNIT 7.7) and synthesize (as described in UNIT 2.11; or order from synthesis facility) three sense and three antisense primers that meet the following criteria: a. By convention, one primer is located in the genomic DNA just adjacent to the shorter homology arm of the targeting vector. b. The other primer is not located in the neor gene (or IRES-neor gene) itself, but is instead located in the genomic region deleted by the targeting vector. Importantly, this priming site will be built into the targeting vector itself at the junction of the short homology arm and the neor gene. Homologous integration of the targeting vector deletes the endogenous site and moves it, altering the size of the PCR product (generally shortening it).

15. Test all nine permutations of these primers on genomic DNA derived from the parental cell line (e.g., HCT116) at several different annealing temperatures, as described in UNIT 15.1. Choose the most robust primer pair for eventual screening of putative gene-targeted clones. As described above (see Strategic Planning), the authors use a strategy for PCR-based diagnosis of knockouts that is slightly different from the conventional strategy used to identify knockouts in mouse embryonic stem cells. This strategy is preferable to the conventional PCR-based screen, since it includes a positive internal control with each PCR reaction.

16. To identify candidate restriction sites for Southern blot–based diagnosis of knockouts, make a computer-generated restriction map of the genomic region using published or Web-based genomic sequences (UNIT 7.7). Identify suitable sites based on the following criteria: a. The site should be present at a defined location in the genomic DNA at least 500 nucleotides outside one homology arm of the completed targeting vector (to allow for placement of a Southern blot probe; also see UNIT 10.8). b. The site should be absent in that homology arm itself. c. The site should be present at a defined location either in the other homology arm or in the genomic DNA adjacent to the other homology arm. d. Furthermore, the site should either be present in the neor gene itself, or should be added to the targeting vector (generally at a junction of a homology arm and neor) during the final step in vector construction. By following these guidelines, homologous integration of the targeting vector will alter the genomic restriction map in a way that should be easily measurable by Southern blot analysis (UNIT10.8).

Replace critical exon(s) with a promoterless neor or IRES-neor gene via homologous recombination in S. cerevisiae 17. Using sequence analysis software (UNIT 7.7), identify candidate restriction sites for linearization of the recombinant yeast shuttle vector created in steps 8 to 13. Confirm location and uniqueness of candidate site(s) via routine restriction analysis (UNIT 3.1). 18. Once an appropriate site has been chosen, perform a larger-scale preparative digest using approximately 2.5 µg of the recombinant shuttle vector plasmid DNA from step 12. Phenol extract and ethanol precipitate the linearized plasmid (UNIT 2.1A).


19. Design (UNIT 7.7) and synthesize (as described in UNIT 2.11; or order from synthesis facility) PCR primers for amplification of the neor or IRES-neor gene with appropriate ends to add the PCR priming sites and restriction sites and to add 40 to 50 nucleotides of sequence homology to direct homologous recombination in S. cerevisiae.



The authors order primers PAGE-purified from Integrated DNA Technologies. If using the neor gene as the template, the authors generally use the pMC1neoPolyA plasmid as the template and use the following PCR priming sites: sense 5′-ATGGGATCGGCCATTGAACAA-3′, antisense 5′-GTCGACGGATCCGAACAAACG-3′. Of note, the first three nucleotides of the sense primer comprise the initiating methionine of neor. If using IRES- neor (available from John Sedivy at Brown University) as the template, the authors use the following PCR priming sites: sense 5′-AACGTTACTGGCCGAAGCCGC-3′, antisense 5′-TCCCAACTCATCCCGGCCTC-3′.

20. Perform high-fidelity PCR with the pMC1neopolyA plasmid or IRES-neor as template, to create the neor PCR product, using the EXPAND High Fidelity PCR System or equivalent (also see UNIT 15.1). 21. Gel purify the PCR product using the QIAX Gel Extraction Kit or equivalent (also UNITS 2.5A, 2.7, or 2.8). 22. Cotransform (UNIT 13.7) 150 ng of linearized recombinant shuttle vector (from step 18) together with an estimated 500 ng of gel-purified PCR product (from step 21) into S. cerevisiae. Include negative controls consisting of vector alone and PCR product alone. The authors generally use either commercially available competent yeast (strain MaV203; Invitrogen) or prepare competent yeast from strain INVSc1 (Invitrogen) using the FrozenEZ Yeast Transformation II kit (Zymo Research). The choice of yeast strain depends on which auxotrophic marker is needed. Specific protocols for transformation are available in each kit. Also see UNIT 13.7 for additional information relating to yeast transformation.

23. Plate the transformation mixes on the appropriate selective medium (available from Qbiogene), and incubate at 30°C for 2 to 4 days. General considerations for the growth of S. cerevisiae are presented in UNIT 13.2. Expect to see ∼5-fold more colonies in the experimental reactions than in vector alone. There should be no colonies in insert alone.

24. Grid approximately 20 colonies from the vector + insert transformation mix (see UNIT 13.2). Also grid some vector-only colonies to serve as negative controls for the subsequent PCR reaction. 25. Design and synthesize (as described in UNIT 2.11; or order from synthesis facility) a set of PCR primers that flank the recombination event. Such a primer pair will create PCR products of different sizes in recombined and nonrecombined plasmids.

26. Test yeast from individual grid positions by PCR to identify those in which a recombination event has occurred. The authors generally use the yeast itself as a PCR template after pretreatment with zymolyase to destroy the cell wall. The authors use the Yeast PCR Kit (Qbiogene) for this purpose.

27. Once yeast-harboring recombinant plasmids have been identified, sequence the ends of the PCR products to demonstrate that the recombination junctions are intact and free of mutations. Design (UNIT 7.7) and synthesize (as described in UNIT 2.11; or order from synthesis facility) sequencing primers for the purpose of sequencing the junctions from “inside out,” making it possible to generate sequence from the very end of the PCR products. Sequence the PCR products (see Chapter 7). Introduction of DNA into Mammalian Cells


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Transfer completed targeting vectors into E. coli for large-scale DNA preparation 28. Prepare a miniprep of the recombinant yeast DNA using the Yeast RPM kit. 29. Electroporate the miniprep DNA into E. coli as described in UNIT 1.8. Plate on the appropriate selective medium (UNIT 1.3). Streak out individual colonies to purify. 30. Perform large-scale plasmid preps as described in UNIT 1.7. The authors generally use Nucleobond kits (Clontech).

31. Confirm the integrity of the resultant plasmid via restriction analysis and resequencing of the critical junctions (see Chapter 7) using the sequencing primers used in step 27 above. 32. Linearize the vector for transfection (Support Protocol). SUPPORT PROTOCOL

LINEARIZATION OF THE COMPLETED TARGETING VECTOR PRIOR TO TRANSFECTION INTO HUMAN CELLS The completed targeting vector must be linearized prior to transfection, since homologous recombination is more efficient with linear DNA. As such, it is necessary to identify a suitable restriction enzyme that cuts only in the vector backbone. There are generally sites for rare-cutting enzymes (e.g., SalI or NotI) in the polylinker of the yeast shuttle vector that are useful for this purpose Materials Completed targeting vectors (Basic Protocol 1) TE buffer (APPENDIX 2) Additional reagents and equipment for restriction digestion (UNIT 3.1), agarose gel electrophoresis (UNIT 2.5A), and phenol extraction and ethanol precipitation of DNA (UNIT 2.1A) 1. Identify an appropriate restriction enzyme (UNIT 3.1) for linearization of the completed targeting vector in the vector backbone. Confirm that the enzyme cuts the targeting vector singly and in the predicted location by performing small-scale restriction digests and analyzing an aliquot by agarose gel electrophoresis (UNIT 2.5A). 2. Perform a preparative digest of 50 to 100 µg of the targeting vector with the chosen enzyme (UNIT 3.1). Phenol extract and ethanol precipitate the digest (UNIT 2.1A), then resuspend in TE buffer to a final concentration of 0.5 µg/µl. 3. Confirm the efficiency of digestion by analyzing an aliquot of the digest by agarose gel electrophoresis (UNIT 2.5A). The authors generally prepare linearized DNA from three identical targeting vectors derived from different yeast colonies, in the unlikely event that one or several of the vectors has a PCR-generated inactivating mutation in the neor (or IRES-neor) gene.

BASIC PROTOCOL 2


TRANSFECTION OF THE LINEARIZED TARGETING VECTOR INTO CULTURED HUMAN CELLS AND IDENTIFICATION OF HETEROZYGOUS KNOCKOUT CELLS The authors generally use HCT116 cells for the initial gene-targeting experiments, since these cells divide rapidly, grow well as individual clones in 96-well plates, and are generally well established as suitable for human somatic cell gene targeting. Once the targeting vector has been shown to work efficiently in HCT116 cells, attempts can be made to target the gene in other cell lines as well.



Two sequential transfections are performed in this step. Initially, the cells are transfected, and colonies are selected directly in 25-cm2 tissue culture flasks and stained with crystal violet. This preliminary experiment demonstrates whether the new targeting vector(s) can create G418-resistant colonies, and identifies any targeting vectors that may harbor inactivating mutations in their neor (or IRES-neor) gene. Also, based on the number of colonies produced, it is possible to estimate the dilutions needed for creation of single colonies via limiting dilution in 96-well plates. Next, the HCT116 cells are retransfected and set up in a large number (50 to 100) of 96-well plates for obtaining individual drug-resistant clones by limiting dilution. Once single colonies have formed (first identifiable ∼10 days post-transfection), they are expanded (at ~20 days post-transfection) for cryopreservation and preparation of genomic DNA. The authors generally work up 200 to 500 colonies in this step. Finally, the genomic DNAs are tested by PCR to identify heterozygous knockout cell lines. Knockouts are then thawed, and are grown up to create more frozen vials and to reprepare genomic DNA. The genotype of the cell lines is then reconfirmed by PCR and Southern blotting. Materials Complete McCoy’s 5A medium (see recipe) with and without 0.6 µg/ml G418 (geneticin, Life Technologies; see UNIT 9.5) HCT116 cells (ATCC #CCL 247) Lipofectamine reagent (Invitrogen; see UNIT 9.4) Linearized targeting vector (Support Protocol) Plasmid unrelated to targeting vector and lacking neor gene (negative control for colony formation) PMC1neopolyA plasmid (Stratagene) CMV β-gal plasmid (Clontech) Hanks’ Balanced Salt Solution (HBSS, Life Technologies; also see APPENDIX 2) 0.05% (w/v) trypsin/0.53 mM tetrasodium EDTA (Life Technologies) 25-cm2 and 75-cm2 tissue culture flasks 8-channel pipettor (e.g., Easy Step with 8-port manifold from Continental Laboratory Products) with 50-ml reservoir (Brinkmann 22-49-614-0) 96-well tissue culture plates 24-well tissue culture plates 15-ml conical centrifuge tubes Additional reagents and equipment for culture of mammalian cells (APPENDIX 3F), transfection of mammalian cells (UNIT 9.4), Xgal staining of cultured cells (UNIT 9.10), crystal violet staining of cultured cells (as for plaque assay; see UNIT 16.16, Support Protocol), preparation of genomic DNA (UNIT 2.2), PCR (UNIT 15.1), and Southern blotting (UNIT 2.9A) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Determine if target vector is capable of creating neor clones 1. Grow HCT116 cells (see APPENDIX 3F for culture techniques) to 60% to 70% confluence in complete McCoy’s 5A medium (without selective agent). Transfect HCT116 cells with linearized targeting vectors using Lipofectamine reagent as described in



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and in the manufacturer’s instructions, using 3 µg DNA per 25-cm2 tissue culture flask of 60% to 70% confluent cells. Also transfect:

UNIT 9.4

a. A plasmid harboring a neor gene driven by a heterologous promoter (e.g., pMC1neopolyA) as a positive control for colony formation. b. An unrelated plasmid lacking a neor gene as a negative control for colony formation. c. CMV β-Gal plasmid as a control for transfection efficiency. Do not add selective medium to the cells yet. The authors recommend transfecting ∼10 to 20 25-cm2 flasks in this step.

2. The day after transfection, stain the CMV β-Gal transfected cells with Xgal (UNIT 9.10; expect to see 20% to 40% blue cells). Change the medium on the remaining flasks to complete McCoy’s 5A medium containing 0.6 µg/ml G418. The authors generally use the β-Gal Staining Kit (Roche).

3. Replace the medium with fresh complete McCoy’s 5A medium containing 0.6 µg/ml G418 every 3 days (colonies should begin to appear at approximately day 10). Stain the colonies with crystal violet (see UNIT 16.16, Support Protocol) on approximately day 15. In the authors’ experience, high-efficiency promoterless targeting vectors generally create between 30 and 200 colonies per transfected 25-cm2 flask of HCT116 cells, whereas vectors harboring neor genes driven by heterologous promoters generally produce thousands of colonies per transfected 25-cm2 flask.

Perform selection on neor clones 4. Once the pilot experiment described in steps 1 to 3 above demonstrates that targeting vector is capable of creating neor clones, repeat to generate hundreds of individual neor clones. To do this, repeat the transfection strategy described above, including the positive and negative controls. At 18 to 24 hr after transfection, stain the CMV β-Gal–transfected control cells and feed the other control cells with complete McCoy’s 5A medium containing 0.6 µg/ml G418. The remainder of the flasks will be trypsinized and transferred to 96-well plates, as described is step 5.

5. Wash the cell monolayers in the 25-cm2 flasks with 5 ml HBSS, then add 1 ml of 0.05% trypsin/0.53 mM EDTA and swirl gently to detach the cells. In a sterile, disposable beaker, mix the trypsinized cells with complete McCoy’s 5A medium containing 0.6 µg/ml G418. The volumes to be used depend on the dilution and the number of 96-well plates to be created from the cell suspension (25 ml of medium per 96-well plate). Initially, try three different “dilutions” of cells to increase the likelihood of obtaining wells with single colonies: 1, 1/5, and 1/20th of a 25-cm2 flask in a 96-well plate.

6. Place an aliquot of the cell suspension in each well of the appropriate number of 96-well plates using an 8-channel pipettor with a sterile 50-ml reservoir. Stack plates, cover with plastic wrap to prevent evaporation, and begin incubating. 7. Identify wells containing single colonies as early as possible (after 10 days to 2 weeks). Human Somatic Cell Gene Targeting

As colonies become larger, cells tend to detach and form “satellite” colonies. Also, over time, two initially distinct colonies may merge together and appear to be a single colony.



8. When colonies cover at least 50% of the surface area of the well, transfer to 24-well plates. To do this, wash the well once with 250 µl HBSS, then add 100 µl of 0.05% trypsin/0.53 mM EDTA. Transfer each colony to a separate well of a 24-well plate containing 1 ml of complete McCoy’s 5A medium containing 0.6 µg/ml G418. 9. As the wells become confluent, expand the cultures to 25-cm2 flasks. To do this, wash the well once with 1 ml HBSS, then add 250 µl 0.05% trypsin/0.53 mM EDTA. Transfer the trypsinized cells to a 25-cm2 flask containing 5 ml of complete McCoy’s 5A medium containing 0.6 µg/ml G418. Harvest cells and identify knockouts 10. As the 25-cm2 flasks become confluent, harvest the cells: use 1/3 to prepare genomic DNA; cryopreserve the remaining 2/3. To harvest: a. Wash the monolayer once with 2 ml HBSS, then add 1 ml of 0.05% trypsin/0.53 mM EDTA. b. Transfer 1/3 and 2/3 of the cells to separate 15-ml conical tubes, each containing 5 ml of of complete McCoy’s 5A medium (without selective agent). c. Centrifuge 7 min at 250 × g, room temperature, to create a cell pellet. 11. Prepare genomic DNA from the tube containing 1/3 of the cells (UNIT cryopreserve the cells in the tube containing 2/3 of the cells (APPENDIX 3F).

2.2)

and

12. Test genomic DNAs by PCR as described in Basic Protocol 1 and UNIT 15.1 to identify knockouts. 13. Thaw PCR-positive clones into a 75-cm2 tissue culture flask containing 20 ml of complete McCoy’s 5A medium with 0.6 µg/ml G418. Expand into a 225-cm2 flask for cryopreservation of nine cryovials (APPENDIX 3F) and repreparation of genomic DNA (UNIT 2.2). 14. Retest by PCR (UNIT 15.1) and confirm via Southern blotting (UNIT 2.9A). RESTORATION OF G418 SENSITIVITY VIA Cre-MEDIATED EXCISION OF THE neor GENE

BASIC PROTOCOL 3

In order to recycle the targeting vector to disrupt the remaining allele, it is necessary to excise the neor gene (or IRES- neor gene) from heterozygous knockouts via expression of Cre recombinase. Generally, a recombinant Cre-expressing adenovirus is used for this purpose. Materials High-titer Cre-expressing adenovirus (e.g., Microbix Biosystems) Heterozygous knockout cell lines (see Basic Protocol 2) growing at 50% confluence in 25-cm2 tissue culture flasks Complete McCoy’s 5A medium (see recipe) with and without 0.6 µg/ml G418 (geneticin, Life Technologies; see UNIT 9.5) Hanks’ Balanced Salt Solution (HBSS, Life Technologies; also see APPENDIX 2) 0.05% (w/v) trypsin/0.53 mM tetrasodium EDTA (Life Technologies) 25-cm2 tissue culture flasks 96-well tissue culture plates 24-well tissue culture plates Additional reagents and equipment for culture of mammalian cells (including counting and cryopreservation; APPENDIX 3F), plating cells at limiting dilution (as for hybridoma cells; UNIT 11.8), and crystal violet staining of cultured cells (as for plaque assay; see UNIT 16.16, Support Protocol)



Supplement 63

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. 1. Add 1 µl recombinant adenovirus to heterozygous knockout cells at 50% confluence in a 25-cm2 tissue culture flasks containing 5 ml complete McCoy’s 5A medium without selective agent (G418). Incubate 24 hr. 2. Wash cells twice with HBSS, then trypsinize (see Basic Protocol 2) and count cells (APPENDIX 3F). 3. Plate out at limiting dilution (UNIT 11.8) in 96-well plates in complete McCoy’s 5A medium without selective agent, to obtain single colonies. To be sure to obtain single colonies, set up a variety of 96-well plates, e.g., with 1 cell/well, 5 cells/well, and 10 cells/well. 4. Identify single colonies after 10 days, and trypsinize at ∼2 weeks as described in Basic Protocol 2. Expand each clone to a well in a 24-well plate, and then to two 25-cm2 tissue culture flasks. One of the 25-cm2 flasks will be used to test the G418 sensitivity, and the other will be used for cryopreservation.

5. Add complete McCoy’s 5A medium containing 0.6 µg/ml G418 to one of the 25-cm2 flasks when it reaches 50% confluence. Continue incubating for ∼10 to 14 days, feeding every 3 days. Cryopreserve the cells in the other 25-cm2 flask when it reaches confluence, as described in APPENDIX 3F. 6. After ∼10 to 14 days, stain the 25-cm2 flask containing the cells exposed to selective agent with crystal violet as described in UNIT 16.16. Identify those cell lines that are now completely G418-sensitive, and use them to disrupt the remaining allele (see Basic Protocol 4). ALTERNATE PROTOCOL

SWAPPING SELECTABLE MARKERS IN THE TARGETING VECTOR An alternative to recycling the neor targeting vector (see Basic Protocol 3) is to modify the targeting vector to switch selectable markers, replacing neor with hygr. It should be noted that the authors have been consistently unable to generate drug-resistant colonies when using puror, IRES-hygr, or IRES-puror (C. Lee, J. S. Kim, T. Waldman, unpub. observ.). To do this, repeat Basic Protocol 1 with PCR amplification of a hygr gene. Perform Basic Protocol 2 as written, except substitute 0.1 µg/ml hygromycin for G418.

BASIC PROTOCOL 4


TARGETING OF THE REMAINING ALLELE The principles and protocols needed for targeting the remaining allele are virtually identical to those already described for targeting the first allele. If recycling the neor targeting vector, one should repeat Basic Protocol 2, using G418-sensitive heterozygous knockout cell lines (see Basic Protocol 3) as the “parental” cells. If swapping selectable markers in the targeting vector, use neor heterozygous knockout cell lines as the “parental” cells. Of the homologous integration events obtained, approximately half should represent homozygous knockouts, and the other half should have targeted the already targeted allele. After confirmation of the homozygous knockout with PCR (UNIT 15.1) and Southern blotting (UNIT 2.5A), the authors generally perform an immunoblot (UNIT 10.8) to confirm the absence of protein.




Complete McCoy’s 5A medium McCoy’s 5A medium (Life Technologies cat. no. 16600-082) 10% fetal bovine serum (FBS, Life Technologies; also see APPENDIX 3F) 1× penicillin/streptomycin (Life Technologies cat. no. 15140-122) Store up to 4 months at 4°C COMMENTARY Background Information History of human somatic cell gene targeting The first descriptions of successful gene targeting in cultured human cells appeared in 1991, when two groups reported heterozygous targeting of different genes. Heterozygous targeting of the 6-16 gene, an interferon-inducible transcript, was reported by Itzhaki and Porter (1991). The frequency of homologous integration was extremely low (estimated at 1 per 1550 colonies screened), leading these authors to employ an unusual screen involving integration of a human growth hormone gene (hGH) and the screening of conditioned media from putative clones via hGH ELISA. That same year, heterozygous targeting of the HPRT gene was reported (Zheng et al., 1991). That publication was a proof-of-principle study in which the authors were able to directly select for heterozygous knockout cell lines. In 1993, the first homozygous (or complete) knockout of a gene in human cells was reported (Porter and Itzhaki, 1993). Using a promoterless targeting vector, Porter and Itzhaki were able to disrupt the remaining allele of 6-16 in the heterozygous gene-targeted cells described in their 1991 paper. These three early studies demonstrated the feasibility of gene targeting in human cells; they also demonstrated that it was a very laborintensive process. In 1993, a landmark paper in the history of human somatic cell gene targeting was published, reporting the heterozygous gene targeting of the K-ras oncogene in two human colon cancer cell lines (Shirasawa et al., 1993). Although this was not the first manuscript to describe human somatic cell gene targeting, it pointed out the potential utility of somatic cell gene targeting for studying human cancer. In particular, these authors created derivatives of both HCT116 and DLD1 cells in which either the wild-type or the oncogenic allele of K-ras

had been deleted. They went on to show that the K-ras oncogene was absolutely required for the tumorigenic properties of both cell lines. They reported an extremely high frequency of knockouts (8/93 in HCT116; 7/24 in DLD1), attributing it to the use of a promoterless neor gene. Furthermore, they introduced the use of HCT116 and DLD1 cells for gene targeting. In 1995, a careful study was published that examined the merits of promoterless versus conventional targeting vectors for somatic cell gene targeting (Hanson and Sedivy, 1995). Their conclusion bolstered the work done by Shirasawa et al. (1993) showing that promoterless targeting vectors were virtually essential to achieve high frequencies of homologous recombination. They also reported that negative selectable markers commonly used for gene targeting in embryonic stem cells were of limited utility for somatic cell gene targeting. Furthermore, they pointed out that while the neor gene was extremely useful for gene targeting, the hygr gene seemed to work poorly. Subsequent studies, mostly unpublished, confirmed these seminal observations. Also in 1995, homozygous deletion of the p21 cdk inhibitor in human HCT116 cells was reported (Waldman et al., 1995). This study brought together several of the most recent technical advances, such as the use of promoterless targeting vectors, the lack of a need for negative selectable markers, and the value of HCT116 cells. Importantly, this study demonstrated the feasibility and utility of human somatic cell gene targeting for the study of tumor suppressor gene pathways. Other important incremental technical gains have been made as well. Advances have been made that simplify the construction of human promoterless targeting vectors, such as the use of homologous recombination in Saccharomyces cerevisiae (see UNIT 13.10). An important modification to the standard PCR screen for knockouts has made identification of gene tar-



Supplement 62

geted cell lines simpler and more robust (see UNIT 13.10). Cre-loxP recombination (see UNITS 9.5 & 13.3) has been successfully employed to overcome difficulties associated with creation of homozygous knockouts by making it possible to recycle the same targeting vector for both alleles of a gene. Today, through the implementation of these advances, human somatic cell gene targeting has emerged as an important tool for the study of gene function in human cells.



The frequency of homologous recombination in human cells is 3 to 5 orders of magnitude lower than the frequency of nonhomologous recombination. As such, the vast majority of drug-resistant clones formed after transfection of a conventional targeting vector represent random integration events. Identification of the tiny fraction of gene-targeted clones can be extremely challenging. It should be noted that the frequency of homologous integration in human cells is thought to be significantly lower than the frequency of homologous integration in murine embryonic stem (ES) cells. This fact, coupled with the fact that creation of homozygous knockouts in human cells requires sequential targeting of both alleles (unlike in mice, in which heterozygous knockouts are mated to homozygosity), renders promoter-containing targeting vectors unsuitable for human somatic cell gene targeting. Instead, promoterless targeting vectors are required. In such targeting vectors, the selectable marker gene (generally neor) lacks its own promoter and is instead fused in-frame with the gene being targeted. The most important critical parameter is the efficiency with which the targeting vector creates gene-targeted cell lines. Vectors vary in their efficiency from 30°C. This makes 15 ml acrylamide solution, sufficient for twenty 1.5 × 160–mm gels. Although the BDH pH 4 to 8 Resolytes are highly recommended, one may wish to use a blend of ampholytes as a substitute. For a broad-range blend, mix 1 part ampholytes, pH 2 to 11 (Serva) and 2 parts ampholytes, pH 3.5 to 10 (Amersham Biosciences). For an alternative broad-range blend, use equal volumes of ampholytes, pH 3 to 10 (Bio-Rad) and pH 2 to 11 (Serva). Mix enough ampholytes at one time to last ≥1 year. This will improve run-to-run reproducibility. This alternative can be used if Resolytes are unavailable or for a custom pH gradient.

7. Deaerate the solution by applying a strong vacuum for 2 to 3 min. 8. Add 0.3 ml NP-40 and swirl until dissolved. Do not add the NP-40 before deaeration because it will foam.

9. Pour solution into a 20-ml syringe fitted with a 0.2- or 0.45-µm filter capsule and force through the filter. 10. Add 10 ml TEMED and swirl. Add 70 ml of 10% ammonium persulfate and swirl. Immediately pipet the gel solution into the space between the gel tubes and the large glass tube. Work quickly, as the gel solution will start to polymerize in ∼3 min.

11. Gently run water down the outside of the gel tubes using a wash bottle. Add water until the level of the acrylamide solution inside the tubes reaches the desired height. As water is layered on top of the gel solution it will force the solution up the tubes from the bottom. There is no need to overlay the gel solution inside the tubes. Allow ≥1 hr for polymerization. Analysis of Proteins


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Set up the gels in the tube cell 12. Remove the Parafilm from the bottom of the gel casting tube and push the gel tubes, containing the polymerized gel, out the bottom. Cut across the gel-tube bottoms to remove excess acrylamide with a single-edge razor blade. Rinse the bottom of the gel tubes under running deionized water to remove residual acrylamide. 13. Place rubber grommet on the top of each tube, making sure that the top surface of the gel is visible below the grommet; ∼5 mm of the gel tube should be visible above the grommet. 14. Seat the tube and grommet assemblies in the holes of the upper buffer reservoir of the tube cell. Plug any unused holes with rubber stoppers. 15. Fill the lower reservoir with ∼3 liters of 0.085% phosphoric acid. 16. Place the upper reservoir into lower reservoir and adjust lower buffer level if needed. The buffer should cover the entire gel for good heat dissipation.

17. Fill the upper buffer reservoir with 250 ml of 0.02 M NaOH. 18. Fill the gel tubes to the top with 0.02 M NaOH using a 1-ml syringe equipped with a 22-G hypodermic needle. Be careful to eliminate any air bubbles in the gel tubes. 19. Connect the tube cell to the power supply. The black (−) lead goes to the upper reservoir. Prefocus the gel 1 hr at 200 V constant voltage (see UNIT 10.2A introduction for a discussion of electricity and electrophoresis). Disconnect the tube cell from the power supply. Load the samples and run the gels 20. Layer protein samples on top of the gels through the upper buffer with a 50-µl syringe. The maximum amount of total protein that can be loaded onto a first-dimension gel varies depending upon the nature of the sample. Samples such as whole-cell lysates which contain a large number of proteins of widely varied isoelectric point (pI) can contain much more total protein than a sample of a single highly purified protein. As much as 100 to 150 ìg of a protein mixture can be loaded on a first-dimension gel 1.5-mm in diameter, while a tenth that amount of highly purified protein would probably be an overload. For gels 1.5-mm in diameter, 10 to 20 ìl of sample is preferred; however, up to 30 ìl can be applied. Alternatively, place the samples directly on the gel, then overlay with 10 to 20 ìl half-strength solubilization buffer (diluted 1:2 with water). This will protect samples from exposure to the basic upper buffer. Fill the remainder of the tube with 0.02 M NaOH to eliminate any bubbles.

21. Place the lid on the upper reservoir and attach the electrical leads to a power supply. 22. Turn on the power supply and adjust to the desired settings at constant voltage. For 1.5-mm-i.d., 16-cm-long gels, 700 to 800 V (constant voltage) for 16 hr (11,000 to 13,000 V hr) works well for most samples.

23. Reduce the voltage setting to zero and turn off the power supply to end the run. Add ∼1 µl concentrated bromphenol blue to the top of each gel with a 50-µl syringe. The bromphenol blue will quickly diffuse into the gel. At the end of the run, the gels may be noticeably shorter and the bromphenol blue may stop short of the top of the gel. In this case, the power can be applied for a few minutes and the dye will migrate into the top of the gels. The blue dye will mark the basic end of the first-dimension gel and serve as the tracking dye in the second-dimension separation. Two-Dimensional Gel Electrophoresis



Extrude the gels 24. Extrude the gels from the tubes using water pressure from a 1-ml syringe fitted with a 200-µl pipettor tip (cut off ∼1 cm of the large end of the tip so it fits on the syringe). Gels with a diameter >2.0 mm may have to be loosened by inserting a blunt 26-G, 9-cm-long needle between the gel and the wall of the tube while injecting water. Rotate the tube so the needle will pass over the entire circumference of the gel while continuously injecting water. Repeat this operation at the opposite end of the tube and the gel should slide out easily.

25. Place each gel in a labeled gel vial. The gels can be stored at −70°C for many weeks or used immediately. 26. Soak gel tubes overnight in chromic acid cleaning solution, then rinse thoroughly under running deionized tap water for 15 min. Remove excess water from the gel tubes with suction and allow them to dry. SOLUBILIZATION AND PREPARATION OF PROTEINS IN TISSUE SAMPLES

SUPPORT PROTOCOL 1

Tissue samples are solubilized in either SDS or urea solubilization buffer (see commentary) by Dounce homogenization. After centrifugation, the protein sample can be loaded onto the first-dimension gel. Materials Tissue samples SDS or urea solubilization buffer (see recipes) Dounce homogenizer with pestles A and B 200-µl centrifuge tubes Beckman 42.2-Ti rotor (or equivalent) 1. Weigh and place tissue samples in a Dounce homogenizer. 2. Add 1.5 to 2.0 ml SDS or urea solubilization buffer per 100 mg tissue. 3. Homogenize using 50 strokes with pestle B, then 50 strokes with pestle A. Cultured cells and body fluids do not normally require homogenization.

4. Let stand a few minutes, then transfer an aliquot to a 200-µl centrifuge tube. Centrifuge ≥2 hr at 100,000 × g, or 1 hr at >200,000 × g, 20°C. Save the supernatant (protein sample) to load onto the first-dimension gel as described in Basic Protocol 1. If using SDS solubilization buffer, place the sample in boiling water 5 min before centrifugation. Never heat samples in urea solubilization buffer. Heating proteins in a urea solution will cause carbamylation of proteins, and thus result in charge variants. Centrifugation should be done just prior to loading the sample onto the first-dimension gel. Failure to centrifuge tissue samples may cause plugging of the first-dimension gels, resulting in poor entry of proteins. The Beckman 42.2-Ti rotor, which accommodates seventy-four 200-ìl tubes, is recommended.

SECOND-DIMENSION GELS A wide variety of equipment suitable for casting and running second-dimension gels is available. Nearly any SDS-PAGE slab-gel system will work. For the best possible resolution over a wide molecular-weight range, a 10% to 20% acrylamide gradient gel is recommended (UNIT 10.2A); however, for many applications, a conventional slab gel as described below is sufficient.The procedure described uses the Bio-Rad PROTEAN II system, but it can be easily adapted to similar equipment, such as the vertical slab-gel units from Hoefer.

BASIC PROTOCOL 2

Analysis of Proteins


Supplement 67

The second-dimension gel is poured between glass plates. The first-dimension gel is then loaded onto the second-dimension gel and sealed in place with agarose. The gels are mounted in the cell and run. After disassembling the electrophoresis unit, the gel is stained and analyzed. Materials 30% acrylamide/0.8% bisacrylamide (see recipe for acrylamide/bisacrylamide solutions) Gel buffer (see recipe) 10% (w/v) SDS TEMED 10% ammonium persulfate (see recipe) Isobutyl alcohol, H2O-saturated Stacking gel buffer (optional; see recipe) First-dimension gel (see Basic Protocol 1) Equilibration buffer (see recipe) Hot 0.5% and 1% agarose (see recipe; keep in boiling water bath) Protein molecular weight standards (Table 10.2A.2; available as kits Bio-Rad or Amersham Biosciences) SDS solubilization buffer (see recipe) Reservoir buffer (see recipe), prechilled to 10° to 20°C Coolant (from running tap water or circulating refrigerated water bath) Gel plates, one long and one short 1.5-mm spacers (∼14 cm × 14 cm × 0.75 mm) Casting stand Gel identification tag (e.g., typed consecutive numbers on filter paper) Nylon screen 5 × 15–cm glass plate PROTEAN II electrophoresis cell (Bio-Rad) Prepare and pour the gel 1. Assemble the gel plates by placing 1.5-mm spacers vertically between a long and short gel plate. The side where the long plate protrudes is the top. Gel thickness is usually equal to the diameter of the isoelectric focusing gels, but this is not absolutely necessary.

2. Position clamps on each side of the gel sandwich over the spacers and place on the casting stand. Be sure the plates and spacers are properly aligned, then tighten the clamps and cams to get a leak-proof seal. Make adjustments so that plates are level and vertical. 3. Place the gel identification tag between the glass plates so that it rests in the lower right hand corner. 4. Prepare the gel solution by combining 30% acrylamide/0.8% bisacrylamide, gel buffer, and water in a vacuum flask (Table 10.4.1). 5. Deaerate the solution by applying vacuum for 5 min. 6. Add 10% SDS and TEMED and swirl, then add 10% ammonium persulfate and swirl (Table 10.4.1).

Two-Dimensional Gel Electrophoresis

7. Fill the gel sandwich to 5 mm below the top of the short plate and overlay with H2O-saturated isobutyl alcohol or water.



Table 10.4.1

Solutions for Second-Dimensional Gelsa

Final acrylamide concentration (%) Stock solutions

7.5

10

12.5

15

30% acrylamide/ 0.8% bisacrylamide Gel buffer H 20 10% SDS TEMED 10% ammonium persulfate

12.5

16.7

20.8

25.1

12.5 24.6 0.5 0.026 0.185

12.5 20.3 0.5 0.026 0.185

12.5 16.2 0.5 0.026 0.185

12.5 12.0 0.5 0.026 0.185

aValues in table body are milliliters of stock solution. Preparation of solutions is described in steps 4 to 6 of protocol.

A stacking gel is not usually required when running second-dimension gels. If a stacker is desired, stop the gel 1.5 cm from the top of the short plate, overlay with H2O-saturated isobutyl alcohol, and allow to polymerize 1 hr. Remove the overlay and fill the remaining space with stacking gel solution made by combining 4.8 ml stacking gel buffer, 3.8 ml water, and 1.4 ml 30% acrylamide/0.8% bisacrylamide (all deaerated) with 200 ìl of 10% ammonium persulfate and 8 ìl TEMED. Pipet onto the separating gel, overlay with H2O-saturated isobutyl alcohol, and allow to polymerize 30 min.

8. Allow the second-dimension gel to polymerize ≥1.5 hr. Load first-dimension gel onto second-dimension gel 9. If first-dimension gel is frozen, thaw at room temperature. Add equilibration buffer to completely cover the gel. The time the gel is in equilibration buffer containing SDS can vary from seconds to several minutes depending upon the sample. Most proteins run well without the introduction of SDS from equilibration buffer; however, a few proteins do not. Leaving gels in equilibration buffer for >10 to 15 min can result in loss of proteins.

10. Pour the gel and equilibration buffer onto a nylon screen placed over a beaker. 11. Place the first-dimension gel on a 5 × 15–cm glass plate (Parafilm is not rigid enough). Using a spatula, lay the gel out straight along one edge of the glass plate. 12. Pipet a very thin layer of hot 0.5% agarose on the top of the slab gel to be loaded. This can easily be done by putting ∼0.1 ml hot agarose on the upper left corner of a gel and quickly tilting the gel to the right so that the agarose will flow across the gel surface.

13. Using a spatula, slide the first-dimension gel off the glass plate and place it across the top of the slab gel. Orient the first dimension gel with the blue (basic) end to your right (Fig. 10.4.2). Take care that no air bubbles get trapped between the first- and second-dimension gels and that the first-dimension gel does not get stretched or compressed.

14. Pipet a thin layer of hot 0.5% agarose over the first-dimension gel to seal it in place. Allow the agarose to solidify. Protein molecular-weight markers may be run as one-dimensional separations on the sides of the second-dimension gel. Solubilize marker proteins in SDS solubilization buffer by boiling for 5 min, then dilute 1:1 with hot 1% agarose solution and draw up the hot solution in a glass tube the same diameter as the first-dimension gel. A short piece of the solidified agarose can be applied to one or both sides of the second-dimension gel and held in place with 0.5% agarose.



Supplement 67

protein-molecular-weight standards in agarose

firstdimensional gel seconddimensional gel gel identification tag

Figure 10.4.2 Assembly of second-dimension gel.

Run and analyze the gel 15. Mount the gels on the PROTEAN II electrophoresis cell (the short plate goes toward the center of the unit). 16. Fill the upper and lower reservoirs with prechilled reservoir buffer. 17. Attach tubing for coolant to the in and out ports and start the flow of coolant to maintain the temperature of the tank buffer at 10° to 20°C during the run to ensure that the gels are adequately cooled. The tank buffer should be the same temperature for each run.

18. Attach the electrical leads to the power supply (the upper reservoir is connected to the negative lead). Electrophorese at 15 to 20 mA/gel until the tracking dye reaches the end of the gel (or 3 to 5 mA/gel overnight). 19. Reduce the voltage setting to zero and turn off the power supply at the end of the run. Remove the gels from the electrophoresis unit and take off the clamps. Pry the glass plates apart with a spatula. 20. Stain the gels (UNIT 10.6), process the gels for immunoblotting (UNIT 10.8), or autoradiograph the gels (APPENDIX 3A). ALTERNATE PROTOCOL 1

ISOELECTRIC FOCUSING OF VERY BASIC PROTEINS USING NEPHGE First-dimension IEF gels run in the standard way using broad-range ampholytes will resolve proteins with isoelectric points (pI) between 3.8 and 8. More basic or more acidic proteins may be resolved by modifying the first-dimension gels using nonequilibrium pH gradient electrophoresis (NEPHGE) as described below (Anderson, 1988). Additional Materials (also see Basic Protocol 1) Ampholytes, pH 2 to 11 (Serva) 0.01 M phosphoric acid, deaerated 4 M urea, deaerated To analyze very basic proteins, the procedure is the same as described in Basic Protocol 1 for first-dimension gels with the following exceptions in the indicated steps: 6a. Use 0.75 ml ampholytes, pH 2 to 11, in gel solution.


15a. Fill the lower reservoir with ∼3 liters deaerated 0.02 M NaOH.



17a. Fill the upper reservoir with 250 ml deaerated 0.01 M phosphoric acid. 19a. Do not prefocus gels. 20a. Overlay samples with deaerated 4 M urea to protect proteins from phosphoric acid. 21a. Attach the negative lead to the lower tank and the positive to the upper tank (this is the reverse of the usual setup). 22a. Focus 1 hr at 400 V and then 4 to 5 hr at 800 V for a total of 4000 V-hr. Because the bromphenol blue applied to the top of the gel at the end of the run now marks the acidic end, load first-dimension gels onto the second-dimension gel with the blue end to the left.

ISOELECTRIC FOCUSING OF VERY ACIDIC PROTEINS USING NEPHGE Additional Materials (also see Basic Protocol 1) Ampholytes, pH 2.5 to 4 (Pharmacia LKB) Concentrated sulfuric acid Water, deaerated


To analyze very acidic proteins, the procedure is the same as described in Basic Protocol 1 for first-dimension gels with the following exceptions in the indicated steps: 6b. Prepare the gel solution as follows: 8.25 g urea 5.5 ml H2O 2.0 ml 30% acrylamide/1.8% bisacrylamide 1.0 ml ampholytes, pH 2.5 to 4 (Pharmacia LKB) 0.3 ml ampholytes, pH 2 to 11 (Serva). 10b. Add 90 µl of 10% ammonium persulfate and swirl, then add 10 µl TEMED and swirl. 15b. Prepare lower buffer by adding 4.5 ml concentrated sulfuric acid to 3 liters of water and fill the lower reservoir. 17b. Prepare upper buffer by adding 3 ml ampholyte, pH 2 to 11, to 120 ml deaerated water and place in the upper reservoir. 22b. Run at 800 V for 4.5 to 5.0 hr or 250 V for 16 hr for a total of 3600 to 4000 V-hr. TWO-DIMENSIONAL MINIGELS Though limited in resolving area, small two-dimensional SDS-PAGE is a quick way to separate proteins for a variety of applications. These include peptide sequencing, purity checks, and protocol development. Where a large-format two-dimensional SDS-PAGE separation might take two days, a minigel separation (UNIT 10.2A) takes 4 to 5 hr, including the first-dimension (IEF) gel. The solutions and procedures in the basic protocols directly translate to the small format, except that much less reagent is required and electrophoresis times are much shorter. To perform two-dimensional SDS-PAGE in the small format, a few simple changes from the basic protocol are needed. Tube gel adaptors for IEF, such as the Hoefer SE 220 or the Bio-Rad Mini-Protean II Tube Cell, fit in the minigel unit (UNIT 10.2A). Because the tube gels are much shorter (6 to 8 cm), the isoelectric focusing time is less; 2 to 4 hr at 500 V is usually adequate. Furthermore, less protein is generally required. Stacking gels




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are not normally required, so the mini slab gel should be cast to within 0.5 to 1 cm of the top. The second-dimension gel is processed in the same way as a first-dimension minigel (UNIT 10.2A). ALTERNATE PROTOCOL 4

TWO-DIMENSIONAL (2-D) ELECTROPHORESIS WITH IMMOBILIZED pH GRADIENTS In Basic Protocol 1, first-dimension separation is performed in carrier ampholyte–containing polyacrylamide gels cast in narrow tubes. In this protocol, the carrier ampholyte– generated pH gradients have been replaced with immobilized pH gradients and the tube gels have been replaced with thin rectangular IEF gels supported by a plastic backing (Westermeier et al., 1983; Berkelman and Stenstedt, 1998; Görg et al., 2000). Such gradients produce superior resolution and reproducibility (Bjellqvist et al., 1982). This protocol describes the use of the Ettan IPGphor isoelectric focusing system (Amersham Biosciences), but the PROTEAN IEF System form Bio-Rad is also popular. The cost is is higher than when Basic Protocols 1 and 2 are used, but many laboratories have adopted these systems. The use of commercial quality-controlled and standardized reagents and instrumentation greatly simplifies the process of 2-D analysis and makes it possible to obtain much more consistent results. Materials Protein sample for analysis Sample solution: default sample solution, hydrophobic sample solution, or tissue sample solution (see recipes) Tissue homogenization solution (see recipe) Rehydration stock solution (see recipe) IPG buffer or Pharmalyte (same range as the IPG strip; Amersham Biosciences) Cleaning solution (e.g., Ettan IPGphor Strip Holder Cleaning Solution; Amersham Biosciences) IPG Dry Strip Cover Fluid (Amersham Biosciences) Vertical gel for SDS-PAGE (Basic Protocol 2 in UNIT 10.2A) SDS equilibration buffer (see recipe) SDS electrophoresis buffer (UNIT 10.2A) Molecular weight markers (UNIT 10.2A) Agarose sealing solution (see recipe) Isoelectric focusing system (Ettan IPGphor, Amersham Biosciences; PROTEAN IEF System, Bio-Rad) IPG strips (Amersham Biosciences) Toothbrush Platform rocker 100°C heating block Thin plastic ruler Additional reagents and equipment for second-dimension gel electrophoresis (see Basic Protocol 2) Prepare the sample 1a. For an unknown protein: Dissolve sample in default sample solution. 1b. For large and more hydrophobic proteins: Dissolve sample in hydrophobic sample solution.


1c. For proteins from tissues that are dilute sources of protein and contain high levels of interfering substances (e.g., plant tissues): Grind tissue in mortar and pestle with liquid nitrogen. Suspend powder in tissue homogenization solution. Keep at −18°C



Table 10.4.2

Suitable Sample Loads for Silver and Coomassie Staining Using Rehydration Loading

Immobiline DryStrip Size 7 cm

11 cm

13 cm

18 cm

pH range

Silver stain

Coomassie stain

4–7 6–11

4-8 8-16

20-120 40-240

3–10, 3–10 NL 4–7

2-4 10-20

10-60 50-300

6–11 3–10 L

20-40 4-8

100-600 20-120

4–7 6–11

15-30 30-60

75-450 150-900

3–10, 3–10 NL 4–7

8-15 30-60

40-240 150-900

60-120 15-30

300-1500 75-450

4–7, 3–7 6–9, narrow intervala

45-90 80-170

200-1300 400-2000

3–10, 3–10 NL

20-40

100-600

6–11, 6–9, narrow 3–10, 3–10 NL 24 cm

Suitable sample load (µg of protein)

intervala

Rehydration solution volume, per strip 125 µl

200 µl

250 µl

340 µl

450 µl

aImmobiline DryStrip narrow intervals pH: 3.5–4.5, 4.0–5.0, 4.5–5.5, 5.0–6.0, and 5.5–6.7.

overnight and centrifuge. Wash pellet with acetone. Dry and suspend in tissue sample solution. This method produces protein solutions substantially free of salts, nucleic acids, and other contaminants.

Rehydrate IPG strips IPG strips must be rehydrated prior to IEF. With integrated IEF systems such as the Amersham IPGphor Isoelectric Focusing and Bio-Rad PROTEAN system, both rehydration of the IPG strip and IEF occur in individual strip holders or trays. Different strip holder lengths are available for different IPG strip lengths. If the strip holders are in use for IEF, rehydration may be carried out in the Immobiline DryStrip Reswelling Tray (see Support Protocol 2). 2. Just prior to use, slowly thaw a 2.5-ml aliquot of rehydration stock solution. Add the appropriate amount (see Reagents and Solutions) of IPG Buffer or Pharmalyte, if it is not already included in the rehydration stock solution, and add 7 mg DTT. Sample may be applied now (“rehydration loading”), or at step 10. If rehydration loading is desired, refer to Table 10.4.2 for suitable sample loads. DTT and the sample must be added fresh, just prior to use.

3. Select the strip holder(s) corresponding to the IPG strip length chosen for the experiment. Handle the ceramic strip holders with care, as they are fragile.

4. Wash each strip holder with detergent to remove residual protein, using a neutral-pH detergent (e.g., Ettan IPGphor Strip Holder Cleaning Solution) as follows. a. Rinse off the strip holder. A mild liquid soap may be used to remove any residual DryStrip cover fluid. Analysis of Proteins


Supplement 67

Table 10.4.3

Recommended Final Concentrations and Volumes of IPG Buffer/Pharmalyte for the Rehydration Solutiona

Strip holder Cup loading

Rehydration and sample well loading Vertical gels, flatbed Cup holder Vertical gels, flatbed

0.5% IPG Buffer (12.5 µl per 2.5 ml) 2% IPG Buffer (50 µl per 2.5 ml)

aThe recommended IPG Buffer / Pharmalyte concentration for the IPGphor system is 0.5%, but up to 2% can be added if sample solubilization remains

a problem.

b. Squeeze a few drops of Ettan IPGphor Strip Holder Cleaning Solution into the strip holder slot. Use a toothbrush and vigorous agitation to clean the strip holder. c. Rinse well with distilled or deionized water. Thoroughly air dry the strip holders or dry well with a lint-free tissue prior to use. The Ettan IPGphor Strip Holder Cleaning Solution has been specifically formulated for removing protein deposits and will not damage the strip holder. Clean strip holders after each first-dimension IEF run. Do not let solutions dry in the strip holder. Cleaning may be more effective if the strip holders are first soaked a few hours overnight in a solution of 2% to 5% Ettan IPGphor Strip. Recalcitrant or dried-on protein deposits may be removed with hot (up to 95°C) 1% (w/v) SDS. Add 1% (w/w) DTT for complete removal of sticky proteins. Rinse completely with distilled or deionized water after cleaning. Handle clean strip holders with gloves to avoid contamination. Strip holders may be baked, boiled, or autoclaved. Do not expose them to strong acids or bases, including alkaline detergents.

5. Pipet the appropriate volume of rehydration solution into each strip holder as indicated in Table 10.4.3. Deliver the solution slowly at a central point in the strip holder channel away from the sample application wells. Remove any larger bubbles. To ensure complete sample uptake, do not exceed the recommended volume of rehydration solution (see Table 10.4.3).

6. Place the IPG strip in the strip holder as follows. a. To prevent damage to the basic (square) end of the IPG strip, which is generally softer, remove the protective cover foil from the IPG strip starting at the acidic (pointed) end. b. Position the IPG strip with the gel side down and the pointed (anodic) end of the strip directed toward the pointed end of the strip holder. c. Pointed end first, lower the IPG strip onto the solution. To help coat the entire strip, gently lift and lower the strip and slide it back and forth along the surface of the solution, tilting the strip holder slightly as needed to assure complete and even wetting. d. Finally, lower the cathodic (square) end of the IPG strip into the channel, making sure that the gel contacts the strip holder electrodes at each end. The gel can be visually identified once the rehydration solution begins to dye the gel. Be careful not to trap air bubbles under the IPG strip.

7. Apply IPG Dry Strip Cover Fluid to minimize evaporation and urea crystallization. Pipet the fluid dropwise into one end of the strip holder until one half of the IPG strip is covered, then pipet the fluid dropwise into the other end of the strip holder, adding fluid until the entire IPG strip is covered. 8. Place the cover on the strip holder. Pressure blocks on the underside of the cover assure that the IPG strip maintains good contact with the electrodes as the gel swells. Two-Dimensional Gel Electrophoresis



9. Allow the IPG strip to rehydrate. For rehydration loading of sample (see annotation to step 2), there are two possible rehydration conditions: passive rehydration with no electric field applied during rehydration, and rehydration under voltage. In some cases, rehydration under a low voltage (30 to 120 V) facilitates the entry of high-molecular weight proteins. Rehydration can proceed on the bench top or on the Ettan IPGphor unit platform. Ensure that the strip holder is on a level surface. A minimum of 10 hr is required for rehydration; overnight is recommended. The rehydration period can be programmed as the first step of an Ettan IPGphor protocol. This is especially convenient if temperature control during rehydration is a concern.

Perform first-dimension electrophoresis 10. If the sample was not applied as a part of the rehydration solution in step 2, pipet ≥7.5 µl of sample solution prepared in step 1a, b, or c (i.e., 15 µl per well or 30 µl total if both sides of both wells are used) to the lateral wells at either end of the strip holder, below the IPG DryStrip Cover Fluid. Replace cover on strip holder. When using the alternative “cup loading” procedure (also see instrumentation manual) an increased sample concentration will lead to an increased risk of protein precipitation in the sample cup. A maximum concentration of 100 ìg protein/100 ìl sample solution (100 ìl is the volume of the cup) is recommended. This is a general recommendation, which will work for most samples, but the maximum concentration that can be used varies greatly between sample types. For larger sample loads, rehydration loading (see annotation to step 2 and Table 10.4.2) is recommended. The IPG strip backing is impermeable; do not apply the sample to the back of the strip.

11. Perform first-dimension IEF separation per manufacturer’s instructions. See Table 10.4.4 for guidelines on different experimental setups. Equilibrate strips prior to second-dimension electrophoresis The second-dimension vertical gel must be ready for use prior to IPG strip equilibration. See Basic Protocol 2 for preparation of vertical gels for 2-D SDS PAGE. 12. Place the IPG strips in individual tubes with the support film toward the wall of the tube. Add 10 ml SDS equilibration buffer (containing DTT) to each tube. Cap the tube, and place it on its side on a platform rocker. Equilibrate for 15 min. 13. If desired, perform a second equilibration with SDS equilibration buffer containing 250 mg/ml iodoacetamide (without DTT). Add 10 ml of solution per tube. Cap the tube, place it on its side on a platform rocker, and equilibrate for 15 min. This second equilibration step reduces point streaking and other artifacts.

14. Dip the equilibrated IPG strip in SDS electrophoresis buffer to lubricate it. 15. Optional: Apply molecular weight marker proteins to a separate small strip. Best results are obtained when the molecular weight marker protein solution is mixed with an equal volume of a hot 1% agarose solution prior to application to the IEF sample application piece. The resultant 0.5% agarose will gel and prevent the marker proteins from diffusing laterally prior to the application of electric current. Other alternatives are to apply the markers to a paper IEF sample application piece in a volume of 15 to 20 ìl. For less volume, cut the sample application piece proportionally. Place the IEF application piece on a glass plate and pipet the marker solution onto it, then pick up the application piece with forceps and apply to the top surface of the gel next to one end of the IPG strip. The markers should contain 200 to 1000 ng of each component for Coomassie staining and about 10 to 50 ng of each component for silver staining.

16. Melt each aliquot of agarose sealing solution needed (each gel will require 1 to 1.5 ml) in a 100°C heat block (it takes ∼10 min to fully melt the agarose; an ideal time to carry out this step is during IPG strip equilibration). Allow the agarose to cool until



Supplement 67

Table 10.4.4

Guidelines for Ettan IPGphor with Rehydration Loading/IEF for Immobiline Dry Stripsa

pH range(s)

Step and voltage mode

Step durationb (h:min)

Kilovolt-hours (kV-hr)

7 cm

3–10 3–10 NL 4–7

11 cm

3–10 4–7

13 cm

3–10 3–10 NL 4–7

18 cm

3–10 3–10 NL 4–7

1. Step and hold 500 2. Step and hold 1000 3. Step and old 5000c Total 1. Step and hold 500 2. Step and hold 1000 3. Step and hold 8000c Total 1. Step and hold 500 2. Step and hold 1000 3. Step and hold 8000c Total 1. Step and hold 500 2. Step and hold 1000 3. Step and hold 8000c Total 1. Step and hold 500 2. Step and hold 1000 3. Step and hold 1000c Total 1. Step and hold 500 2. Step and hold 1000 3. Step and hold 8000c Total 1. Step and hold 500 2. Step and hold 1000 3. Step and hold 8000c Total

0:30 0:30 1:40 2:40 1:00 1:00 1:50 3:50 1:00 1:00 2:00 4:00 1:00 1:00 4:00 6:00 1:00 1:00 7:30 9:30 1:00 1:00 8:20 10:20 1:00 1:00 10:30 12:30

0.25 0.5 7.5 8.0 0.5 1.0 12.5 14.0 0.5 1.0 14.5 16.0 0.5 1.0 30.5 32.0 0.5 1.0 58.5 60.0 0.5 1.0 62.5 64.0 0.5 1.0 94.5 96.0

Immobiline DryStrip

Narrow intervalsd

24 cm

3–10 3–10 NL 4–7 3–7 Narrow intervalsd

aVoltage step and hold mode, 50 µA/IPG strip; 0.5% IPG buffer; 20°C for both rehydration and IEF. Rehydration time, 12 hr. The total rehydration

time can be adjusted somewhat for convenience, but must be >10 hr. bThe sample entry phase step 1 and 2 should be extended for high protein loads, or for convenience, if the strips are to be run overnight. cThis voltage may not be reached within the suggested step duration. dNarrow intervals, 3.5–4.5, 4.0–5.0, 4.5–5.5, 5.0–6.0, and 5.5–6.7.

the tube can be held with fingers (60°C) and then slowly pipet the amount required to seal the IPG strip in place, avoiding introduction of bubbles. Only apply the minimum quantity of agarose sealing solution required to cover the IPG strip. Allow a minimum of 1 min for the agarose to cool and solidify. In any vertical gel system, the agarose sealing serves to prevent the IPG strip from moving or floating in the electrophoresis buffer. If precast Ettan DALT gels are used for the second-dimension electrophoresis, the agarose sealing also serves to block the narrow gap(s) between the gel edge(s) and the lateral spacer(s) to prevent leakage of the upper buffer.

17. Center the IPG strip between the plates on the surface of the second-dimension gel with the plastic backing against one of the glass plates. With a thin plastic ruler, gently push the IPG strip down so that the entire lower edge of the IPG strip is in contact with the top surface of the slab gel. Ensure that no air bubbles are trapped between the IPG strip and the slab gel surfaces or between the gel backing and the glass plate. 18. Perform second-dimension electrophoresis (see Basic Protocol 2). Two-Dimensional Gel Electrophoresis



REHYDRATION OF IPG STRIPS USING THE IMMOBILINE DRYSTRIP RESWELLING TRAY

SUPPORT PROTOCOL 2

The IPG strips are rehydrated in the Immobiline DryStrip Reswelling Tray if strip holders are in use for IEF. The Immobiline DryStrip Reswelling Tray has 12 independent reservoir slots that can each hold a single IPG strip up to 24 cm long. Separate slots allow the rehydration of individual IPG strips in a minimal volume of solution. Additional Materials (also see Alternate Protocol 4) Immobiline DryStrip Reswelling Tray (Amersham Biosciences) 1. Slide protective lid of reswelling tray completely off the tray and level tray by turning the leveling feet until the bubble in the spirit level is centered. Ensure that tray is clean and dry. 2. Pipet appropriate volume of rehydration solution into each slot as indicated in Table 10.4.3. Mix with sample solution (see Alternate Protocol 2, step 1) for rehydration loading (see Alternate Protocol 4, step 2 annotation). Deliver solution slowly at a central point in the slot. Remove any larger bubbles. To ensure complete fluid (and sample) uptake, do not apply excess rehydration solution.

3. Remove protective cover from IPG strip starting at the acidic (pointed) end. Position IPG strip with gel side down and pointed end of strip against the sloped end of the slot. Lower IPG strip onto solution. To coat the entire IPG strip, carefully lift and lower the strip and slide it back and forth along the surface of the solution. Do not trap bubbles under the IPG strip. 4. Overlay the IPG strip with 3 ml DryStrip Cover Fluid to minimize evaporation and prevent urea crystallization. 5. Slide lid onto reswelling tray and allow IPG strips to rehydrate at room temperature. A minimum of 10 hr is required for rehydration; overnight is recommended.


Acrylamide/bisacrylamide solutions 30% acrylamide/1.8% bisacrylamide (first dimension): 30 g acrylamide 1.8 g bisacrylamide H2O to 100 ml 30% acrylamide/0.8% bisacrylamide (second dimension): 300 g acrylamide 8.0 g bisacrylamide H2O to 1 liter Filter through 0.2- to 0.45-µm filter. Store in tightly capped amber bottles at 4°C. Discard after 30 days, as acrylamide gradually hydrolyzes to acrylic acid and ammonia. CAUTION: Acrylamide monomer is neurotoxic. Gloves should be worn while handling the solution, and the solution should not be pipetted by mouth. Wear a mask when weighing out the solid acrylamide. Analysis of Proteins


Supplement 67

Agarose sealing solution 100 ml SDS electrophoresis buffer (UNIT 10.2A) 0.5 g agarose (NuSieve GTG, Cambrex) 200 µl 1% bromphenol blue stock (see recipe; 0.002% w/v final) Combine all ingredients in a 500-ml Erlenmeyer flask. Swirl to disperse. Heat in a microwave oven on “low” or a stirring hotplate until the agarose is completely dissolved. Do not allow the solution to boil over. Dispense 2-ml aliquots into screw-cap tubes and store up to 1 week at 4°C. Agarose solution, 0.5% 0.25 g agarose (standard low-Mr; Bio-Rad) 50 ml reservoir buffer (see recipe) Heat in a boiling water bath to dissolve the agarose and keep in a boiling water bath. Prepare fresh each time. Agarose solution, 1% Prepare as for hot 0.5% agarose solution (see recipe), substituting 0.5 g agarose for 0.25 g agarose. Ammonium persulfate, 10% 10 g ammonium persulfate (Bio-Rad) H2O to 100 ml Store refrigerated ≤2 weeks Bromphenol blue stock solution, 1% 100 mg bromphenol blue (Sigma; 1% w/v final) 60 mg Tris base (50 mM final) Double-distilled H2O to 10 ml Prepare fresh Chromic acid cleaning solution Add a 25 ml bottle of Chromerge (Fisher) to a 9-lb bottle of concentrated sulfuric acid. Add ∼5 ml at a time and stir. CAUTION: This is very corrosive and toxic. Carefully read and observe package instructions.

Concentrated bromphenol blue 50% aqueous glycerol (v/v) 0.01 mg/ml bromphenol blue Prepare fresh Default sample solution 8 M urea 4% (w/v) CHAPS 60 mM DTT 2% (v/v) Pharmalyte 3–10 (Amersham Biosciences) 0.002% (w/v) bromphenol blue (add from 1% stock; see recipe) Store up to 6 months at −20°C


Equilibration buffer 3.75 g Tris base 25 ml glycerol 5.25 g SDS 333 mg dithiothreitol (DTT) Dissolve Tris base in H2O and adjust pH to 6.8 with 6 M HCl. Add other ingredients. Add H2O to 250 ml final volume. This buffer can be stored for up to 1 week at room temperature.



Gel buffer Dissolve 90.8 g Tris base in 300 ml H2O. Adjust to pH 8.6 with 6 M HCl. Add H2O to 500 ml. This buffer can be stored for several weeks in the refrigerator. Hydrophobic sample solution 7 M urea 2 M thiourea 4% (w/v) CHAPS 60 mM DTT 2% (v/v) Pharmalyte pH 3–10 (Amersham Biosciences) 0.002% (w/v) bromphenol blue (add from 1% stock; see recipe) Store up to 6 months at −20°C NaOH, 0.02 M Just before using, add 0.5 ml of 10 M NaOH to 250 ml freshly deaerated water. Store up to 1 month at room temperature. It is especially important to make 0.02 M NaOH with deaerated water.

Phosphoric acid, 0.085% Just before using, dilute 300 ml of 0.85% phosphoric acid to 3.0 liters with deaerated water. Store up to 1 month at room temperature. Rehydration stock solution 12 g urea (FW 60.06; 8 M final) 0.5 g CHAPS (2% w/v final) 50 µl 1% bromphenol blue stock (see recipe; 0.002% w/v final) Double distilled H2O to 25 ml Store stock with above components in 2.5-ml aliquots up to 1 month at −20°C Just prior to use add 7 mg DTT per 2.5-ml aliquot and IPG Buffer or Pharmalyte (same range as IPG strip) to 0.5% (for IPGphor apparatus) or 2% (for Multiphor II and Immobiline DryStrip kit). Use 125 µl IPG Buffer for a 0.5% concentration and 500 µl IPG Buffer for a 2% concentration. Use Pharmalyte 3–10 for Immobiline DryStrip 3–10 or 3–10 NL and Pharmalyte 5–8 for Immobiline DryStrip 4–7. For rehydration loading (see Alternate Protocol 4, step 2 annotation), also add sample to the 2.5 ml aliquot of rehydration solution just prior to use (see Table 10.4.2 for suitable sample loads). If necessary, the concentration of urea can be increased to 9 or 9.8 M. Other detergents (Triton X-100, NP-40, and other nonionic or zwitterionic detergents) can be used instead of CHAPS.

Reservoir buffer 15.0 g Tris base 72.0 g glycine 5.0 g SDS H2O to 5 liters For convenience, make up as a 10× stock or store the preweighed dry ingredients in packets for future use. The 10× stock can be stored for several weeks in the refrigerator. SDS equilibration buffer 10 ml 1.5 M Tris⋅Cl, pH 8.8 (APPENDIX 2; 50 mM final) 72.07 g urea (FW 60.06; 6 M final) 69 ml 87% (v/v) glycerol (30% v/v final) 4 g SDS (FW 288.38; 2% w/v final) continued



Supplement 67

400 µl 1% bromphenol blue stock (see recipe; 0.002% w/v final) Double-distilled H2O to 200 ml Store solution with above components at −20°C Just before use add: 100 mg DTT or 250 mg iodoacetamide SDS solubilization buffer 0.1 g 2-(N-cyclohexylamino)ethanesulfonic acid (CHES) 0.2 g SDS 0.1 g DTT 1.0 ml glycerol H2O to 10 ml total volume Store aliquots at −70°C Stacking gel buffer 15.0 g Tris base 1.0 g SDS Dissolve Tris base and SDS in 200 ml H2O. Adjust to pH 6.8 with 6 M HCl. Add H2O to 250 ml. The 5× stock can be stored for several weeks in the refrigerator. Tissue homogenization solution Prepare in acetone: 10% (w/v) trichloroacetic acid (TCA) 0.3% (w/v) DTT Prepare fresh Tissue sample solution 9 M urea 2% (w/v) CHAPS 1% (w/v) DTT 2% (v/v) Pharmalyte 3–10 Store up to 1 month at −20°C Urea solubilization buffer 5.4 g urea 0.4 ml NP-40 1.0 ml 20% (w/v) stock ampholyte (pH 9 to 11) 0.2 ml 2-ME (Kodak) H2O to 10 ml total volume; if pH is 1000 V). When the proteins have reached their final positions in the pH gradient, there is very little ionic movement in the system, resulting in a very low final current (typically below 1 mA). IEF of a given sample in a given electrophoresis system is generally performed for a constant number of volt-hours (V-hr), this unit being the integral of the volts applied over the time. IEF performed under denaturing conditions gives the highest resolution and the cleanest results. Complete denaturation and solubilization is achieved with a mixture of urea and detergent, ensuring that each protein is present in only one configuration and aggregation and intermolecular interaction is minimized. The original method for first-dimension IEF depended on carrier ampholyte-generated pH gradients in polyacrylamide gel rods in tubes (O’Farrell, 1975). Carrier ampholytes are small, soluble, amphoteric molecules with a high buffering capacity near their pI. Commercial carrier ampholyte mixtures are comprised of hundreds of individual polymeric species with pIs spanning a specific pH range. When a voltage is applied across a carrier ampholyte mixture, the carrier ampholytes with the highest pI (and the most negative charge) move toward the anode and the carrier ampholytes with the lowest pI (and the most positive charge) move toward the cathode. The other carrier ampholytes align themselves between the extremes, according to their pIs, and buffer their environment to the corresponding pHs. The result is a continuous pH gradient. Although the technique described in Basic Protocol 1 is commonly used in 2-D electrophoresis studies, it has several limitations that have prevented its more widespread application. Carrier ampholytes are mixed polymers that are not well characterized and suffer from batch-to-batch manufacturing variations. These variations can reduce the reproducibility of the first-dimension separation. Carrier ampholyte pH gradients are unstable and have a tendency to drift, usually toward the cathode, over time. Gradient drift adversely affects reproducibility by introducing a time variable. Gradient drift also causes a flattening of the pH gradient at each end, particularly above pH 9, rendering the 2-D technique less useful at pH

extremes. Finally, the soft polyacrylamide tube gels used in this procedure have low mechanical stability. The gel rods may stretch or break, affecting reproducibility. Results are often dependent on the skill of the operator. As a result of the limitations and problems with carrier ampholyte pH gradients, immobilized pH gradients (Alternate Protocol 4) were developed and Immobiline chemicals were introduced for the generation of this type of pH gradient. Westermeier (1983) and Görg et al. (2000) pioneered the development and use of IPG IEF for the first-dimension of 2-D electrophoresis. An immobilized pH gradient (IPG) is created by covalently incorporating a gradient of acidic and basic buffering groups into a polyacrylamide gel at the time it is cast. The buffers are composed of a single acidic or basic buffering group linked to an acrylamide monomer. The general structure of Immobiline reagents is: CH = CH–C–NH–R2O

where R = weakly acidic or basic buffering group. Immobilized pH gradients are formed using two solutions, one containing a relatively acidic mixture of acrylamide buffers and the other containing a relatively basic mixture. The concentrations of the various buffers in the two solutions define the range and shape of the pH gradient produced. Both solutions contain acrylamide monomers and catalysts. During polymerization, the acrylamide portion of the buffers copolymerize with the acrylamide and bisacrylamide monomers to form a polyacrylamide gel. To help in handling and performance, the IPG gel is cast onto a plastic backing. It is then washed to remove catalysts and unpolymerized monomers, which could otherwise modify proteins and interfere with separation. Finally the gel is dried and cut into 3-mm-wide strips. The resulting IPG strips can be rehydrated with a rehydration solution containing the necessary components for first-dimension IEF. In the Immobiline systems, IEF is performed horizontally on a flatbed electrophoresis unit. Because IEF requires high field strengths to obtain sharply focused bands, high voltages must be applied. A flatbed design is the most economical way to meet the necessary safety standards required to operate at such high voltages. It also allows for cooling and close temperature control, which can be effectively achieved on a horizontal ceramic cooling plate



connected to a thermostatic circulator or a Peltier cooling plate. The first-dimension separation is more reproducible because the covalently fixed gradient cannot drift.

Critical Parameters Two-dimensional PAGE is not difficult to perform but it is easy for problems to arise. Careful attention to detail is essential. Several steps in the procedure need to be optimized for particular samples. Sample solubilization, for example, may require some experimentation to determine the best solubilization buffer and the optimum ratio of buffer to sample. For most tissue samples, 1.5 to 2.0 ml of urea solubilization buffer per 100 mg of tissue will give good results; however, the ratio of tissue to buffer may have to be altered in some instances. If solubilized samples are stored frozen (−70°C), avoid thawing and refreezing. In general, keep the sample preparation strategy as simple as possible to avoid protein losses. Additional sample preparation steps may improve the quality of the final 2-D result, but at the possible expense of selective protein loss. The cells or tissue should be disrupted in such a way as to minimize proteolysis and other modes of protein degradation. Cell disruption should be done at as low a temperature as possible and with a minimum of heat generation. Cell disruption should ideally be carried out directly by placing the cells into a strongly denaturing solution containing protease inhibitors (UNIT 10.2). Preserve sample quality by preparing the sample just prior to IEF or storing samples in aliquots at −80°C. Do not expose samples to repeated thawing. Remove all particulate material by ultracentrifugation. Solid particles and lipids must be removed because they will block the gel pores. The sample solubilization solutions are in general optimized for either Basic Protocol 1 or Alternate Protocol 4. For Basic Protocol 1, either urea- or SDS-based sample solubilization may be used. Most proteins are soluble in either SDS or urea solubilization buffer. However, some proteins are more soluble in one or the other; therefore, it may be necessary to try both types of solubilization for a particular sample. In general, urea is more effective than SDS for tissue solubilization. Samples that have been solubilized in SDS can only be used with isoelectric focusing (IEF) gels containing urea and NP-40. Otherwise, the SDS remains associated with the proteins, giving them a strong negative charge and causing them to

migrate to the acidic end of the gel. The IPG technique outlined in Alternate Protocol 4 is particularly sensitive to residual SDS, which must be removed or reduced in concentration to 2% SDS. Samples in SDS solubilization buffer may be heated in a boiling water bath for 5 min to aid in protein solubilization. Heating is not always required and should never be done with samples in urea solubilization buffer. The quality of the first-dimension separation depends on the ampholytes used, voltage, volt-hours, and reagent quality. For good resolution over a broad pH range, blends of ampholytes from different suppliers generally give better results than broad-range ampholytes from a single supplier. However, ISO-DALTgrade Resolyte, pH 4 to 8, made by BDH Chemicals and sold in the United States by Hoefer, works very well. Optimum voltage and volt-hours must be determined experimentally. Once suitable conditions have been established, the same conditions must be used for each run to achieve reproducibility. When using homogeneous slab gels, the acrylamide concentration of the gel must be matched to the sample. Low-molecular-weight proteins can be resolved on high-percentage (18% to 20% acrylamide) gels while high-molecular-weight proteins must be run on low-percentage (7.5% to 12.5% acrylamide) gels. Gradient gels, such as 10% to 20% acrylamide gradients, are able to resolve proteins over a wide size range (250 to 10 kDa) on a single gel. Pouring gradient gels is more difficult than pouring homogeneous slabs, but with some practice the technique can be mastered (UNIT 10.2). One alternative is to buy precast gradient gels from a commercial supplier, such as Integrated Separation Systems. IEF in the Ettan IPGphor system is conducted at very high voltages (up to 8000 V) and very low currents (typically 1000 silver-stained protein spots distributed throughout the second-dimension gel, with only a few spots overlapping. Purified protein samples such as immunoprecipitation may have only one major protein and a few minor ones. The following checklists will help the beginner locate common problems associated with two-dimensional electrophoresis. General. (1) Were all of the reagents fresh? (2) Were all of the reagents properly prepared? (3) Were the samples properly solubilized? First dimension. Spots that are elongated horizontally (poorly focused), or a spot pattern in which most of the proteins are crowded into a narrowed zone along x axis of gel, should trigger the following questions: (1) Were samples ultracentrifuged just prior to loading? (2) Were desired number of volt-hours applied? (3) Were correct ampholytes used? (4) Was salt content of the sample low? Second dimension. Vertically elongated spots and spot patterns that are vertically compressed along the y axis of the gel should trigger the following questions: (1) Was the gel solution properly deaerated? (2) Were the tops of the gels flat and smooth? (3) Was the reservoir buffer at the desired temperature throughout the run? (4) Was the reservoir buffer fresh and the proper concentration?

Anticipated Results A two-dimensional electrophoretic separation of proteins similar to that shown in Figure 10.3.2 will be obtained. In this case, a 10% to 12% acrylamide second-dimension gel was used. Proteins between 10 and 250 kDa and pI 3.8 to 8.0 can be seen on this gel. Protein spots on a two-dimensional gel should be round or elliptical and separated from each other. A complex protein mixture such as mammalian tissue or whole cells should give a two-dimensional pattern of ≥1000 silver-stained spots distributed over most of the gel area. Fewer spots will be seen if stains less sensitive than silver stain are used (e.g., Coomassie blue; UNIT 10.6). Few of the protein spots should touch each other. The 2-D gel procedures described will be suitable for subsequent staining via protocols listed in UNIT 10.6 and Lilley (2002)., which allow both pre- and post-staining of samples for documentation, quantitation, and analysis by tandem mass spectrometry (MS/MS; UNIT 10.22) techniques.



Time Considerations Sample preparation takes 10 to 20 min/sample. First-dimension gels can be used as soon as 1 hr after casting or they can be kept for at least 24 hr before use. It is convenient to start the first-dimension gels in the afternoon, so they can run overnight and be ready for the second-dimension separation early the following day. Second-dimension gels can be poured up to a week before they are to be used, provided they are stored refrigerated in a humid, air-tight container to prevent drying. The second dimension run takes ∼5 hr. The two-dimensional gels will be ready to put in fixative or staining solution at the end of the day and can remain in that solution overnight. Usually, 3 working days are required to complete the entire procedure, including sample preparation and gel staining. Two-dimensional minigels take 4 to 5 hr.

Literature Cited Anderson, N.L. 1988. Two-Dimensional Electrophoresis Operation of the ISO-DALT (R) System. Large Scale Biology Press, Washington, D.C. Berkelman, T. and Stenstedt, T. 1998. 2-D Electrophoresis. Principles and Methods. Amersham Biosciences, San Francisco. Bjellqvist, B., Ek, K., Righetti, P.G., Gianazza, E., Görg, A., Westermeier, R., and Postel, W. 1982. Isoelectric focusing in immobilized pH gradients: Principle, methodology and some applications. J. Biochem. Biophys. Methods 6:317-339. Dunbar, B.S. 1987. Two-Dimensional Electrophoresis and Immunological Techniques. Plenum, New York. Görg, A., Obermaier, C., Boguth, G., Harder, A., Scheibe, B., Wildgruber, R., and Weiss, W. 2000. The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 21:1037-1053. Hirano, H. 1989. Microsequence analysis of winged bean seed proteins electroblotted from two-dimensional gel. J. Prot. Chem. 8:115-130. Lilley, K.S. 2002. Protein profiling using two-dimensional difference gel electrophoresis (2-D DIGE). In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, D.W. Speicher, and P.T. Wingfield, eds.) pp. 22.2-22.14. John Wiley & Sons, Hoboken, N.J. Ochs, D.C., McConkey, E.H., and Sammons, D.W. 1981. Silver stains for proteins in polyacrylamide gels: A comparison of six methods. Electrophoresis. 2:304-307.

O’Farrell, P.H. 1975. High-resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:4007-4021. Westermeier, R., Postel, W., Weser, J., and Görg, A. 1983. High-resolution two-dimensional electrophoresis with isoelectric focusing in immobilized pH gradients. J. Biochem. Biophys. Methods 8:321-330. Wilkins, M.R., Pasquali, C., Appel, R.D., Ou, K., Golaz, O., Sanchez, J.C., Yan, J.X., Gooley, A.A., Hughes, G., Humphery-Smith, I., Williams, K.L., Hochstrasser, D.F. 1996. From proteins to proteomes: Large scale protein identification by two-dimensional electrophoresis and amino acid analysis. Biotechnology (N.Y.) 14:61-65.

Key References Berkelman and Stenstedt, 1998. See above. Manual for the GE/Amersham family of 2-D electrophoresis equipment; details the background and protocols for use of immobilized pH gradient (IPG) strips in high-resolution analysis. Celis, J.E. and Bravo, R. (eds.) 1984. Two-Dimensional Gel Electrophoresis of Proteins. Academic Press, San Diego, Calif. Provides detailed information on two-dimensional gels and their applications. Dunbar, 1987. See above. A comprehensive guide to two-dimensional electrophoresis. It covers basic principles of electrophoresis, gives instructions for performing twodimensional electrophoresis and associated procedures such as protein detection, photography, and preparation of antibodies from proteins excised from two-dimensional gels. Garfin, D. and Heerdt, L. (eds.) 2000. 2D Electrophoresis for Proteomics. A Methods and Product Manual. Bio-Rad Laboratories, Inc., Hercules, Calif. Manual for the Bio-Rad family of 2-D electrophoresis equipment; details the background and protocols for use of immobilized pH gradient (IPG) strips in high-resolution analysis. Görg et al., 2000. See above. An important update on the latest developments in 2-D electrophoresis.

Contributed by Lonnie D. Adams The Upjohn Company Kalamazoo, Michigan Sean R. Gallagher UVP, Inc. Upland, California



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Overview of Digital Electrophoresis Analysis

UNIT 10.5

Gel electrophoresis has become a ubiquitous method in molecular biology for separating biomolecules. This prominence is the result of several factors, including the robustness, speed, and potential throughput of the technique. The results of this method are traditionally documented using silver-halide-based photography followed by manual interpretation. While this remains an excellent method for qualitative documentation of single-gel results, digital capture offers a number of significant advantages when documentation requires quantitation and sophisticated analysis. Digital images of gel electropherograms can be obtained rapidly using an image-capture device, and the images can be easily manipulated using image analysis software. REASONS FOR DIGITAL DOCUMENTATION AND ANALYSIS There are several reasons to consider digital documentation and analysis of electrophoresis results. These justifications can usually be categorized into issues of ease of handling, accuracy, reproducibility, and cost. Ease of Handling A major advantage of the digital revolution has been in storage and retrieval of information. Storage in notebooks and filing cabinets previously meant that searching for specific data or experiments was a tedious manual process. With digital information, modern search engines can quickly find specific information in a fraction of the time usually required for a manual search. Making backup copies of nondigital data can be difficult, expensive, and time consuming since it requires copying, retyping, or photographic reproduction. Copies of digital data can be generated more easily and at reduced costs. Manipulation of information is also easier when it is in a digital format. While the cut-and-paste analogy comes from physical documentation, it takes on a new perspective when applied digitally. Electrophoresis images can be resized, cropped, and inserted into reports. Data can be passed to spreadsheets and statistical packages for analysis and later insertion into notebooks and reports. These reports can be distributed via the Internet to colleagues throughout the world. A single individual can do all this in a few hours. Digital analysis also provides an easier method for handling the data when comparing large numbers of results or large numbers of separate experiments. Research that requires comparing the banding patterns on 1000 gels containing 50 lanes each can be an undertaking of heroic proportions if the analysis is performed manually. Database software can dramatically speed the analysis and handle the more mundane tasks, leaving the researcher free to interpret the data. Accuracy The human eye is an extremely versatile measuring instrument. It can handle light intensities covering a range of nearly nine orders of magnitude and is sensitive to a fairly wide spectrum of light (Russ, 1995). Yet the eye cannot accurately and reproducibly quantitate density and patterns, nor can it deal with large numbers of bands or spots. Accuracy of measurement is a primary reason for using digital analysis on electrophoretically separated proteins and nucleic acids. Two categories of accuracy are key to digital analysis: positional accuracy, which is important for mobility determinations such as molecular weight, and quantitative accuracy. Contributed by Scott Medberry, Sean Gallagher, and Butch Moomaw Current Protocols in Molecular Biology (2004) 10.5.1-10.5.25 Copyright © 2004 by John Wiley & Sons, Inc.



Positional accuracy is based on both resolution of the recording medium and measuring accuracy. Silver-halide-based recording has a theoretical resolution based on ∼2000 imaging elements (silver grains) per inch. Measurement traditionally occurs using a ruler, with an accuracy of ∼20 to 40 elements (50 to 100 elements per inch). In comparison, typical digital systems have 200 to 600 picture elements (pixels) per inch. The advantage that digital systems have is in measuring accuracy, which can occur at the level of a single imaging element. Quantitative accuracy is also an issue. The amount of material represented by a band or spot is difficult to determine accurately from an image of a gel unless it is a digital image. On a digital image, the amount present is directly correlated with the derived volume of the band or spot—the volume is calculated using the intensity values of the pixels within the object. Reproducibility Any technique or measurement is only as good as its ability to be faithfully replicated. With software-defined routines, measurements are performed in the same manner every time. Allowing the computer to do repetitive tasks and complicated calculations minimizes the chance for individual errors. This does not imply that such measurements are correct, just that they are reproducible. An incorrect routine or algorithm can also invalidate data. Cost A consideration when evaluating any laboratory method is cost. Digital electrophoresis analysis equipment can be expensive. In many cases however, it offers the only method for achieving acceptable analysis performance. In other cases, equal performance can be achieved using silver-halide technology. However, traditional photography can also be expensive when the costs of consumable supplies such as film and developers and other expensive requirements such as developing tanks and dark rooms are included. Often, digital methods can be a good choice when all costs are considered. KEY TERMS FOR IMAGING There are several specialized terms encountered during digital image analysis. The most commonly encountered are contrast, brightness, gamma, saturation, resolution, and dynamic range. They describe controls on how the light detectors report a range of light intensities. Below is a brief description of each. Contrast


Contrast describes the slope of the light intensity response curve. An increase in the contrast increases the slope of the curve. The result is a more detailed display over a narrowed range of intensities with less detail in the remaining portions of the intensity range. This is depicted in Figure 10.5.1A and 10.5.1B, where a normal, unadjusted image and a contrast-adjusted image are displayed, respectively. The contrast was increased on midrange intensity values in Figure 10.5.1B to highlight band intensity differences at the expense of background information. Images with a narrow range of informative intensities can benefit from increasing the contrast since it effectively increases the scale and improves detection of minor differences in intensity. Contrast settings should be lowered if information is being lost outside of the contrast range. For example, in Figure 10.5.1B, loss of background information between peaks indicates that this image should not be used for quantitation.



A

normal

Output

100

0 0

Input

B

100 200 300 400 500

contrast

Output

100

0 0 100 200 300 400 500

Input

C

brightness

Output

100

0 0 100 200 300 400 500

Input

D

gamma

Output

100

0 0

Input

E

100 200 300 400 500

saturation 100

0 0

100 200 300 400 500

Figure 10.5.1 Examples of how altering image capture settings affects the image and the analysis. The graph on the left displays the light intensity response curve used for image capture while the image and resulting lane profile on the right display how the setting affects the image. The lane profile displays pixel position versus normalized pixel intensity (A). In this case, the output has not been altered, giving a straight line with a slope of 1 on the response curve. (B) The image acquisition was adjusted to increase the contrast of the displayed image. Although useful for images with a narrow range of informative intensity values, increasing the contrast can lead to a loss of low and high values. (C) Decreasing the brightness reduces peak values but also leads to a loss of the weak bands and original background. (D) Gamma adjusts raw data to appear more visually accurate. Note that this leads to a loss of fidelity between the adjusted image and the original. (E) Saturation indicates that the detector is reporting its maximum value or that the dynamic range for the visualization method has been exceeded.



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Brightness While brightness can have many different definitions, only one will be considered here. Brightness shifts the light intensity response curve without changing its slope as is shown in Figure 10.5.1C. Another name for brightness is black level since it is commonly used to control the number of black picture elements (pixels) in an image. Incorrect brightness levels can lead to either high background and potential image saturation or, as is illustrated in Figure 10.5.1C, to loss of background information entirely and partial loss of band information. Gamma Nonlinear corrections are often applied to images to compensate for how the eye perceives changes in intensity, how display devices reproduce images, or both. The most common correction is an exponential one with the exponent in the equation termed the gamma. A typical gamma value is 0.45 to 0.50 for camera-based systems and is illustrated in Figure 10.5.1D. This is a compromise value that compensates for the 2.2 to 2.5 gamma present in most video monitors and the print dynamics of most printers. Since it is a nonlinear correction, special care must be taken if quantitation is desired. Unless otherwise directed by the manufacturer, gamma values other than 1.0 should be avoided when quantitating. More information on gamma correction can be found on Poynton’s Gamma FAQ (http://www.inforamp.net/∼poynton/Poynton-color.html). Dynamic Range Dynamic range describes the breadth of intensity values detectable by a system and is usually expressed in logarithmic terms such as orders of magnitude, decades, or optical density (OD) units. A large dynamic range is important when trying to quantitate over a wide range of concentrations. The most accurate quantitation occurs in the linear part of the dynamic range, which is usually not the complete dynamic range of the system. An additional consideration is the dynamic range of the visualization method. Many popular visualization methods have linear dynamic ranges of 1 to 2.5 orders of magnitude. An imaging system with greater dynamic range analyzing the results of such a visualization method will not improve the dynamic range. Saturation Saturation occurs when a detector or visualization method receives input levels beyond the maximum end of the dynamic range. This results in a loss of detail and quantitative information from those data points that are saturated. For fluorescent and chemiluminescent samples, reduction in the sampling time can sometimes correct saturation problems. Optical density-based visualization techniques can also generate saturated images, as is illustrated in Figure 10.5.1E; this can sometimes be avoided with longer sampling times or increased detection source intensities. More often, it will be necessary to perform another electrophoresis with more dilute samples or to alter the visualization process to generate a less optically dense material. Resolution Resolution is the ability of a system to distinguish between two closely placed or similar objects. Three types of resolution are important for analysis—spatial resolution, intensity resolution, and technique-dependent resolution. Overview of Digital Electrophoresis Analysis

Spatial resolution is the ability to detect two closely placed objects in one-, two-, or three-dimensional space. It is most accurately described as the closest distance two objects can be placed and still be detected as separate objects. In practice, it is often defined



high-resolution peaks

medium-resolution peaks

42 µm

Intensity

168 µm

840 µm

0

100 Pixel position

Figure 10.5.2 The effect of spatial resolution on the ability to detect closely spaced objects. Whole-cell protein lysates from E. coli were separated using SDS-PAGE and visualized with Coomassie blue staining. An image was captured at 42 µm (600 dpi), 168 µm (150 dpi), and 840 µm (30 dpi) from a segment of the lane, and a lane profile was generated for each image. The lane profiles have offset intensities to allow for comparison. Only major bands can be detected with the low-resolution image (840 µm); at higher resolutions more bands are detectable.

nominally in terms of the number of detectors per unit area such as dots per inch (dpi) or the number of detectors present in total or in each dimension such as 512 × 512 (262,144 total detectors). Actual resolution is less than half the nominal resolution due to the need for two detectors for every resoluble object (one for the object and one for the separation space) and the effects of optical resolution. Figure 10.5.2 demonstrates how spatial resolution can affect detection of objects. The 42-µm resolution image allows detection of closely spaced bands, the 168-µm resolution image detects fewer bands, and the 840-µm resolution image detects only major bands. For instruments with on-line detection systems, a pseudo spatial resolution is often reported in units of time from the start of the separation or the time interval between two objects crossing the detection path. Intensity resolution is the ability to identify small changes in intensity. It is a function of both the dynamic range of a detector and the number of potential values that detector can report. Greater dynamic range decreases the intensity resolution of a given detector. The number of potential values a detector reports is described by its bit depth. An 8-bit detector can report 256 (28) different possible values, while a 12-bit detector can report 4,096 (212) values, and a 16-bit detector can report 65,536 (216) values. The higher the bit depth, the greater the intensity resolution. Technique-dependent resolution directly affects the spatial and intensity resolution. Electrophoretic separation techniques that generate overlapping objects or that have object separation distances shorter than the spatial resolution will fail to provide reliable data. Many factors, including the amount of sample loaded, gel pore size, buffer constituents, and electrophoresis field strengths, can dramatically affect separation and resolution of biomolecules. Likewise, detection methods that can only generate a small range of discrete intensity values will not benefit from systems with improved intensity resolution.



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IMAGE CAPTURE Systems Several components make up a digital imaging system (Fig. 10.5.3). These include: (1) a light source for illuminating the sample, either for nonfluorescent white-light imaging or for fluorescent excitation of a sample stained with a fluorescent dye; (2) a filter on the lens to act as a contrast or signal-to-noise (S/N) enhancer for nonfluorescent applications, or an emission wavelength isolation filter for fluorescent detection applications; (3) a fixed or zoom lens; (4) the CCD camera for acquiring the image; and (5) the computer and software for analysis. Filters used for digital imaging have a number of functions depending on the application (Table 10.5.1). For nonfluorescent imaging, colored glass or wavelength-isolation filters are typically used to enhance the contrast and S/N. For example, a Coomassie blue–stained protein gel shows enhanced contrast over background illumination with use of an orange filter. This is very useful for acquiring faintly stained bands on a gel. With fluorescent applications, the lens filter acts to isolate and detect the emission wavelength from the fluorescent sample or gel. These filters are typically very accurate band-pass interference filters. Devices Capturing digital images involves a detection beam or source, a sensor for that beam or source, and some method of assembling a two-dimensional image from the data generated. Most systems use a light source for detection. The light wavelengths used range from ultraviolet (UV) to infrared (IR) and can be broad spectrum or narrow wavelength.


Figure 10.5.3 Typical light-tight desktop system for UV/visible colorimetric, chemiluminescent/bioluminescent, and fluorescent imaging.



Table 10.5.1

Filters for Digital Imaging Applications

Application

Desired effect

Typical filters

Source

Optical protein stains (e.g., Coomassie Blue)

Contrast enhancement

Edmund Scientific, Omega Optical, Schott Glass

Fluorescent nucleic acid and protein stains, green fluorescent protein

Signal to noise (S/N) enhancement Excitation and emission wavelength isolation

Colored glass Broad band-pass Short-pass Long-pass Interference filters Broad and short band-pass Short-pass Long pass

Edmund Scientific, Omega Optical, Schott Glass

Broad-spectrum detection is more versatile since it can often be used for more than one detection wavelength. A typical UV light table will emit an broad excitation light that peaks at 302 nm, useful for excitation of many fluorescent stains. However, when compared to narrow-wavelength sources such as lasers, broad-spectrum detection suffers from reduced sensitivity and reduced dynamic range. While many types of light sensors have been used, including charge-coupled devices (CCDs), charge-injection devices (CIDs), and photon multiplier tubes (PMTs), technology advances in CCDs have led to their dominance. CCDs are semiconductor imaging devices that convert photons into charge. This charge is then read and converted into a digital format via an analog-to-digital converter (ADC). The method of image assembly depends on the light source and detector geometry. One method is to capture the image all at once using a two-dimensionally arrayed CCD detector similar to the detectors found in digital and video cameras (see Advances in CCD Technology). Typically a camera-type sensor is paired with a light source that evenly illuminates the sample. This same sensor is often used with fluorescent and chemiluminescent detection methods as its ability to detect light continuously over the entire sample reduces image capture times. Another method of image assembly is to capture the image a line at a time. This typically involves a linearly arrayed CCD scanning slowly across the sample in conjunction with the detection beam of light. The data from each line are then compiled into a composite image. Spatial resolution in this method can be significantly better on large-format samples compared to the resolution of a camera-based system. This method is also advantageous when OD-based detection is used, since the more focused light beam is usually of higher intensity and can penetrate denser material. A third method of image assembly is to use a point light source and single-element detector on each point on a sample. The image is then compiled from each point sampled. This method is slower than the others but can offer extremely high resolution and sensitivity. A fourth commonly encountered method is that of generating a pseudoimage of electrophoresis results through the use of a finish line type of detection system. This is comprised of a light source positioned at the bottom of the gel (i.e., the end opposite of the site of sample loading) and light detectors positioned next to each lane to detect the transmitted light or emitted fluorescence. A lane trace is generated using time on the x-axis and light intensity on the y-axis. The pseudoimage is then generated from this data (Sutherland et al., 1987). Capture Process Prior to image capture, electrophoretic separation and any visualization steps are performed. To calibrate the separation process, standards are usually run at the edges of the gel and often at internal positions. If quantitation of specific proteins or nucleic acids is to be performed, a dilution series of standards with similar properties to the experimental



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Table 10.5.2 Methodsa

Compatability of Popular Image Capture Devices with Common Visualization

Image capture device Visualization method Optical densityb Fluorescence Chemiluminescence Radioactivity

Silver-halide photography

CCD camera

Desktop scanner

Storage phosphor

Fluorescent scanner

+

+

++

−

−

+ + +

+ ++ −

− − −

− ± ++

++ − −

aThe device with the highest sensitivity and greatest dynamic range for a visualization method is marked with a ++, other devices that can detect this visualization method are indicated with a +, and devices that are not suitable for a visualization method are indicated with a −. A ± indicates that only some devices of this type can be used with this visualization method. bOptical density methods include Coomassie blue staining.

samples should also be included. After separation, the protein or nucleic acid is visualized if necessary. Visualization can include binding of a fluorochrome or chromophore (e.g., Coomassie blue), precipitation of metal ions (e.g., copper, silver, or gold), enzymatic reactions, and exposure of film or phosphor screens to radiant sources. These methods can be grouped based on the type of detection into optical density, fluorescence, chemiluminescence, and radioactivity. The suitability of popular detection devices with these methods is described in Table 10.5.2. Once visualization has occurred, image capture consists of the following steps: previewing the image while adjusting capture parameters, capturing the image, and saving the image for later analysis.


During the preview process, capture parameters are optimized for data content and for ease and rapidity of later processing steps. Typically, the first step is to place the sample so that when the image is captured, the rectangular edges of the gel are horizontal and vertical on the monitor and any lanes are either horizontal or vertical. Since band and spot detection will be much easier if the image is properly oriented, this eliminates the need to later rotate the image digitally. Image rotation is time consuming and can result in spatial linearity errors (a change in the size and shape of objects in image) resulting from rectangular image-capture device geometries. The next step for camera-based systems is to adjust magnification and to focus the sample image. For thicker samples, it might be necessary to reduce the aperture on camera-based systems to get a sufficient depth-of-field to focus the entire sample. Often at this point image imperfections—e.g., dust, liquid, or other foreign objects that will detract from later analysis—are detected, and they need to be removed. Next, image intensity is set. Within the area of interest on the image, band or spot peaks should have values less than the maximum saturated value, and the background should have nonzero values. This is usually accomplished through adjustment of the light-source intensity or the sensor signal integration. If the device allows precapture optimization of other parameters such as spatial resolution, contrast, brightness, gain, or gamma, they are adjusted next. Note that this only applies to controls that affect the response of the sensor or processing of the image prior to a data reduction step and not to controls that affect the image at later stages. The latter process can enhance visualization of specific features but is best left to adjustments in look-up tables (LUTs) in later analysis steps than during image capture since there is a risk of data loss during postacquisition image processing. LUTs are indexed palettes or tables where each index value corresponds to color or gray-scale intensity values present in an image. Many image analysis programs alter LUTs instead of image values directly since it is both faster and does not change the original image data.



Once all the capture parameters are optimized, the image capture process is initiated. This might take less than a second for images captured with camera-based detectors and up to hours for scanning single-point detectors. When the image has been captured, it should be carefully examined for content. It should fully capture the area of interest and the parameters should have been set so that all necessary information is detectable. Furthermore, it should be in a form that will allow for easy analysis. Extra time spent on optimizing the capture parameters will often result in a reduction in total image analysis time and in an increase in data quality. When the best possible image has been captured, it often contains information outside the area of interest. While this is unlikely to cause problems with later analysis, it is often advantageous to crop the image so that the only portion that is saved contains the area of interest. This reduces the amount of disk space necessary to store the image, and the image usually will load and analyze faster with the analysis software. The last step in image capture is to save the image. Several options are available at this point, including choosing the location at which to save the image, what file type or format to use, and whether to use some form of compression. The location where the image is saved is not as trivial a question as it might seem if the image will need to be transferred to another computer at some point. File sizes can easily exceed 15 megabytes on high-resolution images. This is a manageable size for hard drives but exceeds current floppy drive sizes by an order of magnitude. There are software utilities available that will subdivide files into disk-size chunks and then reassemble them at the next computer, but this is an inconvenient and slow method. If the computer used to help capture the image is connected to a network, the image files can easily be transferred this way or potentially saved on a central server. Alternatively, several types of high-capacity removable media are available (e.g., Zip, or Jazz). This usually requires the installation of additional hardware onto two or more computers but does make backing up data easier. Since image files can be very large, compression techniques are sometimes used to reduce disk space requirements. Compression algorithms use several methods, typically by replacing frequent or repetitive values or patterns with smaller reference values and by replacing pixel values with the smaller difference values describing the change in adjacent pixels. When the file is later decompressed, the compressed values are then replaced with the original information. Not all images compress equally, with simple images containing mostly repetitive motifs compressible by ≥90%, while complex images will benefit much less from compression. Because compression is a much slower method of saving files and not every file will benefit from it, compression is not used to save all files. Several different forms of compression are available but are separable into two main classes, lossless and lossy. Lossless methods faithfully and completely restore the image when it is decompressed (no loss of data) but offer only moderate file compression. Compression values range from ∼10% to 90%, depending on the image. Examples of lossless compression include Huffman coding (Huffman, 1952), RLE (Run Length Encoding), and LZW (Lempel, Ziv, and Welch; Welch, 1984). In comparison, lossy methods such as JPEG (Joint Photographic Experts Group), MPEG (Moving Picture Experts Group), or fractal compression schemes can reach compression values of ≥98% (Russ, 1995). The trade-off is that not all information from the original file is recovered during decompression. Lossy compression is sometimes necessary for applications with extremely large image files such as real time video capture, but it usually represents an unacceptable loss of data if used with electrophoresis image capture. Many different file types have been developed to store digital images. Some of these file types are proprietary or are hardware specific. For example, PICT is a Macintosh format



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and BMP is a PC-compatible format (see descriptions of both systems, below). Each file type has its own structure. Some types do not allow compression, for others it is optional, and for some it is mandatory. They vary in the types of images they support, particularly in the number of colors or gray levels. Below is a brief description of a few of the more prevalent file types. TIFF (Tagged-Image File Format) is one of the most commonly used formats. It is particularly versatile since it is an open format that can be modified for specific applications. One reason for its versatility is the ability to attach or tag data to the image. The tags can include information such as optical density calibration, resolution, experimenter, date of capture, and any other data that the application software supports. TIFF images can be monochrome, 4-, 8-, or 16-bit gray scale, or one of many color-image formats. Compression is optional, with LZW, RLE, and JPEG often supported (Russ, 1995). Since TIFF is supported by both Macintosh and PC computers, it is a good choice for multiple-platform environments. The versatility of TIFF can also be a weakness. Since there are many different tagging schemes and since not all programs support all possible compression and color schemes, it is sometimes not possible for one program to access the information in a TIFF file generated by a different program. GIF (Graphics Interchange Format) is a file format that is widely encountered on the Internet due to its compactness and standardization. Its compactness is attributable to a mandatory modified LZW compression. Another feature of GIF is the use of a LUT to index the values in the image. One interesting ability of GIF is that it supports storing multiple images within a single file. This can offer some advantages for applications such as time-lapse image capture. A GIF image can contain no more than 256 individual color or gray levels and does not support intensity resolutions higher than 8 bit. In addition, since the image is implemented as a LUT, it also is not a true gray-scale image. Due to these limitations and others, alternative formats such as PNG have been developed to replace GIF. PICT is a file format and graphics metafile language (it contains commands that can be played back to recreate an image) designed for the Apple Macintosh. It can contain both bitmap images and vector-based objects such as polygons and fonts. It supports a ≤256-gray-level LUT, and monochrome images can be RLE compressed. Because it only offers a 256-gray-level LUT, it has the same weaknesses that GIF does with true gray-scale and high-intensity-resolution images. In addition, any vector objects in the image are difficult to translate on a PC since they are designed to be interpreted by Macintosh QuickDraw routines. BMP is the native bitmap file format present on Windows-based PCs. It supports 2-, 16-, 256-, or 16-million level images. With images of ≤256 gray levels, it implements a LUT, while the highest-resolution image is implemented directly. RLE compression is optional for 16- and 256-gray-level images. Since compression is prohibited on 16 million-graylevel images and there is no intermediate level supported beyond 256 levels, BMP is not a good choice for images with high-intensity resolution requirements. ADVANCES IN CCD TECHNOLOGY


Currently there is a growing emphasis on using CCDs to capture the intensity or spectral data produced in electrophoretic assays. A CCD offers great flexibility in data capture, since it provides both location and intensity information at the same time. Once the data are captured, they must be understood and interpreted. The most difficult component in this scheme is the experimental design necessary to meet the requirements of this type of detector, or the choice of the appropriate detector for a particular experimental design.



Then, it is necessary to know the influence of the detector on the data set. To make good choices in the experimental setup and proper interpretation of the data, a thorough and intimate knowledge of the detector is essential. CCD detectors operate on a simple principle. When a photon of acceptable wavelength interacts with the CCD substrate (a silicon crystal lattice chip), the energy of the photon generates an electron and a “hole pair.” This effect produces a negative charge in that region of lattice, and the accumulation of these electrons and hole pairs is directly proportional to the number of photons that successfully interact with the silicon lattice. Short-wavelength photons interact near the surface of the lattice, and progressively longer wavelengths interact at progressively deeper depths of the lattice. A CCD, substrate is electronically segregated into discrete regions called pixels by creating electrical currents and gates at the surface of a silicon wafer. This is the same process used in other semiconductor manufacturing processes to produce memory chips and other microelectronic devices. When appropriate voltages and timing signals are passed through the microcircuits, a CCD chip becomes divided into pixels and is able to convert incoming photons into an electron signal, create a two-dimensional map of this signal, and then transfer this signal to an amplifier. Each pixel in the CCD has an X and Y value, measured in microns plus a depth in the substrate. These values determine a volume of the pixel and how many electron hole pairs can be generated and segregated in each pixel. This capacity is referred to as the full-well capacity or saturation level of a pixel. When the CCD chip is used to create a camera, the amplified signal is passed to a monitor or recording device for viewing or documentation. If the camera is designated as a digital camera, the analog signal is passed to an analog-to-digital converter (ADC) before being sent to a computer. A key point here is to remember that all CCDs are analog devices. Cameras may be analog or digital depending on how the camera is designed, but the CCD is always analog. Why Use CCDs? CCD technology is currently at the point where the conversion efficiency of photons to electron hole pairs can exceed 90%. This value is called the quantum efficiency of the CCD, and is the key factor in generating the signal in the detector (Fig. 10.5.4). On the opposite side of the signal level is how much noise is generated in the detector and the camera during the signal collection and transfer process. This relationship between the amount of signal collected and the amount of noise generated is called the signal-to-noise ratio, and is one of the key terms necessary to understand the choice of detector and choice of experimental design. In a CCD, this signal-to-noise ratio has reached the point where in most cases the limiting factor of the data accuracy is the signal itself. Few other detectors offer an equivalent signal-to-noise ratio at the data collection speeds of a CCD when a two-dimensional data array is required. CCD imagers range from relatively low cost for simple white light and fluorescent imaging to more sophisticated and costly cooled cameras for chemiluminescent and in vivo imaging (Table 10.5.3). Glossary of CCD Terms CCD Charge-coupled device, the basis of operation for each detector. A CCD is a silicon-based detector in which an electronic charge (an electron and a hole pair) is produced in the silicon lattice by a photon. Pixel An electronically segregated area in the silicon lattice produced by microelectronic circuits on the surface of the silicon substrate, or chip.



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80

Quantum efficiency (%)

70

ER-150 ICX285

60 50 40 30 20 10 0 200

300

400

500

700 600 800 Wavelength (nm)

900

1000

1100

1200

Figure 10.5.4 Quantum efficiency of an extended range (ER-150; solid line) and standard (ICX285; broken line) CCD.

Photon The minimal unit of light, an electromagnetic energy with a particular wavelength (frequency) that interacts with the silicon lattice in a CCD to produce an electron and a hole pair. Full-well capacity The maximum number of electron hole pairs that can be contained in a pixel. Also defined as the upper limit of detection, the point at which additional signals can no longer be detected, and the saturation level of a detector. Noise A statistical description of the fluctuation in an otherwise stable current. Generally, noise is expressed as an RMS (root mean square) value. Noise terms are added or subtracted in quadrature; it is necessary to square each noise term, add or subtract the terms, then take the square root of the result. Shot noise See Signal noise Signal noise The fluctuation in the number of photons in a given amount of time from a stabilized, uniform light source. Under these conditions, the signal noise should be equal to the square root of the signal level, expressed in electrons. Signal noise is not a property of a CCD or a camera, only the signal itself. Readout noise The fluctuation in a nominal zero level signal as it is transferred through the serial register and readout amplifier of a CCD. This noise is related to the speed of the readout (pixel clock speed) and the related circuit design. In modern cameras this is almost always the factor that limits the low-light detection capability.


Dark current The number of electrons that appear in a pixel when no light is falling on the detector. Dark current is caused by migration of electrons from areas outside the sensing area of the CCD into the sensitive area and is reduced by 50% with every 8°C reduction of the CCD temperature. Dark current is expressed as the number of electrons (e)/pixel (p)/sec (s). In modern cameras, this is generally so small that it can be disregarded at short exposure times. Example: Specification for Dark Current = 25e/p/s = 25 electrons per pixel per sec.



Table 10.5.3

CCD Camera Specifications

Price range

Low-end

Mid-range

High-end

Premium

Model/manufacturer

C9260-001, Hamamatsu

C8484-51-03G, Hamamatsu

ORCA AG, Hamamatsu

ORCA 2ER, Hamamatsu

Pixel array/size

1344 × 1024/6.45 × 6.45 µm

1344 × 1024/6.45 × 6.45 µm

1344 × 1024/6.45 × 6.45 µm

1344 × 1024/6.45 × 6.45 µm

CCD size

2/3 in.

2/3 in.

2/3 in.

2/3 in.

Readout noise / frame rate

20 electrons/4 Hz

6 electrons/8.9 Hz

6 electrons/8.9 Hz

6 electrons at 5.6 Hz; 3 electrons at 0.8 Hz

Dark current (electrons/pixel/sec) at indicated temperature

8 electrons/pixel/sec at 20°C

0.1 electrons/pixel/ sec at –10°C

0.03 electrons/pixel/sec at 30°C

0.0045 electrons/pixel/sec at –60°C

Camera noise

21.5 electrons

6 electrons

6 electrons

6 electrons/3 electrons

Full-well capacity (1×1)/(2×2)

18,000/23,000 electrons

15,000 electrons

18,000 electrons

18,000 electrons/40,500 electrons

Dynamic range (1×1)/(2×2) 832:1/1070:1

2500:01:00

3000:01:00

3000:1/13,500;1

Bit depth/gray levels

12 bit/4096

12 bit/4096

12 bit/4096

12 bit/4096 and 14 bit/16,384

Digitizer gain (1×1)/(2×2)

4.3 electrons/5.6 electrons

3.7 electrons

4.4 electrons

4.4 electrons and 2.5 electrons

Maximum quantum efficiency (QE) at indicated wavelength Binning

50% at 525 nm

72% at 550 to 570 nm

72% at 550 to 570 nm

72% at 550 to 570 nm

1 × 1, 2 × 2, 4 × 4

1 × 1, 2 × 2, 4 × 4, 8×8

1 × 1, 2 × 2, 4 × 4, 8×8

1 × 1, 2 × 2, 4 × 4, 8×8

Dark noise The amount of fluctuation in the dark current. This should equal the square root of the dark current. Example 1: dark current (D) = 25 e/p/s; exposure time 1 sec: dark noise ( N D ) = 25e = 5e

Example 2: dark current (D) = 25 e/p/s; exposure time 0.5 sec: dark noise ( N D )

25e = 12.5e = 3.53e 0.5

=

Camera noise The sum of the readout noise and the dark noise. This term refers only to a camera, not to an image. Example: camera with read noise = 10e and dark noise = 5e: camera noise = CN =

( N R )2 + ( N D )2

=

(10e )2 + (5e )2

= 100e + 25e = 11.18e Analysis of Proteins


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Readout noise The fluctuation in a nominal zero level signal as it is transferred through the serial register and readout amplifier of a CCD. Often referred to as simply “read noise,” the noise is related to the speed of the readout (pixel clock speed) and the related circuit design. In modern cameras, this is almost always the factor that limits the low-light-detection capability. Total noise The sum of the camera noise and the signal noise. This term only applies to an image.

( N R )2 + ( N D )2 + ( NS )2

total noise =

where NR is read noise, ND is dark noise and NS is signal noise. Example: 1-sec exposure, camera with 10e read noise, 5 e/p/s dark current, average gray value in one region of the image equals 3800, digitizer gain equals 4 electrons per count. To find the camera noise, the signal noise, and the total noise in the image: 1. read noise (NR) = 10e (from camera data sheet or measured); 2. dark noise (ND) = Ds = 25e = 5e

(from camera data sheet or measured); 3. camera noise (NC) =

( RN )2 + ( DN )2

=

(10e )2 + (5e )2 =

100e + 25e = 11.18e

4. signal noise (NS) = S = 3800 × 4e = 15, 200e = 123.2e

5. total noise (NT) =

( N R )2 + ( N D )2 + ( NS )2

=

(10e )2 + (5e )2 + (123e )2

= 15,325e = 123.7e

Please note the difference the camera noise made in this image, only 0.5 electrons! Dynamic range A calculation of the biggest difference in brightness detectable by a device. To obtain this value, divide the Full-Well Capacity (FWC) of a detector by the camera noise to get this range as a ratio. Example: Camera with 20,000 electrons FWC and 10 electrons camera noise: 20,000e/10e = 2000:1 dynamic range. For dynamic range in decibels (dB), take the log of 2000 = 3.3 × 20 to get 66 dB. Bit depth The number of gray levels that the digitizer will create from the maximum signal level of a device. Bits refers to the exponent of 2 used to create the number of gray levels. Overview of Digital Electrophoresis Analysis



Examples: 10-bit = 210 = 1024 gray levels, bit = 212 = 4096 gray levels. A 10-bit digitzer will divide the maximum signal (full-well capacity of a CCD) by 1024 gray levels to produce 1024 gray levels in the image. Bit depth is determined by the digitizer in a digital camera; not the CCD. See example under “digitizer gain,” below. Digitizer gain/digitizer count The number of electrons in the signal used to represent each gray level in an image. This is an essential number for making quantitative calculations and comparisons of digital camera data. A close approximation may be determined by dividing the full-well capacity by the number of gray levels in the digitizer. Camera data sheets should include the exact number. Example: Camera with FWC = 20,000 electrons; what is the digitizer gain? 10-bit = 210 = 1024 gray levels = 20,000 (electrons)/1024 (gray levels) = 19.5e 12-bit = 212 = 4096 gray levels = 20,000 (electrons)/4096 (gray levels) = 4.9e 14-bit = 214 = 16384 gray levels = 20,000 (electrons)/16384 (gray levels) = 1.2e Always compare data and results by number of electrons in the signal, not by gray level, since different cameras have different digitizer gains. In many cases, larger gray values may actually represent smaller data values when making comparisons or calculating the signal noise in an image. Digitizer offset A value established in the analog-to-digital converter that represents the average zero value in an analog signal. Since a digitizer only has positive numbers but an analog signal will fluctuate both above and below an average value, a digitizer must have some positive values that can represent the negative portion of the analog signal to prevent clipping of the full data set. This range of positive values in the digitizer is called the digitizer offset. A digitizer with an offset of 200 means that the values from 1 to 199 are used to represent values less than zero in the analog signal. Gray level 200 represents the zero level signal and values above 200 represent signal levels above zero. List of Typical Camera Features and Benefits CCD camera features and their advantages and disadvantages that should be considered are listed below. Number of pixels A CCD camera has more spatial resolution when compared to another camera with the same size pixel, but fewer pixels. This means there is less sensitivity when compared to another camera with fewer, but larger, pixels. In addition, a camera has a potentially slower frame rate when compared to another camera with fewer pixels. Size of pixels Smaller pixels provide higher spatial resolution. Cameras with larger pixels provide larger full-well capacity, bigger dynamic range, and better accuracy, and collect signal faster than small pixels. Size of detector Larger detector dimensions mean a larger field of view, but require more expensive optics.



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Cooling temperature Temperature alone is not meaningful. The dark current at that temperature must be determined. Cooling reduces the dark current. Cooling temperature may not be relevant for fast exposures Readout noise specification Lower values improve low-light sensitivity, but slower readout speeds may be required to achieve lower values. Always be sure the readout noise specification is valid at the desired frame rate. Quantum efficiency (QE) value Higher QE is better if it is in the same wavelength range as the signal, but is a problem if it is in a range other than the signal—i.e., near-infrared! QE value must be considered along with other factors, such as camera noise. Dark current Lower dark current is better for dim signals and for dynamic range calculations. The cost of necessary cooling may not be worthwhile for very short exposures. Dark noise Dark noise is the square root of the dark current. It is important to use the dark noise to calculate camera noise, not dark current. Major Considerations in Camera Selection Detectability Detectability is determined by the camera noise and QE as illustrated in the Figure 10.5.3. Once the camera noise is minimized, the largest gain in this area is obtained through greater QE. Accuracy of a single measurement Accuracy is determined by the noise of the signal itself in most applications. Since this noise is equal to the square root of the signal level, several points are worth noting. The signal level must always be calculated in electrons, using the digitizer gain value to multiply the gray levels in the image. The more signal that is collected, the better. Since the square root of at larger signal is a larger number, this concept is not always obvious. The importance is found in the signal-to-noise calculation (S/N). As the signal becomes bigger, the noise becomes a smaller percentage of the signal. This percentage is a good approximation of the limit of accuracy. Examples: 100e = 10e = 10% accuracy 500e = 22e = 4% accuracy


The accuracy of data in different areas of an image is dependent upon the signal level in each area. The full-well capacity is the limit to accuracy for the CCD. In absorption studies (measuring dark regions in a bright background), the need for accuracy means that devices with extremely large full-well capacities are needed. The dark regions need to have high signal and the background areas must still not be saturated. It is not uncommon for a typical CCD camera to have 20,000 electrons full-well capacity and a 12-bit digitizer. Under these conditions, each of the 4096 gray values will represent



∼5 electrons of signal. With a digitizer offset of 200 gray levels, any signal from a sample will be superimposed on this 200 level. In this example, we will assume that some region of the image has an average intensity of 500 gray levels and an adjacent region has an intensity of 600 gray levels. We want to find out how much confidence we can place in each of these measurements and how much different they are. We first must subtract the digitizer offset from the indicated signal which leaves us with gray levels of 300 and 400 respectively. Then we multiply each data set by 5 electrons to get the actual signal values: 1500e in one region and 2000e in the adjacent region. Now we take the square root of each to get 39e and 45e. These values represent the total noise in each region of the image. This is the amount of fluctuation we can expect in each signal. We can now make these numbers a percentage of the signal (2.6% and 2.3%) or convert them back to gray levels (8 gray levels and 9 gray levels) to establish the range of fluctuation, or accuracy, that we can expect from this data. If the expectation is accuracy of ∼2%, we have enough data to make this statement. If the need for accuracy is 1%, we will have to find some way to improve our data set. We would need about 10,000 electrons in one image to get close to 1% accuracy. Improving accuracy Accuracy can be improved in two ways. First, one should collect more signals by increasing the QE, exposing longer, or increasing the signal intensity. These are the most common choices, but are not always possible. Second, measurements should be averaged. If it is possible to repeat the data collection process and mathematically average more than one image of the same data set, then the accuracy can be improved by the square root of the number of frames averaged. Even averaging two frames will increase the accuracy by 1.4 times. ANALYSIS Once the image has been captured, the data must be analyzed and distilled into information about the results of the electrophoresis experiment. Through the use of standards and experimenter input, this software-driven process can estimate mass and quantity of objects in an image and detect relationships between objects within one image and between similar images. The type of software used depends on the analysis to be performed. Images from single electrophonetic separations are examined by one-dimensional analysis software optimized for lane-based band detection. Images from two-dimensional electrophoresis are best handled by specific programs designed to detect spots and to assign two mobility values and a quantity value to the spot. After the initial characterization of bands and spots, comparisons are often made between bands or spots from different experiments through the use of database programs and matching algorithms. Software for One-Dimensional Analysis Table 10.5.4 lists popular sources of electrophoresis imaging software. Lane positioning For one-dimensional analysis, the first activity is to detect the lanes on the image. One of three different methods is commonly employed for this. For images with straight, well-defined lanes with a large number of bands, automatic lane-detection algorithms can quickly and accurately place the lanes. On images with very well-defined lanes, such as



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Table 10.5.4 Popular Sources of One- and Two-Dimensional Electrophoresis Analysis Software

Company

Web site

Software

1-D gels UVP, Inc. Nonlinear Dynamics MediaCybernetics Scanalytics, Inc. NIH (free, with gel tutorial)

http://www.uvp.com http://www.nonlinear.com http://www.mediacy.com http://www.scanalytics.com http://rsb.info.nih.gov/nih-image/

LabWorks, DocIt Phoretix series Gel-Pro, ImagePro 1Dscan EX NIH Image

2-D gels Genebio Nonlinear Dynamics Amersham Scanalytics, Inc. Bio-Rad Compugen

http://www.genebio.com http://www.nonlinear.com http://www.amersham.com http://www.scanalytics.com http://www.biorad.com http://www.2Dgels.com

Melanie Phoretix series Melanie Gellab II+ PDQuest Z3, Z4000

pseudoimages from finish-line type electrophoresis equipment, automated lane calling based on image position is possible. For images with “smiling,” bent, or irregular lanes, manual positioning of the lanes is often the fastest and most accurate method of lane definition. Regardless of the method of identifying the lanes, the lane boundaries need to be carefully set for accurate quantitation and mass determinations. Lane widths should be wide enough so that the entire area of all bands in that lane are included, but they should not be so wide as to include bands from adjacent lanes. To accomplish this, curved or bent lanes might need to be used in order to follow the electrophoresis lane pattern. Lane length and position also must be adjusted as necessary so that all bands of interest are included. If mass determinations are necessary, the sample loading point should probably also be included in the lane or be the start of the lane. At this point, lines of equal mobility (often called Rf or iso-molecular-weight lines) are added to the image as necessary. These lines allow for correction of lane-to-lane deviations in the mobility of reference bands and generate more accurate measurements of mass. A similar form of correction is also possible for within-lane correction of mobilities. This correction is important for accurate detection and quantitation of closely spaced bands.


Band detection Once the lanes have been defined, the bands present in each lane need to be detected. There are many methods for detecting bands. One method is to systematically scan the lane profile from one end to the other, identifying regions of local maxima as bands. Another common method is to use first and second order derivatives of the lane image or lane profile in order to find inflection points in the change of slope in pixel intensity values (Patton, 1995). Regardless of the method used, it is often necessary to alter the search parameters so that they perform reliably under a given experimental condition. Typical search parameters include ones for detection sensitivity, smoothing, minimum interband gaps, and minimum or maximum band peak size. Smoothing reduces the number of bands detected due to noise in the image. A minimum interband gap is often used to avoid detection of false secondary bands on the shoulders of primary bands. Limitations on peak sizes, especially for within-lane comparison to the largest band’s peak, can be a useful way to allow sensitive detection of bands in underloaded lanes without detecting false bands in overloaded lanes.



Band edges are often detected in addition to band peaks in order to further define bands or to quantitate band amounts. This can be accomplished by using local minima, derivatives of the lane profile, or fixed parameters such as image distances or a percentage of band peak height. The band edges can be applied as edges perpendicular to the long axis of the lane or as a contour of equal intensity circling the band. The perpendicular method is advantageous for bands with uneven distribution of material across the face of the band, while the latter method is better for “smiling” or misshapen bands. Background subtraction With nearly all electrophoresis procedures, the most informative images have a low level of signal intensity at each pixel that does not result from protein or nucleic acid. Instead, this background intensity is attributable to the gel medium, the visualization method, electronic noise, and other factors. Since this background tends to be nonuniformly distributed throughout the image, failure to subtract it can make band detection and quantitation less accurate. Many methods of background subtraction are possible. Sometimes it is possible to generate a second image under conditions that do not detect the protein or nucleic acid. The second image is then digitally subtracted from the data-containing image to remove the background. More often, background information must be obtained from a single image. If the background varies uniformly across the image, a line that crosses the variation can be defined at a point where no bands are present. The intensity values at each point on the line can be used as the background value for the pixels perpendicular to the line at that point. Commonly, background is also present as variations in intensity along the long axis within each lane. One simple method is to take the lowest point in the lane profile as the background. Another method is to use an average value of the edge of each band as the background for that band. More complicated methods such as valley-to-valley and rolling-disk use local minima points in the lane profile to define a variable background along the length of the lane. Because there can be many different causes and distributions of background, no single method of background determination can be recommended for all experiments. Characterization Once lanes and bands have been detected, it is possible to interpret the mobility of the nucleic acid or protein bands. Depending on the method of electrophoretic separation, information on mass (length or size), pI, or relative mobility (Rf) can be inferred from mobility information. The mobility is characterized using a standard curve with internal standards of known properties. The type of curve depends on several parameters. By definition, with Rf-based separation, a linear first-order curve is used since it represents the linear relationship between mobility and Rf. Similarly, pI and mobility are generally linear in isoelectric focusing separations. For separations based on size, a curve generated from mobility versus the log of the molecular weight provides a relatively good fit as measured by the correlation coefficient (R2). Several other curves have been suggested for size-based separations, including modified hyperbolic curves and curves of mobility versus (molecular weight)2/3 that have good correlation coefficients (Plikaytis et al., 1986). In some cases, no single curve equation can adequately represent the data, and methods of fitting smooth contiguous curves using only neighboring points such as a Lagrange or spline fit (described in Hamming, 1973) are necessary. This is most common for size separations with a very large range of separation sizes and with nonlinear gradient gels. Care must be taken with multiple-curve techniques since they rely on only a few data points for any one part of the composite curve, and outlying data points can drastically affect the outcome. For size and Rf determination, a uniform position must be found in each lane as a point from which to measure the mobility of each band. Many software analysis packages use



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the end of the lane as the measuring start point, so for them it is important to position each lane start point at an iso-molecular-weight or iso-Rf point. A convenient point is the well or sample-loading position since it is usually easily detectable and at an equal mobility position in each lane. A consistent point on each band must also be chosen to measure mobility. A band’s peak is easily defined in digital image analysis and is commonly used. Since peak positions are harder to detect visually than edges on silver-halide images, the leading band edge is sometimes used when comparing digital results with silver-halidebased results. Once lanes and bands have been detected, it is also possible to quantify the amount or at least relative amount of nucleic acid or protein present in each band. The amount in a band is related to the sum total of the intensity values of each pixel subtracted by the background value for each pixel in a band. For absorptively detected bands, intensity values are converted to OD values. The total value that is calculated is equivalent to the volume of the band and can be directly compared to other bands that are within the linear range for the visualization method. If standards of known amounts are loaded onto the same gel, they can be used to generate a standard curve that converts band volume into standard units such as micrograms. For greatest accuracy, it is important to be able to generate multiple standard curves when using visualization methods, such as Coomassie blue staining, that are affected by band or spot composition. Quantitation becomes more complicated when bands are not fully resolved. In this case, material from one band is contributing to the volume measurement of an adjacent band and vice versa. The simplest method for handling this is to partition into each band only the volume within its edges. Alternatively, a Gaussian curve can be fitted to each band and the volume contained within the curve used to estimate the amount of the band. Since most electrophoresis bands have a pronounced skew towards the leading edge of the band, modified Gaussian curves have also been used (Smith and Thomas, 1990). In either case, the curve-fitting process is calculation intensive and can significantly increase analysis times for images with many bands. Software for Two-Dimensional Analysis In two-dimensional analysis, the first-dimension separation is performed in a single column or lane followed by a second separation performed perpendicular to the first. The result after visualization is a rectangular image of up to 10,000 spots. The most common two-dimensional gel type is one in which protein is separated first by apparent pI and second by molecular weight, although two-dimensional separation of nucleic acids is also possible. While many of the concepts and analysis techniques used with one-dimensional gels are applicable to two-dimensional gels, the complex nature of most two-dimensional gels requires somewhat different methodology. For example, spots are more difficult to detect since they are not conveniently arranged in lanes and can vary more in shape and overlap than bands. In addition, two-dimensional experiments usually require some method of comparing between two images whereas one-dimensional images usually contain all of the information from an experiment. See Table 10.5.4 for sources of two-dimensional electrophoresis imaging software.


Spot detection Probably the most difficult aspect of two-dimensional analysis is efficient and accurate spot detection. If it is incorrectly done, it can lead to hours of manual editing. Due to the complexity and computational intensity of some algorithms, the detection process itself can last hours on relatively fast desktop computers. One theoretically effective but computationally intense method is to treat the image as essentially a three-dimensional



image with spots treated as hills and background as valleys. A large number of Gaussian curves are then combined to describe the topology of the image. Many other methods make use of a digital-imaging technique known as filtering. In essence, filtering is a way to weight the value of a pixel and its neighbors in order to generate a new value for a pixel. By passing a filter across an image pixel by pixel, a secondary image is generated. Filters can be designed for many tasks, including sharpening an image or removing high-frequency noise. Filters can also be generated to help detect spots by making images that are first and second derivatives of the original image. The derivative images indicate inflection points in the intensity pattern and can be used to detect spot centers and edges. In a different method, called thresholding, filters can be used to detect the edges of objects. Instead of looking for inflection points, threshold filters identify intensities above a set level or at ratios between central and edge pixels above a set value. Since the edges on two-dimensional spots tend to be diffuse, sharpening filters are sometimes used prior to the thresholding filter. In some cases, multiple techniques are used to detect spots (Glasbey and Horgan, 1994). Unlike one-dimensional detection, detection on images of two-dimensional experiments usually requires secondary processing to get acceptable performance. One example of a secondary process is to discard spots with sizes below a set minimum or above a set maximum. Another is to analyze spots that are oval for possible splitting into two spots. Even after secondary processing, it is likely that a small amount of manual editing will be necessary. When manually editing an image, care must be taken to use as objective criteria as possible, especially if two or more images are to be matched and spot volumes compared. Characterization In a two-dimensional system, determination of protein or nucleic acid mobility is complicated by the fact that there are two mobilities to account for and that the seconddimension separation tends to make estimation of the separation which occurred in the first dimension more difficult. One method for dealing with this is to have a series of markers in the sample that, after both separations are completed, are evenly distributed within the gel and image. It is also possible to estimate separation characteristics from calibration points located at the periphery of the gel. For example, distance measurements can be used to pass calibration data from the first dimension separation, and standards can be separated at the ends of the gel to calibrate for mobility in the second dimension. Regardless of the method used, in many instances, a series of related images will be examined and similar spots in each image will be matched. When this occurs, it is possible to calibrate one image and then pass the calibration information via the matches to the related images. Quantity determination is similar in many regards to that which occurs in one-dimensional analysis, but there are some differences. If spot edges are detected, a simple method of determining spot volume is to take the sum of the intensity value of each pixel in the spot reduced by a background intensity value. Multiple Gaussian curves can also be fitted to the spot to approximate the volume (Garrels, 1989). More difficult is attempting to compensate for a skewed distribution in a size-separating dimension while trying to use a regular Gaussian fit for a pI separation, such as is encountered with the most common form of two-dimensional protein separations. The distribution of background makes quantitation more difficult in two-dimensional gels. There is no lane-dependent component, so it is necessary to use other methods such as image stripes, finding local minima values or using values derived from the spot edges to determine background values. Analysis of Proteins


Supplement 66

Matching Matching is the process in which proteins or nucleic acids with similar separation properties are linked or clustered together. Matching can occur within one image or between multiple images as long as a frame of reference is established. Matching allows for comparisons between samples. It also makes annotation and data entry easier, since if one spot or band is matched to others and is characterized or annotated, this information is easily passed to all the other matches. An underlying assumption of matching is that objects with similar separation properties are actually similar. Care must be taken to confirm the identity of matched spots or bands by other methods on critical experiments. A simple form of matching is to link bands or spots at similar positions on the gel images. This works well when separation and imaging conditions are uniform. This is very seldom the case, since slight differences in the electrophoresis, visualization, and imaging conditions across a gel and between gels generates incorrect matching with this method. Since bands on one-dimensional gels are relatively easy to calibrate for mobility, matching can occur along contours of equal mobility. This dramatically decreases but does not eliminate the variability in detecting similar bands. Much of the remaining variability can be attributed to calibration errors. This error can often be compensated for by allowing a small tolerance in mobility values in determining whether a band is matched or not. Because of the difficulties in calibrating mobility in two-dimensional gels, it is often more practical to use matched spots for calibrating mobility than vice versa. Spot matching between two two-dimensional images starts by finding a small number of landmark spots that are used as seeds for subsequent matches (Appel et al., 1991; Monardo et al., 1994). There are many methods for finding the landmarks in both images, including finding the highest intensity spots, spots in unique clusters, and manual positioning. The most common procedure from this point is to derive a vector that describes the direction and extent of the path from one matched spot to the other when the two images are superimposed. The vector is used as the basis for finding more matches near the landmark matches. To allow for error, the area within a small radius is searched extending from the end of the vector. Once another match is found, its vector is computed and used as the starting point for finding neighboring matches. From this progression, the entire gel is matched. If all vectors are displayed graphically when matching is complete, questionable matches can often be identified as vectors that are significantly different from neighboring vectors. One specialized use of matching is as an estimator of the similarity and potential genetic relatedness of organisms. For example, on a one-dimensional gel image, a ratio of the matched to unmatched bands for each pairwise combination of lanes can be calculated. This ratio can be used as an indicator of similarity, with values near 1 indicating a pair of highly similar lanes, and values near zero indicating very dissimilar lanes. Assuming that the contents of the lanes are valid samples of the originating organism’s genetic makeup, the information on lane similarity can be converted to estimates of genetic similarity. A convenient way to display this similarity data graphically is to generate a dendrogram with similar objects close to each other and less similar objects more distantly placed. An example of such a dendrogram is presented in Figure 10.5.5, where samples from Listeria isolates are arranged based on banding pattern. Databases


In many cases, image analysis is not the last step in the process. The image and analysis data need to be archived in a searchable format. There may be a need to analyze the data from multiple experiments conducted at different sites or in laboratories around the world. Bioinformatic links to diverse data sources might be desired to help develop a unified understanding of the biology behind particular phenomena. When these situations arise, database programs can be utilized to store, link, and search image analysis results.



Figure 10.5.5 Example of a dendrogram generated from similarity data on band matching between lanes. DNA samples from 22 isolates of Listeria were subjected to Random Amplification of Polymorphic DNA (RAPD) analysis and the resulting electrophoresis image was analyzed with ImageMaster software (Amersham Pharmacia Biotech). Clustering was performed using the Dice coefficient with a tree structure based on the Unweighted Pair-Group Method using Arithmetic Averages (UPGMA). Similarity values between isolates can be determined by locating the node that connects the isolates and reading the value from the scale on the lower left edge of the dendrogram.

As the number of images that are captured and analyzed grows, it becomes increasingly more difficult to find particular information from the large number of files that are stored. Relatively simple databases can be used if the major requirement is to find previously analyzed images and associated data. Such databases often display a miniaturized version of each image to aid in visual scanning for the file as well as simple searching for image-specific information such as date of analysis, file name, or other information that was entered at the time of image capture. More powerful database products are also available that can perform complex searches on data generated during the analysis. For an example, a search on a two-dimensional database might include finding proteins exhibiting a specific expression profile and having a molecular weight >20 kDa with a pI between 3 and 5 or 8 and 10 with an amount 800 Ci/mmol) can be obtained from several suppliers of radiolabeled amino acids. Protein hydrolyzates of Escherichia coli grown in the presence of [35SO4]2− (e.g., Tran35S-label from ICN Biomedicals or Expre35S35S from NEN Life Sciences) can be used as substitutes for [35S]methionine in metabolic labeling techniques. These preparations contain 35S distributed among several different compounds. In a typical batch, ∼70% of the radioactivity will be present as [35S]methionine and ∼15% as [35S]cysteine; the remainder are other 35S-labeled compounds. Labeling with these radioactive protein hydrolyzates in methionine-free medium will result in incorporation of label only in methionine residues; use of methionineand cysteine-free medium will result in labeling of both methionine and cysteine residues.



Purified [35S]methionine should be used whenever certainty of labeling with only [35S]methionine is required. Another reason for using purified [35S]methionine is that it tends to be more stable and emit less volatile decomposition products than 35S-labeled protein hydrolyzates. In most cases, however, the relatively inexpensive radioactive protein hydrolyzates (∼1/3 the cost of [35S]methionine) can be used instead of purified [35S]methionine. [35S]methionine preparations should be stored frozen at −80°C. Under these conditions, they are stable for at least 1 month; the half-life of 35S is 88 days. For proteins with one or no methionine residues but several cysteine residues, labeling with [35S]cysteine (available with specific activities >800 Ci/mmol) is a good option. 35Slabeled protein hydrolyzates are not good sources of [35S]cysteine for radiolabeling because only ∼15% of the 35S-labeled compounds are [35S]cysteine. Purified [3H]leucine (available with specific activities of up to 190 Ci/mmol) is a good alternative to 35S-labeled amino acids. The halflife of 3H is ∼12 years. The specific activities of other 3H-labeled amino acids range between 5 and 140 Ci/mmol (Table 10.18.1). Several problems can arise when using certain 3H-labeled amino acids due to their participation in metabolic pathways. The following problems must be considered when using tritiated amino acids for metabolic labeling of proteins. Nonessential versus essential amino acids. Cells are able to synthesize nonessential amino acids from other compounds. If the radiolabeled amino acids used are nonessential, their specific activity will be reduced by dilution with the endogenously synthesized amino acids. Therefore, essential amino acids (E in Table 10.18.1) should be favored for metabolic labeling. Transamination. The α-amino groups of many amino acids can be removed in reactions catalyzed by transaminases (T in Table 10.18.1). Deamination of the radiolabeled amino acids can be prevented by addition of the transaminase inhibitor (aminooxy)acetic acid during starvation and labeling of the cells (Coligan et al., 1983). Interconversion. Certain radiolabeled amino acids can be converted by cells into other amino acids (I in Table 10.18.1). This problem is of particular importance in methods used to determine the amino acid composition or sequence of radiolabeled proteins.

Labeling time The experimental purpose, the turnover rate of the protein of interest, and the viability of the cells should all be considered when determining the length of time for labeling. If the purpose of the experiment is to identify or characterize a protein precursor, pulse-labeling (10 to 30 min) must be used (see Basic Protocol and Alternate Protocol 1). If the protein of interest has a low turnover rate or if it is necessary to accumulate a labeled mature product, a protocol for long-term labeling (6 to 32 hr) is more appropriate (see Alternate Protocol 3). If the biosynthesis, posttranslational modification, intracellular transport, or fate of newly synthesized proteins is being analyzed, a pulsechase protocol should be used (see Alternate Protocol 2). The turnover rate of the protein of interest is an important parameter to be considered when determining the labeling time. Proteins with a high turnover rate are optimally labeled for short times. Longer times will result in increased labeling of other cellular proteins, causing a decrease in the relative abundance of the labeled protein of interest in the cell lysate. This could result in increased detection of nonspecific proteins in immunoprecipitates (see UNIT 10.16). Conversely, proteins that turn over slowly should be labeled for longer times. Incorporation of radiolabeled amino acids into proteins is directly proportional to their length of labeling time for a certain period, after which it tends to plateau. When all the limiting amino acid is consumed, protein synthesis ceases. The length of the initial phase of linear incorporation will vary with the concentration of the labeling amino acid and the density and metabolic activity of the cells. Concentration and specific activity of the radiolabeled amino acid Because radiolabeled amino acids are used in limiting amounts, their incorporation into proteins is directly proportional to their concentration in the labeling medium. If very short pulses are required, the concentration of labeled amino acid can be increased to compensate for the shorter labeling time. The incorporation of radiolabeled amino acids is also directly proportional to their specific activity. Addition of unlabeled amino acids, as is required in long-term labeling protocols, will result in reduced incorporation over short periods. Analysis of Proteins


Supplement 44

Cell density At low cell densities, the amount of labeled protein synthesized is directly proportional to the cell concentration. At high densities, however, incorporation will increase nonlinearly. Very concentrated cell samples (>5 × 107 cells/ml) can only be labeled for short periods (≤5 min) as rapid loss of metabolic activity and cell viability will occur due to acidification of the medium and accumulation of toxic metabolites. Conditions for very short pulses Very short pulses (2 to 10 min) are required to study posttranslational modifications that occur shortly after synthesis. This includes folding, disulfide bond formation, and early carbohydrate modifications of newly synthesized proteins. Proteins that are rapidly degraded after synthesis are also best studied with very short pulses. The protocol for very short pulses is similar to the Basic Protocol for pulselabeling, with the following modifications: (1) the pulse-labeling medium contains higher concentration of radiolabeled amino acid (up to 1 mCi/ml), (2) labeling is stopped by addition of ice-cold PBS containing 20 mM freshly prepared N-ethylmaleimide (NEM) to prevent oxidation of free sulfhydryls, and (3) if a chase is necessary, 1 mM cycloheximide is added to the chase medium to quickly stop the elongation of nascent polypeptide chains. For additional details on very short pulses, see Braakman et al. (1991). Temperature and pH Unless otherwise required for special conditions (e.g., heat shock, temperature-sensitive mutants), metabolic labeling should be performed at 37°C. For this reason, it is important that the labeling medium be warmed to 37°C before adding it to the cells. A dramatic reduction in incorporation occurs at room temperature. It is also important that the pH of the labeling medium be ∼7.4.

Anticipated Results A typical incorporation of labeled amino acid precursor after a 30-min pulse under the conditions described in this unit (see Basic Protocol and Alternate Protocols 1 and 2) is 5% to 20%. In the case of long-term labeling, the incorporation typically reaches 30% to 60%. If the labeled cells are used for immunoprecipitation (UNIT 10.16), followed by SDS-PAGE (UNIT 10.2) and autoradiography (APPENDIX 3A), specific bands should be visible within two hours to two months of exposure.

Time Considerations It takes 1 to 2 hr to prepare cells and materials for labeling. The actual labeling time depends upon the protocol chosen—pulse-labeling takes 10 to 30 min and long-term labeling takes 6 to 32 hr. Washing and processing cells may take an additional 1.5 hr.

Literature Cited Braakman, I., Hoover-Litty, H., Wagner, K.R., and Helenius, A. 1991. Folding of influenza hemagglutinin in the endoplasmic reticulum. J. Cell Biol. 114:401-411. Coligan, J.E., Gates, F.T. III, Kimball, E.S., and Maloy, W.L. 1983. Radiochemical sequence analysis of metabolically labeled proteins. Methods Enzymol. 91:413-434. Lathe, R. 1985. Synthetic oligonucleotide probes deduced from amino acid sequence data. Theoretical and practical considerations. J. Mol. Biol. 183:1-12. Meisenhelder, J. and Hunter, T. 1988. Radioactive protein labelling techniques. Nature 335:120.

Key Reference Coligan et al., 1983. See above. Contains a detailed description of conditions used to metabolically label proteins with different radiolabeled amino acids.

Contributed by Juan S. Bonifacino National Institute of Child Health and Human Development Bethesda, Maryland

Metabolic Labeling with Amino Acids



Isolation of Proteins for Microsequence Analysis

UNIT 10.19

The first basic protocol can be used to determine the amino-terminal sequence of polypeptides or proteins. It is particularly appropriate for large fragments of insoluble or hydrophobic proteins or proteins that cannot be purified to >90% molar purity without electrophoresis. Although the efficacy of this technique varies with the protein, it is possible to obtain useful sequence information starting with ≤50 pmol of the protein of interest. If the protein is blocked at the amino terminus, chemical cleavage or partial enzymatic digestion must be performed prior to electrophoresis. Upon isolation, the internal amino acid sequence is analyzed as described in the second basic protocol. This method requires ∼200 pmol of protein for analysis. It is understood that both of the analyses will be done in association with an expert operator of an automated protein sequencer. In addition, the second basic protocol requires expertise with reversed-phase HPLC to separate peptides. DETERMINATION OF AMINO ACID SEQUENCE BY SDS-PAGE AND TRANSFER TO PVDF MEMBRANES

BASIC PROTOCOL

A minigel is prepared and preelectrophoresed. A sample containing the desired protein is loaded onto the minigel and fractionated by SDS-PAGE at neutral pH. The proteins are then electrophoretically transferred to a sheet of polyvinylidene difluoride (PVDF) and stained with Coomassie blue. The separated bands are excised, then analyzed in an automated protein sequencer. Materials Separating and stacking gel solutions (Table 10.19.1) 4× gel buffer Glutathione, reduced powder (Sigma #G4251) 10× lower reservoir buffer 10× upper reservoir buffer Mercaptoacetic acid, sodium salt Protein sample in sample buffer (see support protocol) Methanol Transfer buffer 0.1% Coomassie blue in 50% methanol (v/v) 10% acetic acid in 50% methanol (v/v) Vertical minigel unit (e.g., Bio-Rad Mini-Protean II; Hoefer Mighty Small SE 250/280 is not recommended for this procedure) Power supply (constant voltage and constant current) Microvolume syringe or gel-loading pipet tip Powder-free plastic gloves PVDF membranes (e.g., Immobilon-P or -PSQ, Millipore; ProBlott, Applied Biosystems) Small-format transfer apparatus (Midget MultiBlot, Hoefer or Pharmacia LKB; Mini Trans-Blot, Bio-Rad) Automated protein sequencer (Applied Biosystems) Additional reagents and equipment for minigel preparation (UNIT 10.2) Analysis of Proteins Contributed by Malcolm Moos, Jr. Current Protocols in Molecular Biology (2000) 10.19.1-10.19.12 Copyright © 2000 by John Wiley & Sons, Inc.


Table 10.19.1

Recipes for Polyacrylamide Separating and Stacking Gels

SEPARATING GEL Final acrylamide concentration in the separating gels (%)

Stock solutions 5 30% acrylamide monomera H2O TEMEDb

6

7

8

9

10

11

12

13

14

15

3.33 4.00 4.67 5.33 6.00 6.67 7.33 8.00 8.67 9.33 10.00 11.49 10.83 10.16 9.50 8.84 8.17 7.51 6.84 6.18 5.51 4.85 0.040 0.033 0.029 0.025 0.025 0.020 0.018 0.017 0.015 0.014 0.013

Mix the above ingredients (listed in milliliters of stock solution) with 5 ml of 4× gel buffer and 0.14 ml of 5% potassium persulfate or 70 µl of 10% ammonium persulfate. STACKING GEL 0.666 ml 30% acrylamide monomera 3.033 ml H2O 0.025 ml 10% ammonium persulfate or 0.05 ml 5% potassium persulfate 0.025 ml TEMEDb aGas-stabilized monomer solution (containing 37.5:1 acrylamide/N,N′-methylene-bisacrylamide) from which acrylic acid and carbonyl-containing compounds have been removed (Protogel, National Diagnostics; or PAGE1 protein gel mix, Boehringer Mannheim) bTEMED may have to be altered to facilitate proper polymerization. Values given are reasonable approximations.

Pour and preelectrophorese the minigels 1. Pour denaturing minigels as in UNIT 10.2, substituting the separating and stacking gel solutions listed in Table 10.19.1. Deaerate the gel polymerization mixtures prior to adding persulfate and TEMED and be careful not to draw in air through the polymerization mixtures when transferring or pipetting the solutions. After removal of combs, the wells should be rectangular and firm. If they are not, prepare a fresh gel; poorly polymerized stacking gels are the most common cause of low sequencing yields.

2. Assemble vertical minigel unit. 3. Dilute 80 ml of 4× gel buffer to 320 ml (to 1× gel buffer). Pour 200 ml of 1× gel buffer into lower buffer reservoir. Add reduced glutathione (powder) to remaining 1× gel buffer to 1.0 mM final. Pour this into upper buffer reservoir. Attach gel to a constant voltage/constant current power supply (see UNIT 10.2 introduction for a discussion of electricity and electrophoresis). Glutathione acts as a scavenger to eliminate the by-products of acrylamide polymerization.

4. Preelectrophorese by applying 10 mA per minigel for 45 min. Voltage applied for >45 min may impair resolution.

5. Turn off power supply. Allow gel to stand overnight. Gels may be stored this way for several days.

Load and electrophorese the sample 6. Pour off gel buffer and blot wells with tissue or filter paper. Isolation of Proteins for Microsequence Analysis

7. Dilute 10× lower and upper reservoir buffers to 1×. Pour 200 ml of 1× lower reservoir buffer into lower reservoir. Add 0.1 g mercaptoacetic acid (sodium salt) to 150 ml of 1× upper reservoir buffer and pour into upper reservoir.



Mercaptoacetic acid is used as a substitute for glutathione at this step because it is less likely to interfere with sequence analysis.

8. Load protein sample dissolved in sample buffer with a microvolume syringe or gel-loading pipet tip. When using 5-well combs and 0.75-mm gels, results are best with sample volumes 60 kDa, reduce the amount of methanol in transfer buffer to 1%.

Stain the blots and excise the bands 15. Disassemble transfer apparatus. Immerse PVDF membrane blots in 0.1% Coomassie blue in 50% methanol and agitate 5 min. 16. Destain blots in 10% acetic acid prepared in 50% methanol by agitating until bands become clearly visible (5 to 10 min). 17. Transfer to water and photograph (optional; UNIT 10.6). 18. Excise band of interest with a razor blade (use a new razor blade for each band). Place each band in a microcentrifuge tube and allow to air dry at room temperature (do not heat). Store excised bands at −20°C. Sequence the proteins 19. Insert excised band into sequencer reaction cartridge, protein side facing the solvent delivery (see manufacturer’s instructions). All pieces of PVDF must fit in a single layer in the reaction cartridge; they may be cut or trimmed to fit as required. When using conventional chemistry (“NORMAL” program on Applied Biosystems instruments), two alternate configurations have proven optimal, depending on the sequencer. For many gas-phase instruments, placing the PVDF pieces between the upper-cartridge block and the TFA-etched glass-fiber filter gives the best results. For some gas-phase and most pulsed-liquid instruments, placement between the glass-fiber disk and the cartridge seal is better. The results of the cartridge leak test should be normal. The Blott cartridge should be used on Applied Biosystems instruments, if available.

20. Sequence the sample on an automated protein sequencer. The “Blott-2 reaction cycle” should be used if a Blott reaction cartridge is installed. The conversion cycle described by Tempst and Riviere (1989) is recommended. SUPPORT PROTOCOL

PREPARATION OF PROTEIN SAMPLES FOR SDS-PAGE Samples are concentrated and freed of contaminants that interfere with electrophoresis by precipitation. They are then solubilized in a sample buffer containing SDS. Additional Materials Protein samples 1 M NaHCO3 (optional) 100% ethanol, ice-cold (containing no denaturants; USP grade) Sample buffer 0.1% (w/v) pyronin Y Ultrafiltration concentrator (Amicon) or Speedvac evaporator (Savant) Drawn-out Pasteur pipet or gel-loading pipet tip Boiling water bath 1. Adjust the salt concentration of the protein sample to >100 mmol/liter with 1 M NaHCO3, if necessary.


2. Concentrate samples to 50 to 100 µl by ultrafiltration (follow manufacturer’s instructions explicitly) or vacuum centrifugation (it is crucial to avoid introduction of airborne debris when vacuum is released). Transfer to 1.5-ml microcentrifuge tubes.



3. Add 9 vol ice-cold 100% ethanol to the samples. Incubate 1 hr on dry ice or overnight at −20°C. Most samples may be kept indefinitely at this stage.

4. Microcentrifuge 15 min at maximum speed. Aspirate the supernatant with a drawnout Pasteur pipet or a gel-loading pipet tip and save the pellet. A drum rotor, which holds the microcentrifuge tubes at a 90° angle, is more likely to give compact, easily visualized pellets than the commonly used fixed-angle rotor.

5. Dissolve the pellet in 10 µl sample buffer by drawing the sample buffer up and down with a pipettor. Boil 3 min in a boiling water bath. 6. Add 1 µl of 0.1% pyronin Y (tracking dye) to the sample. Load on the minigel in the basic protocol. DETERMINATION OF INTERNAL AMINO ACID SEQUENCE FROM ELECTROPHORETICALLY-SEPARATED PROTEINS

BASIC PROTOCOL

As many as 50% of all eukaryotic proteins are blocked at the amino terminus, making sequence determination of the intact protein impossible. The method presented here allows multiple stretches of internal amino acid sequence to be obtained from most proteins that can be isolated by any one- or two-dimensional gel electrophoresis and transfer method. About 200 pmol of protein is an adequate sample size; smaller amounts have sufficed in several cases. The protein(s) of interest is separated by electrophoresis and transferred to nitrocellulose. Protein bands or spots are visualized with Ponceau S and excised. After destaining, the nitrocellulose is treated with polyvinylpyrrolidone (PVP) to prevent adsorption and inactivation of proteolytic enzyme, which is added to cleave the protein. The resulting peptides, which elute from the membrane, are separated by reversed-phase HPLC. Each peptide is analyzed individually in an automated microsequencer. Materials Protein sample 0.1% (w/v) Ponceau S (Sigma) in 1% (v/v) acetic acid 1% acetic acid (v/v) 0.2 mM NaOH 0.5% (w/v) polyvinylpyrrolidone (Sigma # PVP-40) in 0.1 M acetic acid Digestion buffer 1 mg/ml sequencing-grade trypsin (Promega #V511A) Chromatography solvent A: 5% (v/v) acetonitrile in 0.1% (v/v) trifluoracetic acid (TFA) Chromatography solvent B: 70% (v/v) acetonitrile in 0.085% (v/v) TFA 0.22–µm nitrocellulose membrane (e.g., Schleicher & Schuell #BA83) Acid-washed glass plate or petri dish Powder-free gloves Fine-tipped forceps 0.5-ml microcentrifuge tube Bath sonicator (e.g., Bransonic 12) Centrifugal filter device, 0.22-µm membrane, low-protein-binding (e.g., Millipore #UFC3-OGV-00) Reversed-phase HPLC column (e.g., Vydac #214TP52), UV column monitor, and chart recorder Column oven (optional)



Supplement 19

Electrophorese and transfer the proteins 1. Resolve protein(s) of interest by electrophoresis and transfer to nitrocellulose membrane as described in UNIT 10.8. It is important to end up with as much protein as possible bound to the smallest possible area of nitrocellulose. Apart from this constraint, there are few limitations on the design of the separation, as chemical modification of the amino terminus will not interfere with subsequent analyses. Although PVDF membranes may be used, nitrocellulose seems to give better yields (see commentary). The membrane must never be allowed to dry from this point on.

Stain and excise the separated proteins 2. Place nitrocellulose membrane in an acid-washed glass petri dish (or similar vessel) containing 50 ml (for an 8 × 10–cm minigel) of 0.1% Ponceau S prepared in 1% aqueous acetic acid. Agitate gently 1 min. For larger blots, increase the volume of stain in proportion to the area.

3. Transfer to 1% acetic acid for 1 min, changing the solution as necessary to allow easy visualization of band(s). Use enough 1% acetic acid to cover the membrane generously.

4. Using a new razor blade and wearing powder-free gloves, cut out the band(s) of interest. Place in 1.5-ml microcentrifuge tube(s). It is important to remove all areas of blank nitrocellulose to minimize subsequent adsorption of the protease used for digestion. Meticulous technique is crucial at this step to eliminate contamination of the sample by adventitious proteins. A clean work area, scrupulously clean glassware and instruments, and powder-free gloves are minimum precautions; tryptic fragments of human keratin have been identified in some samples! To distinguish UV-absorbing peaks corresponding to the protein from those derived from the proteolytic enzyme or other reaction constituents, cut a piece of nitrocellulose from a blank area of the blot to use as a control. The bands may be transferred to microcentrifuge tubes containing 0.5 ml water and stored at −20°C at this step.

Destain the membrane pieces 5. Transfer membrane pieces to 1 ml of 0.2 mM NaOH and vortex 1 min. Aspirate the NaOH and immediately proceed to step 6. Proteins are more apt to be lost from the nitrocellulose under alkaline than acidic conditions. Because release of the peptides is desired at a later stage, the length of time for destaining should be minimized. Residual stain on the blot will not adversely affect the final results.

Block the membrane 6. Add 1 ml of 0.5% PVP-40 in 0.1 M acetic acid and agitate tube gently 30 min at room temperature. 7. Aspirate the PVP-40 and wash membrane five times with 1 ml water. Be sure to remove any liquid droplets caught under tube cap. Isolation of Proteins for Microsequence Analysis



8. Using clean fine-tipped forceps, transfer excised band to a clean glass surface (e.g., acid-washed glass plate or petri dish) and cut it into 1- to 2-mm pieces. Use forceps to collect the pieces and squeeze out excess liquid. Immediately proceed to step 9. Digest the protein with protease 9. Transfer pieces to a 0.5-ml microcentrifuge containing 25 µl digestion buffer. Add 1 µl 1 mg/ml trypsin. Mix so that membrane pieces are evenly coated with solution. 10. Incubate overnight at room temperature. Elute the peptides 11. Microcentrifuge sample 1 sec at high speed, room temperature, to recover liquid that may have condensed on tube walls and cap. 12. Sonicate sample 5 min at room temperature. 13. Microcentrifuge 1 min at top speed, room temperature. 14. Transfer supernatant to a centrifugal filter device. Rinse membrane pieces with 100 µl digestion buffer and add to supernatant. Microcentrifuge sample 20 to 30 sec at top speed. Filtration removes nitrocellulose particles that might clog the HPLC column. Samples should be stored frozen unless they will be fractionated immediately.

Chromatograph and process the samples 15. Equilibrate a 2.1-mm-i.d. × 250-mm reversed-phase HPLC column with 95% chromatography solvent A/5% solvent B at a flow rate of 0.15 ml/min. For optimal resolution, perform the separation at 60°C if a column oven is available. 16. Inject the sample. Wash column 10 to 15 min with 95% solvent A/5% solvent B. 17. Elute the peptides with a gradient between chromatography solvents A and B as follows: 5% to 40% solvent B over 1 hr; 40% to 75% solvent B over 30 min; and 75% to 100% solvent B over 15 min. Monitor elution of the peptides at 215 nm. Elution gradient guidelines are described in Stone and Williams (1986). The TFA concentration should be adjusted to equalize the UV absorbance (215 nm) of the eluants. Alternatives to TFA, such as phosphoric acid or hydrochloric acid, are compatible with the procedure. Buffers containing ammonia or UV-containing impurities should not be used. Trypsin elutes at 60% solvent B and can be used as an internal standard. Characteristically, it elutes in a broader peak than the majority of peptides.

18. Monitor the appearance of peptide peaks with a chart recorder adjusted so that 0.05 to 0.1 AUFS corresponds to a full-scale deflection. Minimize the length and capacity of the capillary tubing between the column and the point of collection as much as possible. Polyethylether ketone (PEEK) tubing (0.005-in. i.d.) is helpful for this purpose.

19. When chart-recorder pen begins a deflection indicative of a peak, wipe tip of capillary tubing against a clean surface (e.g., a Kimwipe or the previous collection tube) and collect this fraction in a microcentrifuge tube. Clean gloves should be worn at this point. Commercial microcentrifuge tubes are sufficiently clean as supplied by the manufacturer if they are stored in the original container and handled with clean gloves.



Supplement 19

20. Immediately cap tube and place it on dry ice. Rapid freezing prevents adsorption of separated peptides to the walls of the microcentrifuge tube. The samples can be stored at −80°C indefinitely.

21. Sequence samples using glass support disks precycled with polybrene as prescribed by Applied Biosystems. Improved reaction (Speicher, 1989) and conversion (Tempst and Riviere, 1989) cycles will reduce background and improve sensitivity. It is very useful to compare chromatograms obtained from the sample and from the blank nitrocellulose to identify peaks corresponding to peptides derived from the protein of interest. UV-absorbing impurities may originate from several sources, including residual PVP-40, contaminated glassware, and the proteolytic enzyme (autolysis of the protease). These can be eliminated easily by inspecting the control chromatogram. Peaks as small as 0.002 AUFS have often been sequenced successfully.

REAGENTS AND SOLUTIONS Digestion buffer 0.1 M Tris⋅Cl, pH 8 1 mM CaCl2 10% (v/v) acetonitrile Store frozen 4× gel buffer 41.24 g bis-Tris (0.493 M) 7.08 ml 37.5% HCl (reagent grade) H2O to 400 ml Final pH should be 6.61; do not adjust the pH with acid or base. This buffer may be stored indefinitely at −20°C or ≤2 months at 4°C. 10× lower reservoir buffer 52.4 g bis-Tris (0.626M) 12.2 ml 37.5% HCl H2O to 400 ml Final pH should be 5.90; do not adjust the pH with acid or base. This buffer may be stored indefinitely at −20°C or ≤2 months at 4°C. 10× upper reservoir buffer 40.28 g N-tris-(hydroxymethyl)-methyl-2-aminoethanesulfonic acid (TES; 0.439 M) 94.64 g bis-Tris (1.131 M) 4.0 g sodium dodecyl sulfate (SDS) H2O to 400 ml Final pH should be 7.25; do not adjust the pH with acid or base. This buffer may be stored indefinitely at −20°C or ≤2 months at 4°C.


Sample buffer 50 mM Tris⋅Cl, pH 6.8 5% (v/v) 2-mercaptoethanol 10% (v/v) glycerol 1% (w/v) SDS Store frozen in small aliquots for ≤2 months (continued)



Transfer buffer 2.21 g cyclohexylaminopropane sulfonic acid (CAPS), free acid 0.5 g dithiothreitol 150 ml methanol H2O to 1 liter Adjust pH to 10.5 with NaOH and chill to 4°C Prepare just before use For proteins >60 kDa, reduce amount of methanol to 1%.

COMMENTARY Background Information The most difficult part of any protein sequencing project is to obtain a sample that is sufficiently free of both protein and nonprotein contaminants. SDS-PAGE is used because of its resolving power and its ability to maintain solubility of almost any protein. Many proteins of interest are identified only by migration on an analytical gel or by immunoreactivity in immunoblot experiments. The sequencing procedures outlined here allow the direct comparison of primary structures. Following the demonstration that blotting procedures could be used to prepare samples for microsequencing (Aebersold et al., 1986; Vandekerckhove et al., 1986; Matsudaira, 1987), acceptance of these techniques was slow due to the inability of many investigators to obtain adequate sequencing signals. Evaluation of several parameters has indicated that preelectrophoresis with a scavenger such as glutathione or mercaptoacetic acid to eliminate reactive by-products of acrylamide polymerization, and lowering the separation pH to reduce the reactivity of the N-terminal amine, can dramatically improve the signals (Moos et al., 1988). Recently, Tris-Tricine gels have become popular, especially for low-molecular-weight proteins and peptides (Schägger and von Jagow, 1987). These gels address the concerns of pH-dependent N-terminal blockage and give good results in sequencing experiments. Their commercial availability in precast format makes them an attractive choice for the novice. The ability to sequence proteins blocked at the amino terminus following electrophoretic separation on gels (second basic protocol; Aebersold et al., 1987; Tempst et al., 1990) is an important improvement in protein analysis. Although meticulous cleanliness and exacting technique are required, the protocol is straightforward and highly reliable. Its particular strength is that it can be used following any type of gel electrophoresis or electrophoretic trans-

fer. It is even possible to combine small spots obtained from several gels (e.g., minor components detected by two-dimensional analysis). Its principal limitation is the relatively low yield of material for sequencing from a given amount of blotted protein; a dark band of Ponceau S–staining material corresponding to 100 to 250 pmol (or more for some proteins) is often required. This problem is obviated somewhat by the extremely high quality of sequence data that can be obtained from minute (e.g., 2 mAUFS) HPLC peaks. Modifications to enhance the sensitivity of the microsequencer 5- to 8-fold have been developed (Tempst and Riviere, 1989); these have proven extremely useful for all microsequencing procedures. A new automated chemistry cycle specifically optimized for PVDF significantly improves results (Speicher, 1989). Applied Biosystems has developed a reaction cartridge designed for PVDF blots that simplifies sample placement and substantially improves sequencing results (Sheer et al., 1990). Finally, a procedure for covalently entrapping blotted proteins in a polymer network may prove beneficial (information available from MilliGen/Biosearch; see APPENDIX 4).

Critical Parameters and Troubleshooting Determination of amino acid sequence by SDS-PAGE and transfer to PVDF membranes In practice, the electrophoresis conditions described in the first basic protocol improve the initial sequencing yield by 2-to 3-fold over those obtained with earlier procedures. Because many proteins are available in very small quantities, this consideration is often crucial. If the sample is not in short supply, it may be reasonable to attempt sequencing with electrophoresis conditions commonly in use (UNIT 10.2; Matsudaira, 1987), especially if the reagents are available in the laboratory. Samples already



Supplement 19


transferred to PVDF by other procedures may also be tried; these should be rewet with methanol and washed with water (3 times, 5 min each) before sequencing to remove dust and materials such as glycine that may have been used in the transfer. Efficient acrylamide polymerization is crucial to obtain good sequencing signals. Even the most efficiently polymerized gels contain 50 to 60 mM acrylamide monomer and possibly other compounds that react in a pH-dependent manner with the N terminus during electrophoresis. Therefore, fresh, high-quality reagents, ultrapure water, and thorough deaeration of the polymerization mixtures are necessary. TEMED is the most likely reagent to deteriorate and a fresh bottle should be opened each month. The buffer system used in this protocol operates at a lower ionic strength than others in common use and therefore is more sensitive to the presence of ionic impurities in the sample. The most efficient way to remove these is by precipitation with organic solvent. Ethanol was chosen for this application (support protocol) because it is least likely to contain impurities that could react with amino termini of proteins (Yuan et al., 1987). Precipitation is most efficient with concentrated samples, so centrifugal ultrafiltration can be useful. Too much SDS in the sample will impair stacking. This is why only 1% SDS, which is sufficient to saturate almost all proteins, is used in the sample buffer. Pyronin Y is used as a tracking dye because it binds to PVDF tightly and unambiguously locates the dye front. The gel buffers and polymerization mixtures should be prepared exactly as specified. No adjustments in pH should be necessary. SDS is not included in the gel polymerization mixtures as this could impair stacking. The same buffer is used in both the stacking and resolving gel. If efficient stacking and good resolution are not observed, the buffers are incorrect, the sample contains too much salt (see support protocol), or the gel is overloaded (10 to 20 µg/band is about right). If poor sequencing yields are obtained with a standard protein known to be unblocked, there are problems with gel polymerization, transfer, or placement of the blots in the reaction cartridge of the sequencer. Optimization of transfer is straightforward, and samples that have been applied directly to PVDF pieces without prior electrophoresis can be used to assess the transfer. It is strongly recommended that β-lac-

toglobulin, which has worked well in many laboratories and is widely available, be used on a test run before using valuable experimental samples. In addition, it is a good idea to use β-lactoglobulin as a positive control in every experiment. This will distinguish between existing N-terminal blockage and blockage that may occur during the procedure. Exposure of the blots to high pH and high temperature should be minimized. When preparing the sample for electrophoresis, the supernatant should be aspirated very carefully—a magnifying lens may be helpful to avoid aspirating the pellet. A 10-µg pellet is usually visible to the naked eye. Determination of internal amino acid sequence from electrophoreticallyseparated proteins Once a sufficiently pure protein has been obtained, the most troublesome aspect of sequencing is chemical modification of the N terminus of the protein, either in vivo or in vitro. Approximately 50% of eukaryotic cellular proteins are thought to have naturally occurring N-terminal blockage. For these proteins, enzymatic or chemical cleavage procedures are required in order to obtain sequence information by automated Edman degradation (Hewick et al., 1981; Tempst and Riviere, 1989). In addition, many reagents commonly encountered in protein isolation procedures can irreversibly modify unblocked N-terminal amino groups and prevent Edman degradation. Because of this, a standard protein, such as β-lactoglobulin, should be analyzed as a control. Failure to obtain any sequence or a low-yield sequence for the standard protein suggests problems with the reagents. The success of this method depends on a number of factors. (1) A reasonably homogeneous band or spot must be obtained. If two or more proteins are overlapping, it will be difficult to resolve individual peptides and impossible to assign sequence data to specific proteins. (2) It is important to maintain meticulous cleanliness throughout the procedure. Keratins from the skin are particularly problematic. (3) UV-absorbing contaminants in polyvinylpyrrolidone (PVP) can obscure the peptide peaks, which are often very small. Therefore, careful washing after the PVP blocking step is necessary. (4) The digestion should be performed in a minimum volume, as proteases are less active when dilute. The high concentrations used (compared to traditional



solution digestion procedures) are not a problem because proteases are generally separated from most of the peptides by HPLC. If an autolysis product should be isolated, this will be apparent when its sequence is compared with databases. (5) Careful manual collection of the HPLC fractions can improve the results; a bit of practice, and paying special attention to changes in the rate of change of the recorder pen’s movement will be helpful. In general, quite closely spaced peaks will give unique amino acid sequences. In the author’s experience, trypsin is the most reliable enzyme for this procedure. Other enzymes—including Lys-C, Asp-N, and GluC—may also be tried to provide overlapping sequences or in case the tryptic pattern does not provide the desired information. In this event, the digestion buffer recommended by the supplier of the protease, supplemented as above with 10% acetonitrile, should be substituted. The most common problems are spurious peaks, no peaks, and only a trypsin peak in the HPLC analysis. Spurious peaks arise from UVabsorbing contaminants, as discussed above. If a trypsin peak is absent, a control reaction containing the buffer and protease but no protein or nitrocellulose should be injected. If there is still nothing, the trypsin concentration should be checked. If a trypsin peak is present in the control samples, blocking was inadequate or the amount of nitrocellulose without adsorbed protein was too high. If only a trypsin peak is detected, either not enough protein was present or the trypsin was inactive. The latter possibility can be assessed by digesting some standard protein with the same lot of protease.

have yielded high-quality sequence data, allowing determination of as many as two hundred residues from a single experiment. Though the efficiency of the precipitation procedure (support protocol) varies with the sample, it is usual to recover 70% to 100% of samples >1 µg from a volume of 50 µl.

Time Considerations Time required for the SDS-PAGE and transfer procedure: casting the gels—20 min handson time and ∼2 hr total, including deaeration and polymerization; preelectrophoresis—just a few minutes hands-on time and 45 min total; electrophoreses—60 to 80 min; setting up the transfer—15 min; transfer—30 to 90 min; staining—15 min; band excision—5 to 30 min; sample preparation—15 min hands-on time and 1 hr for incubation on dry ice. The entire procedure can be completed in a single day, though it is preferable to cast the gels a day in advance. Time required for determination of internal amino acid sequences following electrophoretic separation and transfer of proteins: destaining and PVP blocking—30 min; washing—5 min/sample; mincing the nitrocellulose—2 to 3 min/sample; digestions—overnight 12 to 24 hr; sample recovery and filtratio n—1 0 min ; r ever sed -p hase HPLC separations—2 hr/sample. A set of samples may be separated on a gel, transferred, and processed for digestion in ∼5 hr. If several proteins from a given gel are of interest, a few days of HPLC will be required; at 2 hr per run, only 3 samples per day can be analyzed.

Literature Cited Anticipated Results

In the first basic protocol, if 100 pmol β-lactoglobulin are loaded onto a 12% gel prepared as described, the initial sequencing signal should indicate a yield of 50 to 80 pmol with gas-phase instruments (e.g., Applied Biosystems 470A). In some cases, usable sequence may be obtained from as little as 10 pmol. In general, if a band is easy to see with Coomassie blue staining, there is enough to sequence, but if it is so faint that excision is difficult, it is doubtful that there is enough protein. However, very faint bands, particularly of low-molecular-weight proteins, have yielded sequence. For N-terminally blocked proteins, if enough material is present in a band or spot, usable sequence from 2 to 3 peptides can be obtained for most proteins. In several cases, two dozen or more HPLC peaks from a single band

Aebersold, R.H., Teplow, D.B., Hood, L.E., and Kent, S.B.H. 1986. Electroblotting onto activated glass. J. Biol. Chem. 261:4229-4238. Aebersold, R.H., Leavitt, J., Saavedra, R.A., Hood, L.E., and Kent, S.B.H. 1987. Internal amino acid sequence analysis of proteins separated by oneor two-dimensional electrophoresis after in situ protease digestion on nitrocellulose. Proc. Natl. Acad. Sci. U.S.A. 84:6970-6974. Hewick, R.M., Hunkapiller, M.W., Hood, L.E., and Dreyer, W.J. 1981. A gas-liquid solid phase peptide and protein sequenator. J. Biol. Chem. 256:7990-7997. Matsudaira, P. 1987. Sequence from picomole quantities of proteins electroblotted onto polyvinylidene difluoride membranes. J. Biol. Chem. 262:10035-10038. Moos, M., Nguyen, N.Y., and Liu, T-Y. 1988. Reproducible, high-yield sequencing of proteins electrophoretically separated and transferred to an inert support. J. Biol. Chem. 263:6005-6008.



Supplement 19

Schägger, H. and von Jagow, G. 1987. Tricine-sodium dodecylsulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 KDa. Anal. Biochem. 166:368-379. Sheer, D.G., Yuen, S.W., Yamone, D.K., Mattaliano, R.J., and Yuan, P-M. 1990. A modified reaction cartridge for sequencing samples on polymeric sample matrices. Abstract T139, Fourth Symposium of the Protein Society, San Diego. Speicher, D.W. 1989. Microsequencing with PVDF membranes: Efficient electroblot-ting, direct protein adsorption and sequencer program modifications. In Techniques in Protein Chemistry (T.E. Hugli, ed.) pp. 24-35. Academic Press, San Diego. Stone, K.L. and Williams, K.R. 1986. High-performance liquid chromatographic peptide mapping and amino acid analysis in the subnanomole range. J. Chromatogr. 359:203-212. Tempst, P. and Riviere, L. 1989. Examination of automated polypeptide sequencing using standard phenylisothiocyanate reagent and subpicomole high-performance liquid chromatographic analysis. Anal. Biochem. 183:290-300. Tempst, P., Link, A.J., Riviere, L.R., Fleming, M., and Elicone, C. 1990. Internal sequence analysis of proteins separated on polyacrylamide gels at the picomole level: Improved methods, applica-

tions and gene cloning strategies. Electrophoresis 11:537-553. Vandekerckhove, J., Bauw, G., Puype, M., van Damme, J., and van Montagu, M. 1986. Protein blotting from polyacrylamide gels onto glass microfiber filters. In Advanced Methods in Protein Microsequence Analysis (B. Wittmann-Liebold, J. Salnikow, and V.A. Erdman, eds.) pp. 179-193. Springer-Verlag, Berlin. Yuan, P., Hawke, D., Blacher, R., Hunkapiller, M., and Wilson, K. 1987. Ethanol precipitation of electroeluted-electrodialyzed samples. Applied Biosystems User Bulletin #27.

Key References Moos et al., 1988. See above. Tempst, et al. See above. Address common problems encountered in procedures of this type.

Contributed by Malcolm Moos, Jr. Center for Biologics Evaluation & Research Food and Drug Administration Bethesda, Maryland




Capillary Electrophoresis of Proteins and Peptides

UNIT 10.20

Capillary electrophoresis (CE) is a high-resolution technique for the separation of a wide variety of molecules of biological interest such as metabolites, drugs, amino acids, nucleic acids, and carbohydrates. This unit focuses on the use of CE to separate proteins and peptides. As with polyacrylamide gel electrophoresis (PAGE; UNIT 10.2), CE separations of proteins and peptides are based on charge-to-mass ratios. While PAGE separations are restricted to polyacrylamide matrices and a relatively small number of buffer systems, CE separations can be achieved in a variety of different matrices using a wide range of electrophoresis buffers. Consequently, there is a much greater flexibility in the design of optimal separation protocols. CE employs a fused-silica capillary column, which may or may not be derivatized, either in free solution or in the presence of a fluid matrix. In contrast to PAGE, the resulting protein or peptide bands cannot be fixed or stained. The CE separation is, in essence, a dynamic one and is more analogous to high-performance liquid chromatography (HPLC) than PAGE. However, CE separation is based on electrophoretic parameters, so it can be viewed as orthogonal to HPLC, which is based predominantly on solvent partitioning parameters. The combination of these two techniques is very powerful for analyzing and characterizing proteins and peptides. The initial expectation that CE would play a major role in protein separations has not been realized. However, CE is particularly useful for determining the purity of a sample and for rapid, efficient, and quantitative evaluation of protein purification. For best separation results, it is necessary to optimize the procedure. If some properties of the protein are known—e.g., its isoelectric point (pI)—the selection of a suitable running buffer is relatively straightforward. In fact, CE itself can be used to determine the isoelectric point of a protein, either in purified form or in a mixture, by focusing the sample in a pH gradient that is generated within the capillary during electrophoresis (Basic Protocol 1). After refocusing, the sample is mobilized either by changing the anode buffer, or by applying a hydrodynamic force to the column to move the separated components past the detector. However, there is little information available that will predict the potential of a protein to interact with the capillary wall. Underivatized silica capillaries have a strong tendency to adsorb proteins, thereby immobilizing them and making protein separations impractical. If such interactions occur, then an additive or a coated capillary column is required. A certain amount of trial and error is involved, and it may be necessary to perform several CE runs to optimize the separation conditions for a given protein. One approach to optimization is described in Basic Protocol 2. CE is most useful for separations of peptides (Basic Protocol 3), because it offers great flexibility in separation parameters. It can be used to monitor proteolytic digestions and optimize digestion conditions for the production of a representative peptide fingerprint of a protein. This profile can subsequently be used to provide structural information about the protein, especially when used in conjunction with reversed-phase HPLC (RP-HPLC; UNIT 10.14). It can be used to screen peptide fractions that are obtained from a preparative RP-HPLC separation of a protease digestion. Fractions that contain single or major components are suitable candidates for protein sequence analysis. Similarly, CE can be used to assess the purity of synthetic peptides. In the presence of an internal standard, it can provide quantitative information about the various components that are present in a peptide mixture. CE can also be used as a micropreparative technique—with either multiple separations that are pooled (Basic Protocol 4) or a single, larger-scale separation Contributed by Dean Burgi and Alan J. Smith Current Protocols in Molecular Biology (2000) 10.20.1-10.20.13 Copyright © 2000 by John Wiley & Sons, Inc.



to data acquisition

detector

temperature control region capillary column

high voltage

buffer/sample

buffer

Figure 10.20.1 Schematic of a CE instrument.

(Alternate Protocol)—for the isolation of peptides from a protease digestion (in much the same way that RP-HPLC is currently used). In most of these examples the same capillary column can be used for all the separations. Only changes in buffer composition, ionic strength, and the presence or absence of additives are required for each specific application. INSTRUMENTATION CE separation in its simplest form can be achieved by passing a high voltage between two buffer reservoirs that are joined by a liquid-filled fused-silica capillary (Fig. 10.20.1). This results in the generation of electroosmotic flow (EOF; see Separation Theory), which allows the molecules of interest to be carried from one end of the capillary to the other. The capillaries are generally 30 to 50 cm long with 50 to 75 µm i.d. The net total volume of these capillaries is in the low microliter range. For comparison, the volume of a slab gel lane is ∼1000 µl. The capillaries are thin-walled; this allows rapid and efficient exchange of the Joule heating that results from the high voltages (20 to 25 kV) that are necessary for electrophoretic separations. This heat dissipation minimizes the negative convective effects that could result in band-broadening during electrophoresis. The fused-silica capillary is coated on the outside with a polymide layer that confers excellent tensile strength to the otherwise fragile capillary. The polyimide sheathing is carefully burned from a small portion of the capillary to expose a section of the silica. This clear section of the capillary is inserted into the light path of a UV detector and becomes the flow cell. As the protein and peptide molecules are swept through the capillary by EOF, they pass through the detector light path and are registered on the UV monitor. In effect the capillary becomes a very-low-volume flow cell. IMPORTANT NOTE: The removal of the polyimide coating makes the capillaries susceptible to breakage. Capillaries that are not provided in cartridges by the manufacturer should be handled with care to avoid breakage. Capillary Electrophoresis of Proteins and Peptides

The combination of high field strength and large surface-to-volume ratios results in rapid and very efficient separations of both proteins and peptides. Sample loading volumes are routinely in the nanoliter range, with starting sample concentrations of ∼0.1 µg/µl for UV detection. On-capillary preconcentration protocols are available when starting concentra-



Table 10.20.1

Commercially Available CE Instruments

Manufacturer

Model

Applied Biosystems Beckman

270-HT

Detection method

Reversal of polarity

UV: 190-700 nm Controlled by software P/ACE 5000 UV: filter; Manual P/ACE 5510 Diode-array: 190-600 nm; P/ACE 5510 Fluorescence: argon laser; MS interface Bio-Rad Biofocus 3000 UV: 190-800 nm Controlled by software Hewlett-Packard HP 3D CE Diode-array: Controlled 190-600 nm by software Waters Quantra 4000E UV: filter Manual

Capillary Cooling of heating sample/buffer

Capillary format

Software available

FAC

Sample

Liquid (Peltier)

Sample/buffer

Free-hanginDA g Cartridge C, DA

Liquid

Sample/buffer

Cartridge

C, DA

FAC (Peltier) FAC (Peltier)

Sample/buffer

Cartridge

C, DA

Sample/buffer

Freehanging

DA

aAll instruments have hydrostatic/electrokinetic sample loading and fraction collecting. bAbbreviations: C, CE system control software; DA, data acquisition; FAC, forced-air convection; MS, mass spectrometry.

tions are below these levels. Sample loading can be achieved either hydrodynamically or electrokinetically. Hydrodynamic loading can be performed by pressure injection, vacuum injection, or gravity injection, e.g., pressure loading at 0.5 lb/in2 for 5 sec. Any of these methods will deliver a small aliquot of sample into the capillary column. Hydrodynamic sample loading has limitations where the capillary contains a viscous matrix or the sample itself has a high viscosity. Electrokinetic loading is performed by applying a low voltage across the capillary column for a fixed time, e.g., 7.5 kV for 4 sec. Sample buffer conductivity, run buffer conductivity, and EOF all play a part in determining the amount of material loaded into the capillary column. In general, samples in high salt buffers should be avoided because buffer ions will carry the charge rather than the sample ions. There are several methods for concentrating the sample onto the capillary column (Burgi and Chien, 1992). All instruments listed in Table 10.20.1 have both sample loading capabilities. Clearly, with respect to sensitivity, speed, and versatility, CE can offer some significant advantages over gel electrophoresis for the separation of proteins and peptides. As shown in Figure 10.20.1, CE can require minimal instrumentation. However, in reality, due to the high voltages that are utilized, safety issues require that the capillary column be incorporated into a dedicated CE instrument. These instruments can provide efficient capillary cooling and online detection of analytes. Table 10.20.1 lists a number of commercially available instruments and some of their characteristics. For the purpose of this discussion, it is assumed that a generic CE system possessing the following capabilities is used: an active capillary cooling system, a UV detector capable of monitoring at 200 nm, a thermostatted autosampler, and a chromatographic data package. Given this basic unit a variety of separations can be performed, depending upon the nature of the proteins or peptides that are being separated. SEPARATION THEORY CE is part of the family of electrophoretic techniques that separate species based upon their sizes and ionic properties. An ion (i) placed into an electric field will move in the direction parallel to the field with a velocity (vi) as follows: vi = µi E



Supplement 34

Here µi is the electrophoretic mobility of species i, and E is the electric field, as defined by the following equation, where V is the voltage and L is the total length from electrode to electrode: E=

V L

The electrophoretic mobility of an ion (µi) is: µi =

q 6 π η ai

where q is the charge on the ion, η is the viscosity of the solution, and ai is the radius of the ion. As seen from these three equations, the movement of the molecule in the column is dependent on the applied voltage, the length of the column, the charge on the molecule, and the size of the molecule. In CE, buffer flow is generated inside the column when the electric field is applied. This flow is from the cathode electrode to the anode electrode in a fused-silica capillary column (from left to right in Fig. 10.20.1); this movement of the buffer is called the electroosmotic flow (EOF). In addition, normal electrophoretic separation occurs whereby a positively charged molecule moves in the same direction as the EOF, a negatively charged molecule moves in a direction opposite to the flow, and a neutral molecule is carried along by the flow. Thus, the total velocity of the molecule (µt) is given by the following equation, where µeo is the EOF in the column. µt = ( µeo + µi ) E

The magnitude of the EOF will vary as a function of the pH of the carrier buffer. In uncoated fused-silica columns, a low pH generates a slow EOF, whereas a high pH generates a fast flow. The flow can be suppressed or completely reversed depending on the coating on the column or organic modifier added to the carrier buffer (Landers et al., 1992; Tsuiji and Little, 1992). Thus, EOF is another parameter that can be optimized to aid in difficult separations. Neutral molecules, because they are all carried along by the flow, can be difficult to separate. Addition of a micelle to the carrier buffer results in the partitioning of molecules between the micelle and carrier buffer and can effectively resolve neutral molecules (Khaledi, 1994). STRATEGIC PLANNING Proteins can be separated on coated or uncoated capillary columns, and the choice of separation protocol depends on the specific properties of the target protein. The most important property is the pI, which can be determined by either conventional gel electrophoresis or CE. The use of a buffer ∼2 pH units above the pI is optimal for CE separations. Table 10.20.2 lists a number of buffers and their pH ranges. The presence of high salt concentrations in the sample can interfere with the separation process, and dialysis or dilution may be required. Protein concentrations of microgram per microliter are optimal for CE separations; however, on-capillary concentration protocols exist that allow the separation of proteins with concentrations 1 to 2 orders of magnitude lower.


Peptides can be effectively separated in open-tube fused-silica capillary columns. The EOF generated within the capillary causes separation of both charged and neutral peptides. The respective migration times are dependent upon both the pH of the electrophoretic separation and the charge-to-mass ratios of the peptides. For example, at low pH, peptides with net positive charge migrate towards the anode faster than neutral or negatively charged peptides. Even negatively charged peptides are swept towards the anode by EOF.



Table 10.20.2

Useful Electrolytes for CE of Peptides

Electrolyte

Useful pH range

Phosphate Citrate Acetate MESa Phosphate Tris Borate

1.14-3.14 3.06-5.40 3.76-5.76 5.15-7.15 6.20-8.20 7.30-9.30 8.14-10.14

Minimum useful wavelength (nm) 195 260 220 230 195 220 180

aAbbreviation: MES, 2-(N-morpholino)ethanesulfonic acid.

In practice this migration order is further modified by the mass of the peptides, with small positively charged peptides having faster migration rates. The choice of buffer for CE is of primary importance. The use of buffer systems that separate at high pH significantly alters the migration positions of peptides relative to those of low-pH separations. The selection of a specific buffer is dictated both by its buffering capacity at a selected pH and by its minimum usable UV absorbance wavelength. Characteristics of some useful buffer systems are shown in Table 10.20.2. Small differences in the pH or composition of the buffer can have a significant impact on the absolute mobilities of the peptides, so it is advisable to adjust the pH of buffers accurately and reproducibly. Alternatively, a suitable internal standard can be incorporated into the separation in order to obtain relative mobilities. The protein can be solubilized in water or 40 mM buffer in the presence or absence of urea. The sample buffer does not need to be the same composition as the running buffer, however, use of ionic detergents is not recommended. For the separation of peptides from a tryptic digest of a protein, where almost every peptide carries at least two positive charges, a phosphate buffer of pH 2.0 to 3.0 is commonly used. Likewise, for V8 protease digestions (which cleaves C-terminal to glutamic acid residues), the preferred separation conditions use a borate buffer with a pH range of 8.0 to 9.0. SEPARATION OF PROTEINS BY ISOELECTRIC FOCUSING Isoelectric focusing (IEF) separations can be performed readily by CE; this can be a useful first step in selecting subsequent buffer systems for protein separations. Proteins and peptides are amphoteric in nature, so their charge is dictated by the surrounding carrier buffer. When a protein or peptide is placed in an electric field, it moves to a region where the surrounding pH equals its isoelectric point (pI); at this point the molecule stops moving. A pH gradient is generated in a coated capillary column by filling the column with a sample solution that contains ampholytes. A high-pH solution (sodium hydroxide) is placed in the cathode reservoir and a low-pH solution (phosphoric acid) is placed in the anode reservoir. An electric field is then applied across the column, and the ampholytes and the sample move to a location in the column corresponding to their respective pIs. If the molecule drifts out of the isoelectric region, a charge is induced by the surrounding pH solution, and the molecule moves back to the region of zero charge. Narrow regions are formed with pI resolutions on the order of 0.01 pH units. To determine the pI of a particular protein in the column, markers of known pIs are added to the sample mixture, and extrapolation between markers yields the pI of the protein of interest. After separation of the sample in the absence of EOF, mobilization of the proteins past the detector is necessary. A simple replacement of the anode reservoir with the buffer in

BASIC PROTOCOL 1



Supplement 34

the cathode reservoir establishes EOF and causes the sample zone to migrate pass the detector. The separation discussed here assumes there is no EOF in the column during focusing. However, separations may also be done in the presence of EOF (Mazzeo and Krull, 1992). Materials Ampholyte mixture, pH 3 to 10 (Bio-Rad) Sample containing 0.5 to 1.0 mg protein/ml in water IEF markers (Bio-Rad; optional) 20 mM sodium hydroxide (NaOH; store at 4°C) 10 mM phosphoric acid (store at 4°C) 50-µm-i.d. coated silica capillary column CE instrument (see Table 10.20.1) 1. Add ampholyte mixture to 0.5 ml protein sample to give a final concentration of 2.5% (v/v) ampholytes. If desired, add IEF markers to a final concentration of 0.1 mg/ml to calibrate the column.

2. Add the solution to the sample reservoir. Fill the capillary by pressurizing the reservoir (0.5 lb/in2). 3. Place 10 mM phosphoric acid in the anode reservoir (negative end). Place 20 mM NaOH in the cathode reservoir (positive end). 4. Focus the sample 4 to 6 min at 8 to 10 kV constant voltage. Monitor the current until it reaches steady state. The instrument should be operated in accordance with the manufacturer’s instructions.

5. Mobilize the sample by placing 20 mM NaOH in the anode reservoir and setting the voltage to 10 kV. Monitor the mobilization of the proteins past the detector 15 to 20 min. Another method for mobilizing the proteins past the detector is to apply a hydrodynamic force to one end of the column and push the zones past the detector. Commercially available CE instruments have this capacity as an option. To minimize distortion of the pI regions, a constant electric field (10 kV) must be applied at the same time. The pressure applied is small (0.5 lb/in2) and is maintained for the entire time of mobilization.

6. Wash the column after each run with 10 mM phosphoric acid for 1 min at 0.5 lb/in2. Store the column at room temperature in running buffer (short-term) or water (long-term). BASIC PROTOCOL 2


SEPARATION OF PROTEINS To use CE for protein analyses, the more that is known about the particular protein, the higher the success rate for developing a separation method. Knowing the pI of a protein permits selection of the appropriate run buffer to maximize differences in the charge-tosize ratio. IEF separation performed using CE (Basic Protocol 1) can be a useful first step in selecting subsequent buffer systems for protein separations on an underivatized capillary column. If no protein is detected, it has probably adsorbed to the silica surface of the column. Repeating the separation in the presence of an ionic detergent such as SDS will coat the protein with negative charge and prevent adsorption. However, this means that proteins will separate on the basis of size alone. Alternatively, if the pI of the protein is unknown, the separation can often be achieved by using a coated capillary column in the presence of high-pH, high-ionic-strength buffer.



Materials Sample containing 10 mg protein/ml in water 50 mM and 500 mM sodium borate, pH 8.0 to 9.5 (for separation of proteins with unknown pI) 5 mM and 50 mM buffer at pH >pI (for separation of proteins with known pI; see Table 10.20.2) 50-µm-i.d. coated (unknown pI) or uncoated (known pI) fused-silica capillary column CE instrument (see Table 10.20.1) For separations of proteins with unknown pI 1a. Dilute protein sample 1/10 (v/v) with 50 mM sodium borate buffer to give a final concentration of 1.0 mg/ml. Alternatively, dialyze sample (at a concentration of 1 mg/ml) in 50 mM sodium borate buffer.

2a. Fill a coated column with 500 mM sodium borate buffer. 3a. Inject the sample hydrodynamically, e.g., 3 sec at 1 lb/in2. Consult the manufacturer’s instruction manual for the proper procedure.

4a. Separate using the following conditions: Detector wavelength: 200 nm Temperature: 25°C Run voltage: 10 kV Run time: 30 min. For separations of proteins with known pI 1b. Dilute protein sample 1/10 (v/v) with a 5 mM buffer that has a pH above the pI of the protein to induce a charge on the protein (final protein concentration = 1.0 mg/ml). Alternatively, dialyze sample (at a concentration of 1 mg/ml) in 5 mM buffer.

2b. Fill an uncoated capillary column with the same buffer at 50 mM. 3b. Inject the sample hydrodynamically, e.g., 3 sec at 1 lb/in2. Consult the manufacturer’s instruction manual for the proper procedure.

4b. Separate using the following conditions: Detector wavelength: 200 nm Temperature: 25°C Run voltage: 25 kV Run time: 30 min. If the protein of interest is not detected, add 10 mM SDS to the run buffer to induce a charge on the molecule. If the peaks are too broad, add 0.1% (w/v) methylcellulose to coat the column and reduce adsorption of the molecule onto the wall of the column. The addition of SDS and/or methylcellulose will increase the run time of the separation.



Supplement 34

BASIC PROTOCOL 3

ANALYTICAL PEPTIDE SEPARATIONS CE is particularly useful for separating specific peptides from complex mixtures such as the products of proteolytic digestion. In this procedure, the peptide mixture to be analyzed is placed in a sample vial at the cathode end of the capillary column. The anode reservoir contains running buffer. The sample is loaded into the capillary either electrokinetically or hydrodynamically by pressure or vacuum, then the sample vial is replaced by a vial containing running buffer and electrophoresis begins. The following protocol describes the use of a P/ACE 5000 CE instrument (Beckman) using low-pressure sample injection. However, other instruments (see Table 10.20.1) are capable of similar separations when operated according to the manufacturer’s instructions. Materials Peptide mixture: e.g., tryptic digest of β-lactoglobulin (Applied Biosystems) 0.05 M and 0.25 M sodium phosphate buffer, pH 2.30 (store at 4°C) 0.1 M sodium hydroxide (NaOH) 75-µm-i.d. fused-silica capillary column (Beckman) CE instrument (e.g., P/ACE 5000, Beckman, or equivalent; see Table 10.20.1) 1. Precondition the capillary by flushing with the following solutions: 10 column volumes of 0.1 M NaOH at low pressure (0.5 lb/in2) 10 column volumes water 4 column volumes of 0.25 M sodium phosphate buffer, pH 2.30. Store the column in 0.25 M sodium phosphate buffer, pH 2.30, at 25°C. When changing to a different separation buffer, equilibrate the column 4 hr with the new buffer (Strickland and Strickland, 1990). If a new capillary is being used, it is essential that this preconditioning step be performed prior to attempting separations.

2. Prepare the peptide mixture by dissolving 10 nmol (∼300 µg) in 10 ml of 0.05 M sodium phosphate buffer, pH 2.30. Freeze unused mixture in 100-µl aliquots. 3. Apply 10 to 20 nl sample using low-pressure injection for 10 sec at 0.5 lb/in2. Consult the manufacturer’s manual for the proper procedure.

4. Separate the peptide mixture using the following conditions: Electrolyte: 0.05 M sodium phosphate buffer, pH 2.3 Detector wavelength: 200 nm Temperature: 25°C Voltage: 25 kV. Consult the manufacturer’s manual for proper operating conditions.

5. Rinse the column with the following solutions: 0.5 min with water 1.0 min with 0.1 N NaOH 1.5 min with water 1 min with 0.25 M sodium phosphate buffer, pH 2.30. Store the column in running buffer or water at room temperature. Capillary Electrophoresis of Proteins and Peptides

An example of a separation protocol for a Beckman P/ACE 5000 instrument with a 37-min run time is described in Table 10.20.3.



Table 10.20.3

Operating Program for Peptide Analysisa

Step

Process

Duration

Inletb

Outletb

Control summaryc

1 2 3 4 5 6 7 8 9 10 11

Set Temp Set Detector Rinse Rinse Rinse Inject Separate Rinse Rinse Rinse Rinse

— — 1.0 min 1.5 min 0.5 min 10.0 sec 30.0 min 0.5 min 1.0 min 1.5 min 1.0 min

— — 33 29 29 11 29 33 32 33 34

— — 7 7 9 9 9 7 7 7 9

Temp 25°C UV 200nm Forward: HP Forward: HP Forward: HP Low pressure Voltage 25.00 kV Forward: HP Forward: HP Forward: HP Forward: HP

aFor analysis of proteolytic digests, HPLC fractions, and synthetic peptides. bVial contents: 7, waste vial (water level just to contact capillary effluent); 9, electrolyte (50 mM sodium phosphate buffer,

pH 2.30); 11, sample; 29, electrolyte (50 mM sodium phosphate buffer, pH 2.30); 32, regeneration solution (0.1 N NaOH); 33, water; and 34, 0.25 M phosphate buffer, pH 2.30. cAbbreviation: HP, high pressure.

MICROPREPARATIVE CAPILLARY ELECTROPHORESIS: MULTIPLE SEPARATIONS

BASIC PROTOCOL 4

Although CE has been most commonly used for analytical separations, considerable interest has developed in using its high resolving power in micropreparative applications. Two basic approaches have evolved, one utilizing multiple separations and collections from an analytical capillary (Bergman and Jörnvall, 1992) and the other utilizing a single separation and collection on a much larger-diameter (150-µm-i.d.) capillary (see Alternate Protocol). For the multiple collection approach to be effective, the elution times of the peptides must be reproducible. The following method uses the P/ACE 5000 capillary electrophoresis instrument to separate a mixture of peptides. Materials Peptides: ACTH 4-10, angiotensin I, and angiotensin II (Sigma) 0.05 mM and 0.25 mM sodium phosphate buffer, pH 2.30 (store at 4°C) 0.1 M sodium hydroxide (NaOH) 75-µm-i.d. fused-silica capillary column (Beckman) CE instrument (e.g., P/ACE 5000, Beckman, or equivalent; see Table 10.20.1) Conical microvials (Beckman) 1. Precondition the capillary column by flushing with the following solutions at low pressure (0.5 lb/in2): 10 column volumes of 0.1 M NaOH 10 column volumes of water 4 column volumes of 0.25 M sodium phosphate buffer, pH 2.30. Store in 0.25 M sodium phosphate, pH 2.30. When changing to a different separation buffer, equilibrate the column 4 hr with the new buffer (Strickland and Strickland, 1990).

2. Prepare a peptide mixture by dissolving 0.2 mg each of ACTH 4-10, angiotensin I, and angiotensin II in 1 ml of 0.05 M sodium phosphate buffer, pH 2.30 (final concentration = 0.6 mg peptide/ml). Store 100-µl aliquots at −20°C. 3. Load 10 to 20 nl peptide mixture by low-pressure injection for 10 sec at 0.5 lb/in2.



Supplement 34

Consult the manufacturer’s manual for the proper procedure.

4. Separate the mixture using the following conditions: Electrolyte: 0.05 M sodium phosphate, pH 2.30 Detector wavelength: 200 nm Temperature: 25°C Run voltage: 25 kV Fraction size: 3-min per collection vial. Consult the manufacturer’s manual for proper operating conditions.

5. Replace the standard outlet reservoir (anode) with a series of conical microvials, each containing 10 µl of 0.05 M sodium phosphate buffer, pH 2.30. Collect 3-min fractions into each of the vials for the length of the separation. 6. Repeat the injection and separation (steps 3 to 5) four times and combine the contents from the corresponding fraction numbers. It will take 2 to 3 hr to complete the fractionation.

7. Screen fractions for peptide content using the analytical separation previously described (see Basic Protocol 3). ALTERNATE PROTOCOL

MICROPREPARATIVE CAPILLARY ELECTROPHORESIS: SINGLE SEPARATION In this micropreparative protocol, a larger quantity of the peptide mixture is loaded onto a 150-µm-i.d. capillary column and the fractions are collected from a single separation (Kenny et al., 1993). For the single-collection approach the CE instrument must be able to effectively control the capillary temperature at the elevated power levels that are required for electrophoresis. Forced air cooling is generally inadequate for this purpose. Additional Materials (also see Basic Protocol 4) 4:1 (v/v) 0.5 M sodium phosphate buffer (pH 2.50)/ethylene glycol 150-µm-i.d. fused-silica capillary column (Polymicro) 1. Prepare peptide mixture and precondition the column (see Basic Protocol 4, steps 1 and 2). 2. Load 0.1 µl peptide mixture by low-pressure injection for 10 sec at 0.5 lb/in2. Consult the manufacturer’s manual for the proper procedure.

3. Separate the peptide mixture using the following conditions: Electrolyte: 0.05 M sodium phosphate buffer, pH 2.30 Detector wavelength: 200 nm Temperature: 25°C Run voltage: 7.5 kV Fraction size: 3-min per collection vial. Consult the manufacturer’s manual for proper operating conditions.

4. Replace the standard outlet reservoir (anode) by a series of conical microvials each containing 10 µl of 4:1 (v/v) 0.5 M sodium phosphate buffer/ethylene glycol. Collect 3-min fractions into each vial for the length of the separation. It will take ∼2 hr to fractionate the sample. Capillary Electrophoresis of Proteins and Peptides

5. Screen fractions for peptide content using the analytical separation protocol (see Basic Protocol 3).



COMMENTARY Background Information Protein separation is one of the more difficult types of separation to perform by capillary electrophoresis (CE; Novotny et al., 1990). Each protein has it own set of optimal separation conditions, and one optimized protocol will not necessarily transfer well to the separation of a different protein. The more that is known about the properties of the molecule, the easier it will be to optimize the separation conditions. The most important property is probably the pI, because it will dictate the selection of pH and ionic strength conditions for the initial separation attempt. However, a variety of other properties must also be taken into account when optimizing a separation protocol: shape (globular or fibrous), aggregation tendencies, solubility in dilute solutions, salt requirements, thermal stability, and hydrophobic nature. Several parameters can be varied within CE separations in order to utilize this knowledge—e.g., temperature, buffer/salt selection, ionic and nonionic detergents, and matrix additives (polyacrylamide, methylcellulose). Several protocols have been developed that can serve as useful starting points for most protein separations. Table 10.20.4 lists references for some groups of proteins. Furthermore, many of the manufacturers listed in Table 10.20.1 have developed additional protocols for protein separations. Ultimately, the selection of an appropriate separation protocol for a protein depends on the specific properties of that protein. However, there are separation approaches that utilize specific properties of proteins. IEF-CE separates on the basis of charge alone; CE in the presence of ionic detergents such as SDS separates on the basis of size alone; and CE using underivatized capillaries at low pH or derivatized capillaries at high salt and high pH separates on the basis of both size and charge. Successful CE separations also depend on the behavior of

Table 10.20.4

other contaminants that are present in the mixture. It may also be possible to take advantage of the knowledge gained from previous purification steps, e.g., ion-exchange or size-exclusion chromatography, to select CE separation conditions. Analytical-scale separation of peptides by CE has found application in a variety of areas. Proteolytic digestions can be monitored by placing the digestion mixture in a sample vial on the sample table that is equilibrated to 37°C. The instrument can then be programmed to analyze an aliquot at desired intervals, such as every 4 hr. Digestion is complete when a stable CE profile is obtained. Also, fractions collected from an HPLC separation of a protease digest can readily be screened orthogonally by CE to determine the homogeneity of any given peak. Fractions with a single major component can then be subjected to protein sequence analysis. The sensitivity of UV detection at 200 nm is such that digest levels of ≥100 pmol can be effectively screened by this method. Below these load levels the fractions have to be reduced in volume or concentrated on the capillary to achieve a sufficiently high concentration for visualization with UV detection (Dolnik et al., 1990; Burgi and Chien, 1992). Selection of an appropriate micropreparative protocol will depend on the type of instrumentation that is available. The single-separation micropreparative protocol will only work with a CE instrument that has adequate cooling capabilities. The multiple-separation micropreparative protocol will work on any CE instrument that has reproducible and stable electrophoretic mobilities.

Critical Parameters For protein analysis, the charge on the protein is the most important parameter for separation. However, the charge-to-shape ratio also plays a role in effective separation of proteins.

References for Separations of Different Protein Types

Protein type

Reference

Basic proteins

Wiktorowicz and Colburn (1990); Bullock and Yuan (1991) Tran et al. (1991); Tsuiji and Little (1992) Nielsen et al. (1991) Chen et al. (1992)

Glycoproteins Antibody-antigen complexes Milk proteins



Supplement 34

The charge on the protein is controlled by the buffer and the additives used for the run. The shape is determined by the denaturing conditions of the solutions. The surface charge on a protein may ultimately dictate the extent of wall interactions in addition to electrophoretic mobility, whereas shape has its greatest effect upon mobility alone. The recovery of peptides from underivatized silica capillary columns appears to be very good—even at subpicomole levels. However, proteins are frequently irreversibly bound to these same capillaries. Clearly, at some point a “peptide” must become a “protein” in the context of CE. There is little published data available to clarify this issue, although good recoveries of peptides with 30-40 amino acid residues in length have been obtained from underivatized capillary columns. If the recoveries of higher molecular weight peptides are poor, then the separation should be repeated using a coated capillary column. During peptide synthesis, single-amino-acid-deletion products are frequently produced. The properties of these deletion products are very similar to those of the expected species, so the deletion products may not be readily resolved by reversed-phase HPLC. In this case, CE has proven to be a very useful orthogonal technique because of its high resolution power and separation based on charge-to-mass ratios. However, it must be emphasized that the effective separation and detection of proteins and peptides requires relatively high solute starting concentrations. When using uncoated columns, a sodium hydroxide rinse is required to maintain the capillary surface. However, this rinse is never used with coated capillaries because it will remove the coating. Commercially available coated columns are satisfactory, but linear polymers (e.g., methylcellulose) have been used with more success (Palmieri and Nolan, 1994). If a protocol can be worked out for the molecule of interest, rapid CE analysis with little loss of starting material can augment the tools used now for protein analysis (e.g., HPLC).

Troubleshooting


Mechanical and electrical problems that might be encountered during instrument use are addressed in the troubleshooting sections of the manufacturers’ manuals. One of the common difficulties is the loss of electrical contact during a run. This loss of current can be caused if a small bubble forms in the column during

injection or if the power generated by the run is so great that the solution boils or outgasses. Purging the column after a failed run removes any bubbles in the column. The sample vial must contain enough sample to cover the end of the column during injection to prevent injection of air into the column. The sample vial can dry out during a long run, so sample temperature control or the use of the proper cap on the vial is required to slow evaporation. Reducing the buffer concentration or the run voltage can eliminate bubble formation during a run. Degassing the run buffer is also useful if the outgassing problems continue. The instrument sometimes arcs during a run; this is caused by salt deposits or moisture on the high-voltage electrode stations. Particular attention should be given to keeping the electrode stations salt free. Moisture buildup can be minimized by reducing the separation voltage by 5 kV. Frequent visual inspection is also advised. The column can plug up because of salt deposition if the column dries out. Washing the column with water dissolves the plug; storing the column in water, if it is not going to be used for a long period of time, prevents plugging. Proteins can precipitate during a run and plug the column. Using lower protein concentrations prevents this type of plugging.

Anticipated Results Electrophoretic profiles for both protein and peptide separations should be highly reproducible. However, absolute migration times may vary from run to run, especially if the separation temperature is not adequately controlled. For this reason, the use of an appropriate internal standard is highly recommended (see the manufacturer’s instructions). At load levels 100 kDa were being routinely analyzed with subpicomole sensitivity (Hillenkamp et al., 1991) and molecular weights could be assigned with an accuracy of ∼0.1%. Beavis and Chait (1990) demonstrated that for small proteins (30 kDa that differ only by an initiator Met residue



Supplement 38

Overview of Mass Spectrometry

(e.g., see Fig. 10.21.8). These issues are considered in more detail in the discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution. Several important considerations relating to the production of ions in the MALDI source limit the resolving power of linear TOF instruments. The most important of these is the initial kinetic energy spread of individual ion populations: as a result of this spread, ions of any particular mass will, after reaching final accelerating voltage, exhibit a spread in velocities. This means that various members of the same ion population will arrive at the detector at slightly different times, and that in turn gives rise to a broader peak for each ion than would be observed if there were no initial kinetic energy spread. Because the initial energy spread is mass dependent, peaks for highermass ions will be disproportionally broader than those for lower-mass ions. Resolution in a linear TOF instrument can be improved by increasing the length of the flight tube. This enhances the time dispersion of ions of different m/z; unfortunately, it also increases the spread of arrival times for ions of the same m/z due to the initial kinetic energy distribution. This energy spread is typically reduced in magnitude by increasing the final accelerating voltage. A more effective way to correct for energy distributions is through the use of an ion mirror, or reflectron. A reflectron TOF instrument (Fig. 10.21.2, panel B) corrects for the initial energy spread by acting as an energy-focusing device. The reflector works by slowing an ion down until it stops (the back of the reflector is at a voltage slightly higher than the source-accelerating voltage), turning it around, and then reaccelerating it back out to a second detector. Ions with an initial kinetic energy (and velocity) slightly lower than full accelerating potential will not penetrate the reflector as deeply, and will therefore turn around sooner, catching up to those ions with full kinetic energy. Ions with slightly greater energies (and thus higher velocities) will penetrate more deeply into the reflector, be turned around later, and have their flight times retarded, allowing the other ions to catch up. All of this causes ions of a given mass-to-charge ratio to be spatially focused into packets having flight times that are closer together, thus improving the resolution. This improvement in resolution is illustrated in Figure 10.21.3, which shows the molecular ion region of the linear (panel A) and reflector (panel B) spectra of a synthetic tyrosine phosphorylated peptide TRDIYETDpYYRK. The

isotopically resolved ion cluster in the reflector spectrum shows the contributions of all of the isotopes of C, H, N, O, and P that make up the elemental composition of this peptide. The improvement in resolution is most noticeable at masses of ∼3000 and lower. For larger molecules, where the resolution is no longer sufficient to provide separation of the isotopes, the natural width of the unresolved isotopic envelope has a large influence on the apparent resolution of the instrument, as explained in the discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution. In addition to resolution, mass accuracy is also improved when spectra are recorded in the reflector mode (Vorm and Mann, 1994). Tremendous improvement in the resolution achievable on MALDI-TOF instruments has recently been demonstrated using a focusing technique developed many years ago by Wiley and McLaren (1953). This method, referred to as time-lag focusing or “delayed extraction,” has been shown to provide a resolution of ∼2000 to 4000 (full-width, half-maximum; or FWHM) in linear mode for peptides and resolution of ∼3000 to 6000 FWHM in reflecting mode, on a 2-meter-long instrument (Brown and Lennon, 1995; King et al., 1995; Vestal et al., 1995; Whittal and Li, 1995). In a delayed extraction source, ions are created in a field-free region and allowed to spread out before an extraction voltage is applied and the ions are accelerated into the drift tube. The delayed extraction also probably limits the number of collisions the ions undergo on their way out of the source, thereby reducing additional peak broadening caused by metastable decomposition. Mass accuracy with the use of an internal reference standard may also be improved with the use of the delayed extraction source. Assigning an accurate m/z value to an ion whose flight time has been measured requires that the mass spectrometer be properly calibrated. This is usually accomplished by measuring the flight times of two compounds whose molecular weights are accurately known. In practice, it is usually best to select two compounds with molecular weights that bracket the mass range over which samples are to be examined. For molecules >10,000 Da, the [M+H]+ and the [M+2H]2+ ions of a well-characterized protein standard work very well for calibration. Application of an internal calibration, where the calibrant peaks are present in the same spectrum as the sample, can provide accuracies of 0.01% to 0.05% (e.g., 0.1 to 0.5 Da at m/z = 1000). Internal calibration is sometimes experi-



A

1690 1695 1700 1705 1710 1715

Figure 10.21.3 Molecular ion region from MALDI spectra of the peptide TRDIYETDpYYRK (monoisotopic Mr = 1701.8) recorded in the (+) ion mode. Spectra were recorded in (A) the linear mode, where the average mass is measured at the centroid of the unresolved isotopic cluster, and (B) the reflector mode, where the monoisotopic mass is measured at the centroid of the first peak in the cluster. This peak contains the lightest isotopes of the elements present (see discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution).

B

1690 1695 1700 1705 1710 1715 m /z

mentally difficult owing to the need to adjust the amount of calibrant compounds added so that the calibrant peaks and the analyte peaks are simultaneously observed with reasonable intensity. MALDI mass spectra may also be calibrated externally; this approach is experimentally simpler. In this method calibrant compounds are measured using the same matrix and laser irradiance as were used for sample analysis. A mass accuracy of ∼0.1 to 0.2% (e.g., ±1 to 2 Da at m/z = 1000) can be achieved in linear mode using external calibration (Karas et al., 1989). In reflector mode, one can routinely expect mass accuracies on the order of 0.01% to 0.05% using external calibrations. In reflector mode it is also possible to use either the [M+H]+ ion or the [2M+H]+ ion (the protonated gas-phase dimer) of the matrix as one of the two calibrant peaks, which means that only one calibrant compound needs to be added.

MALDI Post-Source Decay MS for Peptide Sequencing A peptide molecular ion that is sufficiently stable to be transported out of the ion source

but insufficiently stable to survive the flight to the detector will decompose in the flight tube, giving rise to a series of fragment ions that are characteristic of the molecule’s original structure. This process is referred to as metastable decay or post-source decay. It is believed that molecular ions acquire excess internal energy necessary for fragmentation via multiple collisions with matrix ions in the source (Kaufmann et al., 1994). When a precursor ion decomposes in the flight tube, all of the fragment ions will have the same velocity as the precursor, but they will have only a fraction of its kinetic energy. This means that the precursor and metastable fragments will all strike the linear detector at the same time and will be detected at the same apparent mass. Thus metastable or post-sourcedecay fragment ions are never observed in a linear spectrum. In contrast, a reflector TOF is capable of discriminating fragment ions by flight-time dispersion. Although all ions will have essentially the same velocity, the kinetic energy of a post-source-decay fragment ion will be determined by the ratio of its mass to the mass of the precursor. Thus the fragments, which have less



Supplement 38

Relative abundance (%)

A 100

2066

1284

50

804

1300

634

1705 1776 1550

1002 1250 1082

530

2168 2384

2705 2937

0


B

600

800 1000 1200 1400 1600 1800 2000 2200 2400 2600 2800

100 2066 y3 50

15 14 13 12 11 10 9

y4

8

7 6

5

4 3 2 1

I–F–Y–P–E–I–E–E–V–Q–A–L–D–D–T–E–R

yn

PEIE

y1 PEI y2

PEIEE y6 y5

y7

600

800

y8

y9 y10

y11

y12

y14

y15

0 200

400

1000 1200 m /z

1400

1600

1800

2000

Figure 10.21.4 MALDI-MS analysis of a tryptic digest of an 18-kDa protein (Ladner et al., 1996). A total of 30 pmol of protein was separated on an SDS-PAGE gel, the band of interest excised from the gel, and the protein digested in situ with trypsin; then 1⁄20 of the sample was analyzed by MALDI-MS. (A) MALDI mass spectrum of the tryptic peptides recorded in the linear mode; (B) MALDI-PSD spectrum of the [M+H]+ ion at m/z 2066. The peak was selected using an ion gate (R = 100). A nearly complete series of yn ions (see Fig. 10.21.1) makes it possible to deduce the sequence of residues 3 to 17.


kinetic energy than the precursor, will not penetrate as deeply into the ion mirror, and so will have their flight times decreased relative to the precursor. The fragments are therefore detected at a lower mass. By successively lowering the reflector voltage, thereby bringing lower- and lower-mass fragment ions into focus, a complete post-source decay fragment ion spectrum can be acquired in segments. The individual segments are calibrated and assembled afterward automatically by the data system. The energy-focusing features of reflectrontype mass spectrometers make it possible to use post-source decay for peptide sequencing (Kaufmann et al., 1994). Post-source decay spectra resemble low-energy collision-induced dissociation (CID) spectra, similar to those commonly recorded on triple-quadrupole mass spectrometers (Hunt et al., 1986). The spectra contain mainly bn and yn ions, often with very abundant internal fragments and immonium ions (see Fig. 10.21.1 and Table 10.21.2). The

use of an ion gate makes it possible to perform a low-resolution (R = 100) selection of a precursor ion of interest from an unfractionated mixture, such as a protein digest. This approach is very powerful for providing internal sequence information on in situ–digested proteins that have been purified by SDS-PAGE (see Fig. 10.21.4 for an example).

ESI-MS In electrospray ionization (ESI)-MS, ions are formed from peptides and proteins by spraying a dilute solution of these analytes at atmospheric pressure from the tip of a fine metal capillary (Fig. 10.21.5). The spray process, often assisted by pneumatic nebulization, creates a fine mist of droplets (Fenn et al., 1990; Kebarle and Tang, 1993). The droplets are formed in a very high electric field created by applying a high voltage (∼4 kV) to either the spray tip (Fig. 10.21.5) or the counter electrode; in this process, the droplets become highly



possible sample inlets: syringe pump sample injection loop HPLC

triple-quadrupole mass spectrometer

atmosphere (760 Torr)

vacuum (10 – 6 Torr)

liquid 1st quadrupole collision 2nd quadrupole mass analyzer cell mass analyzer

capillary electrophoresis

data system expansion of the ion formation and sampling regions

electrospray needle

nitrogen drying gas vacuum

atmosphere

3–5 kV

liquid nebulizing gas

droplets containing solvated ions

ions sampling orifice counter electrode

Figure 10.21.5 Schematic diagram of a triple-quadrupole mass spectrometer (top) and the electrospray interface (bottom).

charged. Solvent evaporation is rapid from these small droplets. As the droplets evaporate, the peptide and protein molecules in the droplets pick up one, two, or more protons from the solvent to form singly or, more frequently, multiply charged ions (e.g., [M+H]1+, [M+2H]2+, etc.). The number of charges acquired by a molecule is roughly equivalent to the number of possible sites of proton attachment. As the droplets continue to shrink, the charge density on the surface of each one increases to the point where charge repulsion overcomes the forces holding together the droplet and the solvated ions contained within it. Ions are then “emitted” or “evaporated” from the droplet surface. The ions are sampled into the high-vacuum region of the mass spectrometer for mass analysis and detection, most often using a quadrupole (or triple-quadrupole) mass analyzer (Fig. 10.21.5). Usually little or no fragmentation is observed in the normal ESI mass spectra of peptides. The typical solvent for peptides and proteins is a mixture of water, an organic modifier such as CH3CN, and up to a few percent by volume of acetic, trifluoroacetic, or another volatile acid (the latter included to enhance ionization of sample constituents). Because the ions are produced at atmospheric pressure from flowing liquid

streams, ESI is ideally suited for on-line coupling to high-performance liquid chromatography, making it possible to analyze mixtures of peptides and proteins rapidly (see below). As mentioned above, a key feature of the electrospray process is the formation of multiply charged molecular species from analytes that contain more than one possible site of proton attachment (Fenn et al., 1990; Smith et al., 1991). Proteins usually exhibit a characteristic series of multiply charged ions (e.g., see Fig. 10.21.8). As one proton (i.e., one charge) is typically attached for each 1000 Da of molecular mass, the ion series for a protein, even a large protein, will fall in the m/z range of 800 to 3000. This makes it possible for simple instruments like quadrupole mass spectrometers to be used for mass analysis of ions produced by ESI. The molecular mass of the protein can easily be calculated using the observed masses of any two adjacent ions in the series (Covey et al., 1988; Mann et al., 1989; Reinhold and Reinhold, 1992). Normally these calculations are performed automatically by the data system. Because each multiply charged peak provides an independent measure of the molecular mass of the protein, an ion series from a single experiment provides a measure of the mass measurement precision. Most data sys-



Supplement 38


A

100 75

2+ 673.8

2+ (– H, +Na) 2+ (– H, +K)

50

1+ 805.4

25

1+ 1346.6 600

B Relative abundance (%)

1000

800 b3 y2

100

y4 75 50

b2

y6

1200 2+

10 9 8 7 6 5 4 3 2

D G D G pY I S A A E L R

y7

b5

bn

2 3 4 5 6 7 8 9 10

y3 b7 b4

200

yn

y5

y1

25

1400

400

b9

b6

600

y8 b 10 y9

b8 800

y10 1000

1200

1400

m /z

Figure 10.21.6 Electrospray analysis of an HPLC fraction from the tryptic digest of a 17-kDa phosphoprotein. The protein was purified by SDS-PAGE, electroblotted onto PVDF membrane, eluted, digested with trypsin, and the peptides analyzed by on-line LC-ESI-MS using a phosphopeptide-selective scanning method (Huddleston et al., 1993b). The column flow was split after UV detection, with 10% going to the mass spectrometer and 90% to a fraction collector. (A) Nanoelectrospray mass spectrum of a putative phosphopeptide-containing fraction. Peaks labeled (+Na) and (+K) correspond to [M + H + Na]2+ and [M + H + K]2+, respectively. (B) Collision-induced decomposition (CID) spectrum of the doubly charged ion ([M+2H]2+) at m/z = 673.8. This spectrum confirms the sequence of the peptide and identifies the site of phosphorylation as Tyr5.


tems also provide algorithms for transforming the mass-to-charge axis into a molecular mass axis for simplifying interpretation and presentation of the data. Proteins with molecular masses of up to 150,000 Da have been successfully analyzed by ESI using commercially available quadrupole instrumentation. Mass assignment accuracies of 0.01% are routine (see Table 10.21.1 and discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution). ESI has also been successfully interfaced with ion-trap mass spectrometers (Korner et al., 1996), Fourier-transform ion-cyclotron resonance mass spectrometers (Winger et al., 1993; O’Conner et al., 1995; Wu et al., 1995; Smith et al., 1996), and TOF mass spectrometers (Mirgorodskaya et al., 1994; Verentchikov et al., 1994).

In the case of peptides, the maximum number of charges observed often correlates reasonably well with the number of amino acid side chains that can readily accept a proton at the low pH of the analyte stream (i.e., Arg, Lys, and His, plus the free amino terminus). In the case of tryptic peptides, at least two charge sites are normally present, and tryptic peptides with molecular weights of 800 to 3000 typically exhibit a major, if not dominant, [M+2H]2+ ion. For example, the ESI mass spectrum of an HPLC fraction from a tryptic digest of a phosphoprotein shows a simple mixture of peptides, in which doubly charged forms of the major peptide predominate (Fig. 10.21.6). In this spectrum, in addition to proton attachment, doubly charged ions due to Na and K adduction are also abundant. In general, the smaller the pep-



tide, the more likely it is that the singly charged [M+H]+ ion will also be observed. As the size of the peptide increases, triply and highercharged states of the peptide will increase in abundance in the spectra while the abundance of ions with lower charge states will decrease (Fig. 10.21.7). Electrospray mass spectra are keen reminders that mass spectrometers measure the ratio of mass to charge (m/z) and not molecular mass directly. The peak at m/z = 1346.6 in Figure 10.21.6 (panel A) is the monoisotopic [M+H]+ of a peptide of molecular weight 1345.6 (see section on Fundamentals of Mass Measurement Accuracy and Mass Resolution for a discussion of monoisotopic mass). Evidence that this ion carries a single charge (1+) as opposed to a higher number of charges is obtained from the spacing of the isotope peaks, which are one m/z unit apart. The doubly charged form of this same peptide is observed on the m/z scale at [1345.6 +2]/2 = 673.8. The isotope peaks in the [M+2H]2+ isotope cluster are spaced 0.5 m/z units apart, indicating that these ions are doubly charged. Quadrupole mass analyzers can routinely achieve resolution sufficient to distinguish singly from doubly charged ions up to m/z = 3000 (if sufficient sample is available and the instrument is scanned slowly). This capability is particularly relevant to tryptic digests of proteins where, as noted above, the doubly charged ion may be the only ion observed for a given peptide. In the absence of the peak spacing information, the analyst could have difficulty determining if such an ion is singly or doubly charged. For ions with three or more charges, a charge series (e.g., 2+ and 3+) is usually present that permits the analyst to determine the charge state and mass of any given ion in the spectrum without the need to determine the isotopic peak spacing.

LC-ESI-MS Electrospray ionization has made on-line liquid chromatography (LC)-MS a practical and robust method for sample introduction because gas-phase ions, as opposed to liquid droplets, are what is mainly being sampled by the instrument. The ideal milieu for presenting a sample to any mass spectrometer, regardless of the ionization mechanism, is in mixtures of highly pure water and volatile organic solvents containing low levels of volatile acid or base. Fortunately, such mixtures (e.g., water, acetonitrile, and trifluoroacetic acid at 0.05% to 0.1% by volume) are also those that provide optimal separation of peptides and proteins by reversed-

phase liquid chromatography. From a practical standpoint, LC-MS reduces or eliminates the need to preparatively fractionate complex mixtures prior to MS, thereby saving valuable instrument time and preventing possible sample losses. LC-MS data may also be searched retrospectively for components present at very minor levels that may not have been considered worthy of collection based on a weak UV response. In an LC-MS experiment, mass spectra are recorded continuously as the components elute from the HPLC column. If the mass spectrometer is made to scan once every 4 sec, the LC-MS data file will contain ∼900 mass spectra. To identify regions of data that are likely to contain useful spectra, the mass spectrometer’s data system produces a plot of the total number of ions detected during each mass spectrum scan versus time (the scan number). This plot, which is called a mass chromatogram or a total ion current (TIC) trace, looks very much like the UV chromatogram (Fig. 10.21.7). If it is of interest to know whether a particular peptide is present in the digest and where it elutes, the analyst simply requests the data system to display a selected ion-current trace for the specific masses of the most likely charge states (e.g., 1+ to 4+). Figure 10.21.7 illustrates the type of data typically acquired during the LC-ESI-MS analysis of a protein digest. In this example, the protein is heterogeneously glycosylated at a single site, and the multiplicity of peaks evident is due to differing carbohydrates attached to a single tryptic peptide (Fig. 10.21.7, panel C). Because an ESI mass spectrometer behaves as a concentration-dependent detector, much like a UV detector, only a few percent of the total column effluent (∼1 to 5 µl/min) needs to be directed into the mass spectrometer. The remainder can be diverted, via a flow splitter, through an in-line UV detector to a fraction collector. This arrangement makes it possible to perform straightforward correlation of the UV trace with the total ion current trace and then analyze individual fractions further by MS/MS (see discussion of ESI MS/MS for Peptide Sequencing) or some other method. The UV response is also useful for estimating the amount of a peptide that has been injected on the column. This is generally not possible to do by MS without appropriate internal standards (see section entitled Is MS Data Quantitative?). LC-ESI-MS has become a key analytical method for the analysis of natural and recombinant proteins, and data from these experiments is now often included in the regulatory



Supplement 38

Relative intensity (%)

A

125 54.6

100 75 10.8

50

19.8

25 0 1 0.0

101 7.5

201 14.9

301 22.4

401 29.8

501 37.3

601 44.9

701 52.4

801 59.8

scan/time (min)


B

125

10.8 min (scans 317– 319)

1+

439.2

100

= Mr 438.2 75 50 [2M+H]+ 877.6

25 0


C

[M+2H]2+

125 100 75 50 25

19.8 min (scans 261– 271)

2+

E292

1317.8 1398.8 x, y = 4 (Hex)x (HexNAc)y ( dHex) – N296 x = 3 y=4 204.0 366.2 R300 [M+3H]3+ x=5 933.0 y=4 1480.0

= Mr 2795.6

0

D

3+

54.6 min (scans 731–74 4)


950.0 2+

= Mr 2344.4

1173.2

= Mr 2846.8

4+

712.8

200

400

600

2+

1424.4 1564.4

1+

2345.2

800 1000 1200 1400 1600 1800 2000 2200 m/z


Figure 10.21.7 On-line LC-ESI-MS analysis of a tryptic digest of reduced and alkylated heavy chain (51 kDa) from a recombinant monoclonal antibody. (A) Total ion current trace showing the distribution of tryptic peptides. (B) ESI spectrum of the peak at 10.8 min showing a small singly charged peptide and its gas-phase dimer. (C) ESI spectrum of the peak at 19.8 min corresponding to a tryptic glycopeptide with heterogenous carbohydrate. At least five different carbohydrate structures are indicated. Abundant low-m/z ions at m/z = 204 and m/z = 366 are characteristic fragment ions produced from the carbohydrate (Roberts et al., 1995). (D) ESI spectrum of the peak at 54.6 min showing two large tryptic peptides, each represented by several different charge states.



filings for new biopharmaceutical products (Carr et al., 1989; Palczewski et al., 1994; Stults et al., 1994; Watts et al., 1994; Resing et al., 1995; Roberts et al., 1995; Rush et al., 1995; Schindler et al., 1995; Hemling et al., 1996).

ESI MS/MS for Peptide Sequencing The principles of MS/MS using triple-quadrupole mass spectrometers have been reviewed (Yost and Boyd, 1990). Briefly, the first quadrupole (Fig. 10.21.5) acts as a mass filter to select specifically the [M+nH]n+ ion (where n is typically 1, 2, 3, or 4) of a peptide of interest and isolate it from other ions produced in the source. The selected precursor ion is then fragmented with a neutral gas such as nitrogen or argon in a quadrupole collision cell. This process is known as collision-induced decomposition. The fragment or product ions produced are then transmitted into the quadrupole second mass analyzer, which is being used as a scanning mass analyzer. The product ions are separated based on m/z and detected. The result is a product-ion mass spectrum of the selected precursor that is free of interferences from other components present in the mixture. An example of this process is shown in Figure 10.21.6. The fragmentation of this peptide reveals its sequence and the location of the phosphorylated residue. Fragmentation can also be induced in the ESI source by increasing the energetics of the ion sampling process. This is done by adjusting the lens potentials in the high-pressure region upstream from the first quadrupole mass filter. Ions with increased kinetic energy undergo decomposition via collisions with residual gas molecules (Katta et al., 1991). Structurally useful fragmentation can thus be obtained on a single-quadrupole instrument, with the caveat that because this method has no ability to preferentially select precursor ions, it requires a highly purified sample. Recently strategies have been developed that take advantage of this ability of the ESI interface to generate fragment ions prior to mass analysis. Selective detection of glycosylated peptides (Carr et al., 1993; Huddleston et al., 1993a), phosphorylated peptides (Huddleston et al., 1993b; Till et al., 1994), and sulfated peptides (Bean et al., 1995) during on-line LC-ESI-MS analysis of protein digests can be accomplished by monitoring the total ion current trace for highly diagnostic low-m/z fragment ions produced in the ESI interface. These procedures are equally applicable to the detection of these modifications in unseparated mixtures.

IS MS DATA QUANTITATIVE? A question that frequently arises in the analysis of peptide or protein mixtures is whether the relative peak heights (or areas) are quantitatively representative of the relative solution concentrations of the sample components. Unfortunately, the answer to this question is no, unless appropriate internal reference standards are used. Due to differing ionization and desorption efficiencies, the absolute yields of different ions from one peptide in a mass spectrometer can vary by perhaps as much as 2 orders of magnitude, depending on the sequence, composition, and size of the peptides (Dunayevskiy et al., 1995; Jesperson et al., 1995; Cohen and Chait, 1996). Furthermore, strongly ionizing components can frequently suppress the ionization of more weakly ionizing components present in the same sample or chromatographic fraction. In MALDI, the matrix itself can strongly influence the detectability of specific sample constituents. Quantitation by mass spectrometry requires that the response of a specific sample component be determined relative to that of a reference compound added to the sample and analyzed under identical conditions. In certain cases, semiquantitative estimates of relative amounts of components may be made if the components are of nearly identical structure and their chemical differences do not significantly affect the overall charge or hydrophobic/hydrophilic character of the molecules. For example, the relative ratios of the variants of β-casein observed in the ESI mass spectrum (Fig. 10.21.8) are probably a good reflection of the relative solution concentrations of these variants, as they differ from one another by only one or two amino acid substitutions out of more than 200 amino acids. The smaller the molecule, the more likely it is that such estimates will be in error.

SAMPLE PREPARATION The ideal method for preparing samples for any type of MS analysis is reversed-phase highperformance liquid chromatography (RPHPLC). If purification by RP-HPLC is not an option because of known problems with sample loss, then the cleanup protocol employed should produce a sample composition that resembles as closely as possible that of a sample which has been purified by RP-HPLC. That is, the sample should contain the least amount possible of buffers, salts, detergents, and anything else besides water, organic modifier, and volatile acid or base.



Supplement 38

A

B 23,984 2

14+ 13+


[M + H]1+ 24,010 30

24,024 2

+

18+ 15 16+

24,093 2 12+

1400

1420

11+ 10+ 9+

20,000 22,000 24,000 26,000

800

1200

1600

8+

2000

2400

m/z

Figure 10.21.8 Molecular weight determination of β-casein, a 24-kDa phosphoprotein with known sequence heterogeneity. (A) Linear MALDI-TOF spectrum. Molecular weight was determined from the centroid of the smoothed peak using an external calibration. (B) ESI spectrum. The inset, an expansion of the region around the 14+ charge state, reveals the presence of at least three variants. The molecular weights shown are the averages of determinations made from ten individual charge states.


The quality of the data obtained using ESIMS is very dependent on the type and concentration of excipients present in the sample. If buffers are necessary for protein solubility or enzymatic digestion, it is best to use a volatile buffer such as ammonium bicarbonate or ammonium acetate at a concentration of ≤30 mM. In general it is best to avoid using any ionic detergents (surfactants), even though it is possible to obtain spectra for some proteins when these are included as long as the surfactant concentration is kept 3% of the protein in the antigen solution should be the antigen. The total concentration of protein in the antigen solution should be increased for semipurified antigen preparations. Do not raise the total protein concentration in the antigen solution to >10 ìg/ml, since this concentration usually saturates >85% of the available sites on Immulon microtiter plates. For some antigens, coating may occur more efficiently at different pHs.

Coat plate with antigen 3. Using a multichannel pipet and tips, dispense 50 µl antigen solution into each well of an Immulon microtiter plate. Tap or shake the plate to ensure that the antigen solution is evenly distributed over the bottom of each well.

Ab

= detected Ab

coat well with anitgen Ag

Ag

block

incubate with antibody Ab

Ab

Ag

Ag

wash E

E

Ab

Ab

Ab

Ab

Ag

Ag

incubate with antibodyenzyme conjugate

wash E

E

Ab

Ab

Ab

Ab

Ag

Ag

add substrate and observe color change or fluorescence

Figure 11.2.1 Indirect ELISA to detect specific antibodies. Ag = antigen; Ab = antibody; E = enzyme.

Immunology


Supplement 15

4. Wrap coated plates in plastic wrap to seal and incubate overnight at room temperature or 2 hr at 37°C. Individual adhesive plate sealers are sold commercially but plastic wrap is easier to use and works as well. Sealed plates can be stored at 4°C with antigen solution for months.

5. Rinse coated plate over a sink by filling wells with deionized or distilled water dispensed either from a plastic squirt bottle or from the tap. Flick the water into the sink and rinse with water two more times, flicking the water into the sink after each rinse. Block residual binding capacity of plate 6. Fill each well with blocking buffer dispensed as a stream from a squirt bottle and incubate 30 min at room temperature. Residual binding capacity of the plate is blocked in this step. Tween 20 (0.05%) by itself is more effective at blocking than any protein tested, but because the combination of protein and Tween 20 may be more effective than Tween 20 alone in some cases, bovine serum albumin (BSA; 0.25%) is included in the blocking buffer.

7. Rinse plate three times in water as in step 5. After the last rinse, remove residual liquid by wrapping each plate in a large paper tissue and gently flicking it face down onto several paper towels laying on the benchtop. Rinsing with water is cheaper and easier than rinsing with buffered solutions and is as effective.

Add antibody to plate 8. Add 50 µl antibody samples diluted in blocking buffer to each of the coated wells, wrap plate in plastic wrap, and incubate ≥2 hr at room temperature. While enough antibody may be bound after 1 to 2 hr to generate a strong signal, equilibrium binding is generally achieved after 5 to 10 hr. Thus, the specific signal may be significantly increased by longer incubations. For this and all steps involving the delivery of aliquots of many different solutions to microtiter plates with multichannel pipets, such as the primary screening of hybridoma supernatants, the same pipet tips can be reused for hundreds of separate transfers. Wash tips between transfers by expelling any liquid remaining in the tips onto an absorbent surface of paper tissues, rinsing tips five times in blocking buffer, and carefully expelling any residual liquid from tips onto the tissues. Avoid bubbles in the tips; any tip with intractable bubbles should be replaced.

Wash the plate 9. Rinse plate three times in water as in step 5. 10. Fill each well with blocking buffer, vortex, and incubate 10 min at room temperature. Plates are vortexed to remove any reagent remaining in the corners of the wells.

11. Rinse three times in water as in step 5. After the final rinse, remove residual liquid as in step 7. Add developing reagent to plate 12. Add 50 µl developing reagent in blocking buffer (at optimal concentration determined in step 1) to each well, wrap in plastic wrap, and incubate ≥2 hr at room temperature. The strength of the signal may be increased by longer incubations (see annotation to step 8). Enzyme-Linked Immunosorbent Assays



13. Wash plates as in steps 9 to 11. After final rinsing, plates may be wrapped in plastic wrap and stored for months at 4°C prior to adding substrate.

Add substrate and measure hydrolysis 14. Add 75 µl MUP or NPP substrate solution to each well and incubate 1 hr at room temperature. 15. Monitor hydrolysis qualitatively by visual inspection or quantitatively with a microtiter plate reader (see below). Hydrolysis can be stopped by adding 25 µl of 0.5 M NaOH. a. Visually, hydrolysis of NPP can be detected by the appearance of a yellow color. If using a microtiter plate reader to measure NPP hydrolysis, use a 405-nm filter. b. Visually, hydrolysis of MUP can be monitored in a darkened room by illumination with a long-wavelength UV lamp. If using a microtiter plate spectrofluorometer to measure MUP hydrolysis, use a 365-nm excitation filter and a 450-nm emission filter. The fluorogenic system using the MUP substrate is 10 to 100 times faster than the chromogenic system using NPP. Furthermore, the rate of spontaneous hydrolysis of MUP is much lower than that of NPP. To detect bound antibodies that are present at low concentration, measure hydrolysis at a later time. To calculate when to measure hydrolysis the second time, remember that the amount of hydrolysis is almost directly proportional to the time of hydrolysis. For example, if the hydrolysis in the wells of interest reads 200 at 1 hr and a reading of 2000 is desired, incubate the plate ∼10 hr before taking the second reading.

DIRECT COMPETITIVE ELISA TO DETECT SOLUBLE ANTIGENS This assay is used to detect or quantitate soluble antigens and is most useful when both a specific antibody and milligram quantities of purified or semipurified antigen are available (Fig. 11.2.2). To detect soluble antigens, plates are coated with antigen and the binding of specific antibody-enzyme conjugates to antigen-coated plates is inhibited by test solutions containing soluble antigen. After incubation with mixtures of the conjugate and inhibitor in antigen-coated wells, unbound conjugate is washed away and substrate is added. The amount of antigen in the test solutions is proportional to the inhibition of substrate hydrolysis and can be quantitated by interpolation onto an inhibition curve generated with serial dilutions of a standard antigen solution.

ALTERNATE PROTOCOL

The direct assay may also be adapted as an indirect assay by substituting specific antibody for specific antibody-enzyme conjugate. The amount of specific antibody bound is then detected using a species-specific or isotype-specific conjugate as a tertiary reactant. Additional Materials Specific antibody–alkaline phosphatase conjugate (UNIT 11.1) Standard antigen solution Test antigen solutions Round- or cone-bottom microtiter plates 1. Determine the optimal concentration of coating reagent and antibody–alkaline phosphatase conjugate by criss-cross serial dilution analysis in which the concentrations of both the antigen (coating reagent) and the conjugate (developing reagent) are varied (see first support protocol). Prepare a 2× conjugate solution by diluting the Immunology


Supplement 15

specific antibody–alkaline phosphatase conjugate in blocking buffer to twice the optimal concentration. The final concentration is usually 25 to 500 ng antibody/ml. Prepare 3 ml antibody–alkaline phosphatase conjugate for each plate.

2. Coat and block wells of an Immulon microtiter plate with 50 µl antigen solution as in steps 2 to 7 of the basic protocol. 3. Prepare six 1:3 serial dilutions of standard antigen solution in blocking buffer (see first support protocol for preparation of serial dilutions)—these antigen concentrations will be used in preparing a standard inhibition curve (see step 10). Antigen concentrations should span the dynamic range of inhibition. The dynamic range of inhibition is defined as that range of inhibitor concentrations wherein changes of inhibitor concentration produce detectable changes in the amount of inhibition. The dynamic range of inhibition is empirically determined in an initial assay in which antigen concentration is typically varied from the micromolar (10−6 M) to the picomolar (10−12 M) range. For most protein antigens, initial concentration should be ∼10 ìg/ml, followed by nine 1:4 serial dilutions in blocking buffer. These antigen dilutions are assayed for their ability to inhibit the binding of conjugate to antigen-coated plates under standard assay conditions. From this initial assay, six 1:3 antigen dilutions spanning the dynamic range of inhibition are selected for further use as standard antigen-inhibitor dilutions. Prepare ≥75 ìl of each dilution for each plate to be assayed. Inhibitor curves are most sensitive in the region of the curve where small changes in inhibitor concentrations produce maximal changes in the amount of inhibition. This

with inhibitor antigen

without inhibitor antigen Ag

= detected AG

coat well with antigen

Ag

Ag

Ag

Ag

block E Ab Ag Ag

E

E

E

Ab

Ab

Ab

Ag

Ag

Ag

incubate with antibodyenzyme conjugate with or without inhibitor antigen

wash

Ag

Enzyme-Linked Immunosorbent Assays

E

E

E

Ab

Ab

Ab

Ag

Ag

Ag

add substrate and measure inhibition of color change or fluorescence

Figure 11.2.2 Direct competitive ELISA to detect soluble antigens. Ag = antigen; Ab = antibody; E = enzyme.



region of the curve normally spans 15% to 85% inhibition. In most systems, this range of inhibition is produced by concentrations of inhibitor between 1 and 250 ng/ml.

4. Mix and incubate conjugate and inhibitor by adding 75 µl of 2× conjugate solution (from step 1) to each well of a round- or cone-bottom microtiter plate, followed by 75 µl inhibitor—either test antigen solution or standard antigen solution (from step 3). Mix the conjugate and inhibitor solutions by pipetting up and down in the pipet tip three times (see annotation to step 8 in the basic protocol) and incubate ≥30 min at room temperature. For accurate quantitation of the amount of antigen in the test solutions, test antigen solutions should inhibit conjugate binding between 15% to 85%. It may be necessary to assay two or three different dilutions of the test solutions to produce inhibitions within this range.

5. Prepare uninhibited control samples by mixing equal volumes of 2× conjugate solution and blocking buffer. 6. Transfer 50 µl of the mixture of conjugate plus inhibitor (from step 4) or conjugate plus blocking buffer (from step 5) to an antigen-coated plate (from step 2) and incubate 2 hr at room temperature. If samples are to be assayed in duplicate, the duplicates should be in adjacent columns on the same plate. Reserve column 11 for uninhibited control samples (step 5) and column 12 for substrate alone without any conjugate. If the concentration of antigen in the test samples is to be accurately quantitated, dilutions of homologous antigen solutions (step 3) should be included on each plate.

7. Wash plate as in steps 9 to 11 of the basic protocol. 8. Add 75 µl of MUP or NPP substrate solution to each well and incubate 1 hr at room temperature. 9. Read plates on the microtiter plate reader after ≥1 hr, at which time enough substrate has been hydrolyzed in the uninhibited reactions to permit accurate measurement of the inhibition. 10. Prepare a standard antigen-inhibition curve constructed from the inhibitions produced by the dilutions of the standard antigen solutions from step 3. Plot antigen concentration on the x axis, which is a log scale, and fluorescence or absorbance on the y axis, which is a linear scale. 11. Interpolate the concentration of antigen in the test solutions from the standard antigen-inhibition curve. The dynamic range of the inhibition curve may deviate from linearity if the specific antibodies are heterogeneous and possess significantly different affinities or if the standard antigen preparation contains heterogeneous forms of the antigen. Antigen concentration in test samples can be accurately interpolated from the inhibition curve as long as the test antigen is antigenically identical to the standard antigen and the concentration of test antigen falls within the dynamic range of inhibition.

Immunology


Supplement 24

ALTERNATE PROTOCOL

ANTIBODY-SANDWICH ELISA TO DETECT SOLUBLE ANTIGENS Antibody-sandwich ELISAs may be the most useful of the immunosorbent assays for detecting antigen because they are frequently between 2 and 5 times more sensitive than those in which antigen is directly bound to the solid phase (Fig. 11.2.3). To detect antigen, the wells of microtiter plates are coated with specific (capture) antibody followed by incubation with test solutions containing antigen. Unbound antigen is washed out and a different antigen-specific antibody conjugated to enzyme (i.e., developing reagent) is added, followed by another incubation. Unbound conjugate is washed out and substrate is added. After another incubation, the degree of substrate hydrolysis is measured. The amount of substrate hydrolyzed is proportional to the amount of antigen in the test solution. Additional Materials Specific antibody or immunoglobulin fraction from antiserum or ascites fluid, or hybridoma supernatant (UNIT 11.10), or bacterial lysate (second support protocol)

Ag

= detected Ag

coat well with antibody

Ab

Ab

block

incubate with antigen Ag

Ag

Ab

Ab

wash

E

E

Ab

Ab

Ag

Ag

Ab

Ab

incubate with antibody-enzyme conjugate

wash


E

E

Ab

Ab

Ag

Ag

Ab

Ab


Figure 11.2.3 Antibody-sandwich ELISA to detect antigen. Ag = antigen; Ab = antibody; E = enzyme.



1. Prepare the capture antibody by diluting specific antibody or immunoglobulin fraction in PBSN to a final concentration of 0.2 to 10 µg/ml. The capture antibodies can be monoclonal or polyclonal. If the immunoglobulin fraction from an antiserum or ascites fluid is used, the concentration of total protein may need to be increased to compensate for the lower content of specific antibody. Little advantage is gained by increasing the total protein concentration in the capture antibody solution beyond 10 ìg/ml.

2. Determine the concentration of capture antibody and conjugate necessary to detect the desired concentration of antigen by criss-cross serial dilution analysis (see first support protocol). Prepare a capture antibody solution in PBSN at this concentration. 3. Coat wells of an Immulon plate with capture-antibody solution as in steps 3 to 5 of the basic protocol. 4. Block wells as in steps 6 and 7 of the basic protocol. 5. Prepare a standard antigen-dilution series by successive 1:3 dilutions of the homologous antigen stock in blocking buffer (see first support protocol). In order to measure the amount of antigen in a test sample, the standard antigen-dilution series needs to span most of the dynamic range of binding. This range typically spans from 0.1 to 1000 ng antigen/ml. The dynamic range of binding is defined as that range of antigen concentrations wherein small, incremental changes in antigen concentration produce detectable differences in the amount of antigen bound (see annotation to step 3, in the preceding alternate protocol). In most assay systems, the amount of antigen in a test solution is most accurately interpolated from the standard curve if it produces between 15% to 85% of maximal binding. NOTE: While standard curves are necessary to accurately measure the amount of antigen in test samples, they are unnecessary for qualitative “yes/no” answers.

6. Prepare dilutions of test antigen solutions in blocking buffer. It may be necessary to assay one or two serial dilutions of the initial antigen test solution to ensure that at least one of the dilutions can be accurately measured. For most assay systems, test solutions containing 1 to 100 ng/ml of antigen can be accurately measured.

7. Add 50-µl aliquots of the antigen test solutions and the standard antigen dilutions (from step 5) to the antibody-coated wells and incubate ≥2 hr at room temperature. For accurate quantitation, samples should be run in duplicate or triplicate, and the standard antigen-dilution series should be included on each plate (see step 5). Pipetting should be performed rapidly to minimize differences in time of incubation between samples.

8. Wash plate as in steps 9 to 11 of the basic protocol. 9. Add 50 µl specific antibody–alkaline phosphatase conjugate and incubate 2 hr at room temperature. The conjugate concentration is typically 25 to 400 ng specific antibody/ml. When the capture antibody is specific for a single determinant, the conjugate must be prepared from antibodies which recognize different determinants that remain available after the antigen is bound to the plate by the capture antibody.

10. Wash plate as in steps 9 to 11 of the basic protocol. 11. Add 75 µl of MUP or NPP substrate solution to each well and incubate 1 hr at room temperature. Immunology


Supplement 15

12. Read the plate on a microtiter plate reader. To quantitate low-level reactions, the plate can be read again after several hours of hydrolysis.

13. Prepare a standard curve constructed from the data produced by serial dilutions of the standard antigen (step 5). Plot antigen concentration on the x axis which is a log scale, and fluorescence or absorbance on the y axis which is a linear scale. 14. Interpolate the concentration of antigen in the test solutions from the standard curve. ALTERNATE PROTOCOL

DOUBLE ANTIBODY–SANDWICH ELISA TO DETECT SPECIFIC ANTIBODIES This assay is especially useful when screening for specific antibodies in cases when a small amount of specific antibody is available and purified antigen is unavailable (Fig. 11.2.4). Additionally, this method can be used for epitope mapping of different monoclonal antibodies that are directed against the same antigen. Plates are coated with capture antibodies specific for immunoglobulin from the immunized species. The test antibody solution is incubated on the plates coated with the capture antibodies. Plates are then washed, incubated with antigen, washed again, and incubated with specific antibody conjugated to an enzyme. After incubation, unbound conjugate is washed out and substrate is added. Wells that are positive for hydrolysis may contain antibodies specific for the antigen. Additional Materials Capture antibodies specific for immunoglobulin from the immunized species Specific antibody–alkaline phosphatase conjugate 1. Coat wells of an Immulon microtiter plate with 50 µl of 2 to 10 µg/ml capture antibodies as in steps 2 to 5 of the basic protocol. NOTE: Capture antibodies must not bind the antigen or conjugate antibodies. When analyzing hybridoma supernatants or ascites fluid, coat plates with 2 ìg/ml capture antibody. When analyzing antisera, coat plates with 10 ìg/ml capture antibody.

2. Block wells as in steps 6 and 7 of the basic protocol. 3. Prepare dilutions of test antibody solutions in blocking buffer. Add 50 µl to coated wells and incubate ≥2 hr at room temperature. Hybridoma supernatants, antisera, or ascites fluid can be used as the test samples. Dilute hybridoma supernatants 1:5 and antisera or ascites fluid 1:200.

4. Wash plate as in steps 9 to 11 of the basic protocol. 5. Prepare an antigen solution in blocking buffer containing 20 to 200 ng/ml antigen. Although purified antigen preparations are not essential, the limit of detectability for most protein antigens in this type of system is 2 to 20 ng/ml. A concentration of 20 to 200 ng antigen/ml is recommended.

6. Add 50-µl aliquots of the antigen solution to antibody-coated wells and incubate ≥2 hr at room temperature. 7. Wash plate as in steps 9 to 11 of the basic protocol. Enzyme-Linked Immunosorbent Assays

8. Add 50 µl specific antibody–alkaline phosphatase conjugate to the wells and incubate 2 hr at room temperature.



The conjugate antibodies must not react with the capture antibody or the test antibody. The conjugate concentration is typically between 25 to 500 ng specific antibody/ml, and should be high enough to result in ∼0.50 absorbance units/hr at 405 nm when using NPP as a substrate or a signal of 1000 to 1500 fluorescence units/hr when using MUP as a substrate. If no specific antibodies from the appropriate species are available to serve as a positive control, then a positive control system should be constructed out of available reagents. Such reagents can be found in Linscott’s Directory of Immunological and Biological Reagents.

Ab

= detected Ab

coat well with capture antibody

Ab

Ab

block

incubate with antibody Ab Ab

Ab Ab

wash

incubate with antigen Ag

Ag

Ab Ab

Ab Ab

wash E

E

Ab

Ab

Ag

Ag

Ab Ab

Ab Ab


wash E

E

Ab

Ab

Ag

Ag

Ab Ab

Ab Ab


Figure 11.2.4 Double antibody–sandwich ELISA to detect specific antibodies. Ag = antigen; Ab = antibody; E = enzyme.

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9. Wash plate as in steps 9 to 11 of the basic protocol. 10. Add 75 µl of MUP or NPP substrate solution to each well and incubate 1 hr at room temperature. After 1 hr, examine hydrolysis visually or spectrophotometrically (see step 15 of the basic protocol). In order to detect low-level reactions, the plate can be read again after several hours or days of hydrolysis.

11. Check for false positives by rescreening samples that test positive for antigen-specific antibody. For each positive sample, coat four wells with capture antibody and arm the capture antibody with test antibody (steps 1 to 4). Incubate two of the wells with antigen (steps 5 to 7) and two of the wells with blocking buffer. Add conjugate and substrate to all four wells (steps 8 to 10) and measure hydrolysis after 1 hr. This procedure will eliminate false positives resulting from test antibodies that react with the enzyme-antibody complex. ALTERNATE PROTOCOL

DIRECT CELLULAR ELISA TO DETECT CELL-SURFACE ANTIGENS The expression of cell-surface antigens or receptors is measured using existing antibodies or other ligands specific for cell-surface molecules (Fig. 11.2.5). Cells are incubated with enzyme conjugated to antibodies that are specific for a cell-surface molecule. Unbound conjugate is washed away and substrate is added. The level of antigen expression is proportional to the amount of substrate hydrolysis. This procedure can be as sensitive as flow cytometry analysis in quantitating the level of antigen expression on a population of cells (Coligan et al., 1991). Unlike the flow cytometry analysis, however, this method is not sensitive for mixed populations. This assay can be converted to an indirect assay by substituting biotinylated antibody for the enzyme-antibody conjugate, followed by a second incubation with avidin–alkaline phosphatase.

E

E

Ab

Ab

Ag

Ag

C

C

Ag = detected Ag

incubate cells with antibody-enzyme conjugate

wash and centrifuge


E

E

Ab

Ab

Ag

Ag

C

C

add substrate, resuspend cells, and observe color change or fluorescence

Figure 11.2.5 Direct cellular ELISA to detect cell-surface antigens. Ab = antibody; E = enzyme; C = cell.



Additional Materials Cell samples Specific antibody–alkaline phosphatase conjugate (see second support protocol) Wash buffer, ice-cold Cone- or round-bottom microtiter plates Sorvall H-1000B rotor (or equivalent) 1. Determine the optimal number of cells per well and the antibody-conjugate concentration by criss-cross serial dilution analysis (see first support protocol) using variable numbers of positive- and negative-control cell samples and varying concentrations of antibody-biotin conjugate. Titrate cells initially at 1-5 × 105/well and conjugate at 0.5 to 10 ìg/ml. For preparation and handling of cells, consult steps 2 to 5. Because eukaryotic cells express variable amounts of alkaline phosphatase, test cells must be assayed in a preliminary experiment for alkaline phosphatase by incubation with substrate alone. If the test cells express unacceptable levels of alkaline phosphatase, another enzyme conjugate such as β-galactosidase should be used. Both chromogenic and fluorogenic substrates are available for β-galactosidase.

2. Centrifuge cell samples in a table-top centrifuge 5 min in Sorvall H-1000B rotor at 1500 rpm (450 × g), 4°C, in a 15- to 50-ml centrifuge tube. Count cells (APPENDIX 3) and resuspend in ice-cold wash buffer at 1-5 × 106 cells/ml. If the surface antigen retains its antigenicity after fixation, cells may be fixed at the beginning of the experiment—but do not fix cells unless it can be demonstrated that the antigenicity is retained after fixation. Fix cells by suspending in glutaraldehyde (0.5% final; from a 25% stock, EM grade Sigma #G5882), and incubating 30 min at room temperature. Pellet cells, resuspend in PBSLE (see second support protocol), and incubate for 30 min at 37°C. Wash twice in PBSLE and resuspend in wash buffer. Cells can be kept for months at 4°C after fixation.

3. Dispense 100 µl of cell suspension (1-5 × 105 cells) into wells of cone- or round-bottom microtiter plates, and centrifuge 1 min at 450 × g, 4°C. Remove supernatant by vacuum aspiration, and disrupt pellet by briefly shaking microtiter plate on a vortex mixer or microtiter plate shaker. 4. Resuspend pellet in 100 µl of conjugate in ice-cold wash buffer at the optimal concentration (see step 1). Incubate 1.5 hr at 4°C, resuspending cells by gently shaking at 15-min intervals. Be careful not to splash cell suspensions out of wells.

5. Centrifuge cells 1 min at 450 × g, 4°C, remove supernatant by vacuum aspiration, briefly vortex pellet, and resuspend in 200 µl ice-cold wash buffer. Repeat three times. 6. Add 100 µl MUP or NPP substrate solution. Incubate 1 hr at room temperature, resuspending cells by gently shaking at 15-min intervals during hydrolysis. 7. Determine extent of hydrolysis by visual inspection or using a microtiter plate reader.

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ALTERNATE PROTOCOL

INDIRECT CELLULAR ELISA TO DETECT ANTIBODIES SPECIFIC FOR SURFACE ANTIGENS This assay is designed to screen for antibodies specific for cell-surface antigens (Fig. 11.2.6). Antibodies against surface antigens are detected by incubating whole cells with a test solution containing the primary antibody. The unbound antibody is washed away and the cells are then incubated with an enzyme conjugated to antibodies specific for the primary antibody. Unbound enzyme conjugate is washed away and substrate solution added. The level of bound primary antibody is proportional to the amount of substrate hydrolysis. Additional Materials Positive-control antibodies (i.e., those that react with the experimental cells and are from the immunized species) Negative-control antibodies (i.e., those that do not react with the experimental cells) Test antibody solution Antibody or F(ab′)2 (against immunoglobulin from the immunized species) conjugated to alkaline phosphatase Cone- or round-bottom microtiter plates

Ab

= detected Ab

incubate cells with antibody Ab

Ab

Ag

Ag

C

C

wash and centrifuge E

E

Ab

Ab

Ab

Ab

Ag

Ag

C

C


wash and centrifuge


E

E

Ab

Ab

Ab

Ab

Ag

Ag

C

C

add substrate, resuspend cells, and observe color change or fluorescence

Figure 11.2.6 Indirect cellular ELISA to detect antibodies specific for surface antigens. Ab = antibody; E = enzyme; C = cell.



1. Centrifuge and resuspend cell samples as in step 2 of the previous alternate protocol at 1-5 × 106 cells/ml. Because this technique detects antibodies against uncharacterized epitopes, fixation prior to analysis is not recommended. Fixation may destroy the antigenicity of the epitope. All steps must be performed at 4°C in physiological buffers containing NaN3. Because eukaryotic cells express variable amounts of alkaline phosphatase, test cells must be assayed for alkaline phosphatase activity. If the endogenous alkaline phosphatase level is too high, another enzyme should be substituted for alkaline phosphatase in the antibodyenzyme conjugate (see annotation to step 1 of the previous alternate protocol).

2. In preliminary assays, determine the optimal number of cells per well and conjugate concentration by criss-cross serial dilution analysis using positive- and negative-control antibodies instead of test antibodies (see first support protocol). In adapting the criss-cross serial dilution analysis, whole cells replace the solid-phase coating reagent; see techniques for handling cells are outlined in steps 3 to 8. Set up titrations by varying the number of cells between 1 × 105 and 5 × 105/well, the concentration of positive- and negative-control antibodies between 0.1 and 10 µg/ml, and the concentration of antibody-enzyme conjugate between 0.1 and 10 µg/ml. 3. Dispense 100 µl of cell suspension (1-5 × 105 cells) into wells of round- or cone-bottom microtiter plates. Centrifuge 1 min at 1500 rpm, 4°C, remove supernatant by vacuum aspiration, and disrupt pellet by briefly shaking microtiter plate on the vortex mixer. 4. Resuspend cells in 100 µl solutions containing 1 to 10 µg/ml test antibody or control antibodies in ice-cold wash buffer. Incubate 1.5 hr at 4°C, resuspending cells by gently shaking at 15-min intervals. Be careful not to splash cell suspensions out of wells.

5. Centrifuge cells 1 min at 1500 rpm, 4°C, remove supernatant by vacuum aspiration, briefly vortex pellet, and resuspend in 200 µl ice-cold wash buffer. Repeat twice. 6. Resuspend pellet in 100 µl enzyme-antibody conjugate or F(ab′)2-enzyme conjugate diluted in ice-cold wash buffer. The optimal concentration of antibody, determined in step 2, is usually 100 to 500 ng/ml. Incubate 1.5 hr at 4°C, resuspending cells by gently shaking at 15-min intervals. When working with cells that may express Fc receptors, it is best to use enzyme conjugated to F(ab′)2 fragments. F(ab′)2 fragments have had the Fc portion of the antibody enzymatically removed and no longer bind to Fc receptors.

7. Wash cells as in step 5. Repeat three times. 8. Add 100 µl MUP or NPP substrate solution. Allow hydrolysis to proceed until the signal has reached the desired levels; resuspend cells by gently shaking at 15 min intervals during hydrolysis. If desired, stop hydrolysis by adding 25 µl of 0.5 M NaOH. 9. Determine extent of hydrolysis by visual inspection or spectrophotometrically using a microtiter plate reader.

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CRISS-CROSS SERIAL DILUTION ANALYSIS TO DETERMINE OPTIMAL REAGENT CONCENTRATIONS

SUPPORT PROTOCOL

Serial dilution titration analyses are performed to determine optimal concentrations of reagents to be used in ELISAs. In this protocol, all three reactants in a three-step ELISA—a primary solid-phase coating reagent, a secondary reagent that binds the primary reagent, and an enzyme-conjugated tertiary developing reagent that binds to the secondary reagent—are serially diluted and analyzed by a criss-cross matrix analysis (Fig 11.2.7). Once the optimal concentrations of reagents to be used under particular assay conditions are determined, these variables are kept constant from experiment to experiment. The coating (primary), secondary, and developing (tertiary) reagents will vary depending upon which of the previous protocols needs to be optimized. Additional Materials Coating reagent Secondary reagent Developing reagent 17 × 100–mm and 12 × 74–mm test tubes

Secondary reactant

heterologous (antigen)

(ng/ml)

200

50

12.5

3.12

0.78

0

200

50

12.5

3.12

0.78

0 A

500

over

over

over

3200

1000

0

500

120

40

20

10

B

250

over

over

over

2060

560

0

300

80

20

0

0

C

125

over

over

3650

1370

360

0

195

40

10

10

0

D

62.5

3600

4000

2270

790

240

0

120

30

10

10

10

E

31.25

2700

2100

1200

410

120

0

60

10

10

10

0

F

0

0

0

0

0

0

0

0

0

0

0

0

G

1

2

3

4

5

6

7

8

9

10

11

Rows

Tertiary reactant (antibody-alkaline phosphatase)

homologous (antigen)

H 12

Columns

Figure 11.2.7 Results of a criss-cross serial dilution analysis (for optimization of secondary and tertiary reactant concentrations) of an antibody-sandwich ELISA to detect antigen. The numbers in columns 1 to 11 and rows B to G represent relative fluorescence units observed for each well on a 96-well microtiter plate. Plates were coated overnight with the capture antibody at 2 µg/ml. The secondary reactants, 4-fold serial dilutions of the homologous antigen and a non-cross-reactive heterologous antigen, were incubated on the plate 2 hr. The tertiary reactant, 2-fold serial dilutions of specific antibody–alkaline phosphatase conjugates, were incubated on the plate 2 hr. After 1 hr of incubation with the substrate MUP, the fluorescence was read in a microtiter plate spectrofluorometer.

Reagent concentrations depend upon individual assay variables that are set by the investigator. If the time of hydrolysis is set at 1 hr, the relative fluorescence at ∼1000 relative fluorescence units, and the sensitivity at 780 pg/ml of homologous antigen, then 500 ng/ml of enzyme-antibody conjugate must be used in the ELISA. If, however, the assay has to detect only 3.12 ng/ml of homologous antigen, then the concentration of conjugate can be reduced to 125 ng/ml. It should be noted by comparing the homologous with the heterologous reactions (wells B5 versus B11 and D4 versus D10) that both the specificity and the signal-to-noise ratio for this assay are excellent.



Prepare coating-reagent dilutions 1. Place four 17 × 100–mm test tubes in a rack and add 6 ml PBSN to the last three tubes. In tube 1, prepare a 12-ml solution of coating reagent at 10 µg/ml in PBSN. Transfer 6 ml of tube 1 solution to tube 2. Mix by pipetting up and down five times. Repeat this transfer and mix for tubes 3 and 4; the tubes now contain the coating reagent at 10, 5, 2.5, and 1.25 µg/ml. 2. Using a multichannel pipet, dispense 50 µl of the coating reagent solutions into wells of four Immulon microtiter plates (i.e., each plate is filled with one of the four dilutions). Incubate overnight at room temperature or 2 hr at 37°C. 3. Rinse and block plates with blocking buffer as in steps 5 to 7 of the basic protocol. Prepare secondary-reagent dilutions 4. Place five 12 × 75–mm test tubes in a rack and add 3 ml blocking buffer to the last four tubes. In tube 1, prepare a 4-ml solution of secondary reagent at 200 ng/ml in PBSN. Transfer 1 ml of tube 1 solution to tube 2. Pipet up and down five times. Repeat this transfer and mix for tubes 3 to 5; the tubes now contain the secondary reactant at 200, 50, 12.5, 3.125, and 0.78 ng/ml. If possible, prepare and test serial dilutions of a nonreactive heterologous form of the secondary reactant in parallel (Fig. 11.2.7). If the assay is especially insensitive, it may be necessary to increase the secondary reactant concentrations so the tube-1 solution is 1000 ng/ml.

5. Dispense 50 µl of the secondary reagent solutions into the first five columns of all four coated plates. The most dilute solution is dispensed into column 5, while solutions of increasing concentration are added successively into columns 4, 3, 2, and 1. Thus, the fifth column contains 0.78 ng/ml and the first column 200 ng/ml. Incubate 2 hr at room temperature. 6. Wash plates as in steps 9 to 11 of the basic protocol. Prepare developing-reagent dilutions 7. Place five 17 × 100–mm test tubes in a rack and add 3 ml blocking buffer to the last four tubes. In tube 1, prepare a 6-ml solution of developing reagent at 500 ng/ml in blocking buffer. Transfer 3 ml of tube 1 solution into tube 2 and mix. Repeat this transfer and mixing for tubes 3 and 4—the tubes now contain the developing reagent at 500, 250, 125, 62.5, and 31.25 ng/ml. 8. Dispense 50 µl of the developing reagent solutions into the wells of rows 2 to 6 of each plate, dispensing the most dilute solution into row 6 and solutions of increasing concentration successively into rows 5, 4, 3, and 2. Incubate 2 hr at room temperature. 9. Wash plates as in steps 9 to 11 of the basic protocol. Measure hydrolysis 10. Add 75 µl MUP or NPP substrate solution to each well, incubate 1 hr at room temperature, and measure the degree of hydrolysis visually or with a microtiter plate reader. An appropriate assay configuration results in 0.50 absorbance units/hr at 405 nm when using NPP as a substrate or 1000 to 1500 fluorescence units/hr when using MUP as a substrate. These results can be used to adjust optimal concentrations in the basic and alternate protocols.

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SUPPORT PROTOCOL

PREPARATION OF BACTERIAL CELL LYSATE ANTIGENS A culture of E. coli containing proteins expressed from cloned genes is lysed for use as test antigen in any of the first three protocols of this unit. For more extensive discussion on protein expression for antigen production, see UNITS 16.4-16.7 (expression by fusion protein vectors). Materials Escherichia coli culture in broth or agar (UNITS 1.2 & 1.3) Cell resuspension buffer Lysozyme solution Tris/EDTA/NaCl (TEN) buffer (UNIT 11.1) 10% sodium dodecyl sulfate (SDS) 8 M urea (optional) Nylon-tipped applicator (Falcon #2069, Becton Dickinson) 1. For liquid culture, centrifuge 5 ml of cells at 2500 rpm in a tabletop centrifuge for 10 min. Decant supernatant and resuspend pellet in 5 ml cell resuspension buffer by vortexing gently. For agar culture, remove about 10 colonies from the plate using a nylon-tipped applicator and resuspend in 2 ml cell resuspension buffer. Press swab against side of tube to remove as much liquid as possible. Yield of expressed protein may vary with growth phase. Samples should be taken for analysis at various periods of growth (e.g., mid-log and stationary phases). If samples are taken from agar plates, the culture should be grown overnight at 37°C.

2. Place 1 ml of resuspended cells in a microcentrifuge tube on ice. 3. Add 0.2 ml lysozyme solution to the tube and leave 5 min on ice. 4. Microcentrifuge 5 min. Decant supernatant and save. Resuspend pellet in 1.2 ml TEN buffer. Since many expressed proteins are insoluble, it is worthwhile to assay both the pellet and supernatant for activity.

5. Add 0.065 ml of 10% SDS solution to each sample. Incubate 10 min at 37°C. Samples are ready for ELISA at this point. Store frozen if not used within several hours. Alternatively, add urea to a final concentration of 8 M (4.8 g to a final volume of 10 ml) to denature and solubilize proteins.

REAGENTS AND SOLUTIONS Borate-buffered saline (BBS) 0.017 M Na2B4O7⋅10H2O 0.12 M NaCl Adjust to pH 8.5 with NaOH Blocking buffer BBS (see above) containing: 0.05% Tween 20 1 mM EDTA 0.25% bovine serum albumin (BSA) 0.05% NaN3 Store at 4°C Enzyme-Linked Immunosorbent Assays

Gelatin may be substituted for BSA; 5% instant milk has been successfully used but may interfere nonspecifically with antibody binding.



Cell resuspension buffer (10 mM HEPES) 2.38 g HEPES Add H2O to 1 liter Lysozyme solution 5 mg chicken egg white lysozyme (Sigma Grade VI #L2879) 1 ml TEN buffer (UNIT 11.1) Make fresh immediately before use MUP substrate solution 0.2 mM 4-methylumbelliferyl phosphate (MUP; Sigma #M8883) 0.05 M Na2CO3 0.05 mM MgCl2 Store at room temperature NPP substrate solution 3 mM p-nitrophenyl phosphate (NPP; Sigma #104-0) 0.05 M Na2CO3 0.05 mM MgCl2 Store at 4°C Test antibody solution Hybridoma supernatants (UNIT 11.10) can usually be diluted 1:5 and ascites fluid and antisera (UNIT 11.12) diluted 1:500 in blocking buffer and still generate a strong positive signal. Dilutions of nonimmune ascites or sera should be assayed as a negative control. Prepare antibody dilutions in cone- or round-bottom microtiter plates before adding them to antigen-coated plates. Sources of appropriate antibodies and conjugates can be found in Linscott’s Directory of Immunological and Biological Reagents.

Test antigen solution 0.2 to 10 µg/ml antigen, purified or partially purified in PBSN; store at 4°C Wash buffer Hanks balanced salt solution (HBSS; APPENDIX 2) 1% fetal calf serum (FCS; heat-inactivated 60 min, 56°C) 0.05% NaN3 Store at 4°C COMMENTARY Background Information Since their first description in 1971 (Engvall and Perlman), ELISAs have become the system of choice when assaying soluble antigens and antibodies. Factors that have contributed to their success include their sensitivity, the long shelf-life of the reagents (alkaline phosphatase conjugates typically lose only 5% to 10% of their activity per year), the lack of radiation hazards, the ease of preparation of the reagents, the speed and reproducibility of the assays, and the variety of ELISA formats that can be generated with a few well-chosen reagents. Additionally, no sophisticated equipment is necessary for many ELISA applications, including

screening hybridoma supernatants for specific antibodies and screening biological fluids for antigen content. The ELISAs described here combine the special properties of antigen-antibody interactions with simple phase separations to produce powerful assays for detecting biological molecules. The multivalency of antibodies can result in the formation of long-lived antigen-antibody complexes, thus allowing long periods of time during which such complexes can be measured. By designing an assay so that a capture reagent initiates the binding of antigen-antibody complexes and enzyme conjugates onto a solid phase, the unbound reagents can be easily and Immunology


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rapidly separated from the solid phase. The solid phase is washed and the amount of bound conjugate is visualized by incubating the solid phase with a substrate that forms a detectable product when hydrolyzed by the bound enzyme. ELISAs are similar in principle to radioimmunoassays, except that the radioactive label is replaced by an enzyme conjugate. A number of different enzymes have been successfully used in ELISAs, including alkaline phosphatase, horseradish peroxidase, βgalactosidase, glucoamylase, and urease. Alkaline phosphatase—perhaps the most widely used conjugated enzyme—is recommended because of its rapid catalytic rate, excellent intrinsic stability, availability, ease of conjugation, and resistance to inactivation by common laboratory reagents. Additionally, the substrates of alkaline phosphatase are nontoxic and relatively stable. Solutions of p-nitrophenyl phosphate (NPP) are stable for months at 4°C, while solutions of 4-methylumbelliferyl phosphate (MUP) can be kept for months at room temperature without any significant spontaneous hydrolysis. The biggest disadvantage of alkaline phosphatase is that if NPP is used as a substrate, the yellow color of the nitrophenyl product is relatively difficult to detect visually. Using the substrate MUP instead of NPP can greatly enhance the sensitivity of the assay. The fluorogenic system using MUP is 10 to 100 times faster than the chromogenic system using NPP, and appears to be as sensitive as an enhanced chromogenic assay in which alkaline phosphatase generates NAD+ from NADP (Macy et al., 1988). The disadvantage of using fluorogenic substrates is that they require a microplate fluorometer costing twice as much as a high-quality microtiter plate spectrophotometer. Cellular ELISAs have been shown to be as sensitive as flow cytometry analysis in detecting some cell-surface antigens (Bartlett and Noelle, 1987) and are potentially of great value in rapidly screening hybridoma supernatants for antibodies against surface molecules (Feit et al., 1983). Using ELISAs for screening large numbers of hybridoma supernatants has been hindered by the large number of cells required and high background signal. The increased sensitivity of the fluorogenic system should reduce the number of cells needed by a factor of 5, making the system more useful as a screening assay. In addition to the methods described here, hundreds of other ELISA applications have been described, including the determination of

antibody affinities (Beatty et al., 1987; Schots et al., 1988), the detection of antibodies specific for hormone receptors (Quinn et al., 1988; Wang and Leung, 1985), expression cloning of lymphokine receptors (Harada et al., 1990), and homogeneous assays in which a solid phase is not needed because the antigen-antibody interaction itself modifies the enzymatic activity (Rubenstein et al., 1972). A number of books are devoted to ELISAs and can be consulted for further discussion (Maggio, 1981; Kurstak, 1986).

Critical Parameters Sensitive ELISAs require antibodies of high affinity and high specificity. Since the sensitivity of an ELISA depends upon the affinity of the antibodies involved, antibodies with the highest affinities should be used when setting up ELISAs. Antibodies should be screened for unwanted cross-reactions. For instance, capture antibodies must not bind conjugate antibodies and vice versa. There are many commercial sources of reliable reagents. Linscott’s Directory of Immunological and Biological Reagents is an excellent source book for locating reagents used in ELISAs. If reagents from one source are inadequate, try another. When screening for expressed proteins in E. coli, it is important to utilize conjugates with antibodies that recognize nonnative and native molecules. Many foreign proteins expressed in E. coli will not assume their native conformation, and expression of such proteins will not be detected if antibody specific for the native form is used. It is also important to test enzymeantibody conjugates for cross-reactivity or nonspecific binding to host cell molecules. This potential problem can be eliminated by incorporating these as control antigens in the screening procedures used to select the original antibodies in the basic protocol. When coating plates with antigen, the antigen preparation need not be pure, but should generally comprise >3% of the protein in the coating solution. In some situations, dilution of the antigen solution with BSA has greatly improved the sensitivity of the ELISA (Jitsukawa et al., 1989). All steps after coating the microtiter plates should be carried out in solutions containing 0.05% Tween 20 and a carrier protein (0.25% BSA or gelatin). When using ELISAs for quantitative determinations of antigen or antibody concentrations, four guidelines should be followed.



First, it is essential that all experimental conditions up to the final wash after incubation with conjugate—including incubation times, wash times, reagent concentrations, and temperature—be kept constant between experiments. This is especially important in assays using polyvalent antibodies and complex mixtures of antigens. The optimal concentrations of all reagents for each system should be determined in an initial criss-cross serial dilution experiment (see first support protocol). Second, because the efficiency of binding and other micro-environmental conditions can vary from plate to plate, a standard curve should be included on each plate. Third, all samples must be analyzed at least in duplicate. Fourth, the concentration of the reagent being quantitated must lie within the dynamic range of the standard curve.

Anticipated Results Antibody-sandwich assays are generally the most sensitive ELISA configuration and can detect concentrations of protein antigens between 100 pg/ml and 1 ng/ml. ELISAs in which antigen is directly bound to plates are usually an order of magnitude less sensitive than sandwich techniques. Either the direct or sandwich ELISA may be used to detect and quantitate a bacterially expressed antigen or a purified or partially purified antigen in the range of 1 ng/ml to 1 µg/ml. A major disadvantage of the direct ELISA is that when an impure antigen preparation like a bacterial lysate is coated directly onto the surface of the microtiter well, the antigen must compete with all the other macro-molecules in the lysate for binding to the plastic and very little of the desired antigen may be bound. The sandwich ELISA bypasses this problem by relying on selective adsorption of an antigen to an antigen-specific antibody-coated surface.

Time Considerations

These assays are designed to take ∼6 hr, but the incubation times may be abbreviated or expanded as needed. Since equilibrium binding between the soluble and solid phases frequently takes 5 to 10 hr, stronger specific signals can usually be obtained by longer incubations. Fluorogenic ELISAs are generally 10 to 100 times faster than assays using chromogenic substrates.

Literature Cited Bartlett, W.C. and Noelle, R.J. 1987. A cell-surface ELISA to detect interleukin 4–induced class II MHC expression on murine B cells. J. Immunol. Methods 105:79-85. Beatty, J.D., Beatty, B.G., and Vlahos, W.G. 1987. Measurement of monoclonal affinity by noncompetitive immunoassay. J. Immunol. Methods 100:173-179. Coligan, J.E., Kruisbeek, A.M., Margulies, D.H., Shevach, E.M., and Strober, W., eds. 1991. Current Protocols in Immunology, Chapter 5: Immunofluorescence and cell sorting. Greene Publishing and Wiley-Interscience, New York. Engvall, E. and Perlman, P. 1971. Enzyme-linked immunosorbent assay (ELISA): Quantitative assay of immunoglobulin G. Immunochemistry 8:871-879. Feit, C., Bartal, A.H., Tauber, G., Dymbort, G., and Hirshaut, Y. 1983. An enzyme-linked immunosorbent assay (ELISA) for the detection of monoclonal antibodies recognizing antigens expressed on viable cells. J. Immunol. Methods 58:301-308. Harada, N., Castle, B.E., Gorman, D.M., Itoh, N., Schreurs, J., Barrett, R.L., Howard, M., and Miyajima, A. 1990. Expression cloning of a cDNA encoding the murine interleukin 4 receptor based on ligand binding. Proc. Natl. Acad. Sci. U.S.A. 87:857-861. Jitsukawa, T., Nakajima, S., Sugawara, I., and Watanabe, H. 1989. Increased coating efficiency of antigens and preservation of original antigenic structure after coating in ELISA. J. Immunol. Methods 116:251-257. Kurstak, E. 1986. Enzyme Immunodiagnosis. Academic Press, San Diego. Linscott’s Directory of Immunological and Biological Reagents, Santa Rosa, Calif. Macy, E., Kemeny, M., and Saxon, A. 1988. Enhanced ELISA: How to measure less than 10 picograms of a specific protein (immunoglobulin) in less than 8 hours. FASEB J. 2:3003-3009. Maggio, E.T. 1981. Enzyme Immunoassay. CRC Press, Boca Raton, Fla. Quinn, A., Harrison, R., Jehanli, A.M.T., Lunt, G.G., and Walsh, S.S. 1988. An ELISA for the detection of anti-acetylcholine receptor antibodies using biotinylated α-bungarotoxin. J. Immunol. Methods 107:197-203. Rubenstein, K.E., Schneider, R.S., and Ulmann, E.L. 1972. Homogeneous enzyme immunoassay: A new immunochemical technique. Biochem. Biophys. Res. Commun. 47:846. Schots, A., Van der Leede, B.J., De Jongh, E., and Egberts, E. 1988. A method for the determination of antibody affinity using a direct ELISA. J. Immunol. Methods 109:225-233.

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Wang, K.C. and Leung, B.S. 1985. Fluorometric ELISA methods for rapid screening of anti-estrogen receptor antibody production in hybridoma cultures. J. Immunol. Methods 84:279.

Key Reference Linscott’s Directory. See above. Highly recommended publication listing sources of immunological reagents, kits, and cells/organisms, including addresses and phone numbers of commercial suppliers (updated quarterly).

Contributed by Peter Hornbeck University of Maryland Baltimore, Maryland Scott E. Winston (bacterial cell lysate antigens) Univax Biologics Rockville, Maryland Steven A. Fuller (bacterial cell lysate antigens) Allelix Inc. Mississauga, Ontario




Isotype Determination of Antibodies

UNIT 11.3

Frequently it is necessary to know the amount and serological class of antibodies made by an immunized animal, produced by hybridomas, or present in the serum of patients with inflammatory or neoplastic conditions. The immunologist’s approach to such a problem is to consider the antibody or immunoglobulin molecules themselves as antigens and to use anti-immunoglobulin antibodies as the specific and sensitive agents of detection. This unit describes two methods for measurement and classification of supernatant or serum immunoglobulins—an ELISA (first basic protocol) and a method employing electrophoresis and immunofixation (second basic protocol). SANDWICH ELISA FOR ISOTYPE DETECTION The speed and sensitivity of sandwich ELISAs make them the assays of choice for isotype determination. The antibody-sandwich ELISA to measure soluble antigen (UNIT 11.2; Fig. 11.2.3) is adapted in this unit for isotype detection. Microtiter wells are coated with isotype-specific capture antibodies followed by incubation with test solutions containing the antibodies to be isotyped. After test antibodies have been bound to the plate by reacting with capture antibodies, unbound test antibodies are washed out. Developing reagent is added to the wells, followed by another incubation. Unbound conjugate is washed out and substrate is added. Substrate hydrolysis indicates that the test solution contained the appropriate isotype. A lack of hydrolysis indicates that the test solution did not contain the appropriate isotype. UNIT 11.2 should be consulted for additional information.

BASIC PROTOCOL

Materials Capture anti-isotype antibodies: heavy-chain class-specific antibodies (anti-µ, -α, -γ, -δ, -ε), heavy-chain subclass-specific antibodies (anti-γ1, -γ2a, -γ2b, -γ3, -γ4), or light-chain isotype-specific antibodies (anti-κ, -λ) PBS (APPENDIX 2) containing 0.05% NaN3 (PBSN) Test antibodies: hybridoma supernatants, ascites fluid, or antisera Blocking buffer (UNIT 11.2) Standard isotype antibodies (i.e., purified antibodies of known isotypes) Developing reagent: anti-Ig antibody (specific for all heavy-chain classes)–alkaline phosphatase conjugate (see UNIT 11.1; Southern Biotechnology or Linscott’s Directory) MUP or NPP substrate solution (UNIT 11.2) Immulon 2 or 4 microtiter plates (or equivalent; UNIT 11.2) Additional reagents and equipment for ELISA (UNIT 11.2) 1. Prepare the capture anti-isotype antibodies by diluting in PBSN to 2 µg/ml final. Capture antibodies can be monoclonal or polyclonal. When screening hybridoma supernatants for IgG (anti-γ), it is advisable to initially use a capture-antibody preparation that recognizes all IgG isotypes. Subsequent analysis using subclass-specific capture antibodies can determine which IgG subclass is expressed. It is crucial that the capture antibodies do not bind to the antibodies in the developing reagent.

2. Coat each well of an Immulon microtiter plate with 50 µl capture antibody solution as in steps 3 to 5 of the basic protocol in UNIT 11.2. 3. Block wells as in steps 6 and 7 of the basic protocol in UNIT 11.2. 4. Prepare dilutions of the test antibodies in blocking buffer—typically, hybridoma supernatants are diluted 1:5 while sera are diluted 1:500. Immunology Contributed by Peter Hornbeck, Thomas A. Fleisher, and Nicholas M. Papadopoulos Current Protocols in Molecular Biology (1992) 11.3.1-11.3.6 Copyright © 2000 by John Wiley & Sons, Inc.


5. Prepare the standard isotype antibodies at ∼500 ng/ml to serve as positive controls. 6. Transfer 50-µl aliquots of test or standard isotype antibodies into the antibody-coated wells and incubate >2 hr at room temperature. Specificity controls consisting of wells coated with capture antibodies and a series of standard antibodies of different isotypes should be included on each plate. Negative controls should be included on each plate and consist of wells coated with capture antibodies but receive no test or control antibodies.

7. Wash plate as in steps 9 to 11 of the basic protocol in UNIT 11.2. 8. Prepare the developing reagent so that the final conjugate solution contains ∼200 ng anti-Ig/ml. Crucial that antibodies in developing reagent do not bind to capture antibodies.

9. Add 50 µl developing reagent to each well and incubate 2 hr at room temperature. All test wells, positive-control wells, and negative-control wells should receive the developing reagent.

10. Wash as in steps 9 to 11 of the basic protocol in UNIT 11.2. 11. Add 75 µl MUP or NPP substrate solution to each well and incubate at room temperature. Periodically check the plate for substrate hydrolysis. Hydrolysis of NPP results in liberation of a yellow product that can be detected by visual inspection in ambient light. Hydrolysis of MUP results in the liberation of a fluorescent product that can be detected by visual inspection under a long-wavelength UV lamp in a darkened room. For more quantitative estimates of isotype concentrations, plates can be read with microtiter plate reader. Positive-control wells give strong signals by 1 hr. Weaker reactions can be detected by incubation for many hours or overnight. BASIC PROTOCOL

DETECTING AND ISOTYPING ANTIBODIES BY ELECTROPHORESIS AND IMMUNOFIXATION Qualitative identification and quantitative determination of serum (and other biological fluid) proteins provide useful information concerning pathologic conditions of the lymphoid system. High-resolution zone electrophoresis is a simple method to separate serum proteins based on their classification defined by five electrophoretic zones: albumin, α1-globulin, α2-globulin, β-globulin, and γ-globulin. Following electrophoresis, the proteins can be detected by staining with amido black, or immunofixation can be performed first to achieve more precise identification. In immunofixation procedure described below, immunoglobulins within separated protein bands are identified, and clonality is established, using antisera to the α, γ, µ, ε, and δ heavy chains and κ and λ light chains of immunoglobulins. Other serum proteins—including glycoproteins, transferrin, and C3—can be identified in electrophoresed sample by same technique. The serum (or other fluid) is loaded on an agarose gel–covered microscope slide and electrophoresed. Proteins are detected after zone electrophoresis and can be identified by immunofixation.


Materials Serum (or other biological fluid) Normal saline 95% methanol/5% acetic acid 1% amido black (1 g in 100 ml of 2.5% acetic acid) 2.5% (v/v) acetic acid



γ

β

α

albumin

globulins

–

+

Figure 11.3.1 Normal serum protein electrophoretic pattern obtained by agarose gel zone electrophoresis demonstrating the five protein zones.

2 × 2–cm cellulose acetate strip Monospecific anti-Ig, heavy-chain-specific (α, γ, µ, ε, or δ) or light-chain-specific (κ or λ) 0.85% (w/v) NaCl Agarose gel–covered microscope slides Plexiglas electrophoresis cell with two agarose bridges Electrophoresis power supply (e.g., Pharmacia EPS 500/400) Electrophorese the sample 1. Cut a narrow slit with a razor blade in the middle of an agarose gel–covered microscope slide. For standard fixation and staining (steps 3 and 4), place 1.5 µl undiluted serum in the slit. For immunofixation (steps 5 to 8), dilute serum in normal saline to generate 1 to 2 mg/ml of the specific protein being evaluated (e.g., IgG, IgA, or IgM) and place in the slit. 2. Place the slide in an electrophoresis cell with two agarose bridges having the same composition as the gel on the slide. Electrophorese 12 min at 140 V. Proceed to steps 3 and 4 for standard fixation and staining. Proceed to steps 5 to 8 for immunofixation. Standard fixation and staining 3. Fix slides by submerging in 95% methanol/5% acetic acid 15 min. Air dry with filter paper placed on the gel. 4. Stain agarose gel by submerging in 1% amido black 10 min. Destain in 2.5% acetic acid until the background clears, then rinse in water. Dry 5 min at 60°C in a drying oven for visual inspection (Fig. 11.3.1). Immunofixation 5. Transfer the unstained and unfixed gel (from step 2) to a petri dish containing damp filter paper. Overlay the γ-globulin zone of the gel with a cellulose acetate strip impregnated with a monospecific anti-Ig. The γ-globulin zone is that nearest the cathode (Fig. 11.3.1). The cellulose acetate strip is cut from any commercial membrane and dipped into the appropriate antibody solution (see critical parameters). The concentration of the antibody solution provided by supplier (usually 0.5 to 1 mg/ml) is suitable.

6. Allow the antibody to diffuse into the gel 15 min at room temperature. 7. Remove the the slides and immerse in 0.85% NaCl for 2 hr to wash out unfixed proteins. Air dry with filter paper placed on the gel. 8. Stain as described in step 4 and visually inspect for immunoprecipitated protein bands (Fig. 11.3.2).

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REAGENTS AND SOLUTIONS Agarose gel–covered microscope slides Add 0.5 g agarose (Seakem agarose HE, FMC Bioproducts) to 100 ml of 0.05 M barbital buffer (Sigma #B0500); store at room temperature and heat to 100°C. Stir until agarose has dissolved and allow to cool to 70°C. Pour 2.5 ml of the agarose solution on each microscope slide and allow to gel 2 to 3 min at room temperature. Store the prepared slides at 4°C in a humidified chamber. Set up a chamber by half-filling a petri dish with the above agarose solution. Allow to cool and place in a refrigerator. This humidified chamber can be used to store agarose slides up to 2 weeks. COMMENTARY Background Information Sandwich ELISA Antibodies are heteromeric molecules consisting of heavy and light chains, each of which contains a variable and a constant region. Heavy-chain constant regions include µ, α, γ1, γ2a, γ2b, γ3, γ4, δ, or ε, depending upon the species; light-chain constant regions include κ and λ. Immunoglobulin constant regions, commonly referred to as isotypes, determine many of the biological and immunochemical properties of the antibody molecule including complement fixation, binding to Fc receptors, and binding to proteins A and G. Because the iso-

type may influence the method of purification (Andrew and Titus, 1991), it is routine to determine the isotypes of monoclonal antibodies or other specific antibody preparations as part of their initial characterization. The identification of antibody isotypes can easily be performed with an ELISA employing commercially available anti-isotype reagents. Alternatively, isotypes can be determined using electrophoresis/immunofixation (second basic protocol) or a double-immunodiffusion assay (Hornbeck, 1991). See UNIT 11.2 for a full discussion of the ELISA technique.

A

+

+

γ

β

α

– albumin

globulins

–

monoclonal band –

B

anti-µ

+

C


patient

– anti-κ

Figure 11.3.2 Agarose gel zone electrophresis of patient serum demonstrating a monoclonal band in the γ-globuling zone (A; arrow). Immunofixation electrophoresis with anti-µ (B) and anti-κ (C) demonstrate that the band is a µ-κ monoclonal immunoglobulin. There is no reactivity with the antisera to the other heavy (α, γ, δ, ε) and light (λ) chains (data not shown).



Electrophoresis and immunofixation Zone electrophoresis is based on the principle that charged particles migrate at different rates in an electric field based on the net charge of the particle. The application of this method to the evaluation of serum proteins was first described in a seminal paper by Tiselius (1937). In this paper it was shown that serum proteins were separable into defined zones (albumin, α-globulins, β-globulins, and γ-globulins). While protein separation by zone electrophoresis is excellent, protein quantitation using this method is poor. To overcome these shortcomings, various support media have been employed, including cellulose acetate, agar gel, and, more recently, agarose gel. Agarose gel provides improved stability and clarity, as well as greater sensitivity than other materials (Papadopoulos et al., 1982). Immunofixation electrophoresis combines the high resolution of agarose gel zone electrophoresis and the unique specificity of an antigen-antibody reaction (Johnson, 1982). After electrophoretic separation, the antigen of interest is reacted with an overlayed, monospecific antibody to form an immunoprecipitate, which can be easily detected. The location of the immunoprecipitate depends on the electrophoretic migration of the specific protein antigen. The unreacted proteins and antibody reagents are washed out of the agarose gel and the precipitin band is stained for visualization. Using this method, polyclonality, oligoclonality, or monoclonality can be ascertained (Fig. 11.3.2). Commercial sources for complete immunofixation electrophoresis setups are available. Immunoelectrophoresis, a classic method in which diffusion of antibody into the gel is combined with electrophoresis, is an alternative method for evaluation of protein clonality. However, this approach is less sensitive and more difficult to interpret as compared with immunofixation (Johnson, 1986).

Critical Parameters and Troubleshooting Sandwich ELISA Sources of isotype-specific antibodies can be found in Linscott’s Directory of Immunological and Biological Reagents (see key references, UNIT 11.2). Isotype-specific antibodies should have no detectable cross-reactivity against other isotypes and should not cross-react with other antibodies that might be used in the assay. Check isotype-specific reagents against standard isotype proteins to confirm

their specificity. Sources of purified isotypes from the species of interest, to be used as experimental standards, can also be found in Linscott’s Directory. Alternatively, standard isotype proteins can be prepared from myeloma and hybridoma lines of known isotypes using standard techniques (UNIT 11.8). Many myelomas and hybridomas of defined isotypes and specificities are available from ATCC and other sources; see full listing in Knapp et al. (1991). The concentration of the developing reagent should be adjusted so that the positive control gives a strong signal by 1 hr. Since the hydrolysis of MUP is at least 10-fold easier to detect than the hydrolysis of NPP, assays using MUP can be significantly faster than those using NPP. Test solutions should be scored as positive only when they give 3-fold higher signals than the negative controls. While this assay is designed to qualitatively determine the presence or absence of a given isotype in the test solution, it can be easily modified to quantitate the concentration of isotype by including serial dilutions of standard isotype proteins (for details of quantitation using a standard curve, see UNIT 11.2 protocol for antibody-sandwich ELISA to measure soluble antigen). Electrophoresis and immunofixation Antigen and antibody interaction at or near the point of equivalence results in the formation of immune complexes that produce an insoluble precipitate. In the case of a monoclonal (homogeneous) protein, this precipitate is found in a very narrow band, while a polyclonal protein will generate a broad band. Immunoprecipitation is optimal at antigen/antibody equivalence. It is often useful to quantitate total immunoglobulin levels in the sample to allow for the dilution of the serum (see step 1 of basic protocol). Antigen excess will result in clear spots (lack of detectable immunoprecipitate) in the location where the band(s) are anticipated. If this occurs, dilute the sample and repeat immunofixation electrophoresis. Failure to see a clearly discernible electrophoretic protein pattern after step 4 suggests a technical problem at some point during steps 1 through 4. Immunofixation should not be performed and the initial electrophoresis procedure should be repeated. Absence of a detectable immunoprecipitate after step 8 (in the presence of a gamma globulin band after step 4) suggests antigen excess. Quantitate immunoglobulins, dilute serum accordingly, and repeat entire experiment.

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Anticipated Results Sandwich ELISAs are typically sensitive to 0.5 to 2.0 ng/ml of antibody isotype. During electrophoresis and immunofixation, the specific bands observed in immunofixation electrophoresis are dependent on both the presence of particular proteins and the appropriate antisera being used in the detection process. In the case of normal immunoglobulins, all three major isotypes should be observed. Monoclonal or oligoclonal immunoglobulins produce single or multiple darker (clonal) bands (Fig. 11.3.2). The detection limit for a specific protein using this immunofixation technique is 5 to 10 µg/ml.

Time Considerations The sandwich ELISA requires 6 to 8 hr. However, the times allotted for the various incubation steps can usually be reduced by half, so results can be obtained in 3 to 4 hr. For electrophoresis and immunofixation, running and developing the gel takes 2 to 3 hr. Additional time (10 to 15 min) is needed to prepare and titrate the serum sample if immunoglobulin levels are increased.

Literature Cited Andrew, S.M. and Titus, J.A. 1991. Purification of immunoglobulin G. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.7.1-2.7.12. Greene Publishing and WileyInterscience, New York. Hornbeck, P. 1991. Double-immunodiffusion assay for detecting specific antibodies. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeeck, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.3.1-2.3.4. Greene Publishing and Wiley-Interscience, New York.

Johnson, M.A. 1982. Immunofixation electrophoresis. Clin. Chem. 28:1797-1800. Johnson, M.A. 1986. Immunoprecipitation in gels. In Manual of Clinical Laboratory Immunology. (N.R. Rose, H. Friedman, and J.L. Fahey, eds.) pp. 14-24. Am. Soc. Microbiol., Washington, D.C. Knapp, W., Stockinger, H., Majdic, O., and Shevach, E.M. 1991. The CD system of leukocyte surface molecules. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeeck, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. A.4.1A.4.28. Greene Publishing and Wiley-Interscience, New York. Papadopoulos, N.M., Elin, R.J., and Wilson, D.M. 1982. Incidence of γ banding in a healthy population by high-resolution immunofixation electrophoresis. Clin. Chem. 28:707-708. Tiselius, A. 1937. A new apparatus for electrophoretic analysis of colloidal mixtures. Trans. Faraday Soc. 33:524-526.

Key References Johnson, 1986. See above. A concise discussion of the principles of immunoprecipitation with specific reference to immunofixation electrophoresis. Maggio, E.T. 1981. Enzyme Immunoassay. CRC Press, Boca Raton, Fla. A valuable reference describing parameters of ELISA technology.

Contributed by Peter Hornbeck (sandwich ELISA) University of Maryland Baltimore, Maryland Thomas A. Fleisher and Nicholas M. Papadopoulos (electrophoresis and immunofixation) Warren Grant Magnuson Clinical Center Bethesda, Maryland




PREPARATION OF MONOCLONAL ANTIBODIES

SECTION II

The preparation of monoclonal antibodies should be undertaken carefully, since the production of monoclonal antibodies is expensive and time-consuming. Figure 11.4.1 summarizes the experimental procedures that must be carried out to prepare monoclonal antibodies. The various procedures are presented in individual protocols and described in sufficient detail to allow an individual with no prior experience to carry out a cell fusion, to produce monoclonal antibodies in ascites fluid, and to purify antigen-specific monoclonal antibodies.

immunization of mice ELISA screening of sera

isolation of spleen cells

preparation of myeloma cells

feeder cells cell fusion

ELISA screening of hybridoma supernatants

expansion and selection of cultures to be cloned

freeze cells

feeder cells

cloning by limiting dilution

isolation and expansion of clones

freeze cells

recovery of frozen cells production of ascites fluids

purification of monoclonal antibodies

Figure 11.4.1 Flow chart for preparation of monoclonal antibodies.

Immunology

Contributed by Steven A. Fuller, Miyoko Takahashi, and John G.R. Hurrell

11.4.1

Current Protocols in Molecular Biology (1992) 11.4.1-11.4.6 Copyright © 2000 by John Wiley & Sons, Inc.

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UNIT 11.4

Immunization of Mice Antigen is prepared for injection either by emulsifying an antigen solution with Freunds adjuvant or by homogenizing a polyacrylamide gel slice containing the protein antigen. Mice are immunized at 2- to 3-week intervals. Test bleeds are collected 7 days after each booster immunization to monitor serum antibody levels. Mice are chosen for hybridoma fusions when a sufficient antibody titer is reached.

BASIC PROTOCOL

PRODUCTION OF IMMUNE SPLEEN CELLS: IMMUNIZATION WITH SOLUBLE ANTIGEN Materials Phosphate-buffered saline (PBS; APPENDIX 2) Antigen Complete Freunds adjuvant Any strain mice, 6 to 8 weeks old Incomplete Freunds adjuvant 22-G needles 3-ml syringes with locking hubs (Luer-Lok, Becton Dickinson) Double-ended locking hub connector (Luer-Lok, Becton Dickinson) Sterile sharp scissors Sterile razor blades or scalpel blades Wooden applicator sticks 200-µl pipettor Additional reagents and equipment for ELISA (UNITS 11.2 & 11.3) and western blotting (optional; UNIT 10.8) 1. Prepare an emulsion (200 to 400 µl/mouse) of equal volumes PBS containing 25 to 100 µg antigen and complete Freunds adjuvant. Using a 22-G needle, inject mice intraperitoneally. For each antigen, 3 to 5 mice are immunized. Complete Freunds adjuvant contains mycobacteria—incomplete Freunds does not. An emulsion is most readily prepared by linking two locking syringes, one loaded with antigen and the other loaded with adjuvant, using a double-ended locking connector (see Fig. 11.4.2). Press syringe barrels back and forth, transferring contents from one syringe to the other, for 5 to 10 min until a stable emulsion is produced. For best antibody production, inject antigen in as small an emulsion volume as practicable. A stable emulsion is an oil-in-water emulsion which will not disperse when dropped into water. This is a useful check for the emulsification endpoint. Further, at the endpoint the emulsion will thicken noticeably. Mice may be restrained for immunization in the following manner: Place mouse on grilled cage top. Lift mouse by the tail (generally, when mice are lifted by the tail they will grab the bars of the cage top with their front feet, thus stabilizing themselves for restraint). Immobilize the mouse’s head by pinching together the skin at the base of the skull between thumb and forefinger. Turn hand over so that mouse is lying with its back against the palm. Wrap fourth finger around tail and stretch mouse over arched palm for intraperitoneal injection. CAUTION: Handle Freunds adjuvant carefully, since self-injection can cause a positive TB test and lead to a granulomatous reaction.

Immunization of Mice

2. Boost mice 3 weeks later by intraperitoneally injecting an emulsion (200 to 400 µl) of equal volumes PBS containing 10 to 50 µg antigen and incomplete Freunds adjuvant. The emulsion is prepared and injected as in step 1.



3-ml glass syringes double-ended Luer-Lok connector or 3-way stopcock

antigen and adjuvant

Figure 11.4.2 Double-syringe device for preparation of antigen-adjuvant emulsions.

3. Bleed mice 7 days after second immunization by cutting off 0.5 cm of the tail with sterile sharp scissors or a razor blade. Collect 100 to 200 µl blood into a 1.5-ml microcentrifuge tube. After clot formation, rim the clot with a wooden applicator stick to dislodge the clot from the surface of the tube, but do not break up the clot. After clot retraction, transfer the serum into another microcentrifuge tube with a 200-µl pipettor. If test bleeds are collected more than three times, it will be necessary to cut the tail vein to obtain further samples rather than cutting off additional lengths of the tail itself. This is done by nicking one of the lateral tail veins with a razor blade. The collection of blood may be facilitated by using a heat lamp to warm the mouse for 30 sec to 1 min prior to cutting of the tail. Additionally, if blood flow from the cut tail is slow, the tail may be “milked” from base to the cut tip with thumb and forefinger.

4. Determine the antibody titer in the serum by ELISA (UNITS 11.2 & 11.3). If desired, further characterize the antibody specificity by western blotting (UNIT 10.8). Antibody titer is operationally defined as that dilution of serum that results in 0.2 absorbance units above background in the ELISA procedure.

5. If the antibody titer is considered too low (≥1⁄1000) for cell fusion, mice can be boosted every 2 weeks until an adequate response is achieved. Bleed the mice and test the serum with an ELISA. 6. When the antibody titer is sufficient (>1⁄1000), boost mice by injecting 10 to 50 µg antigen in PBS intraperitoneally (200 to 400 µl), or intravenously (50 to 100 µl) via the tail veins, 3 days before fusion but >2 weeks after previous immunization. In general, the higher the serum antibody titer, the more antigen-specific antibody-producing hybridomas are obtained per fusion. If an antibody against a nonimmunodominant epitope is desired, the cell fusion may be done at an earlier or later time, since the percentage of antibody-producing cells in the spleen directed at these less immunogenic regions of the antigen may vary with time in an unpredictable fashion.

7. Perform cell fusion (UNIT 11.7) 3 days after the immunization (step 6). Immunology


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ALTERNATE PROTOCOL

IMMUNIZATION WITH COMPLEX ANTIGENS (MEMBRANES, WHOLE CELLS, AND MICROORGANISMS) 1. Prime the mice and boost intraperitoneally with adjuvant (i.e., complete Freunds for priming and incomplete Freunds for booster immunizations) as described for soluble antigen (see basic protocol, steps 1 and 2) or suspend antigen in PBS and inject. Use 1 to 2 × 107 cells for mammalian species or 108 to 109 bacterial or yeast cells. 2. Bleed the mice and determine the antibody titer of the serum as described for soluble antigen (see basic protocol, steps 3 to 6). 3. Perform cell fusion (UNIT 11.7) 3 days after final immunization.

ALTERNATE PROTOCOL

IMMUNIZATION WITH ANTIGEN ISOLATED BY ELECTROPHORESIS In some instances the antigen under investigation can be purified most conveniently by gel electrophoresis (UNIT 10.2). Mice can be immunized with protein antigens still contained in a polyacrylamide gel slice, as described in this protocol. Additional Materials 0.1 M KCl, cold Tissue grinder Additional reagents and equipment for denaturing (SDS) discontinuous gel electrophoresis (UNIT 10.2) 1. Apply a protein mixture containing 10 to 50 µg of the desired protein antigen to an appropriate denaturing (SDS) discontinuous gel electrophoresis system (e.g., the Laemmli gel system) and complete the electrophoresis as described in UNIT 10.2. 2. Soak gel 5 to 15 min in cold 0.1 M KCl. Protein bands will appear as white precipitates against a clear gel background. 3. Cut out the appropriate bands from the gel with a razor blade or scalpel blade. 4. Prepare gel suspension by homogenizing the gel slice in a minimum volume of PBS using a tissue grinder. Minimum volume is defined by adding successive 100-µl volumes of PBS until the homogenized gel is liquid. Alternatively, the gel may be air dried for 1 to 2 hr, smashed with a glass rod, and suspended in a minimum volume of PBS.

5. Immunize each mouse with 200 to 400 µl gel suspension containing 10 to 50 µg antigen via an intraperitoneal injection. Amount of antigen is estimated from prior observation of the proportion of desired protein antigen to other antigens in the sample as determined by the relative intensity of stained bands on the polyacrylamide gel (see UNIT 10.6 for staining procedures).

6. Boost mice after 3 weeks with 200 to 400 µl gel suspension containing 10 to 25 µg antigen. 7. Bleed the mice and determine the antibody titer of the serum as described for soluble antigen (see basic protocol). Mice immunized repeatedly with polyacrylamide tend to form adhesions that can make aseptic removal of the spleen difficult.

8. Perform cell fusion (UNIT 11.7) 3 days after final immunization. Immunization of Mice



COMMENTARY Background Information The stimulation of an effective humoral immune response in mice is critical to the production of monoclonal antibodies directed at a particular antigen. The variety and quality of the monoclonal antibodies prepared is generally directly proportional to the serum antibody titer in the particular mouse used for cell fusion. Any means of antigen preparation, antigen delivery, or immunization schedule that increases antibody titer in the serum of the immunized mouse will potentiate the isolation of hybridomas secreting monoclonal antibodies of interest. We have described two methods of antigen preparation: (1) antigen emulsified in Freunds adjuvants (probably the most common technique used) and (2) antigen isolated in a polyacrylamide gel slice and homogenized. Other preparation methods (e.g., adsorption of antigen to supports such as aluminum hydroxide or aluminum phosphate, polystyrene beads, or nitrocellulose paper, and alternate sites of injection such as footpads) are discussed in the key references.

polyacrylamide gel slice), site of immunization (intraperitoneal versus footpad or tail vein), antigen dose, and frequency of immunization. Alternate immunization protocols are presented in the key references below.

Anticipated Results Isolation of high-quality monoclonal antibodies correlates with high-serum antibody titers. A serum ELISA titer of 1⁄1000 is the minimum level before attempting a cell fusion. Titers for most antigens (particularly from animals injected with highly purified antigen) will range from 1⁄1000 to 1⁄100,000 after 3 to 4 immunizations. Occasional serum samples will titer at greater than 106. The proportion of monoclonal antibodies of IgG class rather than IgM class generally increases proportionally to the duration of the immunization schedule, although this can vary dramatically among different antigens. [In general, IgG class antibodies are more suitable for immunoassays, western blotting (U NIT 10 .8), immunoaffinity chromatography (UNIT 10.11), and immunoprecipitation (UNIT 10.16)].

Critical Parameters It is desirable to use antigen of the highest available purity for immunizations, particularly for primary immunizations. Contaminants may be more immunogenic than the antigen of interest and as such may result in a low specificity antibody. Mice given primary immunizations of highly pure antigen may be boosted with less pure material (containing as little as one-third specific antigen in a complex protein mixture).

Troubleshooting Poor success in raising an adequate antibody titer to an antigen of interest can be attributed to several factors. Improperly prepared emulsion when using Freunds adjuvant (i.e., the aqueous and oil phases separate upon standing) is ineffective in stimulation of an immune response. Contaminants in an antigen preparation may be more immunogenic, necessitating a more homogeneous preparation of the desired antigen. Other parameters that can be varied in an effort to produce a higher antibody titer and increased specificity include presentation of antigen (Freunds adjuvant emulsion versus

Time Considerations A primary immunization followed by two booster immunizations and test bleeds will occupy 6 weeks. For many antigens, however, an adequate antibody response in the mice is achieved only after several months and multiple immunizations.

Key References Hurrell, J.G.R., ed. 1982. Monoclonal Hybridoma Antibodies: Techniques and Applications. CRC Press, Boca Raton, Fla. Langone, J.J. and Van Vunakis, H., eds. 1986. Immunological techniques, Part I: Hybridoma technology and monoclonal antibodies. Methods Enzymol. 121:1-947.

Contributed by Steven A. Fuller and Miyoko Takahashi ADI Diagnostics Rexdale, Ontario John G.R. Hurrell Boehringer Mannheim Diagnostics Indianapolis, Indiana

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Supplement 18


Preparation of Myeloma Cells Myeloma cells are cultured with 8-azaguanine to ensure their sensitivity to the HAT selection medium (see UNIT 11.6) used after cell fusion (UNIT 11.7). One week prior to cell fusion, myeloma cells are grown in medium without 8-azaguanine. Cell culture conditions are adjusted such that the Sp2/0 cells are in the log phase of growth and exhibit high viability at the time of collection for fusion (UNIT 11.7). Materials Sp2/0 murine myeloma cell line (ATCC #CRL 1581) Complete culture medium 20 µg/ml 8-azaguanine Tissue culture flasks, 25 cm2 or 75 cm2 8% CO2-in-air gas mixture Humidified 37°C, 8% CO2 incubator Inverted microscope 1. Recover frozen cells from liquid N2 storage, as described in UNIT 11.9. 2. Grow Sp2/0 cells overnight in complete medium in tissue culture flasks at 37°C in a CO2 incubator in 8% CO2-in-air atmosphere with 98% relative humidity. 3. Determine that the cells are growing by examining the cell cultures in the flasks with an inverted microscope and return culture flask to CO2 incubator for continuation of cell growth. 4. To ensure that the Sp2/0 cells remain aminopterin sensitive for the selection process following fusion, supplement the complete culture medium with 8-azaguanine at 20 µg/ml during maintenance. One week prior to fusion, culture cells in medium without 8-azaguanine. A seeding cell density of 2.5 to 5 × 104 cells/ml works well with Sp2/0 cells. Sp2/0 cells will grow to a maximum density of 6 to 9 × 105 cells/ml, with a doubling time of 10 to 15 hr. When this density is reached, there is a rapid decline in cell viability. The Sp2/0 cultures are split every 2 to 3 days either by discarding an appropriate volume from the old flask and replacing with fresh medium or by transferring an appropriate volume of cells to a new flask and adding fresh medium. A 1-in-10 or 1-in-20 split is recommended.

5. A total of 1 × 107 Sp2/0 cells (i.e., 1:10 ratio to immune spleen cells) is used for fusion. Cell viability at the time of collection should be greater than 95%. To ensure that cells are collected in log phase of growth, adjust the cell density to 2 × 105 cells/ml the day before the fusion by adding fresh medium. Determine cell viability using the trypan blue exclusion method (see support protocol, below) on cells suspended in serum-free medium or PBS. SUPPORT PROTOCOL

CELL VIABILITY TEST BY TRYPAN BLUE EXCLUSION This procedure is used to determine the number of viable cells present in the cell culture. A non-viable cell will have a blue cytoplasm; a viable cell will have a clear cytoplasm. Additional Materials

Preparation of Myeloma Cells


Phosphate-buffered saline (PBS; APPENDIX 2) or serum-free complete culture medium 0.4% trypan blue solution Contributed by Steven A. Fuller, Miyoko Takahashi, and John G.R. Hurrell Current Protocols in Molecular Biology (1988) 11.5.1-11.5.3 Copyright © 2000 by John Wiley & Sons, Inc.

Binocular microscope Hemacytometer 1. Centrifuge 1 ml cell suspension at 100 × g for 5 min. 2. Resuspend the cell pellet in 1 ml PBS or serum-free complete culture medium. Serum proteins stain with trypan blue and can produce misleading results. Determinations must be made in serum-free solution.

3. Mix 1 part of trypan blue solution and 1 part cell suspension (1⁄2 dilution). 4. Using a binocular microscope, count the unstained (viable) and stained (dead) cells separately in a hemacytometer. Each of the four corner squares (composed themselves of 16 smaller squares) have 1 mm sides and are 0.1 mm deep (0.1 mm3). Count all cells within each of the four corner squares, including those that lie on the bottom and left-hand perimeters, but not those that lie on the top and right-hand perimeters. Count any clumps of cells as one cell. Calculate the mean number of cells per 0.1-mm3 volume. Multiply by 104 to obtain the number of cells/ml (i.e., cells/cm3). Apply dilution factor for trypan blue (2×) to obtain the number of cells per milliliter of culture. 5. Calculate the percentage of viable cells as follows: Number of viable cells Viable cells (%) = × 100 Total number of cells (dead and viable) REAGENTS AND SOLUTIONS Complete culture medium Dulbecco modified Eagle medium (DMEM), high-glucose formula (4.5 g glucose/ liter; GIBCO/BRL #430-2100) supplemented to the indicated concentrations with the following additives: 2.8 g/liter sodium bicarbonate (33.3 mM) 4.8 g/liter HEPES (20 mM) 10% fetal calf serum (v/v) 10 ml/liter L-glutamine (2 mM) 10 ml/liter sodium pyruvate (1 mM) 10 ml/liter penicillin (50 IU/ml) and streptomycin (50 µg/ml) The last four additives are available as 100× solutions from GIBCO/BRL and other major suppliers of cell culture media. Penicillin and streptomycin are combined in one solution. Samples of fetal calf serum lots should be tested for ability to support efficient cell growth and cloning before a large purchase because there is much variability between lots of a given supplier. The fetal calf serum must be mycoplasma free. If low volume usage of fetal calf serum precludes testing of serum lots, purchase of mycoplasma-free, virus-free, low endotoxin sera from suppliers such as GIBCO/BRL, Flow Laboratories, or Sigma will generally provide satisfactory results. Horse or bovine serum is not an adequate substitute!

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Supplement 1

COMMENTARY Background Information The Sp2/0 cell line was chosen as the fusion partner for immune spleen cells because of its good rate of growth, the efficiency with which hybridomas are obtained after fusion, and, most importantly, because it does not synthesize or secrete any immunoglobulin heavy or light chains itself. The Sp2/0 myeloma cell line was developed by Schulman et al. (1978). Other commonly used cell lines are P3X63-Ag8.653 (Kearney et al., 1979), which does not secrete immunoglobulins, and NS-1 (Kohler and Milstein, 1976), which produces only κ light chains.

Critical Parameters Optimal growth of myeloma cells is density dependent. Cultures should be split at regular intervals to maintain >95% viability. Do not culture Sp2/0 cells longer than 1 month to avoid genetic drift and development of antibiotic-resistant contaminants. Maintain several aliquots of Sp2/0 cells in liquid nitrogen storage.

Anticipated Results

myeloma cell culture able to sustain good production of hybridomas upon fusion.

Time Considerations Depending on culture conditions, 105 cells can be expanded to the 107 cells required for fusion in 4 to 6 days.

Literature Cited Kearney, J.F., Radbruch, A., Liesegang, B., and Rajewsky, K. 1979. A new mouse myeloma cell line that has lost immunoglobulin expression but permits the construction of antibody-secreting hybrid cell lines. J. Immunol. 123:1548-1550. Kohler, G. and Milstein, C. 1976. Fusion between immunoglobulin-secreting and nonsecreting myeloma cell lines. Eur. J. Immunol. 6:511-519. Schulman, M., Wilde, C.D., and Kohler, G. 1978. A better cell line for making hybridomas secreting specific antibodies. Nature 276:269-270.

Contributed by Steven A. Fuller, Miyoko Takahashi, and John G.R. Hurrell Allelix Inc. Mississauga, Ontario

Proper care yields a healthy log phase

Preparation of Myeloma Cells

11.5.3 Supplement 1


Preparation of Mouse Feeder Cells for Fusion and Cloning Chilled sucrose solution is injected intraperitoneally into mice. When withdrawn, the solution contains feeder cells (macrophages and other cells) that are placed in the wells of microtiter plates 1 day prior to seeding of hybridomas from cell fusion (UNIT 11.7) or cloning (UNIT 11.8) procedures.

UNIT 11.6

BASIC PROTOCOL

Materials 0.34 M sucrose solution, sterile and chilled Mice (any strain) 70% ethanol HAT medium, chilled Sterile phosphate-buffered saline (PBS; APPENDIX 2) 10-ml syringe, sterile 18-G needle, sterile 50-ml conical centrifuge tube, sterile Dissecting board Forceps, sterile Scissors, sterile 96-well microtiter plates 8% CO2-in-air gas mixture Humidified CO2 incubator Additional reagents and equipment for estimating cell viability by trypan blue exclusion (support protocol, UNIT 11.5) 1. Just prior to sacrificing a mouse, fill a 10-ml syringe with 8 ml chilled sucrose solution and attach 18-G needle. To avoid macrophages adhering to plastic surfaces, it is important to use chilled solutions to optimize cell harvest.

2. Chill the 50-ml conical centrifuge tube in ice. 3. Kill mouse by cervical dislocation. This is accomplished by firmly holding a thick pencil or similar rod-shaped object to the neck of the mouse just behind the skull and quickly and firmly pulling the tail.

4. Immerse the mouse in a 100-ml beaker containing 70% ethanol. 5. Lay out mouse on dissecting board. 6. Snip skin at diaphragm level and pull skin back, exposing the lower part of the rib cage and abdomen. With forceps pull skin from underlying tissue at the diaphragm level and snip with a scissors. With forceps or sterile gloved hands, pull skin back at both sides of the incision to expose the lower part of the rib cage and abdomen. Care must be taken not to tear or cut the peritoneal membrane.

7. Insert the needle into the peritoneal cavity at the base of the sternum and rest the tip of the needle over the liver. Inject sucrose solution. Gently squeeze the abdomen two or three times. 8. Harvest the peritoneal feeder cells by withdrawing as much solution as possible into the syringe. Care must be taken not to puncture the digestive organs, which may lead to fecal contamination of the feeder cells. Contributed by Steven A. Fuller, Miyoko Takahashi, and John G.R. Hurrell Current Protocols in Molecular Biology (1988) 11.6.1-11.6.3 Copyright © 2000 by John Wiley & Sons, Inc.

Immunology

11.6.1 Supplement 1

Enough peritoneal feeder cells can usually be isolated from one mouse to seed ∼100 to 300 wells. However, some mice do not yield effective feeder cells. Depending on the total number of wells that must be seeded with mouse feeder cells, an appropriate number of mice must be killed. Peritoneal exudate feeder cells can be prepared up to 3 days prior to use.

9. Transfer the feeder cell–containing sucrose solution into the 50-ml centrifuge tube. 10. In a sterile fume hood, add 20 ml chilled HAT medium. 11. Centrifuge at 100 × g for 5 min at room temperature. 12. Resuspend the pellet in 1 ml chilled HAT medium and perform cell viability test by trypan blue exclusion as described in the support protocol, UNIT 11.5. 13. Suspend the cell pellet in chilled HAT medium at 1 × 105 cells/ml. 14. Add 100 µl cell suspension to each of the 60 inner wells of the 96-well plates. The peripheral 36 wells are filled with sterile PBS. Plates having 24 wells may be used. If this is the case, add 1 ml cell suspension/well.

15. Incubate plates overnight at 37°C in a CO2 incubator in 8% CO2-in-air with 98% relative humidity. REAGENTS AND SOLUTIONS The following solutions are sterilized by filtration through a 0.22-µm membrane. A suitable sterilization system is a disposable filter unit (e.g., Nalgene #120-0020). Glassdistilled water should be used for all preparations. 0.34 M sucrose solution 58.2 g sucrose H2O to 500 ml Filter sterilize and store at 4°C in 100-ml aliquots HAT (hypoxanthine/aminopterin/thymidine) medium Complete culture medium (see reagents and solutions, UNIT 11.5) supplemented to the indicated concentrations with the following additives: 20% (v/v) fetal calf serum 0.1 mM nonessential amino acids 100 µM hypoxanthine 0.4 µM aminopterin 16 µM thymidine These additives may be purchased in concentrated and sterile solutions from the major suppliers of cell culture media and reagents. Concentrated solutions of hypoxanthine and thymidine (HT) and aminopterin may also be prepared in the laboratory (see following recipes). 100× HT solution Weigh 340.3 mg hypoxanthine and 96.9 mg thymidine; add water to 250 ml. Heat to 70°C to dissolve. Filter sterilize and store in 20-ml aliquots at −20°C. Thaw at 70°C for 10 to 15 min.

Preparation of Mouse Feeder Cells for Fusion and Cloning

1000× aminopterin solution Weigh 17.6 mg aminopterin. Add 60 ml water and dissolve by adding 0.1 M NaOH dropwise. Titrate with HCl to pH ∼8.5. Adjust volume to 100 ml and filter sterilize. Make 100× working solution by diluting stock in complete culture medium. Store in 5-ml aliquots at −20°C. Aminopterin precipitates at low pH and is light sensitive.

11.6.2 Supplement 1


COMMENTARY Background Information To maximize the yield of hybrids from the fusion and cloning procedures, feeder cells are required to be cocultured with the hybrids, while hybrid cell density is low. Mouse peritoneal cells, most of which are macrophages, have been found to be convenient and effective feeder cells which are a source of soluble growth factors for hybridoma cells.

Anticipated Results

From 1 to 3 × 106 peritoneal feeder cells are harvested from one mouse. The number of feeder cells will be enough to seed 100 to 300 wells.

Time Considerations Peritoneal feeder cells from one mouse can be processed in 1 hr or less.

Critical Parameters Feeder cells such as peritoneal cells provide best support of hybridoma growth when used 1 to 3 days after harvest. Use of chilled solutions is necessary for optimum cell harvest, to prevent macrophages adhering to plastic surfaces.


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Supplement 1

UNIT 11.7

BASIC PROTOCOL

Fusion of Myeloma Cells with Immune Spleen Cells Freshly harvested spleen cells and myeloma cells are copelleted by centrifugation and fused by addition of polyethylene glycol solution to the pellet. Cells are centrifuged again and the PEG solution diluted by slow addition of medium. Fused cells are centrifuged, resuspended in selection medium, and aliquoted into 96-well microtiter plates. Hybridomas are grown to 10 to 50% confluence and then assayed for production of antigen-specific antibody. Materials Any strain immunized mouse (UNIT 11.3) Sp2/0 murine myeloma cells in active log phase (Am. Type Culture Collection #CRL 1581; UNIT 11.5) Diethyl ether 70% ethanol Dulbecco modified Eagle medium (DMEM) with supplements Sterile polyethylene glycol (PEG) solution HAT medium (UNIT 11.6) HT medium Phosphate-buffered saline (PBS; APPENDIX 2) 15- and 50-ml centrifuge tubes Glass desiccator or metal can with lid Dissecting board 10.5-cm scissors (Irex #IR-105), sterile 10.5-cm forceps (Irex #IR-1393), sterile 60- and 100-mm petri dishes Stainless-steel strainer (Cellector; GIBCO #1985-8500), sterile 3-cc glass syringes with 26-G needle 5-ml serological pipets 37° C water bath Stopwatch 8% CO2-in-air gas mixture Humidified CO2 incubator Polyvinyl or polystyrene 96-well microtiter plates Inverted microscope Additional reagents and equipment for estimating cell viability by trypan blue exclusion (UNIT 11.5) and for detection of antibodies (UNIT 11.4) Preparation of myeloma and spleen cells 1. Just prior to sacrificing the mouse, transfer 1 × 107 Sp2/0 murine myeloma cells (prepared as described in UNIT 11.5) to a 50-ml centrifuge tube. Check the percentage of viable cells using the trypan blue exclusion method (support protocol, UNIT 11.5). 2. Sacrifice the mouse by anesthetizing with diethyl ether in a closed container (e.g., a glass desiccator or a metal can with a lid). At this point, a blood sample may be collected from the mouse by severing the blood vessels of one forelimb. Collect the blood with a Pasteur pipet and place the blood in a microcentrifuge tube.

Fusion of Myeloma Cells with Immune Spleen Cells

11.7.1 Supplement 1

3. Immerse the mouse in a beaker containing 70% ethanol and lay out on a dissecting board. Contributed by Steven A. Fuller, Miyoko Takahashi, and John G.R. Hurrell Current Protocols in Molecular Biology (1988) 11.7.1-11.7.4 Copyright © 2000 by John Wiley & Sons, Inc.

4. Using sterile forceps, lift skin over the thorax area and snip with sterile scissors. Peel skin over both sides to expose left side of the rib cage. 5. Using another set of sterile forceps and scissors, remove the spleen from the left upper abdomen of the mouse. The spleen is a small dark red organ. 6. Place the spleen in a 60-mm petri dish containing 3 ml supplemented DMEM. 7. Take spleen in petri dish to sterile hood and carefully dissect away surface fat and other adhering tissue by using a sterile forceps and scissors. 8. Transfer spleen to sterilized, stainless-steel strainer in 100-mm petri dish with 10 ml DMEM. 9. Fill the 3-cc syringe with 2 ml supplemented DMEM. Using a 26-G needle, fill the spleen with DMEM by injecting at several sites. 10. With sterile scissors, cut the supplemented DMEM–filled spleen in 3 or 4 places. 11. Using circular movements, press the spleen against the screen of the stainless-steel strainer with the glass syringe plunger of the 3-cc syringe until only fibrous tissue remains on top of the strainer screen. The tissue that is forced through the strainer is collected in a sterile petri dish underneath. 12. Rinse the screen with 2 ml supplemented DMEM. 13. Transfer the suspension of spleen cells to a 15-ml centrifuge tube. Using a 5-ml serological pipet, disperse the clumps by drawing up and expelling several times. 14. Let suspension stand for 3 min at room temperature. 15. Transfer the top 95% of the cell suspension to a 15-ml centrifuge tube. 16. Perform a viable cell count using the trypan blue exclusion procedure (support protocol, UNIT 11.5); record the percentage of viable cells. Ignore red blood cells that are substantially smaller than the nucleated cells.

17. Transfer 1 × 108 viable spleen cells into a 15-ml centrifuge tube. 18. Wash myeloma cells (from step 1) twice with supplemented DMEM followed by centrifugation at 200 × g for 5 min and resuspend the cells in 5 ml supplemented DMEM. 19. Add spleen cells to myeloma cells in the 50-ml tube and fill the tube with DMEM. 20. Centrifuge the suspension at 200 × g for 5 min at room temperature. 21. Resuspend the cell pellet in 50 ml supplemented DMEM and centrifuge as above. 22. Warm the cell pellet by placing the tube in a 37°C water bath in a beaker for 2 min. 23. Loosen the pellet by flicking the tip of the tube gently. Cell fusion 24. Fuse spleen and Sp2/0 cells with sterile PEG solution: Over the first 1 min—Add 1 ml PEG solution at 37°C. Mix gently. Over the next 2 min—Spin at 100 × g (2 min, total time). Over the next 3 min—Add 4.5 ml supplemented DMEM. Over the next 2 min—Add 5 ml supplemented DMEM. Fill the tube with supplemented DMEM.

Immunology


Supplement 1

Timing is critical and should be monitored with a stopwatch.

25. Centrifuge at 100 × g for 5 min at room temperature. 26. Aspirate supernatant. 27. Resuspend the cell pellet in 35 ml HAT medium. Do not force the dispersion of small cell clumps.

28. Incubate cell suspension at 37°C in a CO2 incubator in 8% CO2-in-air with 98% relative humidity for a minimum of 30 min. Plating and culture of fused cells 29. Add 100 µl cell suspension to each of the 60 inner wells of six 96-well plates. Peripheral 36 wells are filled with sterile PBS. Twenty-four (24) hr prior to use, the 60 inner wells are conditioned with 1 × 104 mouse peritoneal macrophages per well in 100 ìl HAT medium.

30. Incubate plates at 37°C in CO2 incubator in 8% CO2-in-air with 98% relative humidity. (This is day 1 of the culture). 31. On day 5 of the culture, add 100 µl HAT medium to each well. 32. On day 7 of the culture, remove 100 µl from each well and add 100 µl fresh HAT medium. 33. Repeat step 32 every other day until hybrid cell growth covers 10% to 50% of the surface area of the wells. This is monitored by examining the bottom of the wells with an inverted microscope. At this time, the wells should be screened for antibody (UNIT 11.4). 34. Grow the hybrids in HAT medium for 2 weeks after fusion. 35. After 2 weeks, change the medium to HT medium. 36. The hybrids are grown in the HT medium until the completion of two cloning procedures (as described in the protocol for cloning of hybridoma cells by limiting dilution, UNIT 11.8). REAGENTS AND SOLUTIONS Dulbecco modified Eagle medium (DMEM), high-glucose formula (GIBCO/BRL #430- 2100), supplemented to the indicated concentrations with the following additives: 2.8 g/liter sodium bicarbonate (33.3 mM) 4.8 g/liter HEPES (20 mM) HT medium Prepare as described in UNIT 11.6 for HAT medium but without aminopterin solution. PEG (polyethylene glycol) solution Weigh 10 g PEG 4000 (Merck) into a 100-ml glass bottle. Autoclave PEG for 10 min at 121°C. Cool the molten PEG to 50°C. Mix with 10 ml prewarmed supplemented DMEM (50°C) that contains 5% (v/v) dimethylsulfoxide (DMSO). Aliquot the mixture into small glass bottles (3 ml/bottle). Store at 4°C in the dark. Fusion of Myeloma Cells with Immune Spleen Cells

PEG 4000, Merck’s gas chromatography grade, appears to be the best PEG for cell fusions using Sp2/0 myeloma cells regardless of lot number. Remove PEG from the autoclave as soon as the pressure is down in order to avoid prolonged heating of PEG. Incubate PEG solution at 37°C for 24 hr prior to use to test its sterility.

11.7.3 Supplement 1


Alternatively, if the temperature control of the autoclave is uncertain, melt the PEG at 65°C on the day of fusion. Mix the PEG with supplemented DMEM to make a 45% PEG solution and sterilize by filtration through a 0.22-ìm membrane filter.


Troubleshooting

Murine spleen cells, some of which are involved in production of the desired antibodies, are fused with a murine myeloma cell line to form a stable antibody-producing hybridoma cell line. The myeloma cells (Sp2/0) are hypoxanthine–guanine phosphoribosyltransferase deficient (HGPRT−) and therefore are unable to use the purine salvage pathway when de novo purine synthesis is blocked by aminopterin, which is included in the HAT selection medium. See discussion of selectable markers in UNIT 9.5. Hypoxanthine and thymidine in the HAT selection medium allow HGPRT+ cells, the spleen cell–myeloma hybrids, to survive and grow. Unfused spleen cells eventually die.

When poor fusion results (i.e., poor hybridoma growth) are obtained despite close adherence to the protocol, the likely causes are the PEG, the myeloma cells used as fusion partner, or the culture conditions. Most critical is the PEG (see critical parameters). The quality of the fusion partner (i.e., the myeloma culture; see UNIT 11.5 for details) is also very important. The CO2 incubator must provide a stable temperature, pH, and humidity for optimal hybridoma growth.

Literature Review Kohler and Milstein (1975) first demonstrated that somatic cell fusion could be used to generate a hybridoma cell line producing a monoclonal antibody of predetermined specificity. Cell fusion was initially accomplished by addition of Sendai virus. The fusion procedure used in this protocol is a modification of the method of Gefter et al. (1977), which uses polyethylene glycol (PEG) as the fusogen. A similar procedure is presented by Oi and Herzenberg (1980). Each protocol encompasses careful timing of the PEG addition to the cell pellet and its subsequent dilution after fusion.

Critical Parameters The choice and use of PEG in the fusion protocol is the most critical factor. PEG can vary dramatically in efficiency between manufacturers and among lots of a particular manufacturer. The suggested source (Merck) has provided the most consistent lots of PEG. Care must be taken to autoclave the PEG at 121°C for only 10 min in order to minimize the production of toxic aldehydes. In the fusion procedure itself, it is important to adhere carefully to the time schedule established. Extended incubation of the cells with PEG results in decreased cell viability. Dilution of the PEG by medium must be done carefully to avoid lysis of the cells.

Anticipated Results Hybridoma growth should be observed with the aid of an inverted microscope in nearly all wells after a few days in culture. For wells with viable cells, generally 10 to 50% will contain antigen-specific antibody. Results can vary widely, of course, and it has been observed that 0 to 99% of the supernatants in the wells will contain specific antibody.

Time Considerations The fusion procedure requires 3 to 4 hr to complete. For best results, it should be accomplished without interruption. Wells with hybridoma growth can be assayed for specific antibody 7 to 12 days after fusion.

Literature Cited Gefter, M.L., Margulies, D.H., and Scharff, M.D. 1977. A simple method for polyethylene glycolpromoted hybridization of mouse myeloma cells. Somat. Cell Genet. 3:231-236. Kohler, G. and Milstein, C. 1975. Continuous cultures of fused cells secreting antibody of predefined specificity. Nature 256:495-497. Oi, V.T. and Herzenberg, L.A. 1980. Immunoglobulin-producing hybrid cell lines. In Selected Methods in Cellular Immunology (B.B. Mishell and S.M. Shiigi, eds.) pp. 351-372. W.H. Freeman, San Francisco.


Immunology


Supplement 1

UNIT 11.8

BASIC PROTOCOL

Cloning of Hybridoma Cell Lines by Limiting Dilution Hybridomas to be cloned are diluted to 0.8 cells/well. This dilution provides 36% of wells with 1 cell/well by Poisson statistics. When cultures are 10 to 50% confluent, antibody is assayed by ELISA. Two or more cloning procedures are carried out until >90% of the wells containing single clones are positive for antibody production. Materials HAT medium (UNIT 11.6) HT medium (UNIT 11.7) Complete culture medium (UNIT 11.5) Polyvinyl or polystyrene 96-well microtiter plates 6- and 24-well culture plates 8% CO2-in-air gas mixture Humidified CO2 incubator Cryotubes (Nunc #3-63401) 2-, 5-, and 10-ml serological pipets Multichannel pipet and tips Inverted microscope Additional reagents and equipment for preparing mouse feeder cells for fusion and cloning (UNIT 11.6), for ELISA screening (UNIT 11.4), and for estimating cell viability by trypan blue exclusion (UNIT 11.5) 1. The day before cloning, isolate mouse feeder cells and prepare 96-well plates with feeder cells in appropriate medium, as described in UNIT 11.6. Choice of medium depends on the current stage of the cloning process. The first cloning uses HAT medium, and the second cloning uses HT medium. Any other clonings use complete culture medium.

2. Transfer all of the cells from each well containing antigen-specific antibody in its hybridoma supernatant (as determined by an ELISA, UNIT 11.4) into a separate well of a 24-well plate that has been preincubated with 0.5 ml of an appropriate culture medium (see note in step 1) and culture overnight at 37°C, 8% CO2-in-air, and 98% humidity in a CO2 incubator. If there are greater than 40 to 50 positive wells, it is difficult to manage conveniently the cloning procedure. The ELISA assays (UNIT 11.4) or western blotting (UNIT 10.8) can be performed using the 24-well culture supernatants in order to select the most promising samples to be cloned. The remaining positive samples can be transferred to cryotubes and frozen, as described in UNIT 11.9. Cloning efficiency is always improved when hybrid cells are grown to log phase in 24-well plates. The efficiency is still better if the 24 wells are preconditioned with feeder cells prior to transferring the hybrids.

3. Perform cell viability count using trypan blue exclusion method (support protocol, UNIT 11.5) on the overnight cultures in 24-well plates. 4. Using a 6-well plate, make dilutions of cells from overnight cultures in HT or complete medium. In the first well, make a 1:100 dilution in a total of 3 ml; in the second well, dilute an aliquot of the first dilution to 80 cells/ml in 5 ml; in the third well, prepare 8 cells/ml in 10 ml (i.e., a 1:10 dilution from the second well). Cloning of Hybridoma Cell Lines by Limiting Dilution

11.8.1 Supplement 1

Choice of medium is discussed in step 1. Using 6-well plates is far easier for preparing cell dilutions than using tubes. However, using 6-well plates is expensive.

5. With a multichannel pipet, fill the upper 50 wells of the inner 60 wells of the 96-well plate from step 1 with 100 µl of 8 cells/ml dilution (i.e., 0.8 cells/well) Contributed by Steven A. Fuller, Miyoko Takahashi, and John G.R. Hurrell Current Protocols in Molecular Biology (1988) 11.8.1-11.8.2 Copyright © 2000 by John Wiley & Sons, Inc.

and the 10 wells of the bottom row with 100 µl of 80 cells/ml dilution (i.e., 8 cells/well). 6. Incubate at 37°C in a CO2 incubator in 8% CO2-in-air with 98% relative humidity (day 1). As a precaution, the hybridoma cells remaining in the 24-well plate can be transferred to cryotubes and frozen, as described in UNIT 11.9.

7. On day 6, feed the culture with the addition of 100 µl/well of fresh medium, using a multichannel pipet. Thereafter, if necessary, refeed the culture every other day by removing 100 µl media from each well and adding 100 µl fresh media. 8. When cell growth in the bottom of the wells is 10 to 50% confluent (as monitored using an inverted microscope), assay for specific antibody in the hybridoma supernatants using an ELISA (UNIT 11.4). 9. Transfer 2 to 3 selected positive subclones from each plate into a 24-well plate (as for step 2) and incubate overnight. Expand the subclones and freeze one aliquot for each subclone in a cryotube. This is done as a precaution in case one fails to recover positive clones.

10. Repeat cloning procedure from the beginning until a stable and single hybridoma cell line is established. Hybridomas that yield >90% antibody-positive cultures upon recloning are considered to be stable. Those that yield 40 ml) of ascites can be collected from each mouse.

9. Centrifuge the ascites 10 min in TH-4 rotor at 2700 rpm (1500 × g), room temperature. Harvest supernatants and discard pellet. Store ascites fluid at 4°C until all collection is completed (2) in vivo serial passes of the hybridomas.

13. Dilute >1:10 and filter sterilize through a 0.45-µm filter. Aliquot and freeze at −70°C, avoiding repeated freezing and thawing (Yokoyama, 1991a). Shelf life should be several years. Add sodium azide to 0.02% final if the ascites will not be used for bioassay. It is possible that ascites fluid will not form. If the mice die without any ascites forming, particularly within 2 weeks of inoculation, try fewer cells. If the mice do not form ascites after 2 weeks and they appear healthy, inject those mice—as well as naive, Pristane-primed mice—with more cells. If solid tumors form, tease cells into suspension and inject tumor cells into another Pristane-primed mouse. Even if a little ascites forms, the fluid can be transferred to another mouse (∼0.5 ml/mouse), and large amounts of ascites should accumulate. Once the ascites is formed, the mouse-adapted cells can be frozen and used to reinoculate mice in the future.

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COMMENTARY Background Information Production of MAb supernatant There are three basic preparations that contain monoclonal antibodies: supernatant from a MAb-producing hybridoma, ascites from a mouse inoculated with the hybridoma, or purified MAb. In the first protocol, a MAb-containing supernatant is produced. The hybridoma continues to secrete MAb into the culture fluid until cell death occurs. Because the MAb is not metabolized, it accumulates in the culture supernatant. Hybridoma supernatants are advantageous because small amounts (0.5% may also have significant nonspecific effects. In addition to the appropriate

isotype control MAb ascites, ascites harvested from mice inoculated with the nonsecretory partner cell line (i.e., SP2/0) also provides a useful nonspecific control. The general procedure involves the elicitation of nonspecific inflammation in the peritoneal cavity of an appropriate host mouse, usually with Pristane, and injection of the hybridoma cells. The tumor cells then grow as an ascites tumor and should continue to secrete the MAb. Eventually, the mouse develops a monoclonal gammopathy similar to the human disease, multiple myeloma, in that a monoclonal protein reaches high titer in the serum. Since the combination of the ascites tumor and Pristane results in an inflammatory exudate in the peritoneal cavity, the ascites should contain concentrations of the MAb similar to that in serum.

Critical Parameters Monoclonal antibody production by hybridomas is an unstable phenotype. Hybridoma cells always should be grown under log-phase growth conditions. Prolonged in vitro culture and in vivo passage should be avoided. Thus, the most critical parameter is whether the hybridoma of interest is secreting a high titer of the MAb. This should be checked before a major effort is made to grow large amounts of supernatant or to produce ascites fluid. The MAb titer can be determined by serial dilution of the culture supernatant in the assay appropriate for that MAb, such as ELISA (UNIT 11.2) or flow cytometry (Holmes and Fowlkes, 1991; Yokoyama, 1991b). Titers of ≥1:10 should be saturating if spent culture supernatants are examined. If necessary, the hybridoma can be recloned by limiting dilution (UNIT 11.8) to find high-producing clones. If cells are known to produce MAb at high titers, aliquots frozen immediately after cloning (Yokoyama, 1991a) will retain this phenotype. The most critical parameter in the largescale production of cell lines and hybridomas is the adaptation of the cells to roller flasks. Most hybridomas and other nonadherent cells that grow in suspension can be easily adapted. If the cells (particularly adherent cells) cannot be adapted, other methods should be tried. For example, large-scale production of cells for use in isolating cellular components can be performed using multiple 175-cm2 flasks instead of roller flasks. Unfortunately, the




relative surface area is small, and therefore the number of flasks required can become prohibitive. Adherent cell lines are less easily adaptable to growth in roller flasks. The surface area for growing adherent cells can be increased by the use of dextran beads (e.g., Cytodex beads from Pharmacia). These beads can increase the surface area of a culture flask severalfold. If there is any suspicion that the cells may be mycoplasma contaminated, diagnosis and treatment are indicated (Fitch et al., 1991). Mycoplasma-contaminated lines will produce a much poorer yield of final cell numbers because they do not grow to as high a cell density as normal cells. Supernatants frequently contain 1 to 10 µg/ml of MAb, but the concentration is cell-line dependent. The supernatants could be concentrated by salt precipitation, but this is not generally recommended because large volumes of culture supernatants have to be concentrated to derive the amount of MAb in small amounts of ascites. Moreover, the FCS in the supernatant will also be concentrated. While the hybridoma could be adapted to culture in serum-free medium, this requires additional testing and yields may decrease. Affinity purification of the culture supernatants would take a similar amount of effort and produce purified MAb at high concentrations. Thus, instead of concentrating supernatants (if high concentrations of MAb are desired), it is recommended to produce purified MAb by first growing hybridomas at a larger scale (liters) or to produce ascites. For ascites production, it is important to consider the appropriate host for the hybridoma since an injection of allogeneic or xenogeneic cells may result in rejection. For most mousemouse hybridomas, an F1 hybrid—between the BALB/c strain (origin of the commonly used SP2/0 fusion partner) and the strain from which the normal cells were obtained—could be used. For xenogeneic hybridomas, nude mice or lowdose irradiated normal mice are potential hosts. Outbred nude mice are somewhat more expensive than normal mice but do not require irradiation. It is not necessary to use the prohibitively expensive inbred nude mouse strains. Because the level of normal immunoglobulin in mouse serum is in the same range as ascites fluid (mg/ml), ascites fluid can be only partially purified by salt fractionation or antimouse-Ig- or protein A–affinity chromatography (UNIT 2.7). However, it is a convenient source of raw material from which to affinity purify rat MAb with a mouse anti-rat κ MAb (e.g., MAR 18.5) column (Andrew and Titus, 1991).

Troubleshooting It is possible that ascites fluid will not form. The reasons for this are unclear but are probably related to a property of the individual hybridoma. If the mice die without any ascites forming, particularly within 2 weeks of inoculation, try fewer cells. If the mice do not form detectable ascites after 2 weeks and they appear healthy, inject those mice—as well as naive, Pristane-primed mice—with more cells. If solid tumors form, tease cells into suspension and inject the tumor cells into another Pristaneprimed mouse. Even if a little ascites forms, the fluid can be transferred to another mouse (∼0.5 ml/mouse), and large amounts of ascites should accumulate. Once the ascites is formed, the mouse-adapted cells can be frozen and used to reinoculate mice in the future.

Anticipated Results Most culture supernatants will have saturating MAb titers of ≥1:10 when tested at 100 µl for 106 cells. If the spent culture supernatant is used for MAb purification by affinity chromatography (UNITS 10.9 & 10.10; Andrew and Titus, 1991), 1 to 10 mg of purified MAb/liter can be anticipated. If a much lower titer or yield is achieved, recloning of the hybridoma line may be indicated. Hamster-mouse hybridomas are particularly notorious for instability. Most hybridomas can be grown as ascites tumors. The saturating concentration of the MAb in such fluids should be detected at dilutions of 1:500. If MAb titers are significantly lower, the hybridoma may be a poor producer. If ascites do not form, see troubleshooting above. Most tumor cells that grow in suspension should be amenable to growth in roller flasks and densities of >106 cells/ml should be attained. Careful work should result in no contamination.

Time Considerations For high-titer and large-scale production of MAb supernatants, a flask split 1:10 will be overgrown, with cell viability definitely decreasing by day 5 to 6, at which time the supernatants can be harvested. If several liters of supernatant are required, ∼10 days are required to expand a 25-cm2 flask (10 ml) to 2.4 liters. For production of ascites fluid containing MAb, 4 to 6 weeks are necessary for growth of the cells for inoculation, ascites accumulation, tapping the fluid, centrifugation, and determination of the MAb titer.

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Once the 175-cm2 flasks are seeded for large-scale production of hybridomas and cell lines, 10%, resulting in 108 clones) are still acceptable. Usually, the original phage library will consist of in the order of 106 independent recombinants. A 100-fold excess of clones in the excised plasmid pool will ensure that the complexity of the original library is maintained.

7. Analyze the plasmid content of a number (20 to 40) of random colonies using a miniprep procedure (UNIT 1.6), digestions with restriction enzymes (UNIT 3.1), and gel electrophoresis (UNIT 2.5A). Determine the percentage of clones with insertions and the cDNA size distribution. Process phage-infected E. coli and plate library 8. If the excision frequency was sufficient, divide the previously stored phage-infected E. coli BNN132 culture (step 6) over eight 1.5-ml microcentrifuge tubes and microcentrifuge for 1 min to pellet the bacteria. 9. Decant the supernatant, resuspend each bacterial pellet in 1 ml liquid LB medium, and spread the content of each tube on a 24 × 24–cm LB-Cb plate. Grow overnight at 37°C. Yeast One-Hybrid Screening for DNA-Protein Interactions

A dense field of small colonies will result. The initial growth of the plasmid library on plates avoids underrepresentation of toxic cDNA clones. If the culture is directly grown in liquid medium, bacteria containing such clones could be competed out.



10. Scrape the colonies from the plates using sterile cell scrapers. Resuspend in 1 liter prewarmed LB medium (with 200 mg/liter carbenicillin) and grow for another 4 hr, shaking at 37°C. 11. Harvest the bacteria by centrifuging in a centrifuge with large buckets (e.g., 250- or 500-ml buckets) and isolate the DNA using maxiprep procedures (UNIT 1.7) or commercially available maxiprep kits. Determine the DNA concentration by UV spectrometry (APPENDIX 3D). 12. Proceed with the library transformations, following the instructions for library screening (see Basic Protocol 2). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

3-Amino-1,2,4,-triazole (3-AT), 1 M Prepare a 1 M stock solution of 3-AT (e.g., Sigma). Do not autoclave, but filter sterilize the solution. Store at 4°C. Add the 3-AT to the CM medium after the latter has cooled down to ∼60°C. Store up to 1 year at 4°C. LB-Cb plates Solidify LB medium (UNIT 1.1) with 1.5% agar and supplement with 200 mg/liter carbenicillin (add from 1000-fold stock). Sterilize LB medium with 1.5% agar by autoclaving 20 min. Add carbenicillin after LB medium has cooled to 60°C. Pour into 24 × 24–cm square dishes (Nunc) and standard (9-cm) petri dishes. Store up to 3 months at 4°C. Lithium acetate stock, 10× Prepare 1 M lithium acetate, pH 7.5, adjust with acetic acid; sterilize by autoclaving 20 min. Store indefinitely at room temperature. PEG stock, 50% (w/v) Prepare 50% (w/v) polyethylene glycol (PEG 3,350; e.g., Sigma); sterilize by autoclaving 20 min Store indefinitely at room temperature, in small portions in well-closed bottles, because transformation efficiencies are critically dependent on the correct PEG concentration.

TE buffer stock, 10× 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 10 mM EDTA Sterilize by autoclaving 20 min Store indefinitely at room temperature COMMENTARY Background Information To gain insight into the regulation of a gene, it is necessary to identify the transcription factors binding to regulatory sequence elements. Such cis-acting elements mostly have been identified in the 5′ promoter regions of genes, but can also be located in introns, exons, or 3′ untranslated regions. Binding of factors can result in activation or silencing of gene repression, depending on the developmental stage of

the cell or on internal or external signals. Using techniques like gel-shift assays, DNA methylation interference, or DNase footprinting, binding sites for transcription factors can be mapped. Often, these are short sequences of only 5 to 10 bp. Almost all promoters contain a TATA box close to the transcription start site. The TATA box is recognized by a complex of factors, the basal transcription machinery. In turn, this complex interacts with the transcrip-

DNA-Protein Interactions


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Yeast One-Hybrid Screening for DNA-Protein Interactions

tion factors that specifically bind to the cis-acting elements. Traditionally, the cloning of transcription factors relied on biochemical strategies such as affinity chromatography. Furthermore, molecular biological methods have been developed that involve the screening of cDNA expression libraries using labeled oligonucleotides. In the past decade, newly developed yeast genetic selection methods have provided particularly useful alternatives for cloning of transcription factors. One of the possibilities is to complement yeast transcription factor mutants. An illustrative example is the cDNA cloning of a human CCAAT-binding protein via complementation of the yeast hap2 mutant (Becker et al., 1991). With the availability of the complete yeast genome sequence, it has become relatively easy to construct yeast transcription factor mutants and to assess the feasibility of a complementation strategy to clone a heterologous factor. Obviously, the possibilities are limited to cloning of functionally conserved transcription factor genes. More generally applicable is the use of a yeast genetic selection strategy that has become known as the one-hybrid system. In fact, the one-hybrid system is a simplification of the two-hybrid approach for detection of protein-protein interactions (Fields and Song, 1989; UNIT 20.1). The one-hybrid concept (Fig. 12.12.1), relies on the detection of the interaction between a protein expressed from a cDNA library (prey) with a DNA sequence of interest (bait) fused to a reporter gene. Activation of this reporter gene is dependent on the interaction and can be selected for. The required cDNA libraries are designed such that the expressed proteins carry a strong transcriptional activation domain from a known factor (hence the term “one-hybrid”). Therefore, an interaction with the bait sequence will usually guarantee reporter gene activation. This makes the selection method suitable to clone all types of DNA-binding proteins, independent of their normal functional properties. It is also possible to apply a “non-hybrid” selection strategy (Grueneberg et al., 1992; Chan et al., 1993), but this limits cloning possibilities to transcription factors with an intrinsic activation domain that functions properly in yeast. One-hybrid libraries are normally constructed with shuttle vectors containing the replication sequences and marker genes necessary for maintenance in both yeast and E. coli. The activation domain (AD) in the expressed hybrid proteins is usually from the

yeast Gal4p transcription factor, but there are also vectors where the Herpes simplex VP16 AD or the acidic B42 domain (UNIT 20.1; Clontech) is used. To facilitate nuclear import of the hybrid proteins, the ADs used in all vectors are equipped with an SV40 nuclear localization signal. As a frequent additional feature, an epitope domain, such as HA (hemagglutinin), is included to facilitate detection on western blots or other applications. Constitutive promoters, such as the ADH1 promoter, are often used to drive cDNA expression, but alternatives are conditional expression systems, e.g., based on the galactose-inducible GAL1 promoter. All AD vectors designed for two-hybrid screenings are exchangeable with one-hybrid systems. Some AD vectors are available as lambda vector systems (Clontech, Stratagene), with which cDNA cloning is more efficient and it is easier to achieve and maintain high complexity. Described lambda phage vectors for AD-library construction can easily be converted into E. coli–yeast shuttle vectors by automatic subcloning. A selection of useful vectors is shown in Table 12.12.3. The authors and others have had good experiences with use of the λACTII vector (Memelink, 1997) for cloning of a wide variety of transcription factors from plants (Menke et al., 1999; Meijer et al., 2000; van der Fits et al., 2000) and human cell lines (P.B.F. Ouwerkerk, unpub. observ.). The most frequently used reporter for genetic selection procedures is the HIS3 gene, operative in the biosynthesis of histidine and encoding the enzyme imidazoleglycerol-phosphate dehydratase. Detection of its activation is through complementation of the auxotrophic his3 mutation. Usually, HIS3 reporters contain the minimal HIS3 promoter, including the TATA box and transcription start site (Grueneberg et al., 1992). There are several examples of successful application of the HIS3 reporter in one-hybrid screenings (e.g., Wilson et al., 1991; Grueneberg et al., 1992; Wang and Reed, 1993). Alternative strategies make use of the lacZ (Li and Herskowitz, 1993) or green fluorescence (GFP) reporter genes (Display Systems Biotech; UNIT 9.7C). Furthermore, the use of MEL1 as an alternative color marker was recently described (Melcher et al., 2000). Although detection of these reporters is easy, the use of HIS3 as a reporter has important advantages in a genetic selection procedure. Because HIS3 is a growth selection marker, detection of positives is very simple since these appear as clear colonies in a field of nongrowing or slowly growing cells. Screening of lacZ, MEL1,



or GFP activity requires that all the transformed cells grow into colonies. Therefore, the plating density must be far lower than when HIS3 is used as reporter, which complicates the setup of saturating library screens. Another advantage is that the choice of HIS3 offers a solution to possible problems of leaky reporter expression, also termed background activity. Leaky expression can occur when bait sequences are recognized by endogenous yeast transcription factors. Since the activity of the His3p enzyme can be competitively inhibited by 3-amino1,2,4-triazole (3-AT), leaky HIS3 expression can often be eliminated by a 3-AT titration, so that it remains possible to detect increased HIS3 expression resulting from the specific binding of a library protein during the screening. With lacZ, MEL1, or GFP reporters, the distinction between specific and background expression is difficult, and other auxotrophic markers that could be used as reporters (LEU2, ADE2) are not titratable. A good alternative for the HIS3 reporter is a dominant antibiotic marker such as G418, hygromycin, or chloramphenicol. Here, leaky expression can be eliminated by increasing the concentration of the selective compound. An example is the cloning of the transcription factor Nrf1 using a neomycin-resistance gene in combination with G418 selection (Chan et al., 1993). With some bait sequences, the occurrence of excessively high leaky expression, which needs to be eliminated, may be observed, which is the most important constraint on the application of the one-hybrid screening system. Initially, one-hybrid screenings involved the use of two replicating vector systems, one for the AD library and the other for the reporter (e.g., Wilson et al., 1991; Li and Herskowitz, 1993; Wang and Reed, 1993). Afterwards, a switch was made to chromosomally integrated reporters, which has the advantage that the reporter gene copy number is constant and that DNA-protein interactions are studied within the context of the nucleosome (e.g., Clontech). The pHIS3/pINT1 vectors described by the authors (Meijer et al., 1998) provide a system for the integration of reporter genes in the yeast genome. In this system, a bait sequence is first cloned upstream of the HIS3 gene in one of the reporter vectors pHIS3NB or pHIS3NX (Table 12.12.1 and Fig. 12.12.3). In the second step, the reporter construct is cloned in the integrative vector pINT1. Finally, the pINT1-HIS3 reporter construct is integrated via double cross-over in the nonessential PDC6 gene (YGR087c, Chr. VII) using selection for the

dominant APT1 antibiotic marker. Advantages of the pHIS3/pINT1 system are discussed in the introduction to this unit, and examples of its application have been published (Menke et al., 1999; Meijer et al., 2000; van der Fits et al., 2000). A further refinement of one-hybrid screenings lies in the use of dual or triple reporter strains to facilitate discrimination between true and false positives. The Y187 strain that the authors recommend be used in conjunction with the pHIS3/pINT1 reporter system provides the opportunity to use a dual reporter system. It contains an additional GAL1UASGAL1TATA-lacZ construct, thus, a reporter gene with an upstream sequence that is unrelated to the bait sequence. Based on the unwanted activation of this second reporter, it is possible to discard a subset of false positives, probably representing nonspecific DNA-binding proteins. The lacZ reporter in Y187 was integrated with the URA3 marker and can be counterselected with 0.1% (v/v) 5-fluoro-orotic acid. If desired, it can therefore be replaced to make alternative dual-reporter strains, or another construct can be added to make a triple reporter. One possibility is addition of a construct containing the bait sequence upstream of another reporter gene that also has a different minimal promoter than that of the HIS3 reporter (see Table 12.12.4). Such a reporter should be activated by true positives, and a negative interaction would select against false positives that activated the bait-HIS3 reporter through sequences outside the bait itself. Another elegant alternative is to add a reporter with a mutant bait sequence, but this is only applicable if it is already known which nucleotides in the bait sequence are critical for its cis-acting function. A good control is also to retransform isolated library clones to a strain containing a mutant bait-HIS3 repo rter integrated via the pHIS3/pINT1 system, so that it is present in an exactly identical chromosomal context as the bait reporter. Furthermore, control transformations can be performed to strains containing pHIS3/pINT1-based reporters lacking the upstream bait sequence or containing an unrelated upstream sequence. The nature of false positives is diverse (see Anticipated Results) and it should be taken into account that even with various different control reporters, not necessarily all the false positives can be eliminated. Some may follow specificity criteria in yeast exactly as expected for true positives (Schouten et al., 2000). Therefore, it remains essential that



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Table 12.12.4 Procedures

A Selection of Useful Cloning, Expression and Reporter Vectors for Modified Screening Applications or

Vectorsa

Features

Reference

pRS300 seriesb,c

Yeast markers URA3, TRP1, LEU2, HIS3, and LYS2. Replication via ARS-CEN

Sikorski and Hieter, 1989; Sikorski and Boeke, 1991

pRS400 seriesb,c

Yeast markers URA3, TRP1, LEU2, HIS3, MET15, ADE2, and LYS2. Integrating, ARS-CEN, and 2-µm versions

Sikorski and Hieter, 1989; Christianson et al., 1992; Brachmann et al., 1998

pYC seriesd

Yeast markers URA3, MET2-CA, G418. ARS-CEN and 2-µm versions. Counterselectable marker PKA3 regulatable by conditional CHAI or MET25 promoters

Olesen et al., 2000

YcpIF seriesc

Yeast markers URA3, TRP1, LEU2, HIS3. Replication via ARS-CEN. Expression cassettes for translational fusions, equipped with inducible GAL1 promoter

Foreman and Davis, 1994

p4XX seriesc

Yeast markers URA3, TRP1, LEU2, HIS3. ARS-CEN and 2-µm versions. Expression cassettes for transcriptional fusions, equipped with the CYC1 terminator and CYC1, ADH1, GPD, TEF, MET25, or GAL1 promoters

Mumberg et al., 1994, 1995

pMELα/β2, YIpMELα/β2e

MEL1 or lacZ reporter vectors. Integrating versions and ARS-CEN-replicating versions with URA3 marker

Melcher et al., 2000

pSLF187K

lacZ reporter vector, URA3 marker, replication via 2-µm

Forsburg and Guarente, 1988

LacZib

lacZ reporter, URA3 marker. Integrative vector

Clontech

aAll vectors replicate high-copy in E. coli and confer carbenicillin resistance. bComplete sequences are deposited in the NCBI database. cAvailable from the American Type Culture Collection (ATCC). dVectors and sequences available from http://www.crc.dk/phys/pyc/index.htm. eVectors and sequences are available from http://www.rz.uni-frankfurt.de/FB/fb16/mikro/euroscarf/data/km_rep.html.


a yeast result be followed up by an independent confirmation of the specific interaction. Finally, several interesting variations of the one-hybrid concept have been developed. One system, which the authors call “bridge one-hybrid,” can, for example, be applied for cloning coactivators of transcription factors (Sieweke et al., 1996; Yu et al., 1997). Here, a transcription factor, specifically interacting with a target sequence upstream of the reporter gene, is used to attract library proteins that increase the reporter gene expression. This type of “interactor hunt” differs from the conventional two-hybrid approach in that there is a specific selection for proteins that interact when the transcription factor is bound to its own target sequence. Therefore, this would more likely reflect authentic interactions. Another system is a reverse one-hybrid approach that can detect library proteins capable of dissociating the interaction between a known factor and its target sequence (Vidal et al., 1996). Both the one-hybrid and reverse one-hybrid systems can also be modified to mutation screens for identifica-

tion of critical amino acid residues in functional domains of the transcription factor (Bush et al., 1996; Vidal et al., 1996). A last and particularly challenging variation is to express an AD hybrid of a known transcription factor and screen for its target sites. This requires a reporter gene library with genomic DNA fragments or random oligonucleotides upstream of the reporter gene, and a procedure to discriminate between true positive target sequences and false-positive sequences that are activated by endogenous yeast factors (Wilson et al., 1991; Liu et al., 1993; Meijer et al., 1998).

Critical Parameters and Troubleshooting A critical step in setting up yeast one-hybrid screenings is how to deal with the leaky HIS3 expression that can occur due to recognition of the bait sequence by endogenous yeast transcription factors. Whether or not a particular reporter construct will be activated by a yeast transcription factor is difficult to predict in advance, but it is known that several cis-acting



sequences of plant or animal genes can also be recognized by yeast factors. Nevertheless, it may still be possible to screen in yeast for the corresponding plant or animal factors. One-hybrid screenings have even been used to clone factors from yeast itself (Li and Herskowitz, 1993; Kunoh et al., 2000). In recent years, a wide variety of transcription factors from plants (e.g., Kim et al., 1997; Menke et al., 1999; Meijer et al., 2000; van der Fits et al., 2000) or animals (e.g., Wang and Reed, 1993; Wei et al., 1999) have been cloned with a range of promoters and binding sites. Therefore, it does not seem that the one-hybrid approach is limited to cloning of particular transcription factor classes. With the lack of knowledge to predict in advance whether a certain promoter will be activated in yeast, the best and fastest strategy is still trial and error. The authors have used promoters over 1 kb in length that needed to be titrated with only 5 mM 3-AT. On the other hand the authors have experienced the situation whereby occasionally even a short sequence could confer so much HIS3 expression that this could not be inhibited with 3-AT up to 50 mM or higher, making its use in a screening impossible. Obviously, with longer promoters, the risk that such a strong enhancer is present becomes increased. Therefore, it is advised that several lengths or subfragments be tested. Screening conditions will be optimal if concentrations of 3-AT up to 25 mM suffice to reduce the HIS3 expression to an acceptable minimum. Higher concentrations of 50 up to 100 mM can still be tried, but the growth of the positives will also be slowed considerably, and it will be more difficult to distinguish these from the background. If the background is unworkably high, it may be useful to test the reporter in other yeast strains (Table 12.12.2), because the level of leaky expression can depend considerably on the genotype. Furthermore, the use of a yeast TF mutant as host strain could be considered, if clues exist about the identity of the endogenous yeast factor that might cause the background problem. If there are no leaky expression problems, it can be useful to make multimers of the bait sequence. This can increase the sensitivity of the detection of positive interactions in the library screening. The authors have designed a stepwise unidirectional multimerization procedure (Ouwerkerk and Memelink, 1997) to tetramerize sequences ranging from 9 to 250 bp, and have used these successfully in one-hybrid screenings (e.g., Meijer et al., 2000).

It is important to keep track of the number of clones that have been screened to be able to decide at what stage the library screening has been saturating. This is dependent on the complexity of the primary cDNA library. Furthermore, discrimination between true and false positives is an important factor. Isolated library clones should be retransformed to the baitHIS3 strain, and preferably also to different strains with negative control reporters (see Background Information). The nature of common false-positive cDNA clones is discussed under Anticipated Results. Finally, it is important to be aware of possible bacterial contamination in a yeast screening. Occasionally, bacterial contaminants can be very misleading because they can appear in close association with the yeast cells. Therefore, seemingly normal yeast colonies can occur on selective medium, while their capacity to grow actually results from bacterial crossfeeding rather than from expression of the auxotrophic marker gene. The presence of bacteria in suspected yeast colonies can easily be spotted with phase-contrast microscopy. Elimination is possible by streaking the colonies on CM −Leu −His plates supplemented with carbenicillin (50 to 200 mg/liter) and kanamycin (100 mg/liter)—antibiotics which do not affect the growth of yeast.

Anticipated Results It is difficult to predict how many positive clones are to be expected from a saturating library screening. This is very much dependent on the bait sequence as well as on the cDNA library source. First of all, the frequency of false positives is extremely variable. With some bait sequences, these can represent the majority of the clones obtained, whereas nearly all positives in other screenings may be specific transcription factors. Furthermore, the frequency of isolation of a certain clone is no indication for its validity. This frequency is determined by factors such as the abundance in the mRNA population from which the library was derived and the stability in E. coli and yeast. If no positives are obtained at all, it is useful to change the cDNA library. However, it is also possible that a library protein that could potentially interact with the bait sequence was present, but is degraded in yeast or sequestered due to interaction with a certain yeast protein. Another cause for an unsuccessful result may be that the bait sequence is masked or repressed due to binding of a yeast transcription factor. For this reason, it may be worthwhile to try



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different lengths or subfragments of the bait sequence. In the case where a very large number of positives are isolated, it is useful to attempt to sort them into classes, e.g., based on restriction or cross-hybridization patterns. Several groups of possible false positives can be discriminated. The first group includes, among others, nucleoside transporters, ribosomal proteins, RNA binding proteins, thymine glycosylases, amino-acyltransferases, and histones (P.B.F. Ouwerkerk, unpub. observ.; van der Fits, 2000). These factors all have in common the fact that their normal function is related to interactions with nucleic acids. As AD hybrids, such proteins are likely to turn into artifactual transcriptional activators. A second group of artifacts are components of signal transduction pathways such as phosphatases, kinases (Schouten, 1999), and GTPases (van der Fits, 2000), or of proteolysis pathways. Strikingly, their possible occurrence is very much dependent on the bait sequence used. Therefore, it is most likely that such proteins interfere with signal transduction pathways in yeast in such a way that specific activation occurs of endogenous yeast transcription factors that recognize the bait sequence. An exemplifying case is the search for plant factors regulating the T-cyt gene of the tumorinducing soil bacterium Agrobacterium tumefaciens (Schouten et al., 2000). Screenings in yeast resulted in several library clones that were able to activate HIS3 expression via a wild-type bait sequence but not via a mutant sequence, suggesting that the interaction was specific. However, instead of DNA-binding proteins, the discovered clones turned out to represent plant proteins homologous with the yeast Skp1 protein that is involved in ubiquitin-mediated protein degradation. The following mechanism was proposed for the interference of the Skp1 homologs with reporter gene activation. The Skp1-like proteins would interact with the yeast F-box protein, Grr1p. This contact would result in the destabilization of Grr1p, which in turn would inactivate the yeast repressor protein Mig1p. The Mig1p repressor interacted with the bait sequence and consequently its inactivation could make this sequence accessible to another yeast factor, resulting in activation of the reporter gene. Because both the Mig1p repressor and the yet unidentified yeast activator appeared to have the same binding specificity as the searched plant T-cyt binding factor, the identification of this plant factor via yeast screening was impossible.

The third group of false positives are His3p homologs, which can complement the mutant his3 locus in the yeast genome. These exist in plants and fungi, but not in animals. In the authors’ experience, their frequency of isolation in one-hybrid screenings is low. The fourth and last identified category of artifacts is miscellaneous and includes proteins involved in metabolic pathways, such as L-ascorbate peroxidase, pectinesterase, and others (van der Fits, 2000). Their occurrence is enigmatic and it is not very likely that these proteins directly activate reporter gene expression. The fact is that somehow they are beneficial for the yeast cells to survive under the selection conditions. It might be that they are involved in sequestering or degrading the His3p inhibitor 3-AT, due to which titration of leaky HIS3 reporter expression would fail and a false positive colony would grow. The protein-coding sequences of isolated cDNAs can be either partial or full-length. Preceding leader sequences can be translated to form a spacer between the AD and the library protein. Unexpectedly, sometimes positive cDNA clones are found for which the encoded protein sequence is not in frame with the AD sequence. Such clones can still represent true positives, as demonstrated by the following example. In a one-hybrid screening in yeast strains that contained reporters with upstream recognition sites for homeodomain proteins, the authors identified cDNAs from rice encoding homeodomain-leucine zipper (HD-Zip) proteins of two different families, I and II (Meijer et al., 2000). Strikingly all the family II members were in frame with the AD, but all the family I members were not. The authors concluded that the reporters must have been activated due to the fact that truncated HD-Zip I proteins were expressed, which possessed an intrinsic activation function. Subsequently, in plant experiments, it was indeed confirmed that HD-Zip I proteins could act as activators, whereas the HD-Zip II proteins turned out to be repressors. Since none of the isolated HD-ZIP I clones were in-frame fusions, in this case AD-hybrids might have been toxic, or the combination of Gal4p AD and intrinsic AD might have been incompatible.

Time Considerations When the required pHIS3NB/NX and pINT1 reporter constructs have been cloned, it will take ∼1 week to transform yeast and to select for G418-resistant pINT1-HIS3 reporter strains. The next week, the strains should be



tested on a range of 3-AT concentrations in order to determine the selective conditions for the actual library screening. Subsequently, it is useful to invest 1 to 2 weeks to fine tune the 3-AT concentration during pilot library transformations. Unless the transformation efficiency is suboptimal, all the transformations necessary to perform a saturating library screening, including the media preparation, can be finished within 1 week. The screening plates are incubated for 1 to 2 weeks during which positives can already be restreaked. As described in the protocol, from this point onward several strategies can be followed to discriminate between true and false positives, and to confirm the specificity of putative positives. Dependent on the numbers of positives and their nature, these analyses can take several weeks to many months.

Literature Cited Becker, D.M., Fikes, J.D., and Guarente, L. 1991. A cDNA encoding a human CCAAT-binding protein by functional complementation in yeast. Proc. Natl. Acad. Sci. U.S.A. 88:1968-1972. Brachmann, C.B., Davies, A., Cost, G.J., Caputo, E., Li, J., Hieter, P., and Boeke, J.D. 1998. Designer deletion strains derived from Saccharomyces cerevisiae S288C: A useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14:115-132. Bush, S.M., Folta, S., and Lannigan, D.A. 1996. Use of the yeast one-hybrid system to screen for mutations in the ligand-binding domain of the estrogen receptor. Steroids 61:102-109.

Gietz, D., St. Jean., A., Woods, R.A., and Schiestl, R.H. 1992. Improved method for high efficiency transformation of intact yeast cells. Nucleic Acids Res. 20:1425. Grueneberg, D.A., Natesan, S., Alexandre, C., and Gilman, M.Z. 1992. Human and Drosophila homeodomains proteins that enhance the DNAbinding activity of serum response factor. Science 257:1089-1095. Hadfield, C., Fordan, B.E., Mount, R.C., Pretorius, G.H.J., and Burak, E. 1990. G418 resistance as a dominant marker and reporter for gene expression in Saccharomyces cerevisiae. Curr. Genet. 18:303-313. Harper, J.W., Adami, G.R., Wei, N., Keyomarsi, K., and Elledge, S.J. 1993. The p21 Cdk-interacting protein Cip1 is a potent inhibitor of G1 cyclindependent kinases. Cell 75:805-16. Kim, S.Y., Chung, H.-W., and Thomas, T. 1997. Isolation of a novel class of bZIP transcription factors that interact with ABA-responsive and embryo-specification elements in the Dc3 promoter using a modified yeast one-hybrid system. Plant J. 11:1237-1251. Kunoh, T., Kaneko, Y., and Harashima, S. 2000. YHP1 encodes a new homeoprotein that binds to the IME1 promoter in Saccharomyces cerevisiae. Yeast 16:439-449. Leuther, K.K. and Johnston, S.A. 1992. Nondissociation of GAL4 and GAL80 in vivo after galactose induction. Science 256:1333-1335. Li, J.J. and Herskowitz, I. 1993. Isolation of ORC6, a component of the yeast origin recognition complex by a one-hybrid system. Science 262:18701874.

Chan, J.Y., Han, X.-L., and Kan, Y.W. 1993. Cloning of Nrf1, an NF-E2-related transcription factor, by genetic selection in yeast. Proc. Natl. Acad. Sci. U.S.A. 90:11371-11375.

Liu, J., Wilson, T.E., Milbrandt, J., and Johnston, M. 1993. Identifying DNA-binding sites and analyzing DNA-binding domains using a yeast selection system. In Methods: A Companion to Methods in Enzymology. Vol. 5 pp. 125-137. Academic Press, New York.

Christianson, T.W., Sikorski, R.S., Dante, M., Shero, J.H., and Hieter, P. 1992. Multifunctional yeast high-copy-number shuttle vectors. Gene 110:119-122.

Meijer, A.H., Ouwerkerk, P.B.F., and Hoge, J.H.C. 1998. Vectors for transcription factor cloning and target site identification by means of genetic selection in yeast. Yeast 14:1407-1415.

Elledge, S.J., Mulligan, J.T., Ramer, S.W., Spottswoo, M., and Davis, R.W. 1991. λYES: A multifunctional cDNA expression vector for the isolation of genes by complementation of yeast and Escherichia coli mutations. Proc. Natl. Acad. Sci. U.S.A. 88:1731-1735.

Meijer, A.H., de Kam, R.J., d’Erfurth I., Shen W., and Hoge, J.H.C. 2000. HD-Zip proteins of families I and II from rice: Interactions and functional properties. Mol. Gen. Genet. 263:12-21.

Fields, S. and Song, O. 1989. A novel genetic selection system to detect protein-protein interaction. Nature 340:245-246.

Melcher, K., Sharma B., Ding W.V., and Nolden, M. 2000. Zero background yeast reporter plasmids. Gene 247:53-61.

Foreman, P.K. and Davis, R.W. 1994. Cloning vectors for the synthesis of epitope-tagged, truncated and chimeric proteins in Saccharomyces cerevisiae. Gene 144:63-68.

Memelink, J. 1997. Two yeast/Escherichia coli lambda/plasmid vectors designed for yeast one- and two-hybrid screens that allow directional cDNA cloning. Technical Tips Online TT01111 (http://research.bmn.com/). Trends Genet. 13:376.

Forsburg, S.L. and Guarente, L. 1988. Mutational analysis of upstream activation sequence 2 of the CYC1 gene of Saccharomyces cerevisiae: A HAP2-HAP3-responsive site. Mol. Cell. Biol. 8:647-654.

Menke, F.L., Champion, A., Kijne, J.W., and Memelink, J. 1999. A novel jasmonate- and elicitor-responsive element in the periwinkle secondary metabolite biosynthetic gene Str interacts with a jasmonate- and elicitor-inducible AP2-domain



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transcription factor, ORCA2. EMBO J. 18:44554463. Mumberg, D., Müller, R., and Funk, M. 1994. Regulatable promoters of Saccharomyces cerevisiae: Comparison of transcriptional activity and their use for heterologous expression. Nucleic Acids Res. 14:7861-7871. Mumberg, D., Müller, R., and Funk, M. 1995. Yeast vectors for the controlled expression of heterologous proteins in different genetic backgrounds. Gene 156:119-122. Olesen, K., Johannesen, P.F., Hoffmann, L., Sørensen, S.B., Gjermansen, C., and Hansen, J. 2000. The pYC plasmids, a series of cassette-based yeast plasmid vectors providing means of counter-selection. Yeast 16:1035-1043.

van der Fits, L., Zhang, H., Menke, F.H.M., Deneka, M., and Memelink, J. 2000. A Catharanthus roseus BPF-1 homologue interacts with an elicitor-responsive region of the secondary metabolite biosynthetic gene Str and is induced by elicitor via a JA-independent signal transduction pathway. Plant Mol. Biol. 44:675-685. Vidal, M., Brachmann, R.K., Fattaey, A., Harlow, E., and Boeke, J.D. 1996. Reverse two-hybrid and one-hybrid systems to detect dissociation of protein-protein and DNA-protein interactions. Proc. Natl. Acad. Sci. U.S.A. 93:10315-10320. Wang, M. and Reed, R.R. 1993. Molecular cloning of the olfactory neuronal transcription factor Olf-1 by genetic selection in yeast. Nature 364:121-126.

Ouwerkerk, P.B.F. and Memelink, J. 1997. A simple method for directional multimerization of DNA s eq u e n ce s. Tec h n i cal Tips Online T01051(http://research.bmn.com/)/Trends Genet. 13:207.

Wei, Z., Angerer, R.C., and Angerer, L.M. 1999. Identification of a new sea urchin ets protein, Spets4, by yeast one-hybrid screening with the hatching enzyme promoter. Mol. Cell. Biol. 19:1271-1278.

Schouten, J. 1999. Identification of Arabidopsis thaliana cDNAs coding for proteins that can activate gene expression in yeast via the cyt-1 element of the Agrobacterium tumefaciens T-cyt gene promoter. Ph.D. Thesis, Leiden University, The Netherlands.

Wilson, T.E., Fahrner, T.J., Johnston, M., and Milbrandt, J. 1991. Identification of the DNA binding site for NGFI-B by genetic selection in yeast. Science 252:1296-1300.

Schouten, J.S., de Kam, R.J., Fetter, K., and Hoge, J.H.C. 2000. Overexpression of Arabidopsis thaliana SKP1 homologues in yeast inactivates the Mig1 repressor by destabilising the F-box protein Grr1. Mol. Gen. Genet. 263:309-319.

Yu, Y., Li, W., Su, K., Yussa, M., Han, W., Perrimon, N., and Pick, L. 1997. The nuclear hormone receptor Ftz-F1 is a cofactor for the Drososphila homeodomain protein Ftz. Nature 385:552-555.

Key References Meijer et al., 1998. See above.

Sieweke, M.H., Tekotte, H., Frampton, J., and Graf, T. 1996. MafB is an interaction partner and repressor of Ets-1 that inhibits erythroid differentiation. Cell 85:49-60.

First description of the pHIS3/pINT1 vector system for yeast reporter strain construction.

Sikorski, R.S. and Boeke, J.D. 1991. In vitro mutagenesis and plasmid shuffling: From cloned gene to mutant yeast. In Methods of Enzymology, Vol. 194: Guide to Yeast Genetics and Molecular Biology (C. Guthrie and G.R. Fink, eds.) pp. 302-318. Academic Press, San Diego.

http://www.atcc.org/

Sikorski, R.S. and Hieter, P. 1989. A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122:19-27. Singh, H., LeBowitz, J.H., Baldwin, A.S., Jr., and Sharp, P.A. 1988. Molecular cloning of an enhancer binding protein: Isolation by screening of an expression library with a recognition site DNA. Cell 52:415-423. van der Fits, L. 2000. Transcriptional regulation of stress-induced plant secondary metabolism. Ph.D. Thesis, Leiden University, The Netherlands.

Internet Resources American Type Culture Collection(ATCC): source for yeast vectors and strains. http://www.ncbi.nlm.nih.gov/ National Center for Biotechnology Information (NCBI, Genbank): public database of genes and vectors sequences, computational tools for sequence analysis http://www.clontech.com http://www.stratagene.com/ Commercial sources for vectors, AD-hybrid libraries, yeast strains, and supplies.

Contributed by Pieter B.F. Ouwerkerk and Annemarie H. Meijer Leiden University Leiden, The Netherlands




CHAPTER 13 Yeast INTRODUCTION Saccharomyces cerevisiae (baker’s yeast) and Schizosaccharomyces pombe (fission yeast) are often considered to be model eukaryotic organisms, in a manner analogous to Escherichia coli as a model prokaryotic organism. Both yeasts have been extensively characterized and their genomes completely sequenced. They are as easy to grow as other microorganisms, and they have a haploid nuclear DNA content only 3.5 times that of E. coli. However, despite the small genomes sizes, these yeasts display most of the features of higher eukaryotes. The fact that many cellular processes are conserved among different eukaryotic species- combined with the powerful genetic and molecular tools that are available- has made these yeasts important experimental organisms for a variety of basic problems in eukaryotic molecular biology. Primarily for historical reasons, most studies on yeast have involved Saccharomyces cerevisiae (hereafter termed yeast). Culturing yeast is simple, economical, and rapid, characterized by a doubling time of ∼90 min on rich medium. In addition, yeast has been well adapted to both aerobic and anaerobic large-scale culture. Cells divide mitotically by forming a bud, which pinches off to form a daughter cell. The progression through the cell cycle can be monitored by the size of the bud; this has been used to isolate a large collection of mutants (called cdc mutants) that are blocked at various stages of the cell cycle. Since yeast can be grown on a completely defined medium (see UNIT 13.1), many nutritional auxotrophs have been isolated. This has not only permitted the analysis of complex metabolic pathways but has also provided a large number of mutations useful for genetic analysis. Yeast can exist stably in either haploid or diploid states. A haploid cell can be either of two mating types, called a and α. Diploid a/α cells—formed by fusion of an α cell and an a cell (UNIT 13.2)—can grow mitotically indefinitely, but under conditions of carbon and nitrogen starvation will undergo meiosis. The meiotic products, called spores, are contained in a structure called an ascus. After gentle enzymatic digestion of the thick cell wall of the ascus, the haploid spore products can be individually isolated and analyzed (UNIT 13.2). This ability to recover all four products of meiosis has allowed detailed genetic studies of recombination and gene conversion that are not possible in most other eukaryotic organisms. The existence of stable haploid and diploid states also facilitates classical mutational analysis, such as complementation tests and identification of both dominant and recessive mutations. The haploid yeast cell has a genome size of about 15 megabases and contains 16 linear chromosomes, ranging in size from 200 to 2200 kb. Thus, the largest yeast chromosome is still 100 times smaller than the average mammalian chromosome. This small chromosome size, combined with the advent of techniques for cloning yeast genes as well as manipulating yeast chromosomes, has allowed detailed studies of chromosome structure. Three types of structural elements required for yeast chromosome function have been identified and cloned: origins of replication (ARS elements), centromeres (CEN elements), and telomeres. The cloning of these elements has led to the construction of artificial chromosomes that can be used to study various aspects of chromosome behavior, such as how chromosomes pair and segregate from each other during mitosis and meiosis. In addition, systems using artificial chromosomes have been designed that allow cloning of Contributed by Victoria Lundblad and Kevin Struhl Current Protocols in Molecular Biology (2003) 13.0.1-13.0.3 Copyright © 2003 by John Wiley & Sons, Inc.

Yeast


larger contiguous segments of DNA (up to 400 kb) than are obtainable in other cloning systems. These structural elements, as well as cloned selectable yeast genes, have permitted the construction of yeast/E. coli shuttle vectors that can be maintained in yeast as well as in E. coli (UNITS 13.4 & 13.6). Procedures for high-efficiency transformation of yeast (UNIT 13.7) have been available for nearly two decades, allowing cloning of genes by genetic complementation (UNITS 13.8 & 13.9). Because yeast has a highly efficient recombination system, DNAs with alterations in cloned genes can be reintroduced into the chromosome at the corresponding homologous sites (UNIT 13.10). This has permitted the rapid identification of the phenotypic consequences of a mutation in any cloned gene, a technique generally unavailable in higher eukaryotes. In addition, homologous recombination permits a wide variety of genetic techniques that have greatly facilitated the analysis of biological processes. Despite its small genome size, yeast is a characteristic eukaryote, containing all the major membrane-bound subcellular organelles found in higher eukaryotes, as well as a cytoskeleton. Yeast DNA is found within a nucleus and nucleosome organization of chromosomal DNA is similar to that of higher eukaryotes, although no histone H1 is present. Three different RNA polymerases transcribe yeast DNA, and yeast mRNAs (transcribed by polymerase II) show characteristic modifications of eukaryotic mRNAs [such as a 5′ methyl-G cap and a 3′ poly(A) tail], although only a few S. cerevisiae genes contain introns. Transcriptional regulation has been extensively studied and at least one yeast transcriptional activator has been shown to function in higher eukaryotes as well. High-molecular-weight yeast DNA and RNA can be prepared fairly quickly (UNITS 13.11 & 13.12). Another characteristic of eukaryotes is the proteolytic processing of precursor proteins to yield functional products, which is often coupled to secretion. Yeast has several well-studied examples of secreted proteins and pheromones, and the large number of genes that have been identified as involved in protease processing and secretion suggests a highly complex pathway. Yeast protein extracts can be prepared using three different protocols (UNIT 13.13); the best choice will depend on the particular application. The ease and power of genetic manipulation in yeast facilitate the use of this organism to detect novel interacting proteins using the two-hybrid system or interaction trap (UNIT 20.1). Although Saccharomyces cerevisiae is the most commonly studied yeast, S. pombe is also an important experimental organism (UNIT 13.14). Although both yeasts are unicellular microorganisms that grow in similar medium, they are evolutionarily quite distant. It has become increasingly clear that, in terms of molecular mechanisms, S. pombe is more similar to higher eukaryotic organisms than S. cerevisiae. Experimental manipulations in S. pombe are broadly similar to those in S. cerevisiae, although the technical details often differ. The chapter includes units on S. pombe relating to strain maintenance and media (UNIT 13.15), growth and genetic manipulation (UNIT 13.16), and introduction of DNA into cells (UNIT 13.17). This chapter is written for the molecular biologist who has not previously worked with yeast. The glossary below introduces the terms of yeast molecular biology. aerobic growth growth in the presence of oxygen, utilizing the Krebs cycle. α and a factor mating type–specific polypeptides secreted by either α or a haploid cells, respectively, which interact with haploid cells of the opposite mating type to stimulate mating. anaerobic growth growth in the absence of oxygen, utilizing fermentation (via glycolysis).

ARS elements DNA sequences present throughout the yeast genome that confer autonomous replication on plasmids in yeast; most of these sequences also function as chromosomal origins of replication as well. ascus thick-walled sac containing the four haploid products, called spores, resulting from meiosis.

Introduction



cdc mutants strains of yeast that exhibit stagespecific blocks of the cell cycle; these mutations define genes important in DNA replication, meiosis, and sporulation. CEN element DNA sequences present at the centromere that ensure proper segregation of chromosomes during mitosis and meiosis, presumably by promoting interaction with the mitotic spindle. cir+ strains of yeast that contain the naturally occurring 2µm plasmid. ciro yeast strains that have lost this endogenous 2µm plasmid. δ element ∼330-bp sequence, present as direct repeats at the ends of the transposable element Ty1, and also found dispersed throughout the genome. gene disruption a mutation constructed in vitro in a cloned gene which, upon reintroduction into the genome at the homologous chromosomal site, results in inactivation of the gene function. glusulase a digestive enzyme isolated from snails that breaks down thick cell walls of either an ascus to allow isolation of spore products, or a yeast cell to produce spheroplasts. heterothallic common laboratory strains of yeast which—due to a mutation in the HO gene—stably maintain a given allele at the MAT locus. homothallic strains of yeast (typically found in the wild) that, in a haploid state, rapidly interconvert the MAT locus, resulting in rapid switching between the α and a mating types; cultures of such strains rapidly diploidize. killer strains strains of yeast that harbor a double-stranded RNA virus; such strains kill sensitive yeast strains via secretion of a protein toxin, to which killer strains are immune. MAT the mating-type locus which is expressed and therefore determines the mating type of a haploid cell; this locus has two alleles—the MATa allele confers the a mating type, while MATα specifies the α mating type. meiosis the process by which the number of chromosomes present in a diploid cell is halved to yield haploid products. mitosis vegetative cell division (of either haploid or diploid cells) in which the chromosome

number stays the same. petites mutants of yeast (either nuclear or mitochondrial) with impaired mitochondria function; they grow as small colonies on fermentable carbon sources and are unable to grow on nonfermentable carbon sources. schmoo a distinctive shape of a haploid cell (pear-shaped), induced by exposure to mating pheromone. sporulation the end product of meiosis, induced by carbon and nitrogen starvation of a diploid cell, which results in four haploid progeny contained as spores within an ascus; this complicated developmental process requires over 200 genes. telomeres DNA sequences found at the end of linear chromosomes that are essential for chromosome stability and complete replication; in S. cerevisiae, telomeres consist of tandem repeats of the sequence 5′dG1-3dT3′. Ty1 elements the primary transposable element found in yeast, which functions as a retrotransposon (via reverse transcription of its RNA and subsequent reinsertion into the genome). UAS element upstream activating sequences in yeast promoters, to which regulatory proteins bind in order to enhance the rate of transcription. zygote a morphologically distinct cellular structure formed by the fusion of two haploid cells of opposite mating type, which results in formation of a diploid. Zymolyase β-glucanase, isolated from Arthrobacter luteus, that hydrolyzes the yeast cell wall and is used to prepare spheroplasts for a variety of purposes.

Key References Watson, J.D., Hopkins, N.H., Roberts, J.W., Steitz, J.A., and Weiner, A.M. 1987. Yeasts as the E. coli of eukaryotic cells. In Molecular Biology of the Gene, Vol. 1, pp. 550-594. Benjamin/Cummings, Menlo Park, Calif. Strathern, J.N., Jones, E.W., and Broach, J.R. (eds.) 1981. The Molecular Biology of the Yeast Saccharomyces: Life Cycle and Inheritance. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Strathern, J.N., Jones, E.W. and Broach, J.R. (eds.) 1982. The Molecular Biology of the Yeast Saccharomyces: Metabolism and Gene Expression. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.

Victoria Lundblad and Kevin Struhl Yeast


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BASIC TECHNIQUES OF YEAST GENETICS

SECTION I

Like Escherichia coli, yeast can be grown in either liquid media or on the surface of (or embedded in) solid agar plates. Yeast cells grow well on a minimal medium containing dextrose (glucose) as a carbon source and salts which supply nitrogen, phosphorus, and trace metals. Yeast cells grow much more rapidly, however, in the presence of protein and yeast cell extract hydrolysates, which provide amino acids, nucleotide precursors, vitamins, and other metabolites which the cells would normally synthesize de novo. During exponential or log-phase growth, yeast cells divide every 90 min when grown in such media. Log phase can be divided into three stages based on the rate of cell division (or the proportion of budded cells within a culture), which is in turn a function of the cell density of the culture. As cell density increases, nutrient supplies drop and the rate of cell division slows (the measurement of cell density, as well as techniques for the propagation and genetic manipulation of yeast, are described in UNIT 13.2). Early-log phase is the period when cell densities are 5 years), or at 4°C on slants consisting of rich medium supplemented with potato starch (viable for 1 to 2 years). Both methods are described below. BASIC PROTOCOL

Preparation and Inoculation of Frozen Stocks Make a solution of 30% (w/v) glycerol. Pipet 1 ml into 15 × 45-mm, 4-ml screwcap vials. Loosely cap the vials and autoclave 15 min. To inoculate vials for storage, add 1 ml of a late-log or early-stationary phase culture, mix, and set on dry ice. Store at −70°C. Revive by scraping some of the cells off the frozen surface and streak onto plates. Do not thaw the entire vial. Cells can also be stored in the same way by adding 80 µl dimethyl sulfoxide (DMSO) to 1 ml cells (8% v/v) and storing at −70°C.

ALTERNATE PROTOCOL

Preparation and Inoculation of Slants 1. For 250 slants, add the following ingredients to a 1-liter flask: 70 g YPD plate premix 20 mg adenine (hemisulfate salt) 20 g potato starch H2O to 500 ml Excess adenine prevents ade− mutations from being lost.

2. Set the flask in a 4-liter beaker filled with 1 liter of water. Place the beaker on a heat-controlled, magnetic stirring apparatus and stir with the heat setting on high. With smooth and continuous stirring, the contents should not burn and should be molten after ∼1 hr.

3. With the entire setup in place, begin pipetting 2-ml aliquots into 15 × 45-mm, 4-ml screwcap vials. When all vials have been filled, put the caps on loosely, pack in the original boxes, and autoclave 15 min. 4. Lean the boxes against a support at an angle of ∼70°. Allow the slants to dry 2 days and screw the caps on tightly. Preparation of Yeast Media

Slants can be stored at room temperature for at least 6 months.



5. To inoculate a slant, smear cells from the flat end of a sterile toothpick onto the agar surface of the slant. Cap loosely and incubate 1 or 2 days at 30°C. After growth, screw the cap on as tightly as possible and store at 4°C. Slants are a convenient way to store and mail strains. They can be mailed immediately after inoculating since sufficient growth will occur in transit. See another method for mailing strains below.

Mailing and Reviving Strains Yeast strains can be conveniently mailed as slants. Alternatively, transfer cells to a piece of sterile Whatman 3MM paper by pressing the paper onto the desired yeast colony using forceps that have been dipped in ethanol and flamed. Wrap the paper in sterile aluminum foil and mail to recipient.

BASIC PROTOCOL

Revive the strain by placing the paper (yeast side down) on the surface of an agar plate. Incubate the plate at 30°C. A thick patch of yeast should be visible after lifting the paper. LITERATURE CITED Boeke, J.D., LaCroute, F., and Fink, G.R. 1984. A positive selection for mutants lacking orotidine-5′-phosphate decarboxylase activity in yeast: 5-fluoroorotic acid resistance. Mol. Gen. Genet. 197:345-346. Brown, P.A. and Szostak, J.W. 1983. Yeast vectors with negative selection. Meth. Enzymol. 101:278-290. Sherman, F., Fink, G.R., and Lawrence, C.W. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.

Contributed by Douglas A. Treco Massachusetts General Hospital Boston, Massachusetts Victoria Lundblad Baylor College of Medicine Houston, Texas

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UNIT 13.2

Growth and Manipulation of Yeast Aside from different media requirements, yeast cells are physically manipulated essentially as described for bacterial cells—i.e., they are grown in liquid culture (in tubes or flasks) or on the surface of agar plates and are manipulated using the basic equipment described in UNITS 1.1-1.3. In addition, a well-equipped yeast laboratory requires static and shaking incubators dedicated to 30°C and a microscope with magnification up to 400×. A second microscope adapted for dissecting yeast tetrads is extremely valuable for the genetic analyses and strain constructions described in this unit. A small electric clothes dryer is indispensable when replica plating is done frequently and large numbers of velvets are regularly used. This unit first presents the necessary details for growing yeast cells. This is followed by a description of replica plating methods for assessing the nutritional requirements and mating types of strains. Yeast genetic experiments often require the construction of strains with specific genotypes, as well as an analysis of the meiotic segregation patterns of newly introduced mutations (see UNIT 13.8). These genetic manipulations are carried out using the protocols presented in the final sections of this unit, which describe the construction and selection of diploids, sporulation, and tetrad analysis.

BASIC PROTOCOL

GROWTH IN LIQUID MEDIA Wild-type S. cerevisiae grows well at 30°C with good aeration and with glucose as a carbon source. When using culture tubes, vortex the contents briefly after inoculation to disperse the cells. Erlenmeyer flasks work well for growing larger liquid cultures, and baffled-bottom flasks to increase aeration are especially good. It is important that all glassware be detergent-free. For good aeration, the medium should constitute no more than one-fifth of the total flask volume, and growth should be carried out in a shaking incubator at 300 rpm. For small-scale preparations of DNA and RNA, yeast can be grown in glass or plastic culture tubes filled one-third full with medium and shaken at 350 rpm in a rack firmly attached to a shaking incubator platform.

BASIC PROTOCOL

BASIC PROTOCOL

GROWTH ON SOLID MEDIA Yeast cells can be streaked or spread on plates as shown for bacteria in the sketches in UNIT 1.3. When a dilute suspension of wild-type haploid yeast cells is spread over the surface of a YPD plate and incubated at 30°C, single colonies may be seen after ∼24 hr but require ≥48 hr before they can be picked or replica plated (see below). Growth on dropout media (UNIT 13.1) is about 50% slower. DETERMINATION OF CELL DENSITY The density of cells in a culture can be determined spectrophotometrically by measuring its optical density (OD) at 600 nm. For reliable measurements, cultures should be diluted such that the OD600 is Brightness/Contrast Process >Subtract Background (this requires loading of a background image that is devoid of any structures of interest) Process > Filters > Gaussian Blur Analyze >Measure 3. Rotate digital sequences by selecting Rotate from the Image menu for export to QuickTime, which can then be played in Powerpoint presentations or uploaded onto the Web in the orientation that makes most sense in comparison to other image sequences or to a model/diagram. In ImageJ, if a processing routine is applied to a digital sequence, one will be stuck with the results (i.e., this is destructive editing). Therefore, one should always keep a file of the original data separate from the sequence being processed, and save often to preserve desired changes in case subsequent processing is not useful.

4. Save the new image sequences by selecting Save As from the File menu. For a TIFF series, select Image Sequence from the submenu that appears. For QuickTime, select options from the popup menu: Compressor: Video Color Quality: Medium to High Timing: Frames per Second 6-10 Keyframe every 10 Further processing and analysis can be performed in ever increasingly complex software applications.

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Image Analysis, Image Importing, and Image Processing Using Freeware

Image analysis to determine the structure of molecules and particles Prior to the advent of digital image processing, various manual manipulations were required to extract quantitative information from structural data. The term “structural data” in this context refers to photographic or digital images of biological specimens, ranging from whole animals to tiny single molecules. For example, diffraction patterns can be obtained by passing laser light through film containing images of subcellular structures displaying repetitive patterns resembling crystalline arrays, such as the catalase crystal in the peroxisome (Bearer and Orci, 1986), the acetylcholine receptor channel (Unwin, 2002), and the protein coat of viruses (DeRosier and Klug, 1972). Reconstruction of the “ideal” crystalline structure is obtained by solving the diffraction pattern in reciprocal space. Another approach superimposes multiple electron microscopic images of the same structure, which “averages” the image, decreasing the intensity of nonrepeating structures, including those of unrelated background particles or of the movable parts within a fluid region of the structure. This technique has commonly been used in the analysis of electron micrographs. Multiple exposures of the same photographic paper with different images or rotations of the same image superimposed upon itself identifies repeating structures and internal symmetries within a structure. This technique was used to discover the eight-fold symmetry of the endothelial fenestral diaphragm, responsible for the passage of nutrients from blood to tissue in human organs (Bearer and Orci, 1985). Statistical analysis of biological specimens, also known as “morphometrics,” involves compiling measurements of multiple specimens of the same structure and then mathematically deriving the average size and other statistical values such as standard deviation within the population studied. Such mathematical compilations rely on the acquisition of a large number of measurements from multiple specimens, which until very recently was performed by hand. However, only relatively small sample sizes are easily analyzed by hand, since this requires a large time investment. Recent advances in structural determination of complex cellular components not amenable to crystallization include tomographic recon-

struction of electron-microscopic images. In this approach, images are captured from thick sections of biological specimens embedded in either plastic or vitreous ice using an intermediate-voltage electron microscope. The section is photographed at various angles of tilt, and the 3-D structure assembled digitally. The structure of the neuronal synapse is yielding to this approach (Harlow et al., 1998), which was recently cited as one of the ten top scientific advances in 2002 by the journal Science (News and Editorial Staff, 2002). New technology exploiting digital image processing includes laser scanning and twophoton confocal microscopy, which allows elimination of the out-of-focus light from fluorescently labeled specimens. Confocal microscopy allows optical sectioning and thereby the acquisition of sharp images of objects deep within thick specimens (White et al., 1987). In confocal microscopy, the image is filtered before capture. Confocal microscopy relies on monochromatic laser excitation of fluorescent tags, and collects only in-focus emission using a pinhole to block out-of-focus emitted light. In two-photon confocal imaging, a recent advance, excitation and collection occurs only from the same optical section of interest, thus prolonging the length of time that a living biological specimen can be observed without laser damage (Kaech et al., 2001; Meyer et al., 2003). New computational approaches (Meta and Multi-Tracker options in the Zeiss 510 LSM package) narrow the bandwidth of detected emitted light, thus providing the possibility of imaging multiple fluorochromes with the same excitation and even only slightly differing emission spectra in the same specimen. An alternative way to eliminate out-of-focus light is deconvolution, which involves digital processing of a series of images obtained by focusing up and down in the same microscopic field. In this approach, out-of-focus light is eliminated digitally after image capture (Hiraoka et al., 1990). Deconvolution requires a motorized stage to collect a series of images in the z-plane (Kam et al., 2001). These images are then processed digitally to extract in-focus structure and detail and eliminate fuzz from out-of-focus or autofluorescent emissions. The equipment is less costly than that used for confocal, but collection times are longer, and thus deconvolution is not as suitable for living specimens undergoing rapid changes. Both of these approaches allow visualization of struc-



tures by fluorescence microscopy that would not be resolved by conventional microscopy. Useful texts on microscopy include Matsumoto (1993) and Inoué and Spring (1997). Microscopy courses at the Marine Biological Laborato ry in Wo od s Hole, Massachusetts (http://www.mbl.edu/) are also excellent for viewing demonstrations of the latest equipment. Video microscopy, pioneered by Robert Allen in the late 1970s using Nomarski optics (differential interference contrast or DIC), is capable of detecting structures that are theoretically below the resolution of the light microscope if lenses with high numerical apertures are used (Allen et al., 1979; Hayden and Allen, 1984; Brady et al., 1985). Such lenses have narrow focal planes, and often have short working distances as well; thus, they have not yet proven useful for imaging objects deep within thick specimens. However, like confocal microscopy, video imaging with either DIC or fluorescent tags can be performed on live specimens. Analysis of such sequence data can provide information about structural changes over time, such as the rates of changes in cellular surface contours (Zhang et al., 2002), the velocities of movement of intracellular particles such as viruses (Bearer et al., 2000), and even the dynamic behavior of purified cytoskeletal filaments during interactions with motors or severing factors (Vale et al., 1985; Bearer, 1991). In this case, microscopy becomes an assay for enzymatic activity. An Open Microscopy Environment (OME) is being developed as an informatics solution for storage and analysis of optical microscope data (Swerdlow et al., 2003). This public resource hopes to provide image analysis automation, modeling tools, and large sets of data for mining biological material. The goal is for data to be stored as an XML-encoded standard file which can be uploaded and processed by any software. The URL for this project http://www.openmicroscopy.org. Image analysis to determine clues of function from subcellular location and behavior In addition to providing information about the exact measurements of the “ideal” for a particular structure, image analysis is also useful to discover expression patterns, location within cells, and dynamic behaviors of molecules both large (proteins and RNA) and small (ATP and calcium ions). The expression patterns and behaviors of proteins inside cells

detected by microscopy complement those detected by biochemical and molecular approaches, and readily provide functional information about a protein or gene product. Because the genome projects have identified a large number of novel genes whose function is not easily predicted from DNA sequence, functional clues derived from structural analysis have become increasingly fashionable. Morphological data are being collected in highthroughput screening assays to identify key proteins from large data sets. Such key proteins can then be selected for analysis by more timeconsuming approaches in subsequent experiments. Strategies using localization to derive functional information about individual proteins and thereby select interesting proteins from large data sets were originally developed using antibodies to localize the protein (Bearer and Alberts, 1988; Miller et al., 1989). Antibody staining provides evidence concerning the type of cell within a tissue that expresses the protein and the subcellular location of that protein in cells. In conjunction with other stains, including antibodies, against other cellular proteins whose function is known, the contribution of the unknown protein to a particular aspect of cellular behavior can be inferred. In addition to antibody staining, several other methods have been developed to detect gene expression. In situ hybridization (ISH) to detect mRNA expression speeds up the analysis because it uses the DNA as a probe and bypasses antibody production, which is time-consuming and not always reliable. However, results from ISH do not produce information about protein function, only about relative expression between cells in a tissue. Another method, transfection and overexpression of fusions of the target protein with tags such as green fluorescent protein (GFP), allows observation of the dynamic behavior of a protein in the living cell. This method has most commonly been applied to individual proteins once they are determined to be of interest and is not a commonly used screen, as construction of the GFP fusion takes time. Another powerful approach is to label a protein whose function is known with GFP and then observe the effect of overexpression, deletion, or mutation of the unknown protein on the GFP-labeled structures. Finally, labeling of a protein within a larger complex of proteins allows microscopic tracking of the complex within cells, a strategy used to study the movement of viruses within cells (Bearer et al., 2000).



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Questions that microscopic analyses usually address include: (1) the timing of expression changes after stimulation, (2) the types of cells within a multicellular tissue in which the gene product is expressed, and (3) the location of the gene product within the cell and its association with other cellular structures of known function. The function of an unknown protein or small molecule can be assessed using substrate cells that are transfected with some other protein whose function is already known, and monitoring the effect of the novel protein on the behavior of the known protein. This approach has proven very useful in identifying proteins whose function affects the secretory pathway, such as Golgi markers (LippincottSchwartz, 2001; Kreitzer et al., 2003) or the actin cytoskeleton (Westphal et al., 1997; Fischer et al., 1998). While all of these types of structural expression analysis have in common the use of the microscope to obtain images for further analysis, the exact method and interpretation of results differ depending on the question being addressed. Microscope assays can be divided into two subsets: those acquired from living specimens in real time or by time lapse, and those acquired from fixed specimens.These assays use a variety of different microscope technologies and experimental methods. This is a fast-moving area of research, and specific applications and equipment are likely to have evolved between the writing of this unit and its appearance in print.

Critical Parameters

Image Analysis, Image Importing, and Image Processing Using Freeware

Image analysis is only as good as the images being analyzed. It is often a fruitless waste of time to use digital image processing to improve poor images. Therefore, before embarking on an analytical or processing project, images should be carefully scrutinized for: (1) focus; (2) good signal-to-noise ratios; (3) reproducibility; and (4) background contamination. Artifacts to be avoided are those from: (1) laser or other light damage; (2) aggregation of fluorochrome marker; (3) nonphysiologic localizations of fusion proteins; (4) movement of the stage during collection of video images; and (5) other effects of the experiment on the cell that might damage it or perturb its normal processes. Scientists engaged in monitor-based digital processing tasks should ensure that the monitor’s colors, brightness, and contrast are set appropriately. Instructions for this come with the Adobe Photoshop package, and must be

tailored to the individual monitor being used. Printing of processed images requires that the printer be coordinated with the monitor as well. This is dependent on the manufacturer’s specifications for the particular printer and monitor being used. Monitor-based processing requires a low-light work area and a maximum-resolution screen.

Time Considerations Digital image processing and analysis are usually more time-consuming than expected. The better the original image(s), the less time is required for processing. The type of data being extracted also define the length of time that must be invested. If software routines are available, then applying them can be fast once a sequence of steps is defined. It usually takes a graduate student a week to learn a routine, then a half-day to a day to make a processed QuickTime movie from a preselected, excellent series of images.

Literature Cited Allen, R.D., Zacharski, L.R., Widirstky, S.T., Rosenstein, R, Zaitlin, L.M., and Burgess, D.R. 1979. Transformation and motility of human platelets: Details of the shape change and release reaction observed by optical and electron microscopy. J. Cell Biol. 83:126-142. Bearer, E.L. 1991. Direct observation of actin filament severing by gelsolin and binding by gCap 39 and CapZ. J. Cell Biol. 115:1629-1638. Bearer, E.L. and Alberts, B.M. 1988. Novel actinbinding proteins associated with platelet activation. FASEB J. 2:A1538. Bearer, E.L. and Orci, L. 1985. Endothelial fenestral diaphragms: A quick-freeze, deep-etch study. J. Cell Biol. 100:418-428. Bearer, E.L. and Orci, L. 1986. A simple method for quick-freezing. J. Electron Microsc. Tech. 3:233241. Bearer, E.L., Breakefield, X.O., Schuback, D., Reese, T.S., and LaVail, J.H. 2000. Retrograde axonal transport of Herpes Simplex Virus: Evidence for a single mechanism and a role for tegument. Proc. Natl. Acad. Sci. U.S.A. 97:81468150. Brady, S.T., Lasek, R.J., and Allen, R.D. 1985. Video microscopy of fast axonal transport in extruded axoplasm: A new model for study of molecular mechanisms. Cell Motil. 5:81-101. DeRosier, D.J. and Klug, A. 1972. Structure of the tubular variants of the head of bacteriophage T4 (polyheads). I. Arrangement of subunits in some classes of polyheads. J. Mol. Biol. 65:469-488. Fischer, M., Kaech, S., Knutti, D., and Matus, A. 1998. Rapid actin-based plasticity in dendritic spines. Neuron 20:847-854.



Harlow, M., Ress, D., Koster, A., Marshall, R.M., Schwarz, M., and McMahan, U.J. 1998. Dissection of active zones at the neuromuscular junction by EM tomography. J. Physiol. Paris 92:75-78. Hayden, J.H. and Allen, R.D. 1984. Detection of single microtubules in living cells: Particle transport can occur in both directions along the same microtubule. J. Cell Biol. 99:1785-1793. Hiraoka, Y., Agard, D.A., and Sedat, J.W. 1990. Temporal and spatial coordination of chromosomal movements. J. Cell Biol. 111:2815-2828. Inoué, S. and Spring, K.R. 1997. Video Microscopy. Plenum, New York. Kaech, S., Parmar, H., Roelandse, M., Barnmann, C., and Matus, A. 2001. Cytoskeletal microdiffentiation: A mechanism of organizing morphological plasticity in dendrites. Proc. Natl. Acad. Sci. U.S.A. 98:7086-7092. Kam, Z., Hanser, B., Gustafsson, M.G., Agard, D.A., and Sedat, J.W. 2001. Computational adaptive optics for live three-dimensional biological imaging. Proc. Natl. Acad. Sci. U.S.A. 98:3790-3795. Kreitzer, G., Schmoranzer, J., Low, S.H., Li, X., Gan, Y., Weimbs, T., Simon, S.M., and Rodriguez-Boulan, E. 2003. Three-dimensional analysis of post-Golgi carrier exocytosis in epithelial cells. Nat. Cell Biol. 5:126-136. Lippincott-Schwartz, J. 2001. The secretory membrane system studied in real-time: Robert Feulgen Prize Lecture, 2001. Histochem. Cell Biol. 116: 97-107. Matsumoto, B. 1993. Cell Biological Applications of Confocal Microscopy. In Methods in Cell Biology vol. 38 (B. Matsumoto, ed.) Academic Press, San Diego. Meyer, M.P., Niell, C.M., and Smith, S.J. 2003. Brain imaging: How stable are synaptic connections? Curr. Biol. 13:R180-182.

Miller, K.G., Field, C.M., and Alberts, B.M. 1989. Actin-binding proteins from Drosophila embryos: A complex network of interacting proteins detected by F-actin affinity chromatography. J. Cell Biol. 109:2963-2975. News and Editorial Staff. 2002. Breakthrough of the year: The runners-up. Science 298:2297-2303. Swerdlow, J., Goldberg, I., Brauner, E., and Sorger, P.K. 2003. Informatics and quantitative analysis in biological imaging. Science 300:100-102. Unwin, N. 2002. Structure of the acetylcholinegated channel. Novartis Found Symp. 245:5-21. Vale, R.D., Schnapp, B.J., Reese, T.S., and Sheetz, M.P. 1985. Movement of organelles along filaments dissociated from the axoplasm of the squid giant axon. Cell 40:449-454. Westphal, M., Jungbluth, A., Heidecker, M., Muhlbauer, B., Heizer, C., Schwartz, J.M., Marriott, G., and Gerisch, G. 1997. Microfilament dynamics during cell movement and chemotaxis monitored using a GFP-actin fusion protein. Curr. Biol. 7:176-183. White, J.G., Amos, W.B., and Fordham, M. 1987. An evaluation of confocal versus conventional imaging of biological structure by fluorescence light microscopy. J. Cell Biol. 105:41-48. Zhang, H., Wessels, D., Fey, P., Daniels, K., Chisholm, R.L., and Soll, D.R. 2002. Phosphorylation of the myosin regulatory light chain plays a role in motility and polarity during Dictyostelium chemotaxis. J. Cell Sci. 115:17331747.

Contributed by E.L. Bearer Brown University Providence, Rhode Island



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CHAPTER 15 The Polymerase Chain Reaction INTRODUCTION

T

he polymerase chain reaction (PCR) is a rapid procedure for in vitro enzymatic amplification of a specific segment of DNA. Like molecular cloning, PCR has spawned a multitude of experiments that were previously impossible. The number of applications of PCR seems infinite—and is still growing. They include direct cloning from genomic DNA or cDNA, in vitro mutagenesis and engineering of DNA, genetic fingerprinting of forensic samples, assays for the presence of infectious agents, prenatal diagnosis of genetic diseases, analysis of allelic sequence variations, analysis of RNA transcript structure, genomic footprinting, and direct nucleotide sequencing of genomic DNA and cDNA. The theoretical basis of PCR is outlined in Figure 15.0.1. There are three nucleic acid segments: the segment of double-stranded DNA to be amplified and two single-stranded oligonucleotide primers flanking it. Additionally, there is a protein component (a DNA polymerase), appropriate deoxyribonucleoside triphosphates (dNTPs), a buffer, and salts.

original DNA PCR primer new DNA

DNA + primers + dNTPs + DNA polymerase

denature and synthesize



etc.

etc.

etc.

Figure 15.0.1 The polymerase chain reaction. DNA to be amplified is denatured by heating the sample. In the presence of DNA polymerase and excess dNTPs, oligonucleotides that hybridize specifically to the target sequence can prime new DNA synthesis. The first cycle is characterized by a product of indeterminate length; however, the second cycle produces the discrete “short product” which accumulates exponentially with each successive round of amplification. This can lead to the many million-fold amplification of the discrete fragment over the course of 20 to 30 cycles.

The Polymerase Chain Reaction

Contributed by Donald M. Coen

15.0.1


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The primers are added in vast excess compared to the DNA to be amplified. They hybridize to opposite strands of the DNA and are oriented with their 3′ ends facing each other so that synthesis by DNA polymerase (which catalyzes growth of new strands 5′→3′) extends across the segment of DNA between them. One round of synthesis results in new strands of indeterminate length which, like the parental strands, can hybridize to the primers upon denaturation and annealing. These products accumulate only arithmetically with each subsequent cycle of denaturation, annealing to primers, and synthesis. However, the second cycle of denaturation, annealing, and synthesis produces two single-stranded products that together compose a discrete double-stranded product which is exactly the length between the primer ends. Each strand of this discrete product is complementary to one of the two primers and can therefore participate as a template in subsequent cycles. The amount of this product doubles with every subsequent cycle of synthesis, denaturation, and annealing, accumulating exponentially so that 30 cycles should result in a 228-fold (270 million–fold) amplification of the discrete product. This chapter consists of protocols that cover some of the more common applications of PCR. For many applications, the first step is simply to get PCR working with a known segment of DNA and a set of primers. Therefore, UNIT 15.1 presents a basic PCR protocol and ways to optimize it for the sequence of interest. PCR permits direct sequencing of nucleic acids without requiring cloning, thus avoiding cloning difficulties and artifacts. Several different protocols for preparing PCR products for sequencing using either dideoxy (Sanger) sequencing methods or chemical (MaxamGilbert) methods are presented in UNIT 15.2. This unit should permit the practitioner to choose a protocol best suited to the problem at hand and to his or her taste. Several PCR methods have been developed that require knowledge of only a small stretch of sequence (30-40 bases) and add sequence to the ends of amplified molecules to facilitate analyses. One of these, ligation-mediated PCR (UNIT 15.3) has broad applications including genomic footprinting and sequencing. PCR can be used to help clone and manipulate sequences. Various methods for generating suitable ends to facilitate the direct cloning of PCR products are detailed in UNIT 15.4. Other protocols for cloning and mutagenesis of DNA using PCR can be found in UNIT 3.7 and UNIT 8.5. An important application of PCR is to detect RNA transcripts, analyze their structure, and amplify their sequences to permit cloning and/or sequencing. UNIT 15.5 presents procedures that adapt PCR to RNA templates, via production of a cDNA copy of the RNA by reverse transcriptase (RT-PCR). Anchored PCR, which, like ligation-mediated PCR, requires little knowledge of sequence and makes use of the ends of nucleic acids, is applied in UNIT 15.6 to analysis of mRNAs. PCR is frequently used because it is the most sensitive assay for rare sequences. A protocol that not only detects rare DNAs but quantitates them as well is presented in UNIT 15.7. The downside of sensitivity is contamination by infinitesimal amounts of unwanted exogenous sequences. Procedures designed to avoid contamination with undesired DNA sequences are emphasized in this unit.

Introduction

In Supplement 56, we have rearranged the order of the units and have updated some of them. Further updates and new units, especially on quantitative approaches to PCR, are forthcoming. We have also moved one unit, formerly 15.8, which describes a powerful application of PCR called differential display to Chapter 25, “Discovery and analysis of differentially expressed genes in single cells and cell populations”, where it is now UNIT



25B.3. Chapter 25 includes other PCR-based protocols as well. Of course, as PCR has become an invaluable tool for nearly every branch of molecular biology, applications of PCR can be found in many other chapters of Current Protocols in Molecular Biology including Chapters 3, 7, 8, 12-14, 16, 21, 22, and 24.

Donald M. Coen Contributing Editor (Chapter 15) Harvard Medical School



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Enzymatic Amplification of DNA by PCR: Standard Procedures and Optimization This unit describes a method for amplifying DNA enzymatically by the polymerase chain reaction (PCR), including procedures to quickly determine conditions for successful amplification of the sequence and primer sets of interest, and to optimize for specificity, sensitivity, and yield. The first step of PCR simply entails mixing template DNA, two appropriate oligonucleotide primers, Taq or other thermostable DNA polymerases, deoxyribonucleoside triphosphates (dNTPs), and a buffer. Once assembled, the mixture is cycled many times (usually 30) through temperatures that permit denaturation, annealing, and synthesis to exponentially amplify a product of specific size and sequence. The PCR products are then displayed on an appropriate gel and examined for yield and specificity.

UNIT 15.1

BASIC PROTOCOL

Many important variables can influence the outcome of PCR. Careful titration of the MgCl2 concentration is critical. Additives that promote polymerase stability and processivity or increase hybridization stringency, and strategies that reduce nonspecific primertemplate interactions, especially prior to the critical first cycle, generally improve amplification efficiency. This protocol, using Taq DNA polymerase (see UNIT 3.5), is designed to optimize the reaction components and conditions in one or two stages. The first stage (steps 1 to 7) determines the optimal MgCl2 concentration and screens several enhancing additives. Most suppliers (of which there are many) of Taq and other thermostable DNA polymerases provide a unique optimized MgCl2-free buffer with MgCl2 in a separate vial for user titration. The second stage (steps 8 to 13) compares methods for preventing pre-PCR low-stringency primer extension, which can generate nonspecific products. This has come to be known as “hot start,” whether one omits an essential reaction component prior to the first denaturing-temperature step or adds a reversible inhibitor of polymerase. Hot-start methods can greatly improve specificity, sensitivity, and yield. Use of any one of the hot-start approaches is strongly recommended if primer-dimers or other nonspecific products are generated, or if relatively rare template DNA is contained in a complex mixture, such as viral nucleic acids in cell or tissue preparations. This protocol suggests some relatively inexpensive methods to achieve hot start, and lists several commercial hot-start options which may be more convenient, but of course more expensive. NOTE: Use only molecular biology–grade water (i.e., DNase, RNase, and nucleic acid free) in all steps and solutions. Materials 10× MgCl2-free PCR buffer (see recipe) 50 µM oligonucleotide primer 1: 50 pmol/µl in sterile H2O (store at −20°C) 50 µM oligonucleotide primer 2: 50 pmol/µl in sterile H2O (store at −20°C) Template DNA: 1 µg mammalian genomic DNA or 1.0 to 100.0 pg of plasmid DNA (UNIT 2.1-2.4) 25 mM 4dNTP mix (see recipe) 5 U/µl Taq DNA polymerase (native or recombinant) Enhancer agents (optional; see recipe) 15 mM (L), 30 mM (M), and 45 mM (H) MgCl2 Mineral oil TaqStart Antibody (Clontech) Ficoll 400 (optional): prepare as 10× stock; store indefinitely at room temperature Tartrazine dye (optional): prepare as 10× stock; store indefinitely at room temperature Contributed by Martha F. Kramer and Donald M. Coen Current Protocols in Molecular Biology (2001) 15.1.1-15.1.14 Copyright © 2001 by John Wiley & Sons, Inc.



Table 15.1.1

Master Mixes for Optimizing Reaction Components

Components

Final concentration

Master mixa (µl)

Per reaction I

II

III

IV

10× MgCl2-free PCR buffer Primer 1

1×

10 µl

40.0

40.0

40.0

40.0

0.5 µM

1 µl

4.0

4.0

4.0

4.0

Primer 2 Template DNAb

0.5 µM Undiluted

1 µl 1 vol

4.0 4 vol

4.0 4 vol

4.0 4 vol

4.0 4 vol

25 mM 4dNTP mixc Taq polymerase

0.2 mM 2.5 U

0.8 µl 0.5 µl

3.2 2.0

3.2 2.0

3.2 2.0

3.2 2.0

DMSOd (20×) Glycerold (10×)

5% 10%

5 µl 10 µl

— —

20.0 —

— 40.0

— —

PMPEd (100×) H 2O

1% —

1 µl To 90 µl

— To 360

— To 360

— To 360

4.0 To 360

aTotal volume = 360 µl (i.e., enough for n + 1 reactions). bTemplate DNA volume (“vol”) is generally 1 to 10 µl. cIf 2 mM 4dNTP mix is preferred, use 10 µl per reaction, or 40 µl for each master mix; adjust the volume of water

accordingly. dSubstitute with other enhancer agents (see recipe in Reagents and Solutions) as available.

0.5 ml thin-walled PCR tubes Automated thermal cycler Additional reagents and equipment for DNA preparation (UNIT 2.1-2.4, agarose gel electrophoresis (UNIT 2.5A), nondenaturing PAGE (UNIT 2.7), or sieving agarose gel electrophoresis (UNIT 2.8), restriction endonuclease digestion (UNIT 3.1), and Southern blotting and hybridization (UNITS 2.9 & 6.4) NOTE: Do not use DEPC to treat water, reagents, or glassware. NOTE: Reagents should be prepared in sterile, disposable labware, taken directly from its packaging, or in glassware that has been soaked in 10% bleach, thoroughly rinsed in tap water followed by distilled water, and if available, exposed to UV irradiation for ∼10 min. Multiple small volumes of each reagent should be stored in screw-cap tubes. This will then serve as the user’s own optimization “kit.” Thin-walled PCR tubes are recommended. Optimize reaction components 1. Prepare four reaction master mixes according to the recipes given in Table 15.1.1. Enhancing agents probably work by different mechanisms, such as protecting enzyme activity and decreasing nonspecific primer binding. However, their effects cannot be readily predicted—what improves amplification efficiency for one primer pair may decrease the amplification efficiency for another. Thus it is best to check a panel of enhancers during development of a new assay.

Enzymatic Amplification of DNA by PCR

2. Aliquot 90 µl master mix I into each of three 0.5-ml thin-walled PCR tubes labeled I-L, I-M, and I-H. Similarly, aliquot mixes II through IV into appropriately labeled tubes. Add 10 µl of 15 mM MgCl2 into one tube of each master mix (labeled L; 1.5 mM final). Similarly, aliquot 10 µl of 30 mM and 45 mM MgCl2 to separate tubes of each master mix (labeled M and H, respectively; 3.0 and 4.5 mM final concentrations respectively).



It is helpful to set the tubes up in a three-by-four array to simplify aliquotting. Each of the three Mg2+ concentrations is combined with each of the four master mixes.

3. Overlay the reaction mixture with 50 to 100 µl mineral oil (2 to 3 drops). To include hot start in the first step, overlay reaction mixes with oil before adding the MgCl2, heat the samples to 95°C in the thermal cycler or other heating block, and add the MgCl2 once the elevated temperature is reached. Once the MgCl2 has been added, do not allow the samples to cool below the optimum annealing temperature prior to performing PCR. Alternatives to mineral oil include silicone oil and paraffin beads. Additionally, certain cyclers feature heated lids that are designed to obviate the need for an oil overlay.

Choose cycling parameters 4. Using the following guidelines, program the automated thermal cycler according to the manufacturer’s instructions. 30 cycles: 30 sec 30 sec

94°C (denaturation) 55° (GC content ≤50%) or 60°C (GC content >50%) (annealing)

∼60 sec/kb product sequence 72°C

(extension)

Cycling parameters are dependent upon the sequence and length of the template DNA, the sequence and percent complementarity of the primers, and the ramp times of the thermal cycler used. Thoughtful primer design will reduce potential problems (see Commentary). Denaturation, annealing, and extension are each quite rapid at the optimal temperatures. The time it takes to achieve the desired temperature inside the reaction tube (i.e., the ramp time) is usually longer than either denaturation or primer annealing. Thus, ramp time is a crucial cycling parameter. Manufacturers of the various thermal cyclers on the market provide ramp time specifications for their instruments. Ramp times are lower with thinwalled reaction tubes. The optimal extension time also depends on the length of the target sequence. Allow ∼1 min/kb for this step for target sequences >1 kb, and as little as a 2-sec pause for targets 40) can reduce the polymerase specific activity, increase nonspecific amplification, and deplete substrate (nucleotides). Many investigators lengthen the time for the last extension step—to 7 min, for example—to try to ensure that all the PCR products are full length. These guidelines are appropriate for most commercially available thermal cyclers. For rapid cyclers, consult the manufacturers’ protocols.

Analyze the product 5. Electrophorese 10 µl from each reaction on an agarose (UNIT 2.5A), nondenaturing polyacrylamide (UNIT 2.7), or sieving agarose gel (UNIT 2.8) appropriate for the PCR product size expected. Stain with ethidium bromide. For resolution of PCR products between 100 and 1000 bp, an alternative to nondenaturing polyacrylamide gels or sieving agarose is a composite 3% (w/v) NuSieve (FMC Bioproducts) agarose/1% (w/v) SeaKem (FMC Bioproducts) agarose gel. SeaKem increases the mechanical strength of the gel without decreasing resolution. An alternative to ethidium bromide, SYBR Gold Nucleic Acid Gel Stain (Molecular Probes), is 25 to 100 times more sensitive than ethidium bromide, is more convenient to use, and permits optimization of 10- to 100-fold lower starting template copy number.



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Table 15.1.2

Master Mixes for Optimizing First-Cycle Reactions

Components 10× PCR buffer MgCl2 (L, M, or H) Primer 1 Primer 2 Additive Template DNA 25 mM 4dNTP mixb Taq polymerase Taq pol + TaqStart H 2O Preparation temperature

Final concentration 1× Optimal 0.5 µM 0.5 µM Optimal —b 0.2 mM 2.5 U 2.5 U To 100 µl

Master mix (µl) A 10 10 1.0 1.0 Va Va 0.8 0.5 — Va Room temperature

B

C

10 10 1.0 1.0 Va Va 0.8 0.5 — Va Ice slurry

10 10 1.0 1.0 Va Va 0.8 — — Va Room temperature

D 10 10 1.0 1.0 Va Va 0.8 — 1.0 Va Room temperature

aV, variable amount (total volume should be 100 µl). bUse undiluted or diluted template DNA based on results obtained in step 6.

6. Examine the stained gel to determine which condition resulted in the greatest amount of product. Minor, nonspecific products may be present even under optimal conditions.

7. To ensure that the major product is the correct one, digest an aliquot of the reaction with a restriction endonuclease known to cut within the PCR product. Check buffer compatibility for the restriction endonuclease of choice. If necessary, add Na+ or precipitate in ethanol (UNIT 2.1A) and resuspend in the appropriate buffer. Electrophorese the digestion product on a gel to verify that the resulting fragments have the expected sizes. Alternatively, transfer the PCR products to a nitrocellulose or nylon filter and hybridize with an oligonucleotide derived from the sequence internal to the primers (UNITS 2.9 & 6.4). With appropriately stringent hybridization and washing conditions, only the correct product (and possibly some minor related products) should hybridize.

Optimize the first cycle These optional steps optimize initial hybridization and may improve efficiency and yield. They are used when primer-dimers and other nonspecific products are detected, when there is only a very small amount of starting template, or when a rare sequence is to be amplified from a complex mixture. For an optimal reaction, polymerization during the initial denaturation and annealing steps should be prevented. Taq DNA polymerase activity can be inhibited by temperature (reaction B), physical separation (reaction C), or reversible antibody binding (reaction D). PCR without hot start is performed for comparison (reaction A). 8. Prepare four reaction mixtures using the optimal MgCl2 concentration and additive requirement determined in step 6. Prepare the mixes according to the recipes in Table 15.1.2. Use the following variations for addition of Taq polymerase. Enzymatic Amplification of DNA by PCR

a. Prepare reactions A and C at room temperature. b. Chill all components of reaction B in an ice slurry before they are combined.



c. For reaction D, combine 1.0 µl TaqStart antibody with 4.0 µl of the dilution buffer provided with the antibody, add 1.0 µl Taq DNA polymerase (for 1:4:1 mixture of these components), mix, and incubate 5 to 10 min at room temperature before adding to reaction mixture D (glycerol and PMPE are compatible with TaqStart antibody but DMSO will interfere with antibody binding). To ensure that the reaction does not plateau and thereby obfuscate the results, use the smallest amount of template DNA necessary for visualization of the PCR product by ethidium bromide staining. Use the results from step 6 to decide how much template to use. If the desired product stains intensely, dilute the starting material as much as 1/100. If only a faint signal is apparent, use undiluted sample.

9. Overlay each reaction mixture with 50 to 100 µl mineral oil. 10. Heat all reactions 5 min at 94°C. It is most convenient to use the automated thermal cycler for this step and then initiate the cycling program directly.

11. Cool the reactions to the appropriate annealing temperature as determined in step 4. Add 0.5 µl Taq DNA polymerase to reaction C, making sure the pipet tip is inserted through the layer of mineral oil into the reaction mix. Time is also an important factor in this step. If the temperature drops below the annealing temperature and is allowed to remain low, nonspecific annealing will occur. Taq DNA polymerase retains some activity even at room temperature.

12. Begin amplification of all four reactions at once, using the same cycling parameters as before. 13. Analyze the PCR products on an agarose gel and evaluate the results as in steps 5 and 6. 14. Prepare a batch of the optimized reaction mixture, but omit Taq DNA polymerase, TaqStart antibody, PMPE, and 4dNTP mix—these ingredients should be added fresh just prior to use. If desired, add Ficoll 400 to a final concentration of 0.5% to 1% (v/v) and tartrazine to a final concentration of 1 mM. Adding Ficoll 400 and tartrazine dye to the reaction mix precludes the need for a gel loading buffer and permits direct application of PCR products to agarose or acrylamide gels. At these concentrations, Ficoll 400 and tartrazine do not decrease PCR efficiency and do not interfere with PMPE or TaqStart antibodies. Other dyes, such as bromphenol blue and xylene cyanol, do inhibit PCR. Tartrazine is a yellow dye and is not as easily visualized as other dyes; this may make gel loading more difficult. Ficoll 400 and tartrazine dye may be prepared as 10× stocks and stored indefinitely at room temperature.


Enhancer agents For a discussion of how to select enhancer agents, see Commentary. 5× stocks: 25% acetamide (20 µl/reaction; 5% final) 5 M N,N,N-trimethylglycine (betaine; 20 µl/reaction; 1 M final) 40% polyethylene glycol (PEG) 8000 (20 µl/reaction; 8% final) The Polymerase Chain Reaction


Supplement 56

10× stocks: Glycerol (concentrated; 10 µl/reaction; 10% final) 20× stocks: Dimethylsulfoxide (DMSO; concentrated; 5 µl/reaction; 5% final) Formamide (concentrated; 5 µl/reaction; 5% final) 100× stocks: 1 U/µl Perfect Match Polymerase Enhancer (Strategene; 1 µl/reaction; 1 U final) 10 mg/ml acetylated bovine serum albumin (BSA) or gelatin (1 µl/reaction; 10 µg/ml final) 1 to 5 U/µl thermostable pyrophosphatase (PPase; Roche Diagnostics; ; 1 µl/reaction; 1 to 5 U final) 5 M tetramethylammonium chloride (TMAC; betaine hydrochloride; 1 µl/reaction; 50 mM final) 0.5 mg/ml E. coli single-stranded DNA-binding protein (SSB; Sigma; 1 µl/reaction; 5 µg/ml final) 0.5 mg/ml Gene 32 protein (Amersham Pharmacia Biotech; 1 µl/reaction; 5 µg/ml final) 10% Tween 20, Triton X-100, or Nonidet P-40 (1 µl/reaction; 0.1% final) 1 M (NH4)2SO4 (1 µl/reaction; 10 mM final; use with thermostable DNA polymerases other than Taq) MgCl2-free PCR buffer, 10× 500 mM KCl 100 mM Tris⋅Cl, pH 9.0 (at 25°C; see APPENDIX 2) 0.1% Triton X-100 Store indefinitely at −20°C This buffer can be obtained from Promega; it is supplied with Taq DNA polymerase.

4dNTP mix For 2 mM 4dNTP mix: Prepare 2 mM each dNTP in TE buffer, pH 7.5 (APPENDIX 2). Store up to 1 year at −20°C in 1-ml aliquots. For 25 mM 4dNTP mix: Combine equal volumes of 100 mM dNTPs (Promega). Store indefinitely at −20°C in 1-ml aliquots. COMMENTARY Background Information


The theoretical basis of the polymerase chain reaction (PCR; see chapter introduction) was probably first described in a paper by Kleppe et al. (1971). However, this technique did not excite general interest until the mid-1980s, when Kary Mullis and co-workers at Cetus developed PCR into a technique that could be used to generate large amounts of single-copy genes from genomic DNA (Saiki et al., 1985, 1986; Mullis et al., 1986; Embury et al., 1987). The initial procedure entailed adding a fresh aliquot of the Klenow fragment of E. coli DNA polymerase I during each cycle because this enzyme was inactivated during the subsequent denaturation step. The introduction of thermostable Taq DNA polymerase from Thermus

aquaticus (Saiki et al., 1988) alleviated this tedium and facilitated automation of the thermal cycling portion of the procedure. Taq DNA polymerase also permitted the use of higher temperatures for annealing and extension, which improved the stringency of primer–template hybridization and thus the specificity of the products. This also served to increase the yield of the desired product. All applications of PCR depend upon an optimized PCR. The basic protocol in this unit optimizes PCR for several variables, including MgCl2 concentration, enhancing additives— dimethyl sulfoxide (DMSO), glycerol, or Perfect Match Polymerase Enchancer (PMPE)— and prevention of pre-PCR mispriming. These and other parameters can be extremely impor-



tant, as every element of PCR can affect the outcome (see Critical Parameters and Troubleshooting for discussion of individual parameters). There are several PCR optimization kits and proprietary enhancers on the market (Table 15.1.3). Optimization kits generally provide a panel of buffers in which the pH, buffer, nonionic detergents, and addition of (NH4)2SO4 are varied, MgCl2 may be added at several concentrations, and enhancers (e.g., DMSO, glycerol, formamide, betaine, and/or proprietary compounds) may be chosen. The protocol presented here is aimed at keeping the costs low and the options broad.

Critical Parameters and Troubleshooting MgCl2 concentration Determining the optimum MgCl2 concentration, which can vary even for different primers from the same region of a given template (Saiki, 1989), can have an enormous influence on PCR success. In this protocol three concentrations are tested—1.5 mM (L), 3.0 mM (M), and 4.5 mM (H)—against three enhancers. Enhancers tend to broaden the MgCl2 optimal range, contributing to the success of the PCR at one of these concentrations. A 10× buffer optimized for a given enzyme and a separate vial of MgCl2 are typically provided with the polymerase, so that the user may titrate the MgCl2 concentration for their unique primertemplate set. Reagent purity For applications that amplify rare templates, reagent purity is the most important parameter, and avoiding contamination at every step is critical. To maintain purity, store multiple small volumes of each reagent in screw-cap tubes. For many applications, simply using highquality reagents and avoiding nuclease contamination is sufficient; however, avoid one common reagent used to inactivate nucleases, diethylpyrocarbonate (DEPC). Even tiny amounts of chemical left after treatment of water by autoclaving are enough to ruin a PCR. Primer selection This is the factor that is least predictable and most difficult to troubleshoot. Simply put, some primers just do not work. To maximize the probability that a given primer pair will

work, pay attention to the following parameters. General considerations. An optimal primer set should hybridize efficiently to the sequence of interest with negligible hybridization to other sequences present in the sample. If there are reasonable amounts of template available, hybridization specificity can be tested by performing oligonucleotide hybridization as described in UNIT 6.4. The distance between the primers is rather flexible, ranging up to 10 kb. There can be, however, a considerable drop-off in synthesis efficiency with distances >3 kb (Jeffreys et al., 1988). Small distances between primers, however, lessen the ability to obtain much sequence information or to reamplify with nested internal oligonucleotides, should that be necessary. Design primers to allow demonstration of the specificity of the PCR product. Be sure that there are diagnostic restriction endonuclease sites between the primers or that an oligonucleotide can detect the PCR product specifically by hybridization. Several computer programs can assist in primer design (see Internet Resources at end of unit). These are most useful for avoiding primer sets with intra- and intermolecular complementarity, which can dramatically raise the effective Tm. Given the abundance of primers relative to template, this can preclude template priming. Computer primer design is not foolproof. If possible, start with a primer or primer set known to efficiently prime extensions. In addition, manufacturers’ Web sites offer technical help with primer design. Complementarity to template. For many applications, primers are designed to be exactly complementary to the template. For others, however, such as engineering of mutations or new restriction endonuclease sites, or for efforts to clone or detect gene homologs where sequence information is lacking, base-pair mismatches will be intentionally or unavoidably created. It is best to have mismatches (e.g., in a restriction endonuclease linker) at the 5′ end of the primer. The closer a mismatch is to the 3′ end of the primer, the more likely it is to prevent extension. The use of degenerate oligonucleotide primers to clone genes where only protein sequence is available, or to fish out gene homologs in other species, has sometimes been successful, but it has also failed an untold (and unpublished) number of times. When the reaction works it can be extremely valuable, but it can



Supplement 56

Table 15.1.3

PCR Optimization Products

Optimization goal

Supplier

Product

Optimization support Optimization support

Perkin-Elmer Promega

Optimization kits

Boehringer-Mannheim, Invitrogen, Stratagene, Sigma, Epicentre Technologies, Life Technologies Amersham Pharmacia Biotech

Technical information in appendix to catalog PCR troubleshooting program on the Internet: http://www.promega.com/amplification/assistant Several buffers, Mg2+, and enhancers which may include DMSO, glycerol, formamide, (NH4)2SO4, and other unspecified or proprietary agents

Quick startup

Quick startup

Fisher

Quick startup

Life Technologies

Quick startup

Marsh Biomedical

Hot-start/physical barrier Fisher, Life Technologies

Hot-start/separate MgCl2 Invitrogen Hot-start/separate MgCl2 Stratagene Hot Start/separate polymerase

Promega

Hot-start/reversible Clontech inactivation of polymerase by antibody binding Hot-start/antibody Life Technologies binding Hot-start/antibody Sigma binding Hot-start/reversible Perkin-Elmer chemical modification Hot-start/reversible Qiagen chemical modification Enhancer Boehringer Mannheim, New England Biolabs Enhancer Clontech Enhancer CPG Enhancer

Fisher

Enhancer Enhancer Enhancer Enhancer Enhancer

Life Technologies Promega Qiagen Stratagene Stratagene

Ready-To-Go Beads “optimized for standard PCR” and Ready-To-Go RAPD Analysis Beads (buffer, nucleotides, Taq DNA polymerase) EasyStart PCR Mix-in-a-Tube—tubes prepackaged with wax beads containing buffer, MgCl2, nucleotides, Taq DNA polymerase PCR SuperMix—1.1× conc.—premix containing buffer, MgCl2, nucleotides, Taq DNA polymerase Advanced Biochemicals Red Hot DNA Polymerase—a new rival for Taq polymerase with convenience features Molecular Bio-Products HotStart Storage and Reaction Tubes—preadhered wax bead in each tube; requires manual addition of one component at high temperature HotWax Mg2+ beads—wax beads contain preformulated MgCl2 which is released at first elevated-temperature step StrataSphere Magnesium Wax Beads—wax beads containing preformulated Mg2+ TaqBead Hot Start Polymerase—wax beads encapsulating Taq DNA polymerase which is released at first elevated-temperature step TaqStart Antibody, TthStart Antibody—reversibly inactivate Taq and Tth DNA polymerases until first denaturation at 95°C

PlatinumTaq—contains PlatinumTaq antibody JumpStart Taq—contains TaqStart antibody AmpliTaq Gold—activated at high temperature HotStarTaq DNA Polymerase—activated at high temperature Tth pyrophosphatase, thermostable GC-Melt (in Advantage-GC Kits)—proprietary Taq-FORCE Amplification System and MIGHTY Buffer—proprietary Eppendorf MasterTaq Kit with TaqMaster Enhancer—proprietary PCRx Enhancer System—proprietary E.coli Single Stranded Binding Protein (SSB) Q-Solution—proprietary Perfect Match Polymerase Enhancer—proprietary TaqExtender PCR Additive—proprietary



also generate seemingly specific products that require much labor to identify and yield no useful information. The less degenerate the oligonucleotides, especially at the 3′ end, the better. Caveat emptor. Primer length. A primer should be 20 to 30 bases in length. It is unlikely that longer primers will help increase specificity significantly. Primer sequence. Design primers with a GC content similar to that of the template. Avoid primers with unusual sequence distributions, such as stretches of polypurines or polypyrimidines, as their secondary structure can be disastrous. It is worthwhile to check for potential secondary structure using one of the appropriate computer programs that are available. “Primer-dimers.” Primer-dimers are a common artifact most frequently observed when small amounts of template are taken through many amplification cycles. They form when the 3′ end of one primer anneals to the 3′ end of the other primer, and polymerase then extends each primer to the end of the other. The ensuing product can compete very effectively against the PCR product of interest. Primer-dimers can best be avoided by using primers without complementarity, especially in their 3′ ends. Should they occur, optimizing the MgCl2 concentration may minimize their abundance relative to that of the product of interest. Template Aside from standard methods for preparing DNA (UNIT 2.1-2.4), a number of simple and rapid procedures have been developed for particular tissues (Higuchi, 1989). Even relatively degraded DNA preparations can serve as useful templates for generation of moderate-sized PCR products. The two main concerns regarding template are purity and amount. A number of contaminants found in DNA preparations can decrease the efficiency of PCR. These include urea, the detergent SDS (whose inhibitory action can be reversed by nonionic detergents), sodium acetate, and, sometimes, components carried over in purifying DNA from agarose gels (Gelfand, 1989; Gyllensten, 1989; K. Hicks and D. Coen, unpub. observ.). Additional organic extractions, ethanol precipitation from 2.5 M ammonium acetate, and/or gel purification on polyacrylamide rather than agarose, can all be beneficial in minimizing such contamination if the simplest method (precipitating the sample with ethanol and repeatedly washing the pellet with 70% ethanol) is not sufficient.

Clearly the amount of template must be sufficient to be able to visualize PCR products using ethidium bromide. Usually 100 ng of genomic DNA is sufficient to detect a PCR product from a single-copy mammalian gene. Using too much template is not advisable when optimizing for MgCl2 or other parameters, as it may obscure differences in amplification efficiency. Moreover, too much template may decrease efficiency due to contaminants in the DNA preparation. Amount of template, especially in terms of the amount of target sequence versus nonspecific sequences, can have a major effect on the yield of nonspecific products. With less target sequence, it is more likely that nonspecific products will be seen. For some applications, such as certain DNA sequencing protocols where it is important to have a single product, gel purification of the specific PCR product and reamplification are advisable. Taq and other thermostable DNA polymerases Among the advantages conferred by the thermostability of Taq DNA polymerase is its ability to withstand the repeated heating and cooling inherent in PCR and to synthesize DNA at high temperatures that melt out mismatched primers and regions of local secondary structure. The enzyme, however, is not infinitely resistant to heat, and for greatest efficiency it should not be put through unnecessary denaturation steps. Indeed, some protocols (e.g., UNIT 15.7 and the “hot start” method described here) recommend adding it after the first denaturation step. Increasing the amount of Taq DNA polymerase beyond 2.5 U/reaction can sometimes increase PCR efficiency, but only up to a point. Adding more enzyme can sometimes increase the yield of nonspecific PCR products at the expense of the product of interest. Moreover, Taq DNA polymerase is not inexpensive. A very important property of Taq DNA polymerase is its error rate, which was initially estimated at 2 × 10−4 nucleotides/cycle (Saiki et al., 1988). The purified enzyme supplied by manufacturers lacks a proofreading 3′→5′ exonuclease activity, which lowers error rates of other polymerases such as the Klenow fragment of E. coli DNA polymerase I. For many applications, this does not present any difficulties. However, for sequencing clones derived from PCR, or when starting with very few templates, this can lead to major problems. Direct sequencing of PCR products (UNIT 15.2),



Supplement 56

sequencing numerous PCR-generated clones, and/or the use of appropriate negative controls can help overcome these problems. Alternatively, changing reaction conditions (Eckert and Kunkel, 1990) or changing to a non–Taq DNA polymerase (with greater fidelity) may be useful. Another important property of Taq DNA polymerase is its propensity for adding nontemplated nucleotides to the 3′ ends of DNA chains. This can be especially problematic in cloning PCR products. It is frequently necessary to “polish” PCR products with enzymes such as other DNA polymerases before adding linkers or proceeding to blunt-end cloning. Conversely, addition of a nontemplated A by Taq DNA polymerase can be advantageous in cloning (UNIT 15.4). Certain PCR protocols may work better with one thermostable polymerase rather than another. Table 15.1.4 lists currently available thermostable DNA polymerases by generic and trade names, the original source of native and recombinant enzymes, the supplier, the end generated (3′A addition versus blunt), and associated exonuclease activities. A 3′ to 5′ exonuclease activity is proofreading. Removal of the 5′ to 3′ exonuclease activity of Taq DNA polymerase (N-terminal deletion) is reported to produce a higher yield. A 5′ to 3′ exonuclease activity may degrade the primers somewhat. Proofreading enzymes synthesize DNA with higher fidelity and can generate longer products than Taq, but tend to generate low yields. Enzyme blends (Table 15.1.5) have been optimized for increased fidelity and length along with sensitivity and yield.


Hot start What happens prior to thermal cycling is critical to the success of PCR. Taq DNA polymerase retains some activity even at room temperature. Therefore, under nonstringent annealing conditions, such as at room temperature, products can be generated from annealing of primers to target DNA at locations of low complementarity or having complementarity of just a few nucleotides at the 3′ ends. The latter would in effect create new templates “tagged” with the primer sequences. Subsequent cycles amplify these tagged sequences in abundance, both generating nonspecific products and possibly reducing amplification efficiency of specific products by competition for substrates or polymerase. Thus conditions preventing polymerization prior to the first temperature-controlled steps are desirable. In this protocol, three

methods of inhibiting polymerization prior to the temperature-controlled step are compared. These include physical separation of an essential reaction component prior to the first denaturation step, cooling reagents to 0°C, and reversibly blocking enzymatic activity with an antibody. Denaturation of the template before Taq polymerase or MgCl2 is added to the reaction provides a dramatic improvement in specificity and sensitivity in many cases (Chou et al., 1992). The main drawback of this method is that it requires opening the reaction tubes a second time to add the essential missing component. This creates both an inconvenience and an increase in the risk of contamination, an important consideration when testing for the presence of a given sequence in experimental or clinical samples. Cooling all components of the reaction mixture to 0°C prior to mixing is more convenient and the least expensive method but is also the least reliable. Transferring the PCR reaction tubes from the ice slurry to a 95°C preheated thermocycler block may improve the chance of success. Reversible inhibition of Taq DNA polymerase by TaqStart antibody (Clontech) is the most convenient method and very effective (Kellogg et al., 1994). Complete reactions can be set up, overlaid with oil, and stored at 4°C for up to several hours prior to thermal cycling with no loss of sensitivity or specificity compared to the other hot start methods (M.F. Kramer and D.M. Coen, unpub. observ.). Cycling is initiated immediately following 5-min denaturation of the antibody at 94°C. DMSO inhibits antibody binding and should not be used with TaqStart. Several hot-start products are now commercially available (Table 15.1.3). Success with each may depend on strict adherence to the manufacturer’s protocols, even on a specific thermocycler. Wax barrier and reversible antibody binding methods are more forgiving, while chemical modifications have more stringent activation temperature requirements. Deoxyribonucleoside triphosphates In an effort to increase efficiency of PCR, it may be tempting to increase the concentration of dNTPs. Don’t! When each dNTP is 200 mM, there is enough to synthesize 12.5 mg of DNA when half the dNTPs are incorporated. dNTPs chelate magnesium and thereby change the effective optimal magnesium concentration. Moreover, dNTP concentrations >200



Table 15.1.4

Thermostable DNA Polymerases

DNA polymerase Generic name

Trade name

Pfu Pfu (exo-) Psp Psp (exo-)

— — Deep Vent Deep Vent (exo-)

Pwo Taq (native and/or recombinant)

—

Biological source

Supplier

Product ends

Exonuclease activity

Pyrococcus furiosus Pyrococcus furiosus Pyrococcus sp.GB-D Pyrococcus sp.GB-D

Stratagene, Promega Stratagene New England Biolabs New England Biolabs

Blunt Blunt Blunt Blunt

3′-5′ (proofreading) No 3′-5′ (proofreading) No

Pyrococcus woesei Thermus aquaticus

Boehringer Mannheim Blunt Ambion, Amersham 3′A Pharmacia Biotech, Boehringer Mannheim, Clontech, Fisher, Life Technologies, Marsh Biomedical, Perkin Elmer, Promega, Qiagen, Sigma, Stratagene Perkin-Elmer, Sigma 3′A

3′-5′ (proofreading) 5′-3′

Taq, N-terminal Stoffel deletion fragment Klen-Taq Tbr DyNAzyme Tfl

Thermus aquaticus

Tli

Vent

Tli (exo-)

Vent (exo-)

Tma Tth

UlTma —

Thermococcus litoralis Thermococcus litoralis Thermotoga maritima Perkin-Elmer Thermus thermophilus Amersham Pharmacia Biotech, Boehringer Mannheim, Epicentre Technologies, Perkin Elmer, Promega

Thermus brocianus Thermus flavus

MJ Research Promega, Epicentre Technologies New England Biolabs (Vent), Promega New England Biolabs

No

—a Blunt

5′-3′ —a

Blunt

3′-5′ (proofreading)

Blunt

No

Blunt 3′ A

3′-5′ (proofreading) 5′-3′

aNo information at this time.

mM each increase the error rate of the polymerase. Millimolar concentrations of dNTPs actually inhibit Taq DNA polymerase (Gelfand, 1989). The protocol in this unit calls for preparing 4dNTPs in 10 mM Tris⋅Cl/1 mM EDTA (TE buffer), pH 7.4 to 7.5. This is easier and less prone to disaster than neutralization with sodium hydroxide. However, EDTA also chelates magnesium, and this should be taken into account if stocks of dNTPs are changed. Alternatively, to lower the risk of contamination, a 4dNTP mix can be made by combining equal volumes of commercially prepared stocks.

Enhancers Enhancers are used to increase yield and specificity and to overcome difficulties encountered with high GC content or long templates. Nonionic detergents (Triton X-100, Tween 20, or Nonidet P-40) neutralize charges of ionic detergents often used in template preparation, and should be used in the basic reaction mixture, rather than as optional enhancers. Higher yields can be achieved by stabilizing/enhancing the polymerase activity with enzyme-stabilizing proteins (BSA or gelatin), enzyme-stabilizing solutes such as betaine or betaine⋅HCl (TMAC), enzyme-stabilizing solvents (glycerol), solubility-enhancing sol-



Supplement 56

Table 15.1.5

Thermostable DNA Polymerase Blends

Product (trade name)

Supplier

Thermostable DNA polymerases and other components

Expand High Fidelity, Expand Long Template, Boehringer Mannheim Taq + Pwo and Expand 20kb PCR Systems KlenTaq LA Polymerase Mix

Clontech, Sigma

Advantage-HF PCR Kit

Clontech

Advantage-cDNA and Advantage-GC cDNA Polymerase Mixes and Kits

Clontech

Advantage Genomic and Advantage-GC Genomic Polymerase Mixes and Kits

Clontech

Tth + unspecified proofreading polymerase + TthStart Antibody; GC Kit contains GC Melt

eLONGase Enzyme Mix

Life Technologies

Platinum Taq DNA Polymerase

Life Technologies

Taq + Psp + unspecified proofreading polymerase(s) + eLONGase Buffer Taq + Psp + Platinum Taq Antibody

Platinum High Fidelity DNA Polymerase DyNAzyme EXT Polymerase

Life Technologies MJ Research

Taq + Psp + Taq Antibody Tbr with unspecified enhancer

GeneAmp XL PCR and XL RNA PCR Kits OmniBase Sequencing Enzyme Mix

Perkin-Elmer Promega

Tth + Tli Unspecified proofreading polymerase(s) with thermostable pyrophosphatase

AccuTaq LA DNA Polymerase Mix Sigma TaqPlus Long and TaqPlus Precision PCR Stratagene Systems Accurase Fidelity PCR Enzyme Mix; Calypso Tetralink High Fidelity Single Tube RT-PCR System


vents (DMSO or acetamide), molecular crowding solvents (PEG), and polymerase salt preferences [(NH4)SO4 is recommended for polymerases other than Taq]. Greater specificity can be achieved by lowering the TM of dsDNA (using formamide), destabilizing mismatchedprimer annealing (using PMPE or hot-start strategies), and stabilizing ssDNA (using E. coli SSB or T4 Gene 32 Protein). Amplification of high-GC-content templates can be improved by decreasing the base pair composition dependence of the TM of dsDNA (with betaine; Rees et al., 1993). Betaine is an osmolyte widely distributed in plants and animals and is nontoxic, a feature that recommends it for convenience in handling, storage, and disposal. Betaine may be the proprietary ingredient in various commercial formulations. For long templates, a higher pH is recommended (pH 9.0). The pH of Tris buffer decreases at high temperatures, long-template PCR requires

KlenTaq-1 (5′-exonuclease deficient Taq) + unspecified proofreading polymerase KlenTaq-1 + unspecified proofreading polymerase + TaqStart Antibody KlenTaq-1 + unspecified proofreading polymerase + TaqStart Antibody; GC Kit contains GC Melt

Taq + unspecified proofreading polymerase Pfu + Taq; TaqPlus Precision Reaction Buffer (proprietary) Thermus sp. + Thermococcus sp.; Calypso also contains AMV-RT

more time at high temperatures, and increased time at lower pH may cause some depurination of the template, resulting in reduced yield of specific product. Inorganic phosphate (PPi), a product of DNA synthesis, may accumulate with amplification of long products to levels that may favor reversal of polymerization. Accumulation of PPi may be prevented by addition of thermostable PPase. When large numbers of samples are being analyzed, the convenience of adding PCR products directly to a gel represents a significant time savings. Some companies combine their thermostable polymerase with a red dye and a high density component to facilitate loading of reaction products onto gels without further addition of loading buffer. Thermal cycling parameters Each step in the cycle requires a minimal amount of time to be effective, while too much



time can be both wasteful and deleterious to the DNA polymerase. If the amount of time in each step can be reduced, so much the better. Denaturation. It is critical that complete strand separation occur during the denaturation step. This is a unimolecular reaction which, in itself, is very fast. The suggested 30-sec denaturation used in the protocol ensures that the tube contents reach 94°C. If PCR is not working, it is well worth checking the temperature inside a control tube containing 100 µl water. If GC content is extremely high, higher denaturation temperatures may be necessary; however, Taq DNA polymerase activity falls off quickly at higher temperatures (Gelfand, 1989). To amplify a long sequence (>3 kb), minimize the denaturation time to protect the target DNA from possible effects, such as depurination, of lowered pH of the Tris buffer at elevated temperatures. Annealing. It is critical that the primers anneal stably to the template. Primers with relatively low GC content (48 hr) are acceptable.

2. Prepare a microcentrifuge tube for PCR containing the following: 10 µl 10× amplification buffer (optimized as in UNIT 15.1) 10 µl 2 mM 4dNTP mix 20 to 40 pmol oligonucleotide primer 2 (not labeled in step 1) DNA template 1.5 to 2.5 U Taq DNA polymerase 10 µl kinase reaction (step 1) H2O to 100 µl. Overlay with 100 µl mineral oil. Carry out PCR under optimized conditions for 5 to 10 cycles (see UNIT 15.1, steps 6 to 8). Complete PCR with a final long extension step of 5 to 7 min. The final extension ensures that all PCR products are complete and blunt-ended. If the DNA template is a single band from a previous PCR reaction that has been cut out and extracted from a gel, >10 cycles of PCR may be necessary. Similarly, if only a small amount of initial template is present, 20 to 25 cycles of PCR may be necessary to generate sufficient labeled product.

3. Remove the mineral oil with a Pasteur pipet. 4. Separate the labeled PCR product from the unincorporated labeled primer and the unincorporated labeled nucleotides by electrophoresis on an agarose or nondenaturing polyacrylamide gel. Using the characteristic mobilities of the loading dyes (bromphenol blue and/or xylene cyanol) as guideposts (UNITS 2.5A & 2.7), run the gel until the unincorporated primers and nucleotides migrate off the gel and into the lower buffer chamber. Visualize the labeled PCR products by autoradiography for 10 to 60 min. Be conservative in running the gel. Avoid running the PCR products off the gel.

5. Cut and isolate the desired labeled PCR product from the gel, using one of the described protocols for fragment purification (UNITS 2.5A, 2.6, or 2.7).

Direct DNA Sequencing of PCR Products

A preparative gel confirms the presence of the desired amplified product. However, amplified DNA can be also be purified using disposable columns, which yield clean double-stranded DNA. Chemical sequencing can also be carried out without gel or column purification. In such cases, the PCR sample should be phenol extracted and ethanol precipitated in the presence of 0.3 M sodium acetate and 2.5 vol cold 100% ethanol (UNIT 2.1A). The resulting pellet should be washed with 70% ethanol.



The columns and extraction procedures described above should not be used instead of a preparative gel unless it is known that only a single, homogeneous product has been amplified during PCR.

6. Resuspend the labeled PCR product in 30 to 50 µl water. 7. Carry out chemical sequencing using 8% to 10% of the product for each sequencing reaction (UNIT 7.5). 8. Resolve sequencing products on a sequencing gel (UNIT 7.6). Wrap gel in plastic wrap and autoradiograph at −20°C for 2 to 48 hr. Read sequence as described in UNIT 7.5. The gel may be dried, but it is generally not necessary to do so.

GENOMIC SEQUENCING OF PCR PRODUCTS This method combines the PCR amplification method with the genomic sequencing technique (Church and Gilbert, 1984). Following PCR, the amplified DNA is chemically sequenced, transferred by electroblotting, and covalently bound by UV crosslinking onto a nylon filter that can be repeatedly probed with short, sequence-specific oligonucleotides. This method is ideally suited to situations where large amounts of sequence are sought, or where more than one region is being amplified. Several amplified fragments can be mixed, simultaneously sequenced, run out on a single set of lanes, and the sequence of the different fragments successively visualized by the use of appropriate probes.


Materials Filter paper (Schleicher & Schuell #410), precut to gel size Additional reagents and equipment for phenol extraction and ethanol precipitation (UNIT 2.1A); transfer by electroblotting, UV cross-linking, and hybridization (UNITS 2.9, 6.3, & 6.4); and labeling with terminal deoxynucleotidyltransferase (UNIT 3.6) 1. Carry out PCR under optimized conditions, ending with a final long extension step (UNIT 15.1). The template concentrations that yield optimum amplifications must be empirically determined. These concentrations range from 20 to 1000 ng for eukaryotic genomic DNA, 10 to 100 ng for bacterial DNA, and 1 to 20 ng for cloned DNA inserts.

2. After the final long extension step, remove the mineral oil with a Pasteur pipet. 3. Phenol extract the amplified product (UNIT 2.1A). Precipitate with 0.1 vol of 3 M sodium acetate, pH 5.2, and 2.5 vol of 100% ethanol for 20 min in a dry ice/ethanol bath. Microcentrifuge 5 min at top speed, 4°C. Wash the pellet twice with 70% ethanol and dry under vacuum (UNIT 2.1A). 4. Resuspend the pellet in 30 to 50 µl water. Electrophorese 5% to 10% of the PCR sample on an agarose or nondenaturing polyacrylamide gel to determine the yield and purity of the amplified product. Because every double-stranded product generated by PCR will give rise to a sequence ladder when chemically sequenced, the homogeneity of the amplified product is critical. If amplification yields at least 2 bands of similar intensity, a gel separation prior to sequencing is necessary, unless the different products can be separately visualized by using different oligonucleotide probes. The use of internal probes (3′ of the PCR primers) to visualize the sequence will prevent spurious amplification product sequences from appearing, but will not allow sequencing of the entire amplified fragment. The presence of short products (“primer-dimers”) will generally not interfere with the majority of the sequence, but will obscure the bottom section of the sequencing gel.



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5. Carry out chemical sequencing using 8% to 10% of the amplified product for each sequencing reaction (UNIT 7.5). 6. Electrophorese the sequenced samples on a denaturing polyacrylamide sequencing gel (UNIT 7.6). The number of resolvable base pairs can be increased by using a gradient or wedge gel, performing multiple loadings of the sequenced sample, and/or using longer gel plates (≤1 meter; UNIT 7.6).

7. Carefully remove one of the glass gel plates. Place the sheet of filter paper carefully on the gel, making sure not to trap air bubbles between the filter paper and gel. Lift the gel (adhered to the filter paper) onto the appropriate transfer apparatus. 8. Wet the gel with a thin layer of TBE electrophoresis buffer and place the nylon filter over the gel, again being careful not to trap air bubbles between the gel and membrane. Transfer the DNA onto the filter membrane by electroblotting (UNIT 2.9). It is necessary to use a large electroblotting apparatus for this purpose.

9. UV cross-link the DNA onto the filter (UNIT 2.9). 10. Hybridize the filter with the appropriate probe. The filter can now be probed, visualized, stripped, and reprobed as many as 40 times. Best results are obtained by probing with the PCR oligonucleotide primers, ensuring that only completed PCR products are visualized. Degenerate probes or probes internal to the amplified fragment can also be used. A high-specific-activity probe can be obtained by “tailing” 3 to 6 pmol of probe using terminal deoxynucleotidyltransferase.

REAGENTS AND SOLUTIONS Annealing buffer, 5× 200 mM Tris⋅Cl, pH 7.5 100 mM MgCl2 250 mM NaCl Store at −20°C PEG/NaCl solution 20%(w/v) polyethylene glycol (PEG) 6000 2.5 M NaCl Store at room temperature COMMENTARY Background Information


The methods presented in this unit permit rapid sequencing of PCR-amplified DNA. Although the cloning of amplified DNA is relatively straightforward, direct DNA sequencing of PCR products facilitates and speeds the acquisition of sequence information. As long as the PCR reaction produces a single amplified product (or a set of discrete products that can be conveniently separated), such products are amenable to direct sequencing. In contrast to methods where the PCR product is cloned and a single clone sequenced, the

direct sequencing of “bulk” PCR product is generally unaffected by the comparatively high error rate of Taq DNA polymerase. Unless a certain feature of the template causes consistent misincorporation at the same point in the sequence, errors are likely to be stochastically distributed throughout the molecule—thus, the overwhelming majority of the amplified product consists of the correct sequence. The only exceptions to this rule may be those cases where only a very small number of template molecules (1000 bp). In such cases, the sequences of interest can be amplified simultaneously in a single PCR reaction, or separately, as a set of discrete adjacent or overlapping fragments. The fragments can then be mixed, simultaneously sequenced, electrophoresed on gels, and transferred onto nylon filters. Each fragment can be successively visualized by probing with the appropriate labeled oligonucleotide. In addition, sequences from both strands can be derived from a single filter, with a consequent increase in sequence accuracy. This “multiplex” approach, however, requires the additional blotting and hybridization steps. The direct chemical sequencing of a labeled strand is a fast alternative if only a single product (and a single strand) is being sequenced. The utility of this direct approach is thus limited to those cases when the PCR yields a single DNA species or when the product of interest can be readily gel purified. Several excellent papers dealing with direct DNA sequencing of PCR products have been published. Detailed discussions of dideoxy se-

quencing approaches can be found in Wrischnik et al. (1987), Gyllensten and Erlich (1988), Innis et al. (1988), Kreitman and Landweber (1989), and Gyllensten (1989). Chemical sequencing of PCR products is described in DiMarzo et al. (1988) and Ohara et al. (1989). Purification methods commonly used to purify PCR products involve extractions and precipitations, the use of commercial filtration or chromatography columns, or enzymatic cleanup. The protocols used for column purification benefit from their simplicity. However, enzymatic purification (Alternate Protocol 4) is as fast as column purification. This method, developed by Hanke and Wink (1994) and Werle et al. (1994), requires relatively little hands-on time and results in a negligible loss of template, as all steps are performed in the very same PCR tube or well. In addition, the principal advantage of enzymatic purification is the relative ease with which it can be scaled up to highthroughput capacity at a mere fraction of the cost.

Critical Parameters and Troubleshooting The majority of problems that may hinder direct DNA sequencing of PCR products are described in the units on dideoxy and chemical sequencing (UNITS 7.4 & 7.5) and in the unit pertaining to optimization of PCR amplification (UNIT 15.1). Regardless of the source of DNA or the sequencing methods employed, the quality of a DNA sequence depends critically on the quality of the starting material. In the case of PCR products, it is imperative (before sequencing) to optimize the reaction conditions (UNIT 15.1) in order to obtain a homogeneous amplification product. For this reason, a final long extension step must be included in any PCR reaction. In certain cases, no amount of optimization will produce a single amplified product, and individual products must be purified by gel electrophoresis. In general, gel purification is sufficient to generate a homogeneous product. On occasion, even a single isolated band contains more than one amplified product (as revealed by the sequence), and more sophisticated methods, such as denaturing gel gradient electrophoresis (Myers et al., 1988), or reamplification with a set of nested internal primers (Ohara et al., 1989), must be used. Because PCR is normally carried out under conditions strongly favoring synthesis (high concentrations of enzyme, primers, and nucleotides), artifact sequences are easily generated.



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This is a particular risk when amplifying in the presence of only one primer. In such cases, DNA synthesis is not bounded by a second primer, and a heterogeneous collection of molecules, including “hybrid” or “artificial” molecules, can easily arise. In addition, snap-back synthesis can take place, again resulting in artificial amplified molecules. Whenever possible, the identity of amplified fragments should be confirmed prior to sequencing by probing of Southern blots, preferably with specific sequences internal to the amplified region (UNIT 2.9), or by restriction endonuclease digestion analyses of amplified products to confirm the expected restriction sites (UNIT 3.1). Conditions for obtaining single-stranded template must be optimized empirically before undertaking dideoxy sequencing. Alternatively, the alternate protocol described here for sequencing double-stranded PCR products can be employed. Many difficulties with the dideoxy method are caused by the secondary structure of the template strand. The inclusion of nucleotide analogs, coupled with the use of a thermostable polymerase (e.g., Taq DNA polymerase) for chain extension, can reduce or resolve many of these problems. In sequencing double-stranded PCR products, the crucial point is to prevent reannealing of the template strands that compete with binding of the sequencing primer. This can be overcome by two critical manipulations—heat denaturation of the template and quick addition of the primer to the template. For the one-step procedure (Alternate Protocol 4), titration of the amount of DNA template used for the cycle sequencing reaction is critical to sequence quality. Most polymerbased capillary sequencers require ∼50 to 500 ng DNA per sequencing reaction. Insufficient DNA will result in low-amplitude peaks on the sequencing chromatograms. Excessive DNA, which is more often the case, will result in the rapid depletion of the fluorophore-dNTPs in the cycle sequencing reaction. The sequencing chromatogram will appear to have extremely high-amplitude peaks that burn out relatively early, resulting in a very short region of usable sequence data. Often, the direct pipetting of undiluted PCR product will result in satisfactory sequences. However, optimal sequence lengths may require dilution of the PCR product prior to enzymatic purification. The shrimp alkaline phosphatase (ShrAP) should be screened for contaminating endonuclease or exonuclease activity. The ShrAP from USB has undetectable or negligible levels of

the above contaminating activities. Exonuclease I (Exo I) preferentially hydrolyzes single-stranded DNA. However, Exo I does have some detectable double-stranded DNA exonuclease activity. Therefore, prolonged incubation with this enzyme may result in minor hydrolysis at the ends of the double-stranded DNA template. For this reason, using a nested primer for sequencing may give improved sequence data over the use of one of the original PCR primers. However, using one of the original PCR primers as the sequencing primer often gives satisfactory results. Further, the enzyme incubation time should be kept to a minimum (i.e., 15 min) prior to heat inactivation. The amount of hydrolyzing enzyme used in this protocol has been titrated as low as possible without sacrificing sequencing quality. There are two reasons for using minimal enzyme quantities. First, large amounts of protein contaminants have been suspected to cause precipitation within the capillary arrays of ABI 3700 analyzers. The small amounts of enzyme used in this protocol have not caused any increased wear on capillary sequencers to date. Second, using less enzyme results in cost savings for high-throughput sequencing efforts.

Anticipated Results These protocols capitalize on the efficiency and specificity of PCR and streamline the process of DNA sequencing. From 150 to 300 bp of sequence can routinely be obtained from overnight exposures of either dideoxy or chemical (or genomic) sequencing gels, provided oligonucleotides or strands have been labeled with 32P. Longer exposures (24 to 72 hr) may be needed when using 35S as a label. With appropriate modifications, including multiple loadings, repeated probings, or a series of sequencing primers, the yield of legible sequence can be substantially increased. Rapid purification of PCR products can be achieved within one PCR tube or well, which can directly proceed to cycle sequencing using conventional dye-terminator reactions. The sequence quality should be comparable to or better than that achieved through column purification or ethanol precipitation. Further, this protocol can be easily scaled for high-throughput use.

Time Considerations Each protocol described here can be completed in a single day or less, with additional time required for sequencing reactions and analysis.



Literature Cited Church, G.M. and Gilbert, W. 1984. Genomic sequencing. Proc. Natl. Acad. Sci. U.S.A. 81:19911995. Church, G.M. and Kieffer-Higgins, S. 1988. Multiplex DNA sequencing. Science 240:185-188. DiMarzo, R., Dowling, C.E., Wong, C., Maggio, A., and Kazazian, H.H. 1988. The spectrum of βthalassaemia mutations in Sicily. Br. J. Haematol. 69:393-397. Gyllensten, U.B. 1989. Direct sequencing of in vitro amplified DNA. In PCR Technology (H.A. Erlich, ed.) pp. 45-60. Stockton Press, New York. Gyllensten, U.B. and Erlich, H.A. 1988. Generation of single-stranded DNA by the polymerase chain reaction and its application to the direct sequencing of the HLA-DQA locus. Proc. Natl. Acad. Sci. U.S.A. 85:7652-7656. Hanke, M. and Wink, M. 1994. Direct DNA sequencing of PCR-amplified vector inserts following enzymatic degradation of primer and dNTPs. Biotechniques 17:858-860.

Werle, E., Schneider, C., Renner, M., Volker, M., and Fiehn, W. 1994. Convenient single-step, one tube purification of PCR products for direct sequencing. Nucl. Acids Res. 22:4354-4355. Wrischnik, L.A., Higuchi, R.G., Stoneking, M., Erlich, H.A., Arnheim, N., and Wilson, A.C. 1987. Length mutations in human mitochondrial DNA: Direct sequencing of enzymatically amplified DNA. Nucl. Acids Res. 15:529-542.

Key References Kusukawa, N., Uemori, T., Asada, K., and Kato, I. 1990. Rapid and reliable protocol for direct sequencing of material amplified by polymerase chain reaction. BioTechniques 9:66-72. Outlines the method for double-stranded sequencing of PCR products. Maxam, A.M. and Gilbert, W. 1980. Sequencing end-labeled DNA with base-specific chemical cleavages. Methods Enzymol. 65:499-559. Sanger, F., Nicklen, S., and Coulson, A.R. 1977. DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. U.S.A. 74:5463-5467.

Higuchi, R.G. and Ochman, H. 1989. Production of single-stranded DNA templates by exonuclease digestion following the polymerase chain reaction. Nucl. Acids Res. 17:5865.

These two papers outline the basic principles and techniques of chemical and dideoxy sequencing.

Innis, T.M.A., Myambo, K.B., Gelfand, D.H., and Brow, M.A. 1988. DNA sequencing with Thermus aquaticus DNA polymerase and direct sequencing of polymerase chain reaction-amplified DNA. Proc. Natl. Acad. Sci. U.S.A. 85:94369440.

Contributed by Robert L. Dorit Yale University New Haven, Connecticut

Kreitman, M. and Landweber L. 1989. A strategy for producing single-stranded DNA in the polymerase chain reaction. Gene Anal. Technol. 6:84-88. Myers, R.M., Sheffield, V., and Cox, D.R. 1988. Detection of single-base changes in DNA: Ribonuclease cleavage and denaturing gel electrophoresis. In Genomic Analysis: A Practical Approach (K. Davies, ed.) pp. 95-139. IRL Press, Oxford. Ohara, O., Dorit, R.L., and Gilbert, W. 1989. Onesided polymerase chain reaction: The amplification of cDNA. Proc. Natl. Acad. Sci. U.S.A. 86:5673-5677.

Osamu Ohara Shionogi Research Laboratories Osaka, Japan Charles B-C. Hwang (double-stranded sequencing) Harvard Medical School Boston, Massachusetts Jae Bum Kim and Seth Blackshaw (one-step purification) Harvard Medical School Boston, Massachusetts



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Ligation-Mediated PCR for Genomic Sequencing and Footprinting

UNIT 15.3

The polymerase chain reaction (PCR) can be used to exponentially amplify segments of DNA located between two specified primer hybridization sites. This unit describes a single-sided PCR method that initially requires specification of only one primer hybridization site; the second is defined by the ligation-based addition of a unique DNA linker. This linker, together with the flanking gene-specific primer, allows exponential amplification of any fragment of DNA. Because a defined, discrete-length sequence is added to every fragment, complex populations of DNA such as sequence ladders can be amplified intact with retention of single-base resolution. Although it is also suitable for other applications, the ligation-based protocol (see Basic Protocol) was specifically designed for genomic footprinting and direct sequencing reactions, and is described in this context. The starting material is genomic DNA that has been cleaved so that it retains a 5′ phosphate at the cleavage site. In the schematic of the protocol (Fig. 15.3.1), only one cleavage product is shown for clarity; in practice, there would be a population of fragments resulting from the partial cleavage of DNA during footprinting or sequencing reactions. Preparation of DNA for footprinting analysis and genomic sequencing is described in the support protocols. Support Protocol 1 details in vivo and in vitro dimethyl sulfate (DMS) treatment and isolation of DNA from monolayer cells and its subsequent piperidine cleavage. Support Protocol 2 describes in vivo DMS treatment and harvesting of DNA from suspension cells. Support Protocol 3 describes how control DNA generated in the first two support protocols can be prepared for genomic sequencing. NOTE: It is essential that fresh, high-quality reagents be used throughout this unit. In addition, all solutions should be prepared with glass-distilled water. LIGATION-MEDIATED SINGLE-SIDED PCR In this protocol, cleaved DNA is denatured and a gene-specific primer (primer 1) is annealed to the region of interest. In the first-strand synthesis, this primer is extended with a processive polymerase (Vent DNA polymerase) to the cleavage site to create a blunt end. DNA ligase catalyzes the attachment of a unidirectional (staggered) linker to this blunt end. The 3′ end of the longer strand of the linker is ligated to the 5′ end of the genomic DNA. The shorter strand of the linker lacks a 5′ phosphate and therefore is not ligated to the extension product of the gene-specific primer. The DNA is denatured and a second gene-specific primer (primer 2) is annealed to the genomic DNA and extended by Vent DNA polymerase through the ligated linker region. (In theory, the first gene-specific primer could be used here, but in practice substituting a second primer greatly reduces background.) The extended product is now a suitable substrate for a PCR reaction; on one end (left in Fig. 15.3.1) there is a linker sequence to which a linker primer can anneal and on the other end (right in Fig. 15.3.1) there is a genomic sequence to which a gene-specific primer can anneal. Only molecules that have both sequences will be exponentially amplified during the subsequent PCR reaction; molecules with only one of the sequences will be linearly amplified. For the last extension, a third gene-specific primer (primer 3) that overlaps the second is used to label the DNA indirectly. These end-labeled extension products are visualized on a sequencing gel. High-quality, reproducible footprinting or sequencing reactions can be obtained starting from 6 × 105 genomes (3 × 105 diploid nuclei). This corresponds to 2 µg of mouse (mammalian) genomic DNA. For other species, the recommended minimum number of Contributed by Paul R. Mueller, Barbara Wold, and Paul A. Garrity Current Protocols in Molecular Biology (2001) 15.3.1-15.3.26 Copyright © 2001 by John Wiley & Sons, Inc.

BASIC PROTOCOL



original DNA primer • • • • • new DNA linker • • • • • • • • PCR DNA * end-labeled primer

randomly cleave DNA

denature DNA, anneal primer 1

(steps 1-5)

primer 1

not part of rest of reactions extend primer 1 with Vent polymerase

(steps 1-5)

primer 1 ••••••••••••• (steps 6-9) ligate linker to blunt end linker ••••••••••••• denature newly made DNA, anneal (steps 10-16) primer 2

primer 2 not part of rest of reactions

••••••••••• extend primer 2 with Vent polymerase

(step 16)

primer 2 ••••••••••••••• denature newly made DNA, (step 16) anneal linker primer and primer 2 primer 2 linker primer •••••••••••••••• extend linker primer and primer 2 with Vent polymerase

(step 16)

primer 2 •••••••••••••••••••••••

linker primer

•••• •••••••••••••••••

••••••••••••••••• repeat for 16 (total of 18 rounds) cycles and exponentially amplify DNA by PCR 4 5 (10 - to 10 -fold) denature amplified DNA, anneal end-labeled (steps 17-19) primer 3, with Vent polymerase ••••••••••••••••••••••••••

••••••••••••••

primer 3 *

•••••••••••••••••••••••••••••

(steps 20-24) visualize extended, end-labeled products on sequencing gel

LigationMediated PCR

Figure 15.3.1 Flowchart of ligation-mediated PCR protocol (see text for details). The steps correspond to those listed in the Basic Protocol.



haploid genomes remains 6 × 105, but different genome sizes lead to corresponding differences in the absolute mass of DNA. Materials 0.4 µg/µl cleaved genomic DNA in TE buffer, pH 7.5 (Support Protocol 1, 2, or 3) First-strand synthesis mix (see recipe), containing oligonucleotide primer 1 (Figs. 15.3.1 and 15.3.2; also see Critical Parameters) 20 µM unidirectional linker mix (see recipe) Ligase dilution solution (see recipe) Ligase mix (see recipe) 2000 to 3000 “Weiss” U/ml T4 DNA ligase (UNIT 3.14; Promega or Pharmacia Biotech) Precipitation salt mix (see recipe) 100% ethanol, ice-cold and room temperature 75% ethanol, room temperature Amplification mix (see recipe), containing linker primer and primer 2 2 U/µl Vent DNA polymerase mix (see recipe; also see UNIT 7.4) Mineral oil End-labeling mix (see recipe), containing end-labeled primer 3 (Fig. 15.3.3) Vent DNA polymerase stop solution (see recipe) 25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol (see recipe) Loading buffer (see recipe) 1.5-ml microcentrifuge tubes, silanized (APPENDIX 3) and with Lid-Loks (optional; Intermountain Scientific) 4° and 17°C water baths Automated thermal cycler or water baths at 95°, 60°, 76°, and 60° to 70°C Additional reagents and equipment for PCR (UNIT 15.1), denaturing gel electrophoresis for DNA sequencing (UNIT 7.6), and autoradiography (APPENDIX 3A) Carry out first-strand synthesis 1. Transfer 5 µl (2 µg) cleaved genomic DNA to a silanized 1.5-ml microcentrifuge tube and chill several minutes in an ice-water bath. The cleaved DNA sample must be clean; contaminants left in the DNA (e.g., piperidine) will interfere with the reaction.

2. Prepare first-strand synthesis mix containing primer 1 and chill several minutes in an ice-water bath. Add 25 µl to DNA sample, gently mix with a pipettor, and return sample to the ice-water bath. Place Lid-Loks on sample to prevent tube from popping open during denaturation.

A

B genomic DNA

genomic DNA primer 1

primer 1 primer 2 primer 3

increasing Tm

primer 2 primer 3

Figure 15.3.2 Two possible arrangements of gene-specific primers (see Critical Parameters).



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HO 5′

OH

GCGGTGACCCGGGAGATCTGAATTC

3′

LMPCR.1

3′ CTAGACTTAAG

5′

LMPCR.2

HO

OH

Figure 15.3.3 Staggered linker used in ligation-mediated PCR (LMPCR). This staggered linker is made by annealing the oligonucleotides LMPCR.1 and LMPCR.2 to each other. LMPCR.1 has 25 bases and a GC content of 60%, whereas LMPCR.2 has 11 bases and a GC content of 36%. Other oligonucleotides could be used in place of these as long as they meet the criteria defined in text. Note the lack of 5′ phosphates on oligonucleotides (i.e., 5′ OH instead) and the unconventional orientation of LMPCR.2 (3′ to 5′) in the figure.

When mixing with the pipettor, three or four careful strokes are usually sufficient. Try to avoid splashing the sample while mixing; if this occurs, microcentrifuge tubes briefly at 4°C to collect droplets.

3. Denature DNA 5 min at 95°C, anneal primer 30 min at 60°C, and extend 10 min at 76°C. As first-strand synthesis proceeds, prepare solutions in step 4. First-strand synthesis is most easily performed in an automated thermal cycler, but can be performed manually by transferring the tubes to different temperature water baths. It is important that the DNA be completely denatured so the primer can anneal. For this reason it is a good idea to check the calibration of the thermal cycler. The authors have found that some machines must be set to 96° or 97°C to actually obtain 95°C. Furthermore, if a thermal cycler is used, put the samples in the machine only after it is close to the denaturation temperature. This will minimize polymerase activity prior to the initial denaturation step. Most thermal cyclers have a “hold” function that allows sample addition followed by continuation of the program for the full denaturation time. If this reaction is performed manually using water baths, covering the tubes with a styrofoam block during denaturation and annealing minimizes condensation. The extension step creates a blunt-end substrate for the subsequent ligation reaction (see Fig. 15.3.1); therefore, success at this step is absolutely crucial because it determines the molecules that will ultimately be amplified (see Critical Parameters).

4. Thaw 20 µM unidirectional linker mix in ice-water bath. Prepare ligase dilution solution and partially prepare ligase mix (minus unidirectional linker and T4 DNA ligase; see reagents and solutions). Chill both in ice-water bath. Do not add unidirectional linker mix and T4 DNA ligase to ligase mix until step 6.

5. When extension in step 3 is complete, immediately transfer sample to the ice-water bath. Microcentrifuge tube briefly at 4°C to collect condensation, then return to ice-water bath. The samples should be kept cold during steps 6, 7, and 8 to minimize Vent polymerase activity (see Critical Parameters).

Carry out ligation 6. Finish preparing ice-cold ligase mix from step 4 by adding unidirectional linker mix, mixing, adding T4 DNA ligase, mixing, and keeping in ice-water bath. This order of addition is important because the unidirectional linker is prepared in 250 mM Tris⋅Cl, pH 7.7, which is required by the ligase.


7. Add 20 µl ice-cold ligase dilution solution from step 4 to sample, gently mix with a pipettor, and return sample to the ice-water bath. Add 25 µl ice-cold ligase mix from step 6 to sample, gently mix with a pipettor, and return to the ice-water bath.



Microcentrifuge tube briefly at 4°C and incubate overnight in a 17°C water bath (see critical parameters). Addition of ligase dilution solution makes the first-strand synthesis mix into a buffer compatible with T4 DNA ligase (the NaCl concentration and, more importantly, the pH, are reduced). Addition of ligase mix provides a convenient way to add ligase and linker to each sample.

8. Prepare sample for precipitation by placing in the ice-water bath for several minutes. Microcentrifuge tube briefly at 4°C, then return to ice-water bath. Placing samples in the ice-water bath is a convenient way of transferring them from the 17°C bath to the 4°C microcentrifuge without warming them.

9. Prepare and chill precipitation salt mix. Add 9.4 µl ice-cold precipitation salt mix and 220 µl of ice-cold 100% ethanol to the sample. Mix thoroughly by inversion and chill ≥2 hr at −20°C. If desired, the experiment may be stopped at this point. Samples under ethanol are stable for weeks at −20°C.

Carry out PCR 10. Microcentrifuge precipitated ligation reaction 15 min at 4°C and discard supernatant. 11. Add 500 µl of room-temperature 75% ethanol and invert several times to wash pellet and walls of tube. Microcentrifuge sample ∼5 min at room temperature and discard supernatant. Remove last traces of ethanol with a pipettor and allow any remaining ethanol to evaporate by air drying or by using a Speedvac evaporator. The precipitated pellet will be rather large and will spread up the side of the tube.

12. Add 70 µl water and leave sample at room temperature to dissolve pellet. Vortex tube occasionally to assist dissolution, and after each vortexing, collect droplets by microcentrifuging 2 to 3 sec. When pellet is dissolved (usually ≤30 min), chill sample in ice-water bath. While pellet is dissolving, prepare and chill amplification mix containing linker primer and primer 2. 13. Add 30 µl ice-cold amplification mix to sample, gently mix with a pipettor, and return sample to the ice-water bath. 14. Add 3 µl (1 U) Vent DNA polymerase mix to sample and mix carefully with pipettor. Return sample to the ice-water bath. The reaction is very sensitive to the amount of Vent DNA polymerase used; excess polymerase results in high background. Surprisingly, this effect has been observed when the amount of Vent polymerase is merely double the optimal level. The basis for the empirically observed threshold for this background is not known. Different lots of Vent polymerase may need to be titrated (see Critical Parameters).

15. Cover sample with 90 µl mineral oil, microcentrifuge briefly at 4°C, and return to the ice-water bath. If the PCR reaction is performed manually using water baths, a Lid-Lock should be used to prevent the tube from popping open during denaturation.

16. Carry out 18 cycles of PCR. Perform first denaturation for 3 to 4 min at 95°C, and subsequent ones for 1 min. Anneal primers 2 min at a temperature 0° to 2°C above the calculated Tms (if primer 2 and linker primer have different Tms, use the lower Tm; see critical parameters for Tm calculations). Extend 3 min at 76°C; for every cycle, add an extra 5 sec to the extension step. Allow final extension to proceed 10 min.



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Transfer sample to ice-water bath, remove Lid-Lock (if applicable), and microcentrifuge briefly at 4°C to collect any condensation. Keep sample in ice-water bath. In this procedure, 18 rounds of amplification yield highly reproducible sequence ladders that can be visualized (after the radiolabeling step) following an overnight exposure without an intensifying screen. Performing additional rounds of amplification could reduce the exposure time required, but this has not been tested exhaustively. A theoretical concern is that if too many amplification cycles are carried out, some reagents (e.g., polymerase, primers, or dNTPs) may become limiting, leading to undesirable lane-to-lane variability (18 rounds of amplification do not show variability). An important parameter in this step is the temperature of denaturation. If it is too low, no signal or shortened sequence ladders will be seen; if it is too high, the polymerase will be destroyed. This may be a may be a “hidden” problem because some automatic thermal cycling machines and thermometers do not accurately measure the denaturation temperature (see Critical Parameters).

Carry out end-labeling 17. Prepare end-labeling mix containing labeled primer 3 and chill several minutes in the ice-water bath. Add 5 µl to the sample. Mix aqueous phase by gently pipetting up and down, keeping the sample on ice as much as possible. Microcentrifuge briefly at 4°C and return to ice-water bath. CAUTION: The label mix contains a significant amount of 32P. Appropriate care should be taken in handling and disposal of mix and samples to which it is added.

18. Carry out two rounds of PCR to label the DNA. Perform first denaturation 3 to 4 min at 95°C; the second for 1 min. Anneal end-labeled primer 3 for 2 min at a temperature 0° to 2°C above its calculated Tm. Extend 10 min at 76°C. When second extension is complete, transfer sample to ice-water bath. Although it is rare, in a few experimental situations, having two cycles of PCR during the end-labeling step has led to increased background and/or unusual primer artifacts such as primer-dimer products. It is best to try two cycles of end-labeling, cutting back to one cycle if this problem is encountered. It is also possible to label the amplification products using a filter-blotting procedure (Pfeifer et al., 1989). If this reaction is performed manually using water baths, a Lid-Lock should be used to prevent the tube from popping open during denaturation.

19. Place sample at room temperature and immediately add 295 µl Vent DNA polymerase stop solution. Mix by vortexing. Microcentrifuge briefly to collect radioactive droplets from top and sides of tube. Add 500 µl phenol/chloroform/isoamyl alcohol and mix either by vigorous shaking or vortexing. Microcentrifuge 3 to 5 min at room temperature. Transfer upper aqueous layer (∼400 µl; avoid interface if any) to a clean, silanized 1.5-ml microcentrifuge tube, and thoroughly mix. Microcentrifuge briefly to collect droplets from sides of tube. EDTA in the Vent DNA polymerase stop solution chelates the magnesium required by Vent polymerase. The organic extraction removes proteins before precipitation and, more importantly, removes the mineral oil (there should be little or no interface). Transferring the aqueous layer to a new tube and mixing it ensures the reaction products will be equally aliquoted for multiple loadings on the sequencing gel (step 20).

20. Set up four clean, silanized 1.5-ml microcentrifuge tubes and add 235 µl of roomtemperature 100% ethanol to each tube. Transfer 94 µl of the aqueous layer into each tube, thoroughly mix by vortexing, and chill ≥2 hr at −20°C. Discard any remaining aqueous layer. LigationMediated PCR



This yields enough material for four samples of each reaction to be run on a sequencing gel. This accommodates multiple loadings to maximize the number of bases that can be read and provides extra samples for use in case a gel problem is encountered. If desired, this precipitation step can be a stopping point. Samples under ethanol are stable for several weeks at −20°C; however, the specific activity of the product decreases with time.

21. Microcentrifuge precipitated samples 15 min at 4°C and discard supernatants. 22. Add 500 µl of room temperature 75% ethanol, vortex, microcentrifuge samples 5 min at room temperature, and discard supernatants. Remove last traces of ethanol by using a pipettor and allow any remaining ethanol to evaporate by air drying or by using a Speedvac evaporator. 23. Add 7 µl loading buffer to each tube and leave at room temperature while pellets are dissolving. Vortex occasionally to assist pellets in dissolving and to recover any of the DNA pellet that is on the side of the tube. After each vortexing, collect droplets by microcentrifuging 2 to 3 sec. Samples usually resuspend rapidly (within 5 min). Check resuspension by removing the sample from the tube with a pipet and using a Geiger counter to ensure that the radioactivity is in the sample and not left behind in the tube. Return sample to same tube. If >10% of the total radioactivity has remained in the tube, vortex, microcentrifuge, and repeat the resuspension check. Do not let the samples sit longer than necessary in loading buffer, as this may result in smeared background and poor band resolution.

Run and analyze sequencing gel 24. Denature samples 5 min at 85° to 90°C. Load entire content of each tube on a 6% sequencing gel. At the completion of the run, fix and dry gel, then autoradiograph 6 to 24 hr without an intensifying screen. The exposure time varies and is dependent upon several factors, including the gene being amplified, the number of amplification and labeling cycles, and the specific activity of primer 3. The ladder will appear 25 bases longer than the original footprinting or sequencing products due to the addition of the 25-base linker. Because a large amount of material is being applied to the gel, use of the usual 0.2-mm-thick sequencing gels may result in smeared bands; therefore, results are usually better if a relatively thick gel (0.35 to 0.56 mm) is used instead. The first readable sequence band will run at ∼50 nucleotides, depending on the length of the labeling primer used. Therefore, to read as much sequence as possible, multiple loadings on 80-cm gels are recommended (60-cm gels also work well if an electrolyte gradient is used; UNIT 7.6), and any fragments shorter than 50 nucleotides are run into the bottom buffer tank. This means that the bottom buffer tank will contain a significant amount of radioactive 32P in the form of unused primer 3; thus, disposal guidelines must be rigorously applied.

PREPARATION OF GENOMIC DNA FROM MONOLAYER CELLS FOR DMS FOOTPRINTING This protocol describes the preparation of in vivo and in vitro dimethyl sulfate (DMS)– treated genomic DNA from monolayer cells for use in conjunction with ligation-mediated PCR. Figure 15.3.4 outlines the parallel processing of duplicate sets of cultured cells: one set is left untreated (control) and the second is treated with DMS (in vivo treatment). Both cultures are lysed and their DNA harvested. The control-cell DNA is then treated with DMS (in vitro treatment). Both in vivo and in vitro DMS-treated DNA are cleaved with piperidine before use in the ligation-mediated PCR assay (Basic Protocol). The protocol can be modified to prepare DNA from suspension cells (Support Protocol 2). The deproteinized control DNA (not treated with DMS) is of high quality and thus can also be used for genomic sequencing as described in Support Protocol 3.

SUPPORT PROTOCOL 1



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obtain duplicate sets of monolayer or suspension cell cultures

treat one culture as control (steps 1-2 of Support Protocol 1 or steps 1-3 of Support Protocol 2)

use control DNA in genomic sequencing (Support Protocol 3) or genomic methyl cytosine analysis (Saluz and Jost, 1987)

treat one culture with DMS (in vivo treatment; steps 1 & 3-4 of Support Protocol 1 or steps 1, 2 & 4-7 of Support Protocol 2)

harvest DNA from both cultures (steps 5-16 of Support Protocol 1 and step 8 of Support Protocol 2)

treat control DNA with DMS (in vitro treatment; steps 17-20 of Support Protocol 1)

cleave both DNA samples with piperidine (steps 21-29 of Support Protocol 1)

use in ligation-mediated PCR (Basic Protocol)

Figure 15.3.4 Flow chart of preparation of DMS-footprinted genomic DNA for use in ligation-mediated PCR.

DMS methylates guanine residues at the N7 position, rendering them susceptible to subsequent cleavage with piperidine. DMS is commonly used for in vivo footprinting analysis because cellular membranes are freely and rapidly permeable to it. Treatment of both control (in vitro–treated) and experimental (in vivo treated) samples with DMS is limited so that only ∼1 in 150 guanines is methylated. On the other hand, piperidine treatment is quantitative so that all of the methylated guanines are cleaved. The extraction and purification of the DNA must be thorough so that all proteins and reagents are removed before the DNA is used in ligation-mediated PCR (Basic Protocol). This procedure works well with ∼5 × 107 cells/set (i.e., one 15-cm plate at ∼70% confluence for many fibroblasts), although more cells can be used by scaling up the extractions and A. The DNA will be quite viscous, so pipet and aliquot it carefully.

2. Treat with DMS as described in Support Protocol 1, steps 18 and 19. Proceed to step 9. Prepare G+A, C+T, and C reactions 3. Place 20 to 40 µg control DNA in each of three 1.5-ml microcentrifuge tubes. Add sufficient TE buffer to bring the total volume in each to 100 µl. Label one tube G+A, one tube C+T, and one tube C. The DNA will be quite viscous, so carefully pipet and aliquot it.

4. Add 11 µl of 3 M sodium acetate and 222 µl of room-temperature 100% ethanol to each tube. Mix thoroughly by gently inverting a few times. The DNA should precipitate immediately.

5. Microcentrifuge precipitated samples 5 min at maximum speed, room temperature, then remove and discard supernatants. Add 500 µl of room-temperature 75% ethanol to each pellet and mix by inversion until pellets are dislodged. Microcentrifuge 3 min at maximum speed, room temperature, then remove and discard supernatants. Try to remove as much of the 75% ethanol as possible with a pipettor, but do not allow the DNA pellets to dry out. Do not use a Speedvac evaporator, as it will be impossible to resuspend the DNA once it is dried.

6. Redissolve samples in water as follows: add 18 µl to G+A tube, 40 µl to C+T tube, and 10 µl to C tube. Incubate overnight at 4°C. Warm redissolved DNA to room temperature. Uncleaved genomic DNA redissolves slowly at these concentrations. Gentle mixing by flicking the tube with a finger a few times during the resuspension can help speed this process. Overnight incubation is usually sufficient to allow resuspension. A small amount of insoluble material is occasionally seen at this step. It does not interfere with subsequent



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reactions and can be ignored; it will be discarded after the piperidine cleavage reaction in step 9.

For the G+A sample: 7a. Add 54 µl of 88% formic acid, mix thoroughly by vortexing 25 sec, and microcentrifuge briefly to collect droplets. Incubate 7 min at room temperature. 8a. Add 164 µl ice-cold G+A stop solution and vortex briefly (3 sec). Add 750 µl of 100% ethanol prechilled on dry ice, mix by vigorously shaking, and plunge tube into powdered dry ice. Leave samples in dry ice for ∼30 min, then proceed to step 9. The 7-min incubation time used here is a good starting condition. The extent of reaction can be changed by varying the incubation time; the average size of the final product after piperidine cleavage will decrease as incubation time is increased.

For the C+T sample: 7b. Add 60 µl hydrazine, mix thoroughly by vortexing 25 sec, and microcentrifuge briefly to collect droplets. Incubate 3 min at room temperature. 8b. Add 150 µl ice-cold C+T/C stop solution and vortex briefly (3 sec). Add 750 µl of 100% ethanol prechilled on dry ice, mix by vigorously shaking, and plunge tube into powdered dry ice. Leave samples in dry ice for ∼30 min, then proceed to step 9. The 3-min incubation time used here is a good starting condition. The extent of reaction can be changed by varying the incubation time, as described in step 8a.

For the C sample: 7c. Add 30 µl of 5 M sodium chloride, mix thoroughly by vortexing 25 sec, and microcentrifuge briefly to collect droplets. Add 60 µl hydrazine, mix thoroughly by vortexing 25 sec, and microcentrifuge briefly to collect droplets. Incubate 3 min at room temperature. 8c. Add 150 µl ice-cold C+T/C stop solution and vortex briefly (3 sec). Add 750 µl of 100% ethanol prechilled on dry ice, mix by vigorously shaking, and plunge tube into powdered dry ice. Leave samples in dry ice for ∼30 min, then proceed to step 9. The 3-min incubation time used here is a good starting condition. The extent of reaction can be changed by varying the incubation time, as described in step 8a.

Carry out cleavage and sequencing 9. Cleave DNA from each of the four reactions with piperidine as described in Support Protocol 1, steps 21 to 29. Proceed to ligation-mediated PCR (Basic Protocol). REAGENTS AND SOLUTIONS 4dNTP mix, pH 7.0, 25mM Prepare as in UNIT 3.4. Alternatively, purchase as individual 100-mM stock solutions from Pharmacia Biotech and combine equal amounts of dATP, dCTP, dGTP, and dTTP to obtain the 25 mM 4dNTP mix. Store at −20°C.


Amplification buffer, 5× 200 mM NaCl 100 mM Tris⋅Cl, pH 8.9 (APPENDIX 2), room temperature 25 mM MgSO4 0.05% gelatin (from bovine skin; Sigma) 0.5% Triton X-100 Store at −20°C



Amplification mix, per reaction (30 ìl): 20.0 µl 5× amplification buffer (see recipe) 1.0 µl 10 pmol/µl oligonucleotide LMPCR.1 (linker primer; Fig. 15.3.3) 1.0 µl 10 pmol/µl oligonucleotide primer 2 (Figs. 15.3.1 & 15.3.2) 0.8 µl 25 mM 4dNTP mix, pH 7.0 (see recipe) 7.2 µl H2O Prepare immediately before use and chill on ice Addition of LMPCR.1 may not be required, but this has not been tested.

C+T/C stop solution 400 mM sodium acetate, pH 7.5 0.14 mM EDTA, pH 8.0 (APPENDIX 2) Prepare fresh and chill on ice before use End-labeling mix, per reaction (5 ìl): 1.0 µl 5× amplification buffer (see recipe) 2.3 µl 1 pmol/µl end-labeled primer 3 (Fig. 15.3.3) 0.4 µl 25 mM 4dNTP mix, pH 7.0 (see recipe) 0.8 µl H2O 0.5 µl 2 U/µl Thermococcus litoralis DNA polymerase (Vent; New England Biolabs) Prepare immediately before use. Mix first four components and chill on ice before adding Vent DNA polymerase and chill on ice. Then add polymerase and keep on ice. 5′ end-labeled primer 3 can be prepared as in UNIT 3.10 (forward reaction) with the following modifications: end-label 20 to 100 pmol of primer 3 with labeling-grade (i.e., less expensive) [γ-32]P ATP (∼6000 Ci/mmol); incubate at 37°C for 30 min instead of 60 min. Remove unincorporated 32P by gel purification (UNIT 2.12) or by using a Nensorb-20 nucleic acid purification cartridge (Du Pont NEN)—elute primer with 50% ethanol instead of 50% methanol. Resuspend primer 3 in TE buffer, pH 7.5, so that its concentration is 1 pmol/µl; the specific activity should be 4–9 × 106 cpm/pmol.

First-strand buffer, 5× 200 mM NaCl 50 mM Tris⋅Cl, pH 8.9 (APPENDIX 2) room temperature 25 mM MgSO4 0.05% gelatin (from bovine skin; Sigma) Store at −20°C First-strand synthesis mix, per reaction (25 ìl): 6.0 µl 5× first-strand buffer (see recipe) 0.3 µl 1 pmol/µl oligonucleotide primer 1 0.24 µl 25 mM 4dNTP mix, pH 7.0 (see recipe) 18.21 µl H2O 0.25 µl 2 U/µl Thermococcus litoralis DNA polymerase (Vent; New England Biolabs #254L) Prepare immediately before use. Mix all components except Vent DNA polymerase and chill on ice. Then add polymerase and keep on ice. Primer 1 should be stored in a silanized tube at a concentration ≥10 pmol, then diluted in TE buffer, pH 7.5, immediately before use. This will minimize primer loss due to sticking to the tube walls.



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G+A stop solution 360 mM sodium acetate, pH 7.5 0.14 mM EDTA, pH 8.0 (APPENDIX 2) Prepare fresh and chill on ice before use Ligase dilution solution, per reaction (20 ìl): 2.2 µl 1 M Tris⋅Cl, pH 7.5, at room temperature (APPENDIX 2; 110 mM final) 0.35 µl 1 M MgCl2 (17.5 mM final) 1.0 µl 1 M DTT (50 mM final) 0.25 µl 10 mg/ml BSA (DNase-free; Pharmacia Biotech; 125 µg/ml final) 16.2 µl H2O Prepare immediately before use and chill on ice Ligase mix, per reaction (25.0 ìl): 0.25 µl 1 M MgCl2 (APPENDIX 2; 10 mM final) 0.50 µl 1 M DTT (APPENDIX 2; 20 mM final) 0.75 µl 100 mM ATP, pH 7.0 (Pharmacia Biotech; 3 mM final) 0.125 µl 10 mg/ml BSA (DNase-free; Pharmacia Biotech; 50 µg/ml final) 17.375 µl H2O 5.0 µl 20 µM unidirectional linker mix (see recipe; 4 µM linker and 50 mM Tris⋅Cl, pH 7.7, final) 1.0 µl 3 “Weiss” U/µl T4 DNA ligase (UNIT 3.14; 3 U final) Prepare immediately before use. First mix MgCl2, DTT, rATP, BSA, and H2O and chill on ice. Next add ice-cold unidirectional linker mix and T4 DNA ligase. Tris⋅Cl added in the linker mix is 50 mM, pH 7.7, at final concentration.

Loading buffer 80% (v/v) formamide, deionized (see UNIT 14.3) 45 mM Tris base 45 mM boric acid 1 mM EDTA 0.05% (w/v) bromphenol blue 0.05% (w/v) xylene cyanol Store in aliquots at −20°C and discard after 3 months Lysis solution 300 mM NaCl 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2), room temperature 25 mM EDTA, pH 8.0 (APPENDIX 2) 0.2% (v/v) SDS (prepare fresh; add just before use) 0.2 mg/ml proteinase K (prepare fresh; add just before use) Lysis solution without SDS and proteinase K can be stored indefinitely at room temperature. Immediately before use, add 10 µl of 20% SDS and 10 µl of 20 mg/ml proteinase K per milliliter lysis solution. Phenol/chloroform/isoamyl alcohol Mix 25 parts phenol (equilibrated in 150 mM NaCl/50 mM Tris⋅Cl (pH 7.5)/1 mM EDTA) with 24 parts chloroform and 1 part isoamyl alcohol. Add 8-hydroxyquinoline to 0.1%. Store in aliquots at −20°C and discard after 6 months.


Precipitation salt mix, per reaction (9.4 ìl): 8.4 µl 3 M sodium acetate, pH 7.0 (2.7 M final) 1.0 µl 10 mg/ml yeast tRNA (∼1 mg/ml final) Prepare immediately before use



Unidirectional linker mix, 20 ìM 20 µM oligonucleotide LMPCR.1 (Fig. 15.3.3) 20 µM oligonucleotide LMPCR.2 (Fig. 15.3.3) 250 mM Tris⋅Cl, pH 7.7 (APPENDIX 2) Prepare this mix in advance as follows: (1) purify the oligonucleotides on denaturing polyacrylamide gels (UNIT 2.12), (2) combine the two oligonucleotides and Tris⋅Cl and heat 5 min at 95°C, (3) transfer to 70°C and gradually cool ∼1 hr to room temperature, (4) leave ∼1 hr at room temperature and then gradually cool ∼1 hr to 4°C, and (5) leave ∼12 hr at 4°C and store in aliquots at −20°C. Thaw linker on ice before use. Hybridization of the linker undoubtedly takes place more rapidly than allowed for in this procedure, but this has reproducibly worked well. Because the monovalent salts (e.g., NaCl) more traditionally used to raise ionic concentration inhibit T4 DNA ligase, and Tris⋅Cl is required by the ligase in any case, Tris⋅Cl is used as the salt in this hybridization reaction. Consequently, it is difficult to calculate the kinetics of oligonucleotide annealing, and so the question of timing has not been studied in detail.

Vent DNA polymerase mix, per reaction (3.0 ìl) Mix 0.6 µl of 5× amplification buffer (see recipe) with 1.9 µl H2O and chill several minutes in an ice-water bath. Add 0.5 µl (1 U) Thermococcus litoralis DNA polymerase (Vent; New England Biolabs), gently mix, and return to ice-water bath. Prepare just before use. Vent DNA polymerase stop solution, per reaction (295 ìl): 25 µl 3 M sodium acetate, pH 7.0 (260 mM final) 266 µl TE buffer, pH 7.5 (APPENDIX 2; 10 mM Tris⋅Cl, pH 7.5, final) 2 µl 0.5 M EDTA, pH 8.0 (APPENDIX 2; ∼4 mM final) 2 µl 10 mg/ml yeast tRNA (68 µg/ml final) Prepare immediately before use COMMENTARY Background Information Ligation-mediated PCR was originally developed to study in vivo protein-DNA interactions at regions of genes important for transcriptional control of expression. Several genomic sequencing strategies have been previously applied to in vivo footprinting. These relied on the principles of Southern blotting (Church and Gilbert, 1984), solution hybridization (Jackson and Felsenfeld, 1985), or primer extension (Huibregtse and Engelke, 1986). Unfortunately, with these strategies, in vivo footprinting of a single-copy regulatory region in large genomes (e.g., mammalian) is technically challenging, requires large cell numbers, and often produces results with unacceptable signal-to-noise ratios. The polymerase chain reaction (Saiki et al., 1988; White et al., 1989) is used to amplify specific fragments of single-copy genes (see Chapter 15 introduction and UNIT 15.1). It relies on two primers that flank the specific fragment of DNA to be amplified and uses repeated

cycles of template denaturation, primer annealing, and extension by a thermally stable DNA polymerase to exponentially amplify that fragment. PCR is not immediately applicable to sequencing or footprinting because it requires that every fragment of DNA have two defined ends (a sequence ladder is composed of a population of related DNA fragments that differ in length; they all have one end in common but the other end varies depending on the sequence). The Basic Protocol in this unit circumvents this problem by attaching a common sequence to the variable end of every fragment, thus creating substrates suitable for PCR and resulting in exponential amplification of each representative of the sequence ladder. One end of every fragment is primed with a gene-specific primer and the other is primed with the common attached sequence. Footprint and sequence information contained in the original sample of genomic DNA is amplified by an additional 104 to 105 copies. When visualized on a sequencing gel, each band in the ladder is



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shifted to a higher position that corresponds to the length of attached sequence. In practical terms, starting with as little as 2 µg of mammalian DNA (equivalent to 3 × 105 cells), a genomic sequence ladder of a single-copy gene can be seen following overnight exposure without an intensifying screen. Ligation-mediated PCR differs significantly from another single-sided PCR procedure that uses terminal transferase to add a homopolymeric tail of guanines or adenines to one end of the substrate DNA (Frohman et al., 1988; Loh et al., 1989; Ohara et al., 1989). This tail is added nonspecifically to all DNA and is expected to be somewhat variable in length. In contrast, ligation-mediated PCR results in the blunt-end ligation-based addition of a sequence of defined length and composition. The blunt end is generated by the extension of a gene-specific primer. The initial hybridization of this primer selects a small subset of specific gene fragments (to be modified in later steps) from the original complex DNA sample. This level of selection is unique to ligation-mediated single-sided PCR. These features, together with the use of multiple gene-specific primers, greatly reduce nonspecific background and provide the single-base resolution required for genomic footprinting, sequencing genomic DNA (Pfeifer et al., 1989; Garrity and Wold, 1992), studying in vivo methylation patterns of cytosine residues (Pfeifer et al., 1989, 1990; Rideout et al., 1990; Garrity and Wold, 1992), and cloning promoter elements from restriction-digested genomic DNA (Fors et al., 1990). This last adaptation uses gene-specific primers selected from known sequences located at the 5′ end of an mRNA to clone the corresponding unknown promoter region. Support Protocols 1, 2, and 3 describe methods for the preparation of genomic DNA for genomic DMS footprinting and genomic sequencing. Both of these methods are based on the Maxam-Gilbert chemistry described in UNIT 7.5. In addition, DNA cleaved randomly by a nuclease such as DNase I (UNIT 12.4; Tanguay et al., 1990; Rigaud et al., 1991), or site-specifically by a restriction endonuclease (UNIT 3.1) can also in principle be utilized in ligation-mediated PCR. In any case, the cleaved DNA must contain a 5′ phosphate to participate in the ligation reaction.

Critical Parameters LigationMediated PCR

Several important parameters are discussed in this section, but two underlying technical

points deserve emphasis at the outset. First, in order to keep background to a minimum, the hybridizations and extensions should be performed at the highest possible temperatures. Hybridizations are incubated at or 2°C above the calculated melting temperature (Tm) of the primers, and extensions are performed at 76°C with Vent DNA polymerase. (In specific cases, these temperatures may need to be adjusted, but they have worked with all combinations of primers tested to date). Second, background can be greatly reduced by using multiple primers, each primer having an end that extends 3′ to the previous one. This produces a level of specificity that cannot be obtained with a single 25-base primer. It is recommended that a test experiment be done before any footprinting or sequencing experiments are undertaken. This should include several identical samples that are processed in parallel. When visualized on a sequencing gel, these samples should appear identical. In addition, this is a convenient way to test for any problems in the operation of the automated thermal cycler or in the execution of ligation-mediated PCR. Whenever possible, prepare solutions as a reaction cocktail; this will minimize sample variation. Oligonucleotides should be gel-purified (UNIT 2.12) to remove contaminants and stored in TE buffer, pH 7.5, at ≥10 pmol/µl and at −20°C. Gene-specific primers Primer position. Figure 15.3.2 shows possible arrangements for gene-specific primers. Several different primer combinations have been tested and the following operating rules were derived (P.R.M. and B.W., unpub. observ.). Primers 1 and 2 may or may not overlap but the extending end of primer 2 must be 3′ to primer 1. If they do overlap, the overlap should be less than half their length. On the other hand, primer 3 can completely overlap primer 2 and extend a few extra bases 3′ to it (Fig. 15.3.2A), and it must be positioned with an overlap of at least half the length of primer 2 (Fig. 15.3.2B). If primers 2 and 3 do not overlap, the labeling extension will not be successful, because it is necessary for these primers to compete for the same binding site. The basis of this phenomenon is not known. It may be caused by the premature extension product of primer 2 excluding the hybridization of primer 3. The hybridization temperatures used during the annealing phase of the labeling cycle(s) are close to the optimal temperature for Vent DNA po-



lymerase. This is done to minimize background, but probably results in the immediate extension of any primer following its binding. Primer composition and melting temperature. Elevated temperatures minimize the formation of imperfect duplexes between genomic DNA and primers. Thus, to minimize background, hybridizations and reactions should be carried out under temperature conditions that demand as much as much specificity as possible. For hybridizations, the optimal temperature is at or even above the calculated Tm of the primer (see below for calculation). This principle is also used to favor the binding of the labeling primer (primer 3) over the amplification primer (primer 2). In general, the Tm of the primers should increase in the order primer 1 < primer 2 < primer 3. Thus, the binding of the first-strand synthesis primer (primer 1) will be less stable under amplification conditions and the binding of the amplification primer (primer 2) will be less stable under labeling conditions. In practice, this doesn’t appear to be essential, but it would probably be unwise to have the relative Tm of the primers reversed. The Tm of the primers can be adjusted by choosing sequences with differing GC content and/or length (see below). The first-strand synthesis primer (primer 1) should have a Tm of ≥60°C; primers of 20 (∼60% GC content) to 25 bases (∼50% GC content) work well. If the Tm of the primer is below 60°C, it may be necessary to reduce the hybridization temperature below the suggested 60°C (see Basic Protocol, step 3). The amplification primer (primer 2) should have a Tm equal to or higher than that of primer 1, and should closely match the Tm of the linker primer; a primer of 25 bases with a GC content of ∼60% works well. The labeling primer (primer 3) should have an even higher Tm. During the labeling extension, the linker primer does not need to extend, and the labeling primer can better compete with the amplification primer for occupancy if it has a higher Tm (recall that the amplification primer and labeling primer must overlap or the labeling primer will be excluded). Overall, the exact GC content or length of the primers is probably not very important. What is important is their relationship to each other and to the temperature of the reactions. Primers must be long enough to be specific to the gene of interest, but they should not be excessively long or have a Tm greater than ∼76°C (this may lead to nonspecific binding). The following formula (Wahl et al., 1987) may be used to approximate the Tm of primers:

Tm = 81.5 + 16.6 (logM) + 0.41(%GC) − (500 ⁄ n )

where n = length of primer and M = molarity of the salt in the buffer. To determine the molarity, ignore the contribution of Mg2+ and multiply the Tris concentration by 0.67 (N. Davidson, pers. comm.). Therefore, for the amplification buffer, the molarity would be 0.040 (NaCl) + 0.67 × 0.020 (Tris) = 0.053 M salt. Hybridizations work best if done at or 2°C above this calculated Tm. As an example, the Tm calculation for a 25-base primer with a 60% GC content is as follows: 81.5 + 16.6(log 0.053) + .41(60) − (500/25) = 65°C. Hybridization with this primer should be carried out between 65° and 67°C. This is only an approximation, and base stacking and near-neighbor effects may be significant for particular primers. Therefore, it may be necessary to determine empirically the optimal hybridization temperatures for each primer. Unidirectional linker The staggered design of the oligonucleotide linker is important because it assures that the ligation event is directional (see Fig. 15.3.3). The 3′ hydroxyl of the long oligonucleotide (LMPCR.1) will be ligated to the 5′ phosphate of the genomic DNA. For cloning purposes, LMPCR.1 contains restriction sites for BstEII, SmaI, BglII, and EcoRI; however, these are not used in sequencing or footprinting experiments. This oligonucleotide will also function as the linker primer in the PCR amplification. On the other hand, the short oligonucleotide (LMPCR.2) functions only in a structural role. When it is hybridized to the long oligonucleotide, it creates a blunt-end duplex at the 3′ end of LMPCR.1. This enables T4 DNA ligase to attach LMPCR.1 to the blunt-end duplex of the genomic DNA. By making the short oligonucleotide small and low in GC content, it is possible to make it unable to serve as a primer in subsequent Vent DNA polymerase reactions that are carried out far above its Tm. To prevent linker self-ligation, neither oligonucleotide has a 5′ phosphate. By default, this prevents LMPCR.2 from ligating to the first-strand synthesis extension products, but this is not an essential design feature (see Fig. 15.3.1). The exact sequence, GC content, length, and type of restriction endonuclease sites in this linker are not thought to be important for the function of the linker. The crucial properties for the linker are that (1) it contain no 5′ phosphates and be staggered to eliminate self-ligation and assure unidirectionality, (2) it be a ligatable



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structure, (3) the duplex between the two olignucleotides be stable during ligation conditions but not during PCR conditions, and (4) the long oligonucleotide have a Tm in Vent DNA polymerase buffer comparable to that of primer 2. Ligation Most products of the first-strand synthesis reaction appear to be ligated after 45 min at 15°C. However, particular bands in some ladders require longer incubation times to reach full intensity (Mueller and Wold, 1991). For this reason, an overnight (6 to 15 hr) ligation at 15° to 17°C is suggested. Although a higher ligation temperature might decrease the time required for complete ligation, this could inadvertently permit undesirable Vent DNA polymerase activity. Prior to ligation, no attempt is made to destroy the Vent DNA polymerase used during the first-strand synthesis. A possible concern is that if the polymerase is active during the ligation, it may extend the short oligonucleotide of the staggered linker (this would compromise the unidirectionality of the linker addition). With ligation at 15° to 17°C, this does not appear to be a problem, but it may be unwise to attempt higher temperatures unless the polymerase is inactivated.


Vent DNA polymerase Earlier versions of ligation-mediated PCR used a combination of Sequenase and Taq DNA polymerase for the first-strand synthesis and amplification/labeling reactions, respectively. Both of these enzymes have been replaced by Thermococcus litoralis DNA polymerase (Vent DNA polymerase). Vent DNA polymerase virtually eliminates the unequal representation of some members of a sequence ladder and occasional spurious bands that were previously observed with Sequenase/Taq polymerase–based ligation-mediated PCR. It also provides an approximate three-fold increase in signal strength to all members of the sequence ladder (Garrity and Wold, 1992). A very important procedural point is that ligation-mediated PCR is very sensitive to the amount of Vent DNA polymerase activity. A small (e.g., two-fold) increase in polymerase can result in an unacceptable background level. The reason for this threshold response to the amount of Vent DNA polymerase activity is unknown, but it does mean that this polymerase must be used carefully. If high background is a problem (or suddenly becomes one), the polymerase should be titrated. Normally it is not necessary to titrate each lot of Vent DNA po-

lymerase, but a change in manufacturing at New England Biolabs occurred between early lots (i.e., prior to the fall of 1991), which were less active, and more recent ones. Thus, originally 1 U of Vent DNA polymerase was needed in the first-strand synthesis and 3 U were needed at the amplification step (Garrity and Wold, 1992), but now 0.5 U in the first-strand synthesis and 1.0 U at the amplification step appears to be the ideal amount (P.G. and B.W., unpub. observ.). The 1.0 U added at the labeling step does not need to be altered. The “exonuclease-minus” version of Vent DNA polymerase has not been tested in this procedure. The buffers recommended here for use with Vent DNA polymerase in ligation-mediated PCR have been developed through empirical testing and differ from the one recommended by the manufacturer (P.G. and B.W., unpub. observ.). KCl has been replaced by NaCl, which gives better extension through certain guaninerich regions. BSA has been replaced by gelatin, because this reduces the interface formed after the organic extraction. (NH4)2SO4 has been eliminated, because this reduces background and the amount of excess salt deposited in the ethanol-precipitation pellets. Finally, the concentration of MgSO4 has been increased, because this increases signal strength. These conditions may need to be adjusted in specific cases, but so far they have worked with all combinations of primers tested. The first-strand synthesis buffer differs slightly from the amplification/labeling buffer (i.e., there is less Tris and no Triton X-100). This change facilitates the conversion of first-strand buffer to ligation buffer without affecting polymerase activity for the single round of extension. Vent DNA polymerase is added last to each sample and then only after the sample has been chilled on ice. This is to minimize the singleand double-stranded exonuclease activities reported by the manufacturer that are inhibited by cold and by the presence of dNTPs. Efficiency and statistical limitations (founder effects) In this procedure, exponential amplification is so effective that the minimum amount of starting DNA needed is not dictated by signal strength; instead statistical considerations become important. Conventional PCR can be successfully performed using the DNA of a single genome (White et al., 1989). Obviously, footprinting or sequencing reactions require much more material than this, but determining exactly how much more is needed is not straight-



forward (UNITS 7.5 & 12.4). It is useful to first estimate the number of starting genomes or founder molecules that would be required in an idealized case in which all steps in the procedure were 100% efficient. To generate a proper sequence ladder, all “rungs” must be represented at least once (i.e., by at least one starting chromosome). Limited cleavage is used to generate DNA fragments that are, on average, ∼600 bases long (UNITS 7.5 & 12.4), and each of these can be used to generate a rung on the ladder. In the specific case of a DMS/piperidine guanine ladder, there are ∼150 possible cleavage sites (guanine residues) in an average 600-nucleotide fragment. Therefore, ≥150 chromosomes are required to generate all of the rungs in a ladder. However, because cleavage is random, ≥10 representatives (on average) must be generated to ensure that each rung is derived at least once. Furthermore, in footprinting, quantitative differences in the intensity of individual bands is important, and the maximum acceptable variation in band intensity due to sampling considerations should be 37°C (J. McCoy and P. Schendel, unpub. observ.). E. coli can synthesize proteins at temperatures ranging from 10° to 43°C, so trying expression at different temperatures is often worthwhile.



2. Change fermentation conditions. Many proteins contain metals as structural and catalytic cofactors. If the protein is being made faster than metals can be transported into the cell, the apoprotein without its metal cofactor will accumulate. This apoprotein will not fold correctly and will likely be insoluble. At the very least, the average specific activity of the expressed protein will be lower than expected. Different media and metal supplements can be tested and the best combination used. Clearly, if there is information about the metal content of the protein, these supplements can be designed more rationally. If no information is available, a more random approach must be tried. 3. Alter the rate of expression by using low-copy-number plasmids. This can be done by using the pACYC family (Chang and Cohen, 1978) or using single-copy chromosomal inserts of the cloned gene into a suitable target gene (Hamilton et al., 1989). Such reductions in gene dosage often reduce the final yield of protein, but the slower kinetics of synthesis they afford can sometimes result in production of soluble proteins. To restate the obvious, protein expression is an inexact science at present. However, most proteins can be made in E. coli in a form that is useful for a variety of functions. The procedures employed are relatively quick and uncomplicated, and the rewards for success are great.

Literature Cited

de Boer, H.A., Comstock, L.J., and Vasser, M. 1983. The tac promoter: A functional hybrid derived from the trp and lac promoters. Proc. Nat. Acad. Sci. U.S.A. 80:21-25. DeLamarter, J.F., Mermod, J.J., Liang, C.M., Eliason, J.F., and Thatcher, D.R. 1985. Recombinant murine GM-CSF from E. coli has biological activity and is neutralized by a specific antiserum. EMBO J. 4:2575-2581. Edman, J.C., Hallewell, R.A., Valenzuela, P., Goodman, H.M., and Rutter, W.J. 1981. Synthesis of Hepatitis B surface and core antigens in E. coli. Nature 291:503-506. Hamilton, C.M. Aldea, M., Washburn, B.K., Babitzke, P., and Kushner, S.R. 1989. New methods for generating deletions and gene replacem en ts in Escherichia coli. J. Bacteriol. 171:4617-4622. Marston, F.A.O. and Hartley, D.L. 1990. Solubilization of protein aggregates. Methods Enzymol. 182:264-276. Robinson, M., Lilley, R., Little, S., Emtage, J.S., Yarranton, G., Stephens, P., Millican, A., Eaton, M., and Humphreys, G. 1984. Codon usage can affect efficiency of translation of genes in Escherichia coli. Nucl. Acids Res. 12:6663-6671. Schein, C.H. 1989. Production of soluble recombinant proteins in bacteria. Bio/Technology 7:1141-1148. Schein, C.H. and Noteborn, M.H.M. 1988. Formation of soluble recombinant proteins in Escherichia coli is favored by lower growth temperatures. Bio/Technology 6:291-294.

Contributed by Paul F. Schendel Genetics Institute Cambridge, Massachusetts

Chang, A.C.Y. and Cohen, S.N. 1978. Construction and characterization of amplifiable multicopy DNA cloning vehicles derived from the P15A cryptic miniplasmid. J. Bacteriol. 134:11411156.

Protein Expression


Supplement 28

Expression Using the T7 RNA Polymerase/Promoter System

UNIT 16.2

This unit describes the expression of genes by placing them under the control of the bacteriophage T7 RNA polymerase. This approach has a number of advantages compared to approaches that rely on E. coli RNA polymerase. First, T7 RNA polymerase is a very active enzyme: it synthesizes RNA at a rate several times that of E. coli RNA polymerase and it terminates transcription less frequently; in fact, its transcription can circumnavigate a plasmid, resulting in RNA several times the plasmid length in size. Second, T7 RNA polymerase is highly selective for initiation at its own promoter sequences and it does not initiate transcription from any sequences on E. coli DNA. Finally, T7 RNA polymerase is resistant to antibiotics such as rifampicin that inhibit E. coli RNA polymerase, and consequently, the addition of rifampicin to cells that are producing T7 RNA polymerase results in the exclusive expression of genes under the control of a T7 RNA polymerase promoter (hereafter referred to as pT7). To use the two-plasmid pT7 system, it is necessary to clone the gene to be expressed into a plasmid containing a promoter recognized by the T7 RNA polymerase. The gene is then expressed by induction of T7 RNA polymerase. The gene for T7 RNA polymerase is present on a second DNA construction. This second construction can either permanently reside within the E. coli cell (basic protocol), or can be introduced into the cell at the time of induction by infection with a specialized phage, such as an M13 vector (mGP1-2; Tabor and Richardson, 1987) or a λ vector (CE6; Studier et al., 1990) containing the T7 RNA polymerase gene (second alternate protocol). In the basic protocol, two plasmids are maintained within the same E. coli cell. One (the expression vector) contains pT7 upstream of the gene to be expressed. The second contains the T7 RNA polymerase gene under the control of a heat-inducible E. coli promoter. Upon heat induction, the T7 RNA polymerase is produced and initiates transcription on the expression vector, resulting in turn in the expression of the gene(s) under the control of pT7. If desired, the gene products can be uniquely labeled by carrying out the procedure in minimal medium, adding rifampicin to inhibit the E. coli RNA polymerase, and then labeling the proteins with [35S]methionine (first alternate protocol). EXPRESSION USING THE TWO-PLASMID SYSTEM The gene to be induced is subcloned into an expression vector containing pT7. Two series of vectors have been developed for this purpose—the pT7 series (Fig. 16.2.1) and the pET series (Studier et al., 1990); see commentary for discussion of choice of vector. The plasmid containing the introduced gene is then used to transform an E. coli strain already containing the plasmid pGP1-2 (Fig. 16.2.2). pGP1-2 contains the gene for T7 RNA polymerase under the control of the λ pL promoter that is repressed by a temperature-sensitive repressor (cI857). pGP1-2 contains a p15A origin of replication that is compatible with the ColE1 origin of replication on the expression vector. The two plasmids are maintained in the same cell by selection with kanamycin (pGP1-2) and ampicillin (the expression vector). Cells containing the two plasmids are grown for several hours at 30°C and then the gene for T7 RNA polymerase is induced by raising the temperature to 42°C. The production of T7 RNA polymerase in turn induces expression of the genes under the control of pT7. (Rifampicin can be subsequently added to inhibit transcription by E. coli RNA polymerase, although this is usually not necessary since T7 RNA polymerase becomes responsible for most of the transcription even in the absence of rifampicin.) After expression Contributed by Stanley Tabor Current Protocols in Molecular Biology (1990) 16.2.1-16.2.11 Copyright © 2000 by John Wiley & Sons, Inc.

BASIC PROTOCOL

Protein Expression


Hin cll

Hincll

Ap

r

Ap

pT7-5 2400 bp Clal Hindlll Pstl Sall Xbal BamHl Smal Sacl EcoRl

ColE1 ori Pvu ll p T7

r

pT7-6 2210 bp

ColE1 ori Pvu ll p T7

EcoRl Sac l Smal BamHl Xbal Sal l Pst l Hindlll

Hin cll

Ap

r

pT7-7 2470 bp ColE1 ori Bgl ll p T7

ATG rbs Xba l

Clal Hindlll Pstl Sall Xbal BamHl Smal EcoRl Ndel p T7

T7 (22,857) CGATTCGAACTTCTCGATTCGAACTTCTGATAGACTTCGAAATTAATACGACTCACTATAGGGAGA

Met Ala Arg lle CCACAACGGTTTCCCTCTAGAAATAATTTTGTTTAACTTTAAGAAGGAGATATACAT ATG GCT AGA ATT rbs

Xbal

Ndel

EcoRl T7 (22,972)

Arg Ala Arg Gly Ser Ser Arg Val Asp Leu Gln Pro Lys Leu lle lle Asp... CGC GCC CGG GGA TCC TCT AGA GTC GAC CTG CAG CCC AAG CTT ATC ATC GAT... Smal

BamHl

Xbal

Sall

Pstl

Hindlll

Clal

Figure 16.2.1 pT7-5, pT7-6, and pT7-7. pT7-5, pT7-6, and pT7-7 are cloning vectors that contain a T7 promoter and are used to express genes using T7 RNA polymerase. All three vectors contain a T7 RNA polymerase promoter, the gene encoding resistance to the antibiotic ampicillin and the ColE1 origin of replication. pT7-7 has a strong ribosome-binding site (rbs) and start codon (ATG) upstream of the polylinker sequence; the sequence of this region is shown below the map of pT7-7. pT7-5 and pT7-6 lack any ribosome-binding site upstream of the polylinker sequence and consequently are only useful when expressing genes that already contain the proper control sequences. pT7-5, pT7-6, and pT7-7 were constructed by S. Tabor and are derivatives of pT7-1 described in Tabor and Richardson (1985).

Expression Using the T7 RNA Polymerase/ Promoter System



of the genes at 37°C, the cells are harvested and the induced proteins are analyzed. An alternative approach is to induce T7 RNA polymerase with IPTG rather than by heat induction. In this method, the expression plasmid containing pT7 can be placed into E. coli BL21 (DE3), which contains the gene for T7 RNA polymerase on the E. coli chromosome under the control of the lac promoter (Studier and Moffatt, 1986; Studier et al., 1990). Materials pT7-5, pT7-6, or pT7-7 vectors (available from author) E. coli JM105, DH1, or equivalent (Table 1.4.5) LB plates and medium containing 60 µg/ml ampicillin (UNIT 1.1) E. coli K38 or equivalent (Table 1.4.5) pGP1-2 (available from author) LB plates and medium containing 60 µg/ml kanamycin (UNIT 1.1) LB plates and medium containing 60 µg/ml ampicillin plus 60 µg/ml kanamycin (UNIT 1.1) Cracking buffer Sorvall SS-34 or GS-3 rotor or equivalent Additional reagents and equipment for subcloning DNA fragments (UNITS 1.4 & 3.16), transformation of competent E. coli cells (UNIT 1.8), minipreps of plasmid DNA (UNIT 1.6), restriction mapping (UNITS 3.1-3.3), and SDS-PAGE (UNIT 10.2). 1. Subclone the fragment containing the gene to be expressed into pT7-5, pT7-6, or pT7-7. Transform a standard E. coli strain (e.g., JM105 or DH1); this strain should not carry a plasmid that directs synthesis of T7 RNA polymerase (i.e., pGP1-2). Plate the transformants on LB/ampicillin plates and grow overnight at 37°C. λ (35,259) T7 (3106)

pL

λ (35,711) Smal Sacl EcoRl

plac

T7 RNA polymerase

c1857 pGP1-2 7140 bp

T7 (5840) BamHI kanr

p15A ori

Pst1

Hindlll Smal

Xhol

Figure 16.2.2 pGP1-2. pGP1-2 enables T7 RNA polymerase to be produced by heat induction in any E. coli host. pGP1-2 contains the gene for T7 RNA polymerase under the control of the λ pL promoter. It also contains the gene for the λ repressor (cI857) that is expressed under the control of E. coli plac promoter. This repressor inhibits transcription from the λ pL promoter at low temperature (30°C); however, at high temperature (42°C) it is inactivated, resulting in induction of the pL promoter, that in turn results in induction of the T7 RNA polymerase. pGP1-2 also contains the gene encoding resistance to the antibiotic kanamycin, and the p15A origin of replication. pGP1-2 is described in Tabor and Richardson (1985).

Protein Expression


Supplement 11

It is important to first transform the plasmid into a strain that contains no T7 RNA polymerase, in case small amounts of the gene product are toxic to the cell (see critical parameters for discussion on toxic genes).

2. Grow individual transformants in LB/ampicillin medium at 37°C and obtain plasmid DNA by a miniprep procedure. Confirm that the gene has been correctly inserted by restriction mapping. 3. Transform E. coli K38 with pGP1-2, plate on LB/kanamycin plates, and grow overnight at 30°C. Grow an individual E. coli K38/pGP1-2 transformant in LB/ kanamycin medium at 30°C. Colonies take ∼24 hr to appear on plates at 30°C. E. coli K38/pGP1-2 can be stored in the absence of the plasmid containing pT7 as a glycerol stock at −80°C (see commentary).

4. Transform the vector containing the gene to be expressed under the control of pT7 into E. coli K38/pGP1-2 grown in LB/kanamycin medium. Plate the transformants (containing both plasmids) on LB/ampicillin/kanamycin plates and grow overnight at 30°C. Cells may be heat-shocked during transformation; the T7 RNA polymerase gene, under the control of a heat-inducible promoter, is not induced by this brief heating step. As a control, transform E. coli K38/pGP1-2 with the parent pT7 vector (without an insert). If the transformation efficiency of the vector containing the insert is significantly lower (by more than a factor of 50) than that of the parent vector, the gene product may be toxic to E. coli cells. This toxicity arises from background expression of the gene product by basal levels of T7 RNA polymerase. In this situation, the transformants that do arise invariably contain deletions or other mutations in one of the two plasmids, and the desired gene product will not be produced. If the expression of the inserted gene is toxic, it is necessary to use an alternative strategy for the repression and induction of the T7 RNA polymerase gene (see discussion on toxic genes in critical parameters).

5. Pick a single E. coli colony that contains the two plasmids with a sterile toothpick or pipet. Inoculate it into 5 ml LB/ampicillin/kanamycin medium and grow overnight at 30°C. 6. Dilute the overnight culture of cells 1:40 into fresh LB/ampicillin/kanamycin medium and grow several hours at 30°C to an OD590 ≅ 0.4. The size of the culture will depend on the amount of cells needed. For an analytical preparation, use ∼1 ml of cells.

7. Induce the gene for T7 RNA polymerase by raising the temperature to 42°C for 30 min, which in turn induces the genes under the control of pT7. To obtain consistent results, raise the temperature relatively quickly. If small cultures (∼1 ml) are being induced, place the cultures into a 42°C water bath. For larger cultures (∼500 ml), place the flask under hot tap water until the temperature of the media reaches 42°C (measured by inserting a thermometer wiped with ethanol into the flask). Once the cells reach 42°C, continue incubating at 42°C for 30 min.


The E. coli RNA polymerase can be inhibited by adding rifampicin to a final concentration of 200 ìg/ml; when used, it should be added after T7 RNA polymerase has been induced at 42°C for 30 min. Although rifampicin reduces the background of host proteins being expressed, in general it does not significantly increase the final accumulation of gene products, and in some cases it decreases the final yield. Thus, as a general rule, rifampicin is only added to cells when the plasmid-encoded proteins are being uniquely labeled with [35S]methionine (see first alternate protocol).



8. Reduce temperature to 37°C and grow the cells an additional 90 min with shaking. 9. Harvest the cells by centrifuging and discarding the supernatant. For 1-ml cultures, microcentrifuge 20 sec at 10,000 rpm (14,000 × g), room temperature. For 2-ml to 100-ml cultures, centrifuge 5 min in a Sorvall SS-34 rotor at 5000 rpm (3000 × g), 4°C. For >100-ml cultures, centrifuge 10 min in a Sorvall GS-3 rotor at 5000 rpm (4000 × g), 4°C. 10. To analyze the induced proteins by SDS-PAGE, resuspend the equivalent of 1.0 ml of cells in 0.1 ml cracking buffer. Heat at 100°C for 5 min immediately prior to loading a 20-µl aliquot of each sample onto an SDS-polyacrylamide gel (UNIT 10.2). To analyze the cells for an induced enzymatic activity, prepare an appropriate cell extract from ∼10 ml of cells. One example of the preparation of an extract for the purification of T7 RNA polymerase is described in Tabor and Richardson (1985).

SELECTIVE LABELING OF PLASMID-ENCODED PROTEINS Plasmid-encoded proteins under the control of a pT7 (see basic protocol) can be exclusively labeled by inducing the T7 RNA polymerase in cells growing in minimal medium, inhibiting the host E. coli RNA polymerase with rifampicin, and labeling the newly synthesized proteins with [35S]methionine. This procedure provides an attractive alternative to maxicells or minicells for labeling of plasmid-encoded proteins (Dougan and Sherratt, 1977; Sancar et al., 1981).

ALTERNATE PROTOCOL

Additional Materials M9 medium (UNIT 1.1) without and with 5% (v/v) of 18 amino acid mixture 20 mg/ml rifampicin in methanol (e.g., Sigma #R-3501; store in dark at 4°C for 2 weeks; Table 1.4.1) 10 mCi/ml [35S]methionine (>800 Ci/mmol) diluted 1:10 in M9 medium Fluorographic enhancing agent (e.g., Enlightning from Du Pont NEN or Amplify from Amersham) 1. Repeat steps 3 to 6 of the basic protocol (using the T7-promoter expression plasmid obtained from steps 1 and 2 of the basic protocol). An alternative to the use of LB/ampicillin/kanamicin medium for growing cells is M9 medium containing 25 ìg/ml ampicillin and 25 ìg/ml kanamycin, and any required nutrients. The addition of one part in twenty of the 18 amino acid mixture (0.1% stock, 0.005% final concentration) stimulates the growth of cells in M9 medium without interfering with the subsequent labeling of the proteins with [35S]methionine. Note that to grow in this medium, the E. coli strain must be Cys+ and Met+.

2. When OD590 ≅ 0.4, remove 1 ml of cells, microcentrifuge 10 sec, and discard supernatant. 3. Wash cell pellet with 1 ml M9 medium, microcentrifuge 10 sec at room temperature, and discard supernatant. Washing the cells after growth in LB medium is very important in order to remove the unlabeled methionine present in LB medium that otherwise dilutes the [35S]methionine during labeling.

4. Resuspend cell pellet in 1 ml M9 medium containing 18 amino acid mixture. Grow cells 60 min at 30°C with shaking. A time of 30 to 180 min is adequate for adapting cells to M9 medium. Although the OD590 may not increase significantly during this step, induction of T7 RNA polymerase and

Protein Expression


Supplement 11

efficient labeling of the plasmid-encoded proteins will occur even in the absence of apparent cell growth.

5. Induce the gene for T7 RNA polymerase by placing the cells in a 42°C water bath for 20 min. 6. Add 20 mg/ml rifampicin to 200 µg/ml final. Keep cells at 42°C for an additional 10 min after adding rifampicin. It is important to incubate the cells at 42°C for an additional 10 min after adding rifampicin, since rifampicin is more effective at inhibiting expression of host proteins at 42°C, possibly because the cells are more permeable to it at this temperature. The temperature of the cells is subsequently reduced for the labeling since in general the labeling is less efficient at 42°C than at 30° or 37°C.

7. Shift cells to a 30°C water bath for an additional 20 min. Remove 0.5 ml of cells for labeling with [35S]methionine. The other 0.5 ml can be used to label the cells at a later time point (e.g., after an additional 30 min) in order to follow the duration of protein synthesis.

8. Label newly synthesized proteins by adding 10 µl (10 µCi) diluted [35S]methionine to 0.5 ml of cells and incubating for 5 min at 30°C. 9. Microcentrifuge cells 10 sec and discard supernatant. (CAUTION: the supernatant is radioactive; discard properly.) Resuspend cell pellet in 100 µl cracking buffer. 10. Heat samples to 100°C for 5 min. Load a 20-µl aliquot onto an SDS-polyacrylamide gel and electrophorese (UNIT 10.2). 11. Treat the gel with a fluorographic-enhancing agent by soaking it in the fluor for 30 min. Dry the gel under vacuum 2 hr at 65°C and autoradiograph (APPENDIX 3). A 1-hr exposure should be adequate to visualize most proteins induced with this system. To determine whether the plasmid-encoded proteins are susceptible to proteases in the E. coli cell, prepare and induce the cells as described above; however, reduce the duration of the labeling step to 1 min (step 8), and follow this with a chase of nonradioactive methionine at 0.5% final concentration. Remove an aliquot for analysis both immediately prior to the chase, and after a chase reaction of 5, 15, and 60 min. After removing each aliquot, immediately pellet the cells by centrifugation, resuspend in cracking buffer, and heat the aliquot to 100°C for 5 min to inactivate the proteases. Analyze as in step 10. ALTERNATE PROTOCOL


EXPRESSION BY INFECTION WITH M13 PHAGE mGP1-2 Whenever the gene for T7 RNA polymerase is present in E. coli cells, low levels of T7 RNA polymerase are constitutively produced. This can be a problem when the gene products under the control of pT7 are toxic. One strategy to avoid this is to keep the gene for T7 RNA polymerase out of the cell until the time of induction. In the protocol presented here, T7 RNA polymerase is introduced into the cell by infection with the M13 phage mGP1-2. This phage contains the gene for T7 RNA polymerase under the control of the lac promoter (Fig. 16.2.3). Host cells for this phage must carry the F factor so that they are susceptible to M13 infection (e.g., JM101 or K38). The cells are transformed with the single plasmid that contains the gene to be expressed under the control of pT7. The cells are grown at 37°C, and induction occurs by infection with a high multiplicity of mGP1-2 in the presence of IPTG. A λ vector, CE6, that contains the gene for T7 RNA polymerase has also been used to express toxic genes (Studier and Moffatt, 1986; Studier et al., 1990).



Additional Materials M13 phage mGP1-2 (available from author) PEG solution (UNIT 1.7) 100 mM IPTG (Table 1.4.2) Additional reagents and equipment for preparing M13 phage (UNIT 1.15) and titering phage (UNIT 1.11) 1. Prepare a stock of M13 phage mGP1-2 and concentrate the phage by precipitation with PEG solution. (DO NOT proceed to add TE buffer or phenol.) Resuspend phage in M9 medium and titer. If the cell proteins are to be labeled, it is important that the phage used to infect the cells are free of unlabeled methionine. In this case, precipitate the phage with PEG twice, each time resuspending the pellet in M9 medium. For long-term storage of the M13 phage mGP1-2, it is best to purify the phage through a CsCl gradient (Nakai and Richardson, 1986).

2. Transform E. coli cells susceptible to M13 infection (e.g., JM101 or K38) with the T7-promoter expression plasmid obtained from steps 1 and 2 of the basic protocol. Plate the transformants on LB/ampicillin plates and grow overnight at 37°C. 3. Pick a single colony and grow in LB/ampicillin medium overnight at 37°C. 4. Dilute the overnight culture of cells 1:100 in LB/ampicillin medium and grow several hours at 37°C with gentle shaking to OD590 ≅ 0.5. It is very important that only gentle shaking is used when growing cells for M13 infection. Vigorous agitation results in shearing of the pili on the surface of the E. coli cells, resulting in inefficient infection.

5. Infect cells with M13 phage mGP1-2 (from step 1) at a ratio of ∼10 phage for each E. coli cell. Add 100 mM IPTG to 1 mM final (a 1:100 dilution) to induce production

T7 (5840) Sall Pst1 Hindlll T7 RNA polymerase

T7 (3133) mGP1-2 9960 bp

Smal Sac l EcoRI

plac

M13 ori

Figure 16.2.3 mGP1-2. M13 phage mGP1-2 contains the gene for T7 RNA polymerase under the control of the E. coli plac promoter. It is especially useful for the production of gene products that are toxic to the E. coli cell. When E. coli cells are infected with this phage, and IPTG is added to induce the plac promoter, T7 RNA polymerase is produced. As a result, any genes within the cell under the control of pT7 will be induced. mGP1-2 is described in Tabor and Richardson (1987).

Protein Expression


Supplement 11

of T7 RNA polymerase. Incubate the cells 2 hr at 37°C. At OD590 ≅ 0.5, the density of E. coli cells will be ∼2 × 108 cells/ml. Thus, it is necessary to add M13 mGP1-2 phage at a final concentration of 2 × 109 phage/ml to obtain a multiplicity of infection of 10. Small cultures (∼50 ml) can be incubated in a water bath without shaking. Larger cultures should be incubated at 37°C with gentle shaking.

6. Harvest cells and analyze induced proteins as in steps 9 and 10 of the basic protocol. REAGENTS AND SOLUTIONS 18 amino acid mixture Prepare a solution containing 0.1% (v/v) of each amino acid except cysteine (minus cysteine) and methionine (minus methionine). Filter sterilize through a 0.2-µm filter. Store at −20°C for several years. Cracking buffer 60 mM Tris⋅Cl, pH 6.8 1% 2-mercaptoethanol 1% sodium dodecyl sulfate (SDS) 10% glycerol 0.01% bromphenol blue COMMENTARY Background Information

Critical Parameters

Bacteriophage T7 and T7-related phage (e.g., SP6, T3) encode their own RNA polymerase (see UNIT 3.8). Compared to other known RNA polymerases, this RNA polymerase is both relatively simple and highly efficient. T7 RNA polymerase is a single polypeptide of 96,000 kDa. It initiates transcription specifically at a 23-nucleotide promoter sequence, a sequence not present on the E. coli genome. Transcription is very processive, producing transcripts that are many thousands of nucleotides in length. Transcription is relatively rapid—five times the rate of E. coli RNA polymerase. All of these properties make T7 RNA polymerase and its promoter an attractive system for controlling the expression of foreign genes in E. coli and in other organisms. Expression systems in E. coli based on the controlled induction of T7 RNA polymerase have been developed by Tabor and Richardson (1985) and Studier and his colleagues (Studier and Moffatt, 1986; Rosenberg et al., 1987; Studier et al., 1990). The vectors described here are those developed by Tabor and Richardson. T7 RNA polymerase/promoter expression systems have also been successfully applied in yeast (Chen et al., 1987) and mammalian cells (Dunn et al., 1988; Fuerst et al., 1986).

Choice of vector Questions that determine what vector to use to express a gene using T7 RNA polymerase include: Is there a ribosome-binding site upstream of the gene? What are the restriction sites available on each end of the gene? Is the gene product toxic to the E. coli cell? Examples of three standard vectors (pT7-5, pT7-6, and pT7-7) are shown in Figure 16.2.1. These vectors are derivatives of pBR322. The β-lactamase gene encoding ampr is in the opposite orientation of pT7; consequently the only plasmid-encoded genes expressed by T7 RNA polymerase are those cloned into the polylinker region. pT7-5 and pT7-6 contain the polylinker region located immediately downstream of pT7 in opposite orientations. There is no ribosomebinding sequence in these two plasmids; they should thus be used either for the production of transcripts without expectation of good translation of the protein, or for the expression of genes that already have strong ribosome-binding sequences. pT7-7 differs from pT7-5 and pT7-6 in that it contains a strong ribosome-binding sequence between pT7 and the polylinker region; it is recommended for the expression of genes that lack a strong ribosome binding sequence or for the production of fusion proteins.




An extensive series of additional vectors containing pT7, the pET series, have been described by Studier et al. (1990). These vectors are particularly useful for applications that require a greater selection of restriction endonuclease sites to insert the gene into, or that involve the expression of a gene that is toxic to the cell (see below). Some of these vectors contain other transcriptional regulatory elements (i.e., terminators, operators, RNase III cleavage sites) that could be of use for specific applications. A large number of commercially available vectors contain a T7 RNA promoter (e.g., pIBI vectors, available from IBI; pSP6/T7-19, available from GIBCO/BRL; pBluescript II vectors, available from Stratagene; and pTZ18R and pTZ19R, available from U.S. Biochemical). These are intended to be used for producing specific transcripts in vitro using T7 RNA polymerase. In principle, they should be useful for the expression of genes using T7 RNA polymerase in vivo as well. In practice, however, the use of some of these vectors can result in some unexpected problems. (1) Most commercial vectors have extremely high copy numbers within the cell; this can accentuate the problems encountered with toxic genes. (2) In most vectors, the β-lactamase gene is oriented in the same direction as pT7, complicating the analysis of radiolabeled proteins. (3) Some commercial vectors have pT7 oriented in a potentially deleterious direction. Derivatives of pBR322 that contain pT7 oriented clockwise with respect to the standard map are inviable in some E. coli strains that contain the gene for T7 RNA polymerase. This is due to the fact that high levels of transcription through the origin region of these plasmids in this orientation interferes with the replication of the plasmids. (4) Most commercial vectors have a lac operator sequence within them. This can titrate out the lac repressor (UNIT 1.4) and cause problems when the plac is used to control the T7 RNA polymerase gene. Toxic genes In some cases the gene to be expressed is toxic to the cells, even when it is not induced. This is due to a low level of constitutive expression present even under uninduced conditions. Although most genes are not toxic when expressed using the two-plasmid pT7 system, it is important to recognize the symptoms of toxicity to avoid selecting for mutations and to allow alternate systems for induction to be tried. The degree of toxicity varies greatly with each gene.

The symptoms encountered with toxic genes are discussed below, in order of increasing toxicity. Some genes are mildly toxic to the cells when expressed using the two-plasmid pT7 system. In such cases, the cells can be stably transformed with the two plasmids and the gene product is produced at a high level. However, after the cells are several days old, they no longer induce the expected gene product even though they remain resistant to ampicillin and kanamycin. To avoid this problem, it is recommended that the E. coli K38/pGP1-2 be stored in the absence of the plasmid containing pT7 as a glycerol stock at −80°C (UNIT 1.3). The plasmid containing pT7 and the gene to be expressed should be stored as DNA at −20°C or −70°C (UNIT 1.6). To prepare the strain for induction, streak K38/pGP1-2 on an LB/kanamycin plate at 30°C, grow up a single colony, transform with the plasmid containing pT7 and the gene to be expressed, and plate the transformants on LB/ampicillin/kanamycin plates at 30°C. A single colony should then be grown at 30°C and induced as described above. This procedure is not necessary for genes that are not toxic. Strains that do not induce toxic genes can be stored in glycerol at −80°C for many months (UNIT 1.3). A more toxic class of genes consists of those that can be successfully cloned into a plasmid under the control of pT7, but that render the resulting plasmid unable to stably transform a cell that contains the gene for T7 RNA polymerase. Genes that are toxic to the cells only in the presence of pGP1-2 (which expresses the T7 RNA polymerase) are relatively common, occurring on the average ∼5% of the time (S. Tabor, unpublished observation). Note that such plasmids will give transformants in E. coli cells containing pGP1-2, but that the frequency of transformation will be greatly reduced (>50fold) compared to the frequency of transformation by the parent vector alone. The cells that do grow in the presence of ampicillin and kanamycin will invariably contain deletions or other mutations in one of the two plasmids, and the desired gene product will not be produced. When genes are toxic at this level, it is necessary to use an alternative strategy that reduces the expression of the gene under uninduced conditions. One strategy is to remove the gene for T7 RNA polymerase from the cell until induction is desired, and then introduce it by a phage infection. Such an alternate protocol is described using an M13 phage harboring the gene for T7 polymerase, mGP1-2. A lambda

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vector, CE6, that contains the gene for T7 RNA polymerase has also been used for this purpose (Studier and Moffatt, 1986; Studier et al., 1990). Another strategy is to retain the gene for T7 RNA polymerase in the cell but reduce the level of transcription by T7 RNA polymerase under uninduced conditions. For example, a system has been developed that expresses an inhibitor of T7 RNA polymerase—the T7 lysozyme—to reduce the activity of T7 RNA polymerase until it is induced (Studier et al., 1990). Another recent modification is the placement of pT7 under the control of the lac repressor, reducing the activity of T7 RNA polymerase until IPTG has been added (Studier et al., 1990). Finally, some genes are difficult to clone in multicopy plasmids even in the absence of a known E. coli promoter. The difficulty in cloning these genes arises from the fact that their products are extremely toxic and that the residual low level of transcription by E. coli RNA polymerase in most plasmids is sufficient to direct the synthesis of small amounts of these proteins. One strategy that can be used to clone such toxic genes is to insert the gene near a strong E. coli promoter that is oriented so that transcription by the E. coli RNA polymerase results in the accumulation of RNA that is antisense to the toxic gene, reducing the level of its gene product. It is important to remember that the amount of a gene product synthesized is a function not only of the level of transcription but also of the efficiency at which translation is initiated. This is determined primarily by the ribosomebinding sequence located upstream of the start codon. Thus, some toxic genes with relatively weak ribosome-binding sequences can be cloned into multicopy plasmids, but not into a multicopy plasmid that also introduces a strong ribosome-binding sequence (S. Tabor, unpublished observations). In summary, the first step in using the T7 RNA polymerase/promoter system is to clone the gene into an appropriate vector containing a pT7 and be certain it has an efficient ribosomebinding sequence. Once this is accomplished, the next step is determining whether the plasmid can stably transform an E. coli cell containing pGP1-2 at an efficiency comparable to that of the parent vector alone. If this is successful, the system is ready to be induced. If unsuccessful, it is necessary to induce the gene either by infection with M13 phage mGP1-2

(see second alternate protocol), or to use one of the more specialized vectors that further reduce the expression of T7 RNA polymerase in the cell under uninduced conditions (Studier et al., 1990).

Troubleshooting For gene expression, one of the major advantages of the T7 RNA polymerase/promoter system over an E. coli RNA polymerase system is the ability to exclusively label the gene products under the control of pT7. If the level of induction of the gene is estimated by inspection of a standard SDS-polyacrylamide gel, and it is difficult to see the expected induced product, then it is recommended that the induced proteins be labeled using [35S]methionine as described in the first alternate protocol. This is a much more sensitive and specific assay for the specific protein production. Be sure that there is at least one methionine codon in the gene other than the one at the start of the protein (which is often removed in E. coli; Kirel et al., 1989); if not, then it is necessary to label with a cysteine or some other amino acid. If it is not possible to detect the expected labeled product, there may be a problem with one of the two plasmids in the cell. One possibility is that the expressed protein is toxic to the cell, and as a result, a mutation has been selected for such that the toxic product is not synthesized. For more information on determining whether a gene is toxic, see the discussion on toxic genes in critical parameters. To determine if the cells and T7 RNA polymerase gene (e.g., pGP1-2) are inducing T7 RNA polymerase, attempt to induce a control protein that has been shown to work well in this system (e.g., the β-lactamase gene in pT7-1; Tabor and Richardson, 1985). If the expressed protein does not accumulate significantly after induction, determine its stability in E. coli cells by pulse labeling with [35S]methionine and chasing for various time periods with unlabeled methionine. If it is rapidly degraded, try to induce the gene in a protease-deficient strain. It should be noted that there are no known mutations that inactivate several very active E. coli proteases, and thus there is a strong probability that the mutant strains available (e.g., lon−) will have no effect on the stability of the gene product. In addition, such mutant strains generally grow poorly, and as a consequence the gene products are poorly produced upon induction of T7 RNA polymerase. The most common reason for poor induc-



tion of a gene is that the translation does not initiate efficiently. Therefore, it is very important that there be an efficient ribosome-binding sequence the proper distance upstream of the gene. If a gene product does not induce well, and the problem is not the stability of the product, try a different ribosome-binding sequence—one that is known to work efficiently. The sequence and spacing between the ribosome-binding sequence and the start codon is critical. Because of this, it is recommended that the gene be inserted into a vector such as pT7-7, without altering any of the sequences between the ribosome-binding sequence and the start codon.

Anticipated Results Under optimal conditions, the gene product expressed by the T7 RNA polymerase/ promoter system can accumulate to >25% of the total cellular protein. However, in most instances the amount of gene product that accumulates is significantly less than this. There are numerous reasons for poor yields of gene product, as discussed in troubleshooting (see above).

Time Considerations

It should take ∼1 week to insert the gene of interest into the pT7 vector, prepare minipreps of the DNA, and characterize the recombinants for the correct size and orientation of the insert. It should then take 3 days to transform the recombinant plasmid into the E. coli strain containing pGP1-2, induce the cells, and test the extracts for the production of the expected gene product.

Literature Cited Chen, W., Tabor, S., and Struhl, K. 1987. Distinguishing between mechanisms of eukaryotic transcriptional activation with bacteriophage T7 RNA polymerase. Cell 50:1047-1055. Dougan, G. and Sherratt, D. 1977. The transposon Tn1 as a probe for studying ColE1 structure and function. Mol. Gen. Genet. 151:151-160. Dunn, J.J., Krippl, B., Bernstein, K.E., Westphal, H., and Studier, F.W. 1988. Targeting bacteriophage T7 RNA polymerase to the mammalian cell nucleus. Gene 68:259-266. Fuerst, T.R., Niles, E.G., Studier, F.W., and Moss, B. 1986. Eukaryotic transient-expression system based on recombinant vaccinia virus that synthe-

sizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 83:8122-8126. Kirel, P.-H, Schmitter, J.-M., Dessen, P., Fayat, G., and Blanquet, S. 1989. Extent of N-terminal methionine excision from Escherichia coli proteins is governed by the side-chain length of the penultimate amino acid. Proc. Natl. Acad. Sci. U.S.A. 86:8247-8251. Nakai, H. and Richardson, C.C. 1986. Interactions of the DNA polymerase and gene 4 protein of bacteriophage T7. J. Biol. Chem. 261:1520815216. Rosenberg, A.H., Lade, B.N., Chui, D., Lin, S., Dunn, J.J., and Studier, F.W. 1987. Vectors for selective expression of cloned DNAs by T7 RNA polymerase. Gene 56:125-135. Russel, M. and Model, P. 1984. Replacement of the flp gene of Escherichia coli by an inactive gene cloned on a plasmid. J. Bacteriol. 159:10341039. Sancar, A., Wharton, R.P., Seltzer, S., Kacinsky, B.M., Clark, N.D., and Rupp, W.D. 1981. Identification of the uvrA gene product. J. Mol. Biol. 184:45-62. Studier, F.W. and Moffatt, B.A. 1986. Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J. Mol. Biol. 189:113-130. Studier, F.W., Rosenberg, A.H., Dunn, J.J., and Dubendorff, J.W. 1990. Use of T7 RNA polymerase to direct the expression of cloned genes. Methods Enzymol. 185. In press. Tabor, S. and Richardson, C.C. 1985. A bacteriophage T7 RNA polymerase/promoter system for controlled exclusive expression of specific genes. Proc. Natl. Acad. Sci. U.S.A. 82:10741078. Tabor, S. and Richardson, C.C. 1987. DNA sequence analysis with a modified bacteriophage T7 DNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 84:4767-4771.

Key References Studier et al., 1990. See above. Gives extensive list of vectors and protocols for expression using T7 RNA polymerase. Tabor and Richardson, 1985. See above. Describes the use of the two-plasmid system for expression of genes using T7 RNA polymerase.


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UNIT 16.3

Expression Using Vectors with Phage ë Regulatory Sequences Many expression systems have been developed that utilize pBR322-based plasmids into which transcriptional and translational regulatory signals have been inserted. In the system described here, however, plasmids (pSKF) utilize regulatory signals—such as the powerful promoter pL—from the bacteriophage λ. Transcription from pL can be fully repressed and plasmids containing it are thus stabilized by the λ repressor, cI. The repressor is supplied by an E. coli host which contains an integrated copy of a portion of the λ genome. This so-called defective lysogen supplies the λ regulatory proteins cI and N but does not provide the lytic components that would normally lead to cell lysis. Thus, cells carrying these plasmids can be grown initially to high density without expression of the cloned gene and subsequently induced to synthesize the product upon inactivation of the repressor. This system also ensures that pL-directed transcription efficiently traverses any gene insert, which is accomplished by providing the phage λ antitermination function, N, to the cell and by including on the pL transcription unit a site necessary for N utilization (Nut site). The N protein interacts with and modifies the RNA polymerase at the Nut site so as to block transcription termination at distal sites in the transcription unit. In order to express the coding sequence, efficient ribosome-recognition and translationinitiation sites have been engineered into the pL transcription unit. Expression occurs after temperature or chemical induction inactivates the repressor (see first and second basic protocols). Restriction endonuclease sites for insertion of the desired gene have been introduced both upstream and downstream from an ATG initiation codon. Thus, the system allows either direct expression or indirect expression (via protein fusion) of any coding sequence, thereby potentially allowing expression of any gene insert. Direct expression generates “authentic” gene products (first support protocol), while expression of heterologous genes fused to highly expressed gene partners generates chimeric proteins that differ from the native form. In the latter case, the fusion partner can be removed to obtain an unfused version of the gene product (second support protocol).

BASIC PROTOCOL

TEMPERATURE INDUCTION OF GENE EXPRESSION Expression from pL-containing vectors can be induced by raising the temperature. The E. coli lysogens used with these vectors are typically defective for phage replication and carry a temperature-sensitive mutation in the phage λ cI gene (cI857). After transformation and growth, induction is accomplished by raising the temperature of the culture from 32° to 42°C. Materials Expression vector (e.g., pSKF series; see support protocols) E. coli AR58 or equivalent (Table 1.4.5) LB plates containing the appropriate antibiotic (UNIT 1.1) LB medium containing appropriate antibiotic (room temperature and prewarmed to 65°C; UNIT 1.1) SDS/sample buffer (UNIT 10.2) Gyrotory air or water shaker, 32° and 42°C Additional reagents and equipment for transformation (UNIT 1.8)

Expression Using Vectors with Phage ëRegulatory Sequences


1. Transform the expression vector into an E. coli λ lysogen (such as AR58) carrying a temperature-sensitive mutation in its repressor gene (λ cI857). Plate on LB/antibiotic plates and incubate transformants at 32°C. Contributed by Allan R. Shatzman, Mitchell S. Gross, and Martin Rosenberg Current Protocols in Molecular Biology (1994) 16.3.1-16.3.11 Copyright © 2000 by John Wiley & Sons, Inc.

Heat-shock at 37° or 42°C for ≤90 sec during transformation is not a problem.

2. Grow the transformed cells overnight at 32°C in LB/antibiotic medium. 3. Dilute the overnight culture ≥1:20 into fresh LB/antibiotic medium. Grow the culture at 32°C in a gyrotory shaker at 250 to 300 rpm until OD650 = 0.6 to 0.8. 4. Add 1⁄3 vol of 65°C LB/antibiotic medium with swirling in order to elevate the culture temperature rapidly to 42°C. In our experience, a rapid increase in temperature favors production. Small shake-flask cultures (≤25 ml) are more easily induced by transfer to a 42°C gyrotory water bath without addition of prewarmed media. This generally raises the culture temperature to 42°C within 3 to 5 min.

5. Continue growing the culture 2 to 3 hr at 42°C. 6. Remove a 1-ml aliquot for analysis and harvest the remainder of cells by centrifuging 15 min in a low-speed rotor at 3000 × g, 4°C. Discard the supernatant. Freeze cell pellet at −70°C until ready to isolate the gene product.

7. Spin the 1-ml aliquot 1 min at top speed in a microcentrifuge, then resuspend the pellet in 50 µl SDS/sample buffer. Boil 5 to 10 min and analyze gene product by SDS–polyacrylamide gel electrophoresis. CHEMICAL INDUCTION OF GENE EXPRESSION Expression using the pSKF system can also be induced chemically in lysogens that carry a wild-type (ind+) repressor gene (cI857 cannot be used as it is ind−). This is accomplished by treating the bacterial host with an agent such as nalidixic acid. Nalidixic acid inhibits DNA gyrase and leads to DNA damage, which induces the SOS response. During the SOS response, wild-type repressor protein is cleaved. In this case, the wild-type repressor protein is cleaved by the RecA protease, which is induced by the SOS response. In contrast to induction by heat (product accumulates in 45 to 90 min) nalidixic acid–mediated induction of protein expression is comparatively slow (product accumulates in 5 to 6 hr).

BASIC PROTOCOL

Materials Expression vector (e.g., pSKF series; see support protocols) E. coli AR120 or equivalent (Table 1.4.5) LB plates containing appropriate antibiotic (UNIT 1.1) LB medium containing appropriate antibiotic (UNIT 1.1) 60 mg/ml nalidixic acid in 1 N NaOH (not necessary to filter sterilize; Table 1.4.1) Additional reagents and equipment for transformation (UNIT 1.8) 1. Transform the expression vector into a replication-defective, E. coli cI+ lysogen (e.g., AR120). Plate on LB/antibiotic plates and incubate the transformants at 37°C. 2. Grow the transformed cells overnight at 37°C in LB/antibiotic medium. 3. Dilute the overnight culture ≥1:20 into fresh LB/antibiotic medium. Grow the culture at 37°C in a gyrotory shaker at 250 to 300 rpm until OD650 = 0.4. 4. Add 1/1000 vol of 60 mg/ml nalidixic acid solution to give 60 µg/ml final concentration. 5. Continue growing the culture 5 to 6 hr at 37°C. 6. Harvest cells and analyze gene product (steps 6 and 7 of first basic protocol).

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SUPPORT PROTOCOL

AUTHENTIC GENE CLONING USING pSKF VECTORS It is often most desirable to express a gene product in a form as similar to the native protein as possible. Such an “authentic” gene product will have the greatest chance of having a structure and activity identical to that of the native protein. Efficient translation of a coding sequence for an authentic gene product is typically accomplished by placing the inserted information immediately adjacent to a ribosome-binding site (a translational regulatory signal that interacts with the 16S rRNA of E. coli and contains an ATG initiation codon; Gold et al., 1981). Strategic Planning The translation-initiation signal utilized here is that of the phage λ cII gene. In order to make the translational information generally useful, the coding region of the gene has been removed from the vectors, leaving only their initiator fMet codon and upstream translational regulatory sequences. Additionally, these vectors have been engineered to provide restriction endonuclease sites on either side of the ATG, such that the initiation codon can be supplied by either the plasmid or the gene being inserted. Finally, restriction sites have also been engineered upstream of the translational regulatory region to permit insertion of other ribosome-binding sites. Those genes that contain restriction sites compatible with the sites on the vector may be inserted directly into the vector. As most genes do not contain appropriately positioned restriction sites, it is often necessary to adapt existing restriction cloning sites within the gene to fuse it to the translation-initiation signals provided by the vectors. For example, pSKF101 (Fig. 16.3.1) and pSKF102 both have a BamHI site adjacent to the initiation codon (ATGgatcc), while pSKF201 has an NcoI site (ccATGg) and pSKF301 (Fig. 16.3.2) has an NdeI site (catATG). The protocol presented below summarizes the steps to obtain an authentic gene clone using pSKF101 as an example. Sample Protocol Materials Appropriate restriction endonucleases and buffers (UNIT 3.1) pSKF101 vector (available from A. Shatzman; Fig. 16.3.1) Competent E. coli AS1 (Table 1.4.5; also known as MM294cI+) Additional reagents and equipment for restriction digestion, (UNIT 3.1), oligonucleotide synthesis and purification (UNITS 2.11 & 2.12), nondenaturing PAGE (UNIT 2.7), isolation, recovery, and quantitation of DNA (UNIT 2.6 & APPENDIX 3), subcloning DNA fragments (UNIT 3.16), transforming, plating, and growing E. coli (UNITS 1.8, 1.1, & 1.3), and DNA miniprep (UNIT 1.6) 1. Identify a unique restriction endonuclease site close to the 5′ end of the coding sequence of the gene to be expressed, as well as another unique site 3′ to this gene’s termination codon. 2. Synthesize two single-stranded DNA oligonucleotides, recreating the coding sequence immediately preceding the unique restriction endonuclease site near the 5′ end of the gene to be expressed. Purify and quantitate the DNA, then anneal in order to obtain double-stranded DNA.


This synthetic DNA sequence is used to link the gene to be expressed to the initiating ATG of the pSKF expression vector. The double-stranded oligonucleotide should be designed to have ends that are complimentary to the restriction sites identified at the 5′ end of the gene to be expressed as well as the chosen restriction site in the expression vector.



Eco Rl 1 Clal 27

Aat ll 5796

Sspl 5678 start 5662 Scal 5354

Hin dlll 32 / border 32, 33 BstEll 552 start 673

Ps tl 5129

rexB

Apr

end 4901

λ pSKF101 5866 bp ori 4044 Afl lIl 3981

pBR322

Pvu ll 3574

end 1107 Bgl ll 1216 p L(–35) 1324-1329 p L(–10) 1338-1343 message start 1353 Nut L1398 -1414

border 1670, 1672 NutR 1785-1801 BstXI 1819 tRI 1822-1842 c ll rbs 1863-1868 c ll start 1880 BamHl 1882 / border 1882, 1883 Sphl 2072 Nrul 2480 Sall 2157

Figure 16.3.1 pSKF101. pSKF101 is a vector used for authentic gene cloning which allows direct expression of the inserted gene. It is a derivative of pBR322 (UNIT 1.5) containing sequences inserted between HindIII and BamHI sites of pBR322. The inserted λ sequences contain the p L promoter and cII ribosome-binding site (rbs); these are the transcriptional and translational requlatory sequences necessary to express heterologous genes in E. coli. Within this region are several unique restriction sites that permit insertion of the gene. The regions derived from pB322 and λ are indicated. This plasmid can be maintained stably in a λ-lysogenized E. coli strain. The selectable marker is ampicillin, encoded by β-lactamase. An alternate name for pSKF101 is pASI (Rosenberg et al., 1983). Alternative names of related vectors are as follows: pSKF102 is pOTSV (Shatzman and Rosenberg, 1987); pSKF201 is pOTS-Nco (Shatzman and Rosenberg, 1987); and pSKF301 is pMG1.

3. Digest 25 to 50 µg plasmid DNA containing the gene to be expressed with the restriction endonucleases identified in step 1. To ensure complete digestion, determine that the restriction endonuclease buffer is appropriate for each enzyme to be used. If the endonucleases require different buffers, then each restriction digestion must be done separately.

4. Electrophorese the doubly digested plasmid DNA on a polyacrylamide gel. If the DNA fragment to be isolated is between 150 and 1100 bp, a 6% gel can be used. Either a borate- or acetate-buffer system can be used. If digestion was done in a large volume, ethanol precipitate the DNA (UNIT 2.1) and resuspend in 40 to 100 ìl TE buffer. Mix with loading dye and load.

5. Locate the fragment of interest by staining with an agent such as ethidium bromide and cut the DNA fragment out of the gel. 6. Recover the DNA by electroelution and quantitate the amount of DNA. Confirm that the correct fragment has been isolated by running a small aliquot on an agarose gel. Be sure to run appropriate size markers in an adjacent lane.

7. Digest 10 µg pSKF101 with BamHI and a restriction endonuclease that generates ends compatible with the 3′ end of the coding sequence (to accommodate the 3′ end of the gene to be expressed). Confirm that complete digestion of the vector has occurred by analysis of digested DNA

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Sspl Aat ll Scal 5000

Pstl

Bgl ll pL

5192

BstXI

Ap r 1000 4000

NS1-81

pSKF301 5192 bp Eagl/Xmalll

ori 2000

Nde l BamHI HindIII

Ncol Ndel Hpal EcoRV Stul 3 reading frame insertion sites Banll/Sac l Xhol Xbal Clal 3 reading frame termination sites Hin dlll

3000 Accl Tth111l Pvull

Nar l

Ball

Figure 16.3.2 pSKF301. pSKF301 is a vector that can be used for both indirect and direct expression. it is similar to pSKF101 in that it contains the same transcriptional and translational regulatory sequences as well as selectable markers; it differs in that it contains a shorter segment of λ DNA than pSKF101. pSKF301 also contains the coding sequence of the first 81 amino acids of the influenza protein, NS1, shown as NS1-81. This region is adjacent to the cll ribosome-binding site (fbs) and contains restriction sites at the 3′ end of NS1-81 that allow construction of translational fusions in any of the three reading frames. Removal of NS1-81 permits direct expression of the cloned gene. (This vector is also known as pmG1.)

on an agarose gel. Compare undigested pSKF101 with digested to make sure that pSKF101 has been completely linearized.

8. Prepare a ligation reaction (using the conditions described in UNIT 3.16) by combining the following ingredients: 1 ng digested pSKF101 vector DNA 10 ng of the gene fragment to be expressed (from step 6) 20 ng synthetic oligonucleotide (from step 2) T4 DNA ligase. Ligate 10 to 12 hr at 4°C. There is no need to dephosphorylate pSKF101 as long as there is at least a 5-fold molar excess of vector DNA to isolated DNA fragment and synthetic DNA.

9. Remove one-third of the ligation reaction and transform 50 to 100 µl competent E. coli AS1. Plate on LB/ampicillin plates and incubate overnight at 37°C. 10. Pick 12 to 24 colonies and transfer with a sterile toothpick to 3 ml LB/ampicillin medium. Grow cells 5 to 18 hr and isolate DNA by a miniprep method. Cells may be harvested once the broth appears turbid. For best results, allow 8 to 12 hr of growth.


11. Perform appropriate restriction endonuclease digests to determine which clones contain the desired construction of the gene to be expressed. 12. Transform an E. coli strain with the DNA and express the gene as in the basic protocols.



CONSTRUCTION AND DISASSEMBLY OF FUSED GENES IN pSKF301 By fusing the gene to be expressed to a coding region of another gene (the fusion partner), a chimeric gene can be constructed in an appropriate vector. Numerous vectors are available for this purpose but most share the common feature of a fusion partner that is a highly expressed gene. When expression of the chimeric gene is induced, the resulting proteins carry additional peptide information at the N terminus. Although the fusion product may have physical and/or functional properties that differ from the “authentic” protein, advantages of the approach include highly efficient expression (up to 30% of total cell protein) without complicated alterations on the gene, and the presence of a “handle” on the expressed protein which can help to identify and purify it. Such proteins are often used to develop antisera to specific proteins that have diagnostic potential, and have been used successfully to identify and define a variety of gene products (Casadaban et al., 1983; Rose and Botstein, 1983; Guarente, 1983).

SUPPORT PROTOCOL

Strategic Planning Plasmid pSKF301 has been constructed to permit initial expression of a gene as a fusion product, followed by removal of the DNA encoding the fusion portion by restriction digestion. Finally, the unfused version of the gene is expressed as an authentic protein. pSKF301 contains an NdeI restriction site adjacent to the ATG following the cII ribosome-binding site (Fig. 16.3.3). This ATG also serves as the translational start (Gold et al., 1981) of the NS1 gene derived from the influenza nonstructural gene. This gene has been truncated to express only its first 81 amino acids. Just beyond the coding sequence for the 81st amino acid is a second NdeI site followed by three unique blunt-ended restriction sites, HpaI, EcoRV, and StuI, which allow for the insertion of genes into any of three reading frames. Immediately following the StuI site are sequences coding for translational stops in any of the three reading frames.

A Ndel/BamHI

cllrbs

NS1-81 amino acids

Ndel Hpal

Eco RV

Stul

ccatg gat cat atg tta aca gat atc aag gcc tga ctg act gag

stop stop stop

B ccatg gat cat atg tt H pal-digested pSKF301 supplies

2 3

of a codon

ccatg gat cat atg tta aca gat Eco RV-digested pSKF301 supplies a complete codon

ccatg gat cat atg tta aca gat atc aag g 1 Stu l-digested pSKF301 supplies 3 of a codon

Figure 16.3.3 Sequence and restriction endonuclease sites (in the region used for cloning) of pSKF301 (A). Restriction endonuclease digestion shows the strategy utilized to obtain pSKF301 as a vehicle for expression in all three reading frames (B). Protein Expression


Supplement 11

The expression of a gene of interest as a fusion protein may be achieved by utilizing any of the following restriction sites in pSKF301: NcoI, HpaI, EcoRV, or StuI. Choice of restriction site depends upon the reading frame necessary for the translation of a specific protein sequence. First, a unique restriction site close to the 5′ end of the gene (or portion of the gene) to be expressed must be identified. Second, the appropriate restriction endonuclease is selected for digesting pSKF301 such that the gene will be expressed. If the chosen restriction site is a blunt-end cutter, no further manipulation of that end is required. In the event the restriction site identified leaves either a 5′ or 3′ protruding end, further manipulation is required. “Filling in” using the Klenow fragment of E. coli DNA polymerase for 5′ protrusions, or T4 DNA polymerase, S1, or mung bean nuclease for 3′ protrusions, are methods of choice (see UNIT 3.16). Sample Protocol Materials Appropriate restriction endonucleases and buffers (UNIT 3.1) Klenow fragment of E. coli DNA polymerase I (UNIT 3.5) pSKF301 vector (available from A. Shatzman; Figs. 16.3.1 & 16.3.2) T4 DNA ligase (UNIT 3.14) Competent E. coli AS1 (Table 1.4.5; also known as MM294cI+) Additional reagents and equipment for large-scale plasmid prep (UNIT 1.7), agarose gel electrophoresis (UNIT 2.5), extraction and precipitation of DNA (UNIT 2.1), transformation of competent cells (UNIT 1.8), and restriction digestion and mapping (UNITS 3.1-3.3) Construct a gene fusion in pSKF301 1. Assume the restriction site identified in the gene is a BamHI site. Digest with BamHI to obtain: GATCC

XXX

XXX

XXX

XXX

G

YYY

YYY

YYY

YYY

2. Treat with Klenow fragment to fill in the unpaired bases to obtain: GATCC

XXX

XXX

XXX

XXX

CTAGG

YYY

YYY

YYY

YYY

As noted above, Klenow fragment is used to fill in for 5′ protrusions. For 3′ protrusions, use T4 DNA polymerase (UNIT 3.5) or S1 or mung bean nuclease (UNIT 3.12).

3. Determine the proper reading frame of the gene. In this example assume XXX XXX XXX XXX is the proper reading frame; therefore, the coding sequence of the filled-in fragment should read: GA

TCC

XXX

XXX

XXX

4. Determine which restriction endonuclease should be used to digest pSKF301 to allow expression of the fusion protein. For this example, StuI is required to yield: ccatg gat cat atg tta aca gat atc aag gGA TCC XXX XXX XXX XXX pSKF301 Expression Using Vectors with Phage ëRegulatory Sequences

fusion gene

5. Prepare the vector and the fragment of the gene to be expressed as in the first support protocol, steps 3 to 12 (except no synthetic DNA is required).



Generate an authentic version of the gene Once a gene has been expressed as a fusion protein, it may be desirable to obtain an unfused version of the gene product. If this is useful, follow steps 6 to 12. To convert a fusion protein to an unfused protein when using pSKF301, be certain that the gene of interest does not contain an NdeI site. The following theoretical fusion construct will be used as an example in these steps: NdeI

NdeI

CATATGGATCC- - -NS1-81- - -CCATGGATCATATGTT- - -fusion gene- - -tga

6. Set up a large-scale plasmid preparation of the fusion construct to yield ∼100 µg plasmid DNA. 7. Digest 10 µg of the construct with NdeI. Verify that all of the vector DNA has been completely digested by taking a small aliquot of the digested material and running it on an agarose gel next to lanes containing uncut plasmid and appropriate size markers. A 280-bp fragment should be observed; this contains the NS1-81 gene sequence being liberated from the construct. Confirmation of complete digestion is extremely important.

8. Purify the digested construct by phenol/chloroform/isoamyl alcohol extraction followed by ethanol precipitation. 9. Add T4 DNA ligase to 1 µg of the NdeI-digested construct and incubate overnight at 4°C. 10. Transform ligated DNA into competent E. coli AS1 cells (or any other suitable cI+ lysogen). 11. Determine that the construct no longer contains the NS1-81 gene sequence by restriction analysis. Consult the restriction map of pSKF301 and the gene to be expressed to determine which endonucleases are diagnostic for identifying the construct devoid of the NS1-81 gene. If the NdeI digestion was complete upon ligation, reclosure is highly efficient. Expect 95% to 100% of the resulting transformants to contain the unfused construct.

12. Transform the DNA and express the gene by temperature or chemical induction as in the basic protocols. COMMENTARY Background Information Expression of a heterologous gene or gene fragment in E. coli requires that the coding sequence be placed under the transcriptional and translational control of regulatory elements recognized by the bacterial cell. The pSKF vectors were designed specifically to direct gene expression by providing regulatory signals from bacteriophage λ. Phage regulatory signals were chosen because of their high efficiency and ability to be tightly regulated. This system uses a promoter that can be tightly controlled, eliminating problems with “leaky”

basal expression sometimes found in other expression systems (see below). This system uses an antitermination mechanism to help assure efficient transcription across any gene insert. The different vectors used with this system offer several choices of antibiotic selection markers, contain elements that optimize plasmid stability, and carry a variety of restriction sites that permit relatively easy insertion of the gene of interest adjacent to the efficient translation regulatory information. The pSKF system offers some advantages that differentiate it from many other expression

Protein Expression


Supplement 11


systems. Perhaps most important is the “tightness” of regulation of the pL promoter. Several other strong regulatable promoters—ptac (de Boer et al., 1982), ptrp (Edman et al., 1981), and pT7 (promoter of T7 gene 10; Studier and Moffatt, 1986)—are also used routinely for optimizing heterologous gene expression in E. coli. These promoters, along with pL, are all of comparable strength and are sufficient to achieve very high levels of mRNA production (UNIT 16.1). In fact, these promoters are so powerful that further enhancement of promoter strength would not be expected to result in an increase of protein production; indeed, these promoters are so strong that it is very difficult to keep them fully turned off even in the “repressed” state. Because of this basal transcription under repressed conditions, use of the ptac, ptrp, or pT7 (coupled with the plac-T7 polymerase) systems often leads to some expression of the cloned gene even under nonpermissive conditions. This may lead to plasmid loss or rearrangement, or possibly cell death, if small amounts of the gene product are lethal to E. coli. In contrast, one does not typically see expression of the cloned gene in the pL system until cultures have been induced. A second advantage of the pL system over other promoter systems is the flexibility gained from having completely different induction systems (thermal and chemical). In contrast, the ptac and pT7 systems mentioned above permit induction only by a chemical route. Different routes of induction lead to completely different cellular states (e.g., different physiology, morphology, and growth patterns) and these variations can lead to significant differences in gene product accumulation and stability (unpub. observ.). It should be pointed out that ptrp also permits dual modes of induction (by β-indolyl acetic acid or Trp starvation) and that a different version of the pT7 system has been developed (UNIT 16.2; Tabor and Richardson, 1985) in which the T7 RNA polymerase is thermally regulated via the pL-cI857 system (however, this system is not chemically inducible as well). The third major advantage of the pSKF system is the availability of a single vector that permits expression of either an authentic or a fusion gene product; furthermore, this vector allows the fusion gene to be converted to an authentic gene by a simple restriction digest followed by self ligation. Thus, a gene may be rapidly expressed at high levels as a protein fusion to give an initial reagent for use in activity studies and antisera preparation. Time

may then be taken to optimize the expression of the authentic (nonfusion) gene product, which will be better suited for functional and structural studies. Most other expression systems do not provide this flexibility.

Critical Parameters Gene expression is not solely a function of message levels. The efficiency of the ribosomebinding region—including the sequences both upstream and downstream of the ATG initiation codon—also play a role in determining the extent to which a protein is made. Alterations in these sequences may affect the secondary structure of a message and the conformational presentation of the initiation signals which, as a result, can alter translational efficiency (Gold et al., 1981). From our experience, the host strain plays a major role in determining the ultimate levels of gene expression. The reasons for the rather dramatic differences seen in product yield from different host strains are poorly understood. Product stability is, however, one determining factor that has been somewhat characterized. Host strains have been developed that are defective in certain proteases (UNIT 16.6). These specialized host strains can have a significant impact on the expression of certain gene products. However, proteases are not the only factor involved in strain-to-strain variations observed in protein expression. Other uncharacterized factors can have equally dramatic effects. It is therefore recommended that expression be tried in a number of different E. coli strains. Following the induction of cultures carrying the desired expression vector, cells may be analyzed in a variety of ways to detect the presence of the cloned gene product. Most typically, the presence of the novel gene product is determined directly by observing in SDS– polyacrylamide gels a new, inducible protein band not present in lanes from control cultures. The expression of any gene insert can also be identified and/or confirmed in several ways related to the activity or function of the protein including: (1) direct detection of a novel function or activity imparted to the living bacterial host; (2) genetic complementation of the appropriate mutant host; (3) assay of whole-cell extracts for the activity of the cloned gene product; and (4) assay after partial or complete purification of the cloned gene product. Immunochemical methods such as immunoprecipitation (UNIT 10.16) or western blotting (UNIT 10.8) are some of the most sensitive meth-



ods available to detect expression of a gene product. These methods, of course, require that an antiserum be available which is specific for the protein to be expressed. These methods, however, are primarily quantitative and do not necessarily indicate anything about the level of expression, homogeneity, or activity of the gene product. If a good antiserum to the protein of interest is not available, purification of sufficient amounts of a gene product allows generation of high-titer, antigen-specific mono- or polyclonal antisera (UNITS 11.3-11.13). One approach to generate an antiserum is to produce the desired heterologous gene product in bacteria as a native protein, as a fusion, or as a protein fragment. The protein may then be purified and used to produce high-titer mono- or polyclonal antisera. Such antisera have been used to (1) map natural expression of the gene product with respect to cell type, subcellular distribution, and temporal regulation; (2) determine relative levels of expression in various cell types; (3) study protein processing and stability; (4) map immuno-dominant domains; (5) purify by immunoaffinity both the native and modified forms of the protein; and (6) provide in vivo diagnostic reagents for examining tissue distribution and expression of the gene product by immunofluorescent methods.

Troubleshooting There is never a guarantee that a gene will be expressed at high levels, but poor expression upon initial trials does not signify defeat. As mentioned earlier, transcription is rarely limiting and is, therefore, not the first parameter to be addressed in attempting to improve expression. Instead, the easiest parameter to change is the host strain being used for production. Typically, five or six different strains (which might or might not be closely related to each other) may have to be tested in pilot experiments to see which gives optimal production. The next parameter to examine in the event of poor expression is translation. Expression may be increased by altering ribosome-binding sites to improve complimentarity to the 16S rRNA, or by increasing the A-T richness of the 5′-end of the gene’s coding region. After steps have been taken to optimize translation, it is often helpful to alter the promoter and repressor system in order to change the induction system and the physiology of the cells during the production phase. For example, inducing the cI857-containing pL system via a temperature shift generates a cellular heat-

shock response and protein synthesis at 42°C. Induction of this system with nalidixic acid leads to a cellular SOS response (see glossary, Chapter 1 introduction) and protein production at 37°C. Induction of the trp system by tryptophan starvation turns on the host stringent response (a generalized response of E. coli to amino acid starvation). Thus, in each case, a different host response leads to induction of a different set of host proteins as well as to greatly different physiological effects (such as changes in respiration, filamentation, and growth rate). Finally, it may be possible to improve expression by optimizing the temperature at which the protein is made, as this parameter has often been shown to affect the proteins’ solubility, stability, and activity.

Anticipated Results Expression of most gene products as fusions with the first 81 amino acids of the NS1 protein (using pSKF301) can be achieved at levels between 5% and 30% of total cellular protein. Expression levels of nonfusion proteins (authentic) are less predictable and may vary from 50 kDa), or when the protein contains regions that are strongly hydrophobic or highly charged (D.B.S. and L.M.C., unpub. observ.). Insoluble fusion proteins can sometimes be coaxed into solution (see Critical Parameters and Troubleshooting) or can otherwise be purified after solubilization in denaturing reagents. If necessary, the GST moiety can be removed from fusion proteins by cleavage with site-specific proteases (UNIT 16.4B). Note however, that often the GST carrier does not compromise the antigenicity or functional activity of the foreign polypeptide. Modified versions of the original pGEX vectors have been produced that simplify cloning, cleavage or detection of fusion proteins (for review, see Smith, 1993).


Expression and Purification of Glutathione-STransferase Protein Fusions

Contamination of fusion proteins by host cell proteins is usually a sign that sonication has been too severe, perhaps because denaturation exposes regions on proteins that are more likely to cause aggregation. Some contaminating species may represent degraded fragments of the fusion protein that bind to glutathione. Such fragments are not easy to eliminate, except by increasing the stability of the fusion protein. The addition of protease inhibitors [e.g., 1% (w/v) aprotinin and 1 mM phenylmethylsulfonyl fluoride (PMSF)] may help in this regard, and inclusion of 50 to 100 mM EDTA in the lysis buffer has also been beneficial. The E. coli host strain can have a major and unpredictable effect on stability, and it is worth testing several different strains, including the protease-deficient lon− strains (D.B.S. and L.M.C., unpub. observ.; UNIT 1.4). Alterna-

tively, degradation of fusion proteins can be minimized by adding isopropyl-1-thio-β-Dgalactoside (IPTG) later in the course of the culture and keeping the induction period to a minimum. The overall yield of fusion protein can sometimes be improved by increasing the quantity of glutathione-agarose beads, minimizing the volume of liquid during absorption, and extending the period of absorption to 1 hr. Insolubility of fusion proteins can be addressed by several means. In some cases, growth of cells at 30°C is sufficient to alter solubility (D.B.S., unpub. observ.), but in other cases it may be necessary to investigate the effect of mild detergent treatment after cell lysis. Examples of conditions under which binding of GST to glutathione-agarose is unaffected are 1% Triton X-100, 1% Tween-20, 1% CTAB, 10 mM DTT, and 0.03% SDS. Alternatively, fusion proteins can be solubilized in 1% to 2% Sarkosyl prior to sonication; they can be solubilized in 2% to 4% Triton X-100 prior to binding to glutathione agarose (Frangioni and Neel, 1993). Even if a fusion protein is largely insoluble, it may be possible to purify the small proportion that is still soluble, with a yield of perhaps 50 µg/liter. Otherwise, a fusion protein that is abundant, but stubbornly insoluble, can be purified by gel filtration after solubilization under denaturing conditions or by electroelution from a gel after SDS-PAGE. A different approach is to express the polypeptide as smaller fragments, particularly if it is possible to express parts of the protein that do not contain severely hydrophobic or highly charged regions. If the fusion protein is not of the expected size, it may be worth sequencing across the cloning site to ensure that the reading frame of the vector and the insert are matched. There is a hairpin in GST sequences near the cloning sites, so a sequencing primer complementary to either the extreme 3′ end of the GST gene or to sequences within the DNA insert should be used.

Anticipated Results Yields of fusion protein vary from more typical yields of 1 to 3 mg/liter up to 10 mg/liter, and can be as low as 50 µg/liter if most of the fusion protein is insoluble. The single affinity chromatography step can generate fusion protein preparations that are >90% pure.

Time Considerations A full day is required to screen colonies of transformants for expression of fusion proteins,



although continuous attention is not required. Large-scale purification and cleavage of fusion proteins require another day of intermittent work. Cells can be stored as pellets at −70°C before lysis, although with fusion proteins that are unstable, this may be undesirable. Purification is best completed in one session.

Literature Cited Frangioni, J.V. and Neel, B.G. 1993. Solubilization and purification or enzymatically active glutathione-S-transferase (pGEX) fusion proteins. Anal. Biochem. 210:179-187. Smith, D.B. 1993. Purification of glutathione-Stransferase fusion proteins. Methods Mol. Cell Biol. 4:220-229. Smith, D.B., Davern, K.M., Board, P.G., Tiu, W.U., Garcia, E.G., and Mitchell, G.F. 1986. Mr 26,000 antigen of Schistosoma japonicum recognized by resistant WEHI 129/J mice is a parasite glutathione S-transferase. Proc. Natl. Acad. Sci. U.S.A. 83:8703-8707.

Key Reference Smith et al., 1986. See above. Original description of the pGEX system. Smith, 1993. See above. Summary of modified pGEX vectors and alternative purification methods. Smith, D.B. and Johnson, K.S. 1988. Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione Stransferase. Gene 67:31-40. First description of GST fusion system.

Contributed by Donald B. Smith University of Edinburgh Edinburgh, Scotland Lynn M. Corcoran Walter & Eliza Hall Institute Victoria, Australia

Protein Expression


Supplement 28

Expression and Purification of Thioredoxin Fusion Proteins

UNIT 16.8

This unit describes a gene fusion expression system that uses thioredoxin, the product of the Escherichia coli trxA gene, as the fusion partner. The system is particularly useful for high-level production of soluble fusion proteins in the E. coli cytoplasm; in many cases heterologous proteins produced as thioredoxin fusion proteins are correctly folded and display full biological activity. Although the thioredoxin gene fusion system is routinely used for protein production, high-level production of peptides—i.e., for use as antigens— is also possible because the prominent thioredoxin active-site loop is a very permissive site for the introduction of short amino acid sequences (10 to 30 residues in length). The inherent thermal stability of thioredoxin and its susceptibility to quantitative release from the E. coli cytoplasm by osmotic shock can also be exploited as useful tools for thioredoxin fusion protein purification. In addition, a more generic method for purification of any soluble thioredoxin fusion employs a modified form of thioredoxin (called “His-patch Trx”), which has been designed to bind to metal chelate resins. Protein fusions to His-patch Trx can usually be purified in a single step from cell lysates (see Strategic Planning). The basic protocol outlines the construction of a fusion of trxA to any desired gene and expression of the fusion protein in an appropriate host strain at 37°C. Additional protocols describe E. coli cell lysis using a French pressure cell and fractionation (first support protocol), osmotic release of thioredoxin fusion proteins from the E. coli cytoplasm (second support protocol), and heat treatment to purify some thioredoxin fusion proteins (third support protocol). STRATEGIC PLANNING The thioredoxin gene fusion expression vectors pTRXFUS and hpTRXFUS, both of which carry the E. coli trxA gene (Fig. 16.8.1), are used for high-level production of C-terminal fusions to thioredoxin. The vector hpTRXFUS differs from pTRXFUS in that it contains a modified E. coli trxA gene which produces a mutant protein (“His-patch” thioredoxin) that can specifically bind to metal chelate matrices charged with nickel or cobalt, otherwise known as native metal-chelate affinity chromatography (MCAC; UNIT 10.11B). The trxA translation termination codon has been replaced in both vectors by DNA encoding a ten-residue peptide linker sequence that includes an enterokinase (enteropeptidase; LaVallie et al., 1993a) cleavage site. This highly specific site can be cleaved with enterokinase following purification of the fusion protein to release the protein of interest from its thioredoxin fusion partner (cleavage of the fusion protein is covered in UNIT 16.4B). Immediately downstream of the DNA encoding the enterokinase site in pTRXFUS and hpTRXFUS lies a DNA polylinker sequence containing a number of unique restriction endonuclease sites that can be used for forming in-frame translational fusions of any desired gene to trxA. Downstream of the DNA polylinker lies the E. coli aspA transcription terminator. Replication of these vectors is controlled by a modified colE1 replication origin similar to that found in pUC vectors (Norrander et al., 1983). Plasmid selection and maintenance is ensured by the presence of the β-lactamase gene on the vector. The vector pALtrxA-781 (Fig. 16.8.1) is very similar to pTRXFUS. However in this plasmid the trxA gene is followed by a translation termination codon, and the sequences encoding the enterokinase-site peptide linker are absent. A unique RsrII site, present in both pALtrxA-781 and pTRXFUS, allows for the easy insertion of short peptide-encoding DNA sequences into trxA within the region that encodes the active-site loop. Contributed by John McCoy and Edward LaVallie Current Protocols in Molecular Biology (1994) 16.8.1-16.8.14 Copyright © 2000 by John Wiley & Sons, Inc.

Protein Expression


ori

BLA

pALtrxA–781 pTRXFUS p L hpTRXFUS

trxA aspA TGGTGCGGTCCGTGCAAA W C G33 P34 C K pALtrxA–781:

Rsr II

Sfi I

Xba I

Sal I

Pst I

AACCTGGCCTAGCTGGCCATCTAGAGTCGACCTGCAG N L A *

aspA terminator

thioredoxin

KpnI BamHI XbaI

pTRXFUS:

Sal I

Pst I

AACCTGGCCGGTTCTGGTTCTGGTGATGACGATGACAAGGTACCCGGGGATCCTCTAGAGTCGACCTGCAG N L A G S G S G D D D D K fusion point thioredoxin

aspA terminator

linker peptide enterokinase site

Figure 16.8.1 Thioredoxin gene fusion expression vectors pTRXFUS, hpTRXFUS, and pALtrxA-781. pALtrxA-781 contains a polylinker sequence at the 3′ end of the trxA gene. pTRXFUS and hpTRXFUS contain a linker region encoding a peptide that includes the enterokinase cleavage site between the trxA gene and the polylinker. The sequence surrounding the active site loop of thioredoxin has a single RsrII site that can be used to insert peptide coding sequence. The asterisk indicates a translational stop codon. Abbreviations: trxA, E. coli thioredoxin gene; BLA, β-lactamase gene; ori, colE1 replication origin; pL, bacteriophage λ major leftward promoter; aspA terminator, E. coli aspartate amino-transferase transcription terminator.


pTRXFUS, hpTRXFUS, and pALtrxA-781 carry the strong bacteriophage λ promoter pL (Shimatake and Rosenberg, 1981) positioned upstream of the trxA gene. Transcription initiation at the pL promoter is controlled by the intracellular concentration of λ repressor protein (cI). UNIT 16.3 describes λ strains that carry either a temperature-sensitive form of cI (cI857) or a wild-type cI repressor protein. cI857-containing strains can be used for heat inductions of pL at 42°C; alternatively, in the strains carrying the wild-type repressor, pL can be induced by a prior induction of the E. coli SOS stress response. However, it is often desirable to express heterologous genes in E. coli at temperatures considerably lower than 42°C, or under conditions where cells are not undergoing a physiological stress. Strains GI698, GI724 and GI723 were designed to allow the growth and induction of pL expression vectors, including pTRXFUS, hpTRXFUS, and pALtrxA-781, under mild conditions over a wide range of temperatures (see Table 16.8.1; Mieschendahl et al., 1986). Each of these strains carries a wild-type allele of cI stably integrated into the E. coli chromosome at the nonessential ampC locus. A synthetic trp promoter integrated into ampC upstream of the cI gene in each strain directs the synthesis of cI repressor only when intracellular tryptophan levels are low. When tryptophan levels are high, synthesis of cI is switched off; therefore, the presence of tryptophan in the growth medium of GI698, GI723, or GI724 will block expression of λ repressor and thus will turn on pL. Because



Table 16.8.1 E. coli Strains for Production of Thioredoxin Fusion Proteins at Varying Temperatures

Strain

Desired production temperature (°C)

Pre-induction growth temperature (°C)

Induction period (hr)

GI698 GI698 GI698

15 20 25

25 25 25

20 18 10

GI724 GI724 GI723

30 37 37

30 30 37

6 4 5

the three strains carry ribosome-binding sequences of different strengths at the 5′-end of their respective cI genes, they maintain intracellular concentrations of λ repressor that increase in the order GI698 < GI724 < GI723. The choice of which strain to use for a particular application is dependent on the desired culture conditions as described below. Although some thioredoxin fusion proteins produced at 37°C are insoluble, expression at lower temperatures can often result in the fusion protein being produced in a soluble form. Each of the three pL host strains GI698, GI723, and GI724 is suitable for the production of thioredoxin fusion proteins over a particular temperature range. Table 16.8.1 indicates the correct strain for expression of thioredoxin fusion proteins at any temperature between 15°C and 37°C. The induction protocol at any of these temperatures is the same as that described in the basic protocol for induction of GI724 at 37°C, except the preinduction growth temperature and the length of the induction period vary according to the strain used and the temperature chosen. Cultures should be grown at the indicated preinduction growth temperature until they reach a density of 0.4 to 0.6 OD550/ml. They should then be moved to the desired induction temperature and induced by the addition of 100 µg/ml tryptophan. Low-temperature inductions are best performed in strain GI698. However, this strain makes only enough cI repressor protein to maintain the vectors in an uninduced state at temperatures below 25°C. GI698 should therefore never be grown above 25°C when it carries a pL plasmid. A nonrefrigerated water bath can be maintained below room temperature by placing it in a 4°C room and setting the thermostat to the desired temperature. It is often a good idea to collect timepoints during the course of a long induction period and to fractionate cells from these timepoints using the procedure in the first support protocol (steps 9 to 13). Although a particular fusion protein may be soluble during the early part of an induction, during the later phases of induction, it may become unstable or its concentration inside the cell may exceed a critical threshold above which it will precipitate and appear in the insoluble fraction.

Protein Expression


Supplement 28

BASIC PROTOCOL

CONSTRUCTION AND EXPRESSION OF A THIOREDOXIN FUSION PROTEIN This protocol describes construction and subsequent expression of a gene fusion between trxA (encoding thioredoxin) and a gene encoding a particular protein or peptide. After a clone carrying the correct fusion sequence is constructed, analyzed, and isolated, cultures are grown and expression is induced. The protocol is described in terms of the E. coli host strain GI724 with expression at 30°C; it may also be applied to strains GI698 and GI723 (also available from Genetics Institute) for expression at other temperatures by using the parameters specified in Table 16.8.1 (see Strategic Planning). Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

DNA fragment encoding desired sequence Thioredoxin expression vectors (Fig. 16.8.1): pTRXFUS or pALtrxA-781 (Genetics Institute or Invitrogen) or hpTRXFUS (Genetics Institute) E. coli strain GI724 (Genetics Institute or Invitrogen), grown in LB medium and made competent (UNIT 1.8) LB medium (UNIT 1.1) IMC plates containing 100 µg/ml ampicillin (see recipe) CAA/glycerol/ampicillin 100 medium (see recipe) IMC medium containing 100 µg/ml ampicillin (see recipe) 10 mg/ml tryptophan (see recipe) SDS-PAGE sample buffer (see recipe) 30°C convection incubator 18 × 50–mm culture tubes Roller drum (New Brunswick Scientific) 250-ml culture flask 70°C water bath Microcentrifuge, 4°C Additional reagents and equipment for subcloning of DNA fragments (UNIT 3.16), transforming competent E. coli cells (UNIT 1.8), preparing miniprep DNA (UNIT 1.6), restriction mapping (UNIT 3.2), direct sequencing of plasmid DNA (UNITS 7.3 & 7.4), SDS-PAGE (UNIT 10.2), and Coomassie brilliant blue staining (UNIT 10.6) Construct the trxA gene fusion 1. Use DNA fragment encoding the desired sequence to construct either an in-frame fusion to the 3′-end of the trxA gene in pTRXFUS or hpTRXFUS, or a short peptide insertion into the unique RsrII site of pALtrxA-781. A precise fusion of the desired gene to the enterokinase linker sequence in pTRXFUS or hpTRXFUS can be made by using the unique KpnI site trimmed to a blunt end with the Klenow fragment of E. coli DNA polymerase. The desired gene can usually be adapted to this blunt-end construct by using a synthetic oligonucleotide duplex ligated between it and any convenient downstream restriction site close to the 5′ end of the gene. When designing the fusion junction, note that enterokinase is able to cleave —DDDDK↓X—, where X is any amino acid residue except proline. Synthetic oligonucleotides encoding short peptides for insertion into the thioredoxin active-site loop at the RsrII site will insert only in the desired orientation, because the RsrII sticky end consists of three bases. Expression and Purification of Thioredoxin Fusion Proteins

2. Transform the ligation mixture containing the new thioredoxin fusion plasmid into competent GI724 cells. Plate transformed cells onto IMC plates containing 100 µg/ml



ampicillin to select transformants. Incubate plates in a 30°C convection incubator until colonies appear. Strains GI698, GI723, and GI724 are all healthy prototrophs that can grow under a wide variety of growth conditions, including rich and minimal media and a broad range of growth temperatures (see Table 16.8.1). These strains can be prepared for transformation with pL-containing vectors by growing them in LB medium at 37°C. LB medium may also be used for these strains during the short period of outgrowth immediately following transformation. This growth period of 30 min to 1 hr is often used to express drug resistance phenotypes before plating out plasmid transformations onto solid medium. Subsequently, however, these strains should be grown only on minimal or tryptophan-free rich media, such as IMC medium containing 100 ìg/ml ampicillin (for expression of the fusion protein) or CAA/glycerol/ampicillin 100 medium (for plasmid DNA preparations). Except during transformation, LB medium should never be used with these three strains when they carry pL plasmids because LB contains tryptophan. The pL promoter is extremely strong and should be maintained in an uninduced state until needed so that expression of the protein will not lead to selection of mutant or variant cells with lower expression due to undesirable genetic selections or rearrangements in the expression strain.

3. Grow candidate colonies in 5 ml CAA/glycerol/ampicillin 100 medium overnight at 30°C. Prepare minipreps of plasmid DNA and check for correct gene insertion into pTRXFUS by restriction mapping. 4. Sequence plasmid DNA of candidate clones to verify the junction region between thioredoxin and the gene or sequence of interest. Induce expression 5. Streak out frozen stock culture of GI724 containing thioredoxin expression plasmid to single colonies on IMC plates containing 100 µg/ml ampicillin. Grow 20 hr at 30°C. Occasionally there is induction of pL plasmids grown in GI698 and GI724 at 37°C, even in medium containing no tryptophan. Such induction appears to be a temperature-dependent phenomenon. If growth at 37°C prior to pL induction is essential, then GI723 should be used as the host strain because GI723 produces higher levels of cI repressor than both GI698 and to GI724. Otherwise, plasmid-containing GI698 should be grown at 25°C and plasmid-containing GI724 should be grown at 30°C prior to induction (see Table 16.8.1).

6. Pick a single fresh, well-isolated, colony from the plate and use it to inoculate 5 ml IMC medium containing 100 mg/ml ampicillin in an 18 × 150–mm culture tube. Incubate overnight at 30°C on a roller drum. 7. Add 0.5 ml overnight culture to 50 ml fresh IMC medium containing 100 µg/ml ampicillin in a 250-ml culture flask (1:100 dilution). Grow at 30°C with vigorous aeration until absorbance at 550 nm reaches 0.4 to 0.6 OD/ml (∼3.5 hr). 8. Remove a 1-ml aliquot of the culture (uninduced cells). Measure the absorbance at 550 nm and harvest the cells by microcentrifuging 1 min at maximum speed, room temperature. Carefully remove all the spent medium with a pipet and store the cell pellet at −80°C. 9. Induce pL by adding 0.5 ml of 10 mg/ml tryptophan (100 µg/ml final) to remaining cells immediately. 10. Incubate 4 hr at 37°C. At hourly intervals during this incubation, remove 1-ml aliquots of the culture and harvest cells as in step 8. Protein Expression


Supplement 28

11. Harvest the remaining cells from the culture 4 hr post-induction by centrifuging 10 min at 3000 rpm (e.g., in a Beckman J6 rotor), 4°C. Store the cell pellet at −80°C. Procedures for further analysis of these cells are outlined in the support protocols.

Verify induction 12. Resuspend the pellets from the induction intervals (steps 8 and 10) in 200 µl of SDS-PAGE sample buffer/OD550 cells. Heat 5 min at 70°C to completely lyse the cells and denature the proteins. Run the equivalent of 0.15 OD550 cells per lane (30 µl) on an SDS-polyacrylamide gel. 13. Stain the gel 1 hr with Coomassie brilliant blue. Destain the gel and check for expression. Most thioredoxin fusion proteins are produced at levels that vary from 5% to 20% of the total cell protein. The desired fusion protein should exhibit the following characteristics: it should run on the gel at the mobility expected for its molecular weight; it should be absent prior to induction; and it should gradually accumulate during induction, with maximum accumulation usually occurring 3 hr post-induction at 37°C. SUPPORT PROTOCOL 1

E. COLI LYSIS USING A FRENCH PRESSURE CELL A small 3.5-ml French pressure cell can be used as a convenient way to lyse E. coli cells. The whole-cell lysate can be fractionated into soluble and insoluble fractions by microcentrifugation. Other lysis procedures may be used—for example, sonication (UNITS 4.4 & 16.6) or treatment with lysozyme-EDTA (UNIT 4.4). Additional Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Cell pellet from 4-hr post-induction culture (basic protocol) 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2), 4°C Lysis buffer: 20 mM Tris⋅Cl (pH 8.0) with protease inhibitors (optional)—0.5 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM p-aminobenzamidine (PABA), and 5 mM EDTA French press and 3.5-ml mini-cell (Fig. 16.8.2; SLM Instruments), 4°C Lyse the cells 1. Resuspend cell pellet from 4-hr post-induction culture in 20 mM Tri⋅Cl, pH 8.0, to a concentration of 5 OD550/ml. Protease inhibitors can be included in the resuspension if desired. Cells can also be resuspended at densities of 100 OD550/ml or greater; however, at high densities cell lysis may be less efficient.

2. Place 1.5 ml resuspended cell pellet in the French pressure cell. Hold the cell upside down with the base removed, the piston fully extended downwards, and the outlet valve handle that holds the nylon ball seal in the open position (loose). Before filling the pressure cell, check that the nylon ball, which seals the outlet port and sits on the end of the outlet valve handle, is not deformed. If it is, replace it with a new one. Both the condition of the nylon ball and its seat in the pressure cell body are critical for the success of the procedure.


3. Bring the liquid in the pressure cell to the level of the outlet port by raising the piston slowly to expel excess air from the cell. With the outlet valve open and at the same time maintaining the piston in position, install the pressure cell base. Gently close the outlet valve.



Figure 16.8.2 French pressure cell, equipped with 3.5-ml mini-cell. piston

cell body

outlet valve

outlet port

base

CAUTION: Do not over-tighten the valve as this will deform the nylon ball and may irreparably damage its seat on the pressure cell body.

4. Turn the sealed cell right side up and place it in the hydraulic press. 5. Turn the pressure regulator on the press fully counter-clockwise to reset it to zero pressure. Set the ratio selector to medium. Turn on the press. CAUTION: The larger (50-ml) pressure cell is usually used with the selector set on high. The small (3.5-ml) cell is only used on medium ratio.

6. Slowly turn the pressure regulator clockwise until the press just begins to move. Allow the press to compress the piston. It will stop moving after a few seconds. 7. Position a collection tube under the pressure cell outlet. Slowly increase the pressure in the cell by turning the pressure regulator clockwise. Monitor the reading on the gauge and increase the pressure to 1000 on the dial, corresponding to an internal cell pressure of 20,000 lb/in2. 8. While continuously monitoring the gauge, very slowly open the outlet valve until lysate begins to trickle from the outlet. The lysate should flow slowly and smoothly, and the cell pressure should not drop more than 100 divisions on the dial. At 20,000 lb/in2 and 5 OD550/ml, cell lysis will be complete after one passage through the press. Lower pressures and/or higher cell densities may require a second passage.

Fractionate the lysate 9. Remove a 100-µl aliquot of the lysate and freeze at −80°C (whole-cell lysate). 10. Fractionate the remainder of the lysate by microcentrifuging 10 min at maximum speed, 4°C. 11. Remove a 100-µl aliquot of the supernatant and freeze at −80°C (soluble fraction). Discard the remainder of the supernatant. Because this is a pilot experiment, it would not produce enough material to warrant saving any remaining supernatant.

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12. Resuspend the pellet in an equivalent volume of lysis buffer. Remove a 100-µl aliquot and freeze at −80°C (insoluble fraction). 13. Lyophilize the 100-µl aliquots to dryness in a Speedvac evaporator. Solubilize in 100 µl SDS-PAGE sample buffer. Analyze 30-µl samples by SDS-PAGE. This crude fractionation provides a fairly reliable indication of whether a protein has folded correctly. Usually proteins in the soluble fraction have adopted a correct conformation and proteins in the insoluble fraction have not. However, occasionally proteins found in the soluble fraction are not truly soluble; instead they form aggregates that do not pellet in the microcentrifuge. Conversely, sometimes a protein found in the insoluble fraction may be there because it has an affinity for cell wall components and cell membranes, and it may not be intrinsically insoluble. Occasionally proteins can be recovered from these insoluble fractions by extracting with agents such as mild detergents. SUPPORT PROTOCOL 2

OSMOTIC RELEASE OF THIOREDOXIN FUSION PROTEINS Thioredoxin and some thioredoxin fusion proteins can be released with good yield from the E. coli cytoplasm by a simple osmotic shock procedure. Additional Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Cell pellet from 4-hr post-induction cultures (basic protocol) 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA/20% (w/v) sucrose, ice-cold 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA, ice-cold 1. Resuspend cell pellet from 4-hr post-induction cultures at a concentration of 5 OD550/ml in ice-cold 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA/20% sucrose. Incubate 10 min on ice. 2. Microcentrifuge 30 sec at maximum speed, 4°C, to pellet the cells. 3. Discard the supernatant and gently resuspend the cells in an equivalent volume of ice-cold 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA. Incubate 10 min on ice and mix occasionally by inverting the tube. Osmotic release from the cytoplasm occurs at this stage.

4. Microcentrifuge 30 sec at maximum speed, 4°C. Save the supernatant (osmotic shockate). Resuspend the cell pellet in an equivalent volume 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA (retentate). 5. Lyophilize 100-µl aliquots of osmotic shockate and retentate to dryness in a Speedvac evaporator. 6. Solubilize each in 100 µl SDS-PAGE sample buffer. Analyze 30-µl aliquots by SDS-PAGE. The osmotic shock procedure provides a substantial purification step for some thioredoxin fusion proteins. This procedure will remove most of the contaminating cytoplasmic proteins as well as almost all of the nucleic acids. However the shockate will contain as contaminants about half of the cellular elongation factor-Tu (EFTu) and most of the E. coli periplasmic proteins.




PURIFICATION OF THIOREDOXIN FUSION PROTEINS BY HEAT TREATMENT

SUPPORT PROTOCOL 3

Wild-type thioredoxin is resistant to prolonged incubations at 80°C. A subset of thioredoxin fusion proteins also exhibit corresponding thermal stability, and heat treatment at 80°C can sometimes be used as an initial purification step. Under these conditions the majority of contaminating E. coli proteins are denatured and precipitated. Additional Materials For recipes, see Reagents and Solutions in this unit (or cross-referenced unit); for common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Cell pellet from 4-hr post-induction cultures (basic protocol) 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA 80°C water bath 10-ml glass-walled tube 1. Resuspend cell pellet from 4-hr post-induction cultures at a concentration of 100 OD550/ml in 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA. It is important to start off with a high protein concentration in the lysate to ensure efficient precipitation of denatured proteins.

2. Lyse the cells at 20,000 lb/in2 in a French pressure cell as described in steps 2 to 8 of the first support protocol. Collect whole-cell lysate in a 10-ml glass-walled tube. 3. Incubate whole-cell lysate 10 min at 80°C. Remove 100-µl aliquots after 30 sec, 1 min, 2 min and 5 min and plunge immediately into ice. At 10 min plunge the remaining heated lysate into ice. A glass-walled tube (not plastic) provides good thermal conductivity to provide a rapid rise in temperature to 80°C and then a rapid drop in temperature to 4°C. A suitable volume to use in a 10-ml glass tube is 1.5 ml lysate. For large-scale work, a glass-walled vessel should be used and the lysate should be mixed well during both heat treatment and cooling.

4. Microcentrifuge the aliquots 10 min at maximum speed, 4°C to pellet heat-denatured, precipitated proteins. 5. Remove 2-µl aliquots of the supernatants and add 28 µl SDS-PAGE sample buffer. Analyze the samples by SDS-PAGE to determine the heat stability of the fusion protein and the minimum time of heat treatment required to obtain a good purification.

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Casamino Acids (CAA), 2% (w/v) 20 g Casamino Acids (Difco certified) H2O to 1 liter Autoclave or filter sterilize through a 0.45-µm filter Store ≤2 months at room temperature Do not use technical-grade Casamino Acids because it has a higher NaCl content.

CAA/glycerol/ampicillin 100 medium 800 ml 2% (w/v) Casamino Acids (see recipe; 1.6% final) 100 ml 10× M9 salts (see recipe; 1× final) 100 ml 10% (v/v) glycerol (sterile; 1% final) 1 ml 1 M MgSO4(sterile; 1 mM final) 0.1 ml 1 M CaCl2 (sterile; 0.1 mM final) 1 ml 2% (w/v) vitamin B1 (sterile; 0.002% final) 10 ml 10 mg/ml ampicillin (sterile; 100 µg/ml final) Prepare fresh IMC medium 200 ml 2% (w/v) Casamino Acids (see recipe; 0.4% final) 100 ml 10× M9 salts (see recipe; 1× final) 40 ml 20% (w/v) glucose (sterile; 0.5% final) 1 ml 1 M MgSO4 (sterile; 1 mM final) 0.1 ml 1 M CaCl2 (sterile; 0.1 mM final) 1 ml 2% (w/v) vitamin B1 (sterile; 0.002% final) 658 ml glass-distilled H2O (sterile) 10 ml 10 mg/ml ampicillin (sterile; optional; 100 µg/ml final) Use fresh IMC plates 15 g agar [Difco; 1.5% (w/v)] 4 g casamino acids [Difco-certified; 0.4% (w/v)] 858 ml glass-distilled H2O(sterile) Autoclave 30 min Cool in a 50°C water bath 100 ml 10× M9 salts (see recipe; 1× final) 40 ml 20% (w/v) glucose (sterile; 0.5% final) 1 ml 1 M MgSO4 (sterile; 1 mM final) 0.1 ml 1 M CaCl2 (sterile; 0.1 mM final) 1 ml 2% (w/v) vitamin B1 (sterile; 0.002% final) 10 ml 10 mg/ml ampicillin (sterile; optional; 100 µg/ml final) Mix well and pour into Petri plates Store ≤1 month at 4°C


M9 salts, 10× 60 g Na2HPO4 (0.42 M) 30 g KH2PO4 (0.24 M) 5 g NaCl (0.09 M) 10 g NH4Cl (0.19 M) H2O to 1 liter continued



Adjust pH to 7.4 with NaOH Autoclave or filter sterilize through a 0.45-µm filter Store ≤6 months at room temperature SDS-PAGE sample buffer 15% (v/v) glycerol 0.125 M Tris⋅Cl, pH 6.8 (APPENDIX 2) 5 mM Na2EDTA 2% (w/v) SDS 0.1% (w/v) bromphenol blue 1% (v/v) 2-mercaptoethanol (2-ME; add immediately before use) Store indefinitely at room temperature Tryptophan, 10 mg/ml Heat 500 ml glass-distilled H2O to 80°C. Stir in 5 g L-tryptophan until dissolved. Filter sterilize the solution through a 0.45 µm filter and store ≤6 months in the dark at 4°C. COMMENTARY Background Information Two significant problems plague researchers who hope to express heterologous proteins in Escherichia coli: inefficient initiation of translation of many eukaryotic mRNA sequences on bacterial ribosomes (Stormo et al., 1982), and proteins that often form insoluble aggregates, called inclusion bodies, that are composed of misfolded or denatured proteins (Mitraki and King, 1989). Although successful protocols for refolding eukaryotic proteins from inclusion bodies can be developed, the process is always uncertain and usually time-consuming; in most instances it is preferable to prevent inclusion-body formation in the first place. The use of trxA fusions provides a solution to both problems. Inefficient initiation of translation of eukaryotic messages in E. coli can often be improved by modifying sequences at the 5′ end of the gene. A more reliable technique that avoids the problem entirely is to use a gene fusion strategy in which the gene of interest is linked in-frame to the 3′ end of a highly translated partner gene. In this case protein synthesis always initiates on the same efficiently translated fusion partner mRNA, thus high-level expression is assured. Some earlier gene fusion expression systems, for example the trpE and lacZ systems described in UNIT 16.5, offer very reliable ways of producing large quantities of any desired eukaryotic protein. However, these gene fusion systems still suffer from the pervasive inclusion-body problem. They are thus mainly useful for the production of antigens, rather than correctly folded, biologically active

proteins. More recently the maltose binding protein (MBP) and glutathione-S-transferase (GST) gene fusion expression systems (see UNITS 16.6 & 16.7) have proven more successful in producing soluble fusion proteins; these systems retain the translation advantage of the earlier fusion systems. Apart from the obvious advantages in making a correctly folded product, the synthesis of soluble fusion proteins also allows for the development of generic purification schemes based on some unique property of the fusion partner. Why would any particular eukaryotic protein produced in the E. coli cytoplasm be more soluble when it is linked to a fusion partner than it would be by itself? It is likely that physical properties of the fusion partner protein are important, with efficient self-folding and high solubility being useful in this role. It is possible that some good fusion partners (proteins that fold efficiently and are highly soluble), by virtue of their desirable physical qualities, are able to keep folding intermediates of linked heterologous proteins in solution long enough for them to adopt their correct final conformations. In this respect the fusion partner may serve as a covalently joined chaperon protein, in many ways fulfilling the role of authentic chaperon proteins (McCoy, 1992), analogous to the covalent chaperon role proposed for the N-terminal pro regions of a number of protein precursors (Silen et al., 1989; Shinde et al., 1993). Many of the known properties of E. coli thioredoxin (Holmgren, 1985) suggested that it would make a particularly effective fusion partner in an expression system. First, thiore-

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doxin, when overproduced from plasmid vectors, can accumulate to 40% of the total cellular protein, yet even at these expression levels all of the protein remains soluble. Second, the molecule is small (11,675 Mr) and would contribute a relatively modest amount to the total mass of any fusion protein, in contrast to other systems such as the lacZ system. Third, the tertiary structure of thioredoxin (Katti et al., 1990) reveals that both the N- and C-termini of the molecule are accessible on the surface and in good position to link to other proteins. The structure also shows that the molecule has a very tight fold, with >90% of its primary sequence involved in strong elements of secondary structure. This provides an explanation for thioredoxin’s observed high thermal stability (Tm 85°C), and suggests that the molecule might possess the robust folding characteristics that could make it a good fusion partner protein. In support of this view, complete thioredoxin domains are found in a number of naturally occurring multidomain proteins, including E. coli DsbA (Bardwell et al., 1991), the mammalian endoplasmic reticulum proteins ERp72 (Mazzarella et al., 1990), and protein disulfide isomerase (PDI; Edman et al., 1985). These proteins can all be considered as natural precedents for thioredoxin fusion proteins. The synthesis of small peptides in E. coli is often difficult, with the products frequently being extensively degraded or insoluble. The thioredoxin tertiary structure revealed that the characteristic active site, —CGPC—, protrudes from the body of the protein as a surface loop, with few interactions with the rest of the molecule. The loop does not seem to contribute to the overall stability of thioredoxin, so the production of peptides as insertions at this site was an attractive possibility. In this location they would be protected from host-cell amino- and carboxypeptidases, and thioredoxin’s high solubility should help keep them in solution. In addition, the conformation of peptides inserted at this position would be constrained, which could be an advantage for applications in which it is desirable for the peptide to adopt a particular form. Thioredoxin has indeed proven to be an excellent partner for the production of soluble fusion proteins in the E. coli cytoplasm (LaVallie et al., 1993b). Figure 16.8.3 demonstrates the production of soluble fusion proteins between thioredoxin and eleven human and murine cytokines and growth factors using the trxA vectors. All of these mammalian proteins had been previously produced in E. coli only as

insoluble inclusion bodies. As thioredoxin fusions, the growth factors are not only made in a soluble form, but in most cases they are also biologically active in in vitro assays. Experience gained while working with these and a number of other trxA fusion proteins shows that two further characteristics of thioredoxin can be exploited as purification tools. The first is the inherent thermal stability of the molecule, a property that is retained by some thioredoxin fusion proteins. This enables heat treatment to be used as an effective purification step. The second additional property relates to thioredoxin’s cellular location. Although E. coli thioredoxin is a cytoplasmic protein, it has been shown to occupy a special position within the cell—it is primarily located on the cytoplasmic face of the adhesion zones that exist between the inner and outer membranes of the E. coli cell envelope (Lunn and Pigiet, 1982). From this location thioredoxin is quantitatively released to the exterior of the cell by simple osmotic shock or freeze/thaw treatments, a remarkable property that is retained by some thioredoxin fusion proteins, thus providing a simple purification step. A more generic method for purification of any soluble thioredoxin fusion employs a modified form of thioredoxin (called “His-patch Trx”), which has been designed to bind to metal chelate resins (E.A. DiBlasio, J.M. McCoy, and E.R. LaVallie, manuscript in preparation).

Critical Parameters Lack of protein solubility leading to inclusion-body formation in E. coli is a complex phenomenon with many contributing factors: simple insolubility as a result of high-level expression, insolubility of protein-folding intermediates, lack of appropriate bacterial chaperon proteins, and lack of glycosylation mechanisms in the bacterial cytoplasm. Fusion of heterologous proteins to thioredoxin or to other fusion partners can help address most of these solubility issues. However, another important factor contributing to inclusion body formation is the inability to form essential disulfide bonds in the reducing environment of the bacterial cytoplasm, which leads to incorrect folding. Thermal lability of even correctly folded heterologous proteins in the absence of these stabilizing disulfide cross-links is a significant problem, so the expression of fusion genes should be attempted over a wide range of temperatures, even as low as 15°C (the limit for E. coli growth is ∼8°C). Thermal denaturation is



1

2

3

4

5

6

7

8

9

10

11

12

MW(kDa) - 97.4 - 66.2

- 45.0

- 31.0

- 21.5

- 14.4

Figure 16.8.3 Expression of thioredoxin gene fusions. The gel shows proteins found in the soluble fractions derived from E. coli cells expressing eleven different thioredoxin gene fusions. Lane 1, host E. coli strain GI724 (negative control, 37°C); lane 2, murine interleukin-2 (IL-2; 15°C); lane 3, human IL-3 (15°C); lane 4, murine IL-4 (15°C); lane 5, murine IL-5 (15°C); lane 6, human IL-6 (25°C); lane 7, human MIP-1a (37°C); lane 8, human IL-11 (37°C); lane 9, human macrophage colony-stimulating factor (M-CSF; 37°C); lane 10, murine leukemia inhibitory factor (LIF; 25°C); lane 11, murine steel factor (SF; 37°C); and lane 12, human bone morphogenetic protein-2 (BMP-2; 25°C). Temperatures in parentheses are the production temperature chosen for expressing each fusion. This is a 10% SDS-polyacrylamide gel, stained with Coomassie brilliant blue.

a time-dependent process, so it is also prudent to monitor the solubility of the expressed fusion protein over the time course of induction. A great many proteins contain distinct structural domains. For example, hormone receptor proteins usually have an extracellular ligandbinding domain, a transmembrane region, and an intracellular effector domain. Sometimes expressing these domains individually as fusion proteins can yield better results than expressing the entire protein. The exact positions chosen for boundaries of the domains to be expressed in the fusion protein are important and can be determined from a knowledge of the tertiary structure of the protein of interest, by homology comparisons with similar proteins, by limited proteolysis or other domain-mapping experiments, or empirically by generating multiple fusions that test different boundary positions. It is important to be consistent in treating samples for loading on gels. For example, using different heating conditions from one experi-

ment to the next can result in a mobility shift for the protein of interest.

Anticipated Results Thioredoxin fusion protein yields are usually in the range of 5% to 20% of total cell protein. At these expression levels a 1-liter induction culture in a shaker flask will yield ∼3 g (wet weight) of cells, 300 mg total protein, and 15 to 60 mg of thioredoxin fusion protein. The final recovered yield will depend on factors such as solubility of the fusion protein and the efficiency of downstream purification procedures.

Time Considerations From a single colony on a plate, the basic induction protocol requires an overnight growth to prepare a liquid inoculum and a 3.5-hr preinduction growth at 30°C the next day, followed by a 4-hr 37°C induction period. These times are significantly longer if lower induction temperatures are required (see Table

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16.8.1). Lysis of a sample in the French pressure cell should require ≤ 5 min, and both the heattreatment and osmotic-shock procedures require 99.9%, that are derived from these cotransfections contain the repaired virus with the target gene, thus minimizing the need to screen and plaque-purify recombinants. Several companies (Pharmingen, Invitrogen, Clontech, and Novagen; see APPENDIX 4) market linearized ORF 1629–deleted AcMNPV DNA. To further facilitate the identification of recombinants, several of these commercially available baculovirus DNAs contain the bacterial lacZ gene, which codes for β-galactosidase in lieu of the AcMNPV polyhedrin gene, thereby allowing lacZ-negative recombinants to be distinguished visually from any residual nonrecombinant viruses via a plaque assay. Nonrecombinant viruses form blue plaques on Xgal plates because they contain a functional lacZ gene, whereas recombinants form colorless, opaque plaques. Recombinant viruses can also be identified by DNA hybridization and polymerase chain reaction (PCR) amplification. Another rapid and efficient method for generating recombinant baculoviruses uses site-specific transposition to insert foreign genes by homologous recombination into a bacmid propagated in E. coli rather than in insect cells. In this case, recombinant viral DNA is isolated from individual bacterial colonies and is free of any wild-type viral DNA. Upon transfection of insect cells, recombinant virus is generated free of parental nonrecombinant virus, thereby eliminating the need for multiple rounds of plaque purification. This is the basis of the Bac-to-Bac

baculovirusexpressionsystem, which is available commercially from Invitrogen.

POSTTRANSLATIONAL MODIFICATION OF PROTEINS IN INSECT CELLS Because baculoviruses infect invertebrate cells, it is possible that the processing of proteins produced by them is different from the processing of proteins produced by vertebrate cells. Although this seems to be the case for some posttranslational modifications, it is not the case for others. For example, two of the three posttranslational modifications of the tyrosine protein kinase, pp60c-src, that occur in higher eukaryotic cells (myristylation and phosphorylation of serine 17) also take place in insect cells. However, another modification of pp60c-src observed in vertebrate cells, phosphorylation of tyrosine 527, is almost undetectable in insect cells (Piwnica-Worms et al., 1990). In addition to myristylation, palmitylation has been shown to take place in insect cells. However, it has not been determined whether all or merely a subfraction of the total recombinant protein contains these modifications. Cleavage of signal sequences, removal of hormonal prosequences, and polyprotein cleavages have also been reported, although cleavage varies in its efficiency. Internal proteolytic cleavages at arginine- or lysine-rich sequences have been reported to be highly inefficient, and alpha-amidation, although it does not occur in cell culture, has been reported in larvae and pupae (Hellers et al., 1991). In most of these cases a cell- or species-specific protease may be necessary for cleavage. Protein targeting seems conserved between insect and vertebrate cells. Thus, proteins can be secreted and localized faithfully to either the nucleus, cytoplasm, or plasma membrane. Although much remains to be learned about the nature of protein glycosylation in insect cells, proteins that are glycosylated in vertebrate cells will also generally be glycosylated in insect cells. However, with few exceptions the N-linked oligosaccharides in insect cell–derived glycoproteins are only high-mannose type and are not processed to complex-type oligosaccharides containing fucose, galactose, and sialic acid. O-linked glycosylations have been even less well characterized in Sf 9 cells, but have been shown to occur. For further information on posttranslational modifications of proteins and protein processing in insect cells, see Davidson et al. (1990), O’Reilly et al. (1992), Jarvis and Sum-

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mers (1992), Grabenhorst et al. (1993), James et al. (1995), Jarvis and Finn (1995), Davis and Wood (1995), and Ogonah et al. (1996).

STEPS FOR OVERPRODUCING PROTEINS USING THE BACULOVIRUS SYSTEM The use of the baculovirus expression system is presented in detail in UNITS 16.10 & 16.11. The following steps comprise a brief overview (also see Fig. 16.9.2). 1. Clone the gene of interest into the appropriate baculovirus expression vector and cotransfect with linearized baculoviral DNA (available from various vendors) or use transposition to create recombinant bacmid DNA in E. coli (BAC-TO-BAC system; Invitrogen). Alternatively, purify circular wild-type baculovirus DNA (UNIT 16.10, Alternate Protocol 1). 2. Cotransfect baculovirus DNA with the recombinant baculovirus plasmid or transfect purified bacmid DNA into Sf 9 insect cells (UNIT 16.10, Basic Protocol 2). 3. Collect the medium, which contains the baculoviral particles (UNIT 16.10, Basic Protocol 2), and plaque the virus on Sf 9 cells to separate recombinant from nonrecombinant virus (UNIT 16.10, Basic Protocol 4). This and subsequent rounds of plaque purification are optional when linearized virus that contains a lethal deletion is used (e.g., BaculoGold from Pharmingen), as in that case >99.9% of all amplified virus particles will be recombinant because of selection pressure. Similarly, when bacmid DNA is used, no purification is required. 4. Amplify the virus stock by infecting fresh insect cells (UNIT 16.10, Basic Protocol 3) and determine titer of the amplified virus stock (UNIT 16.10, Basic Protocol 4). 5. Express the protein of interest by infecting a new batch of insect cells with the hightiter baculovirus stock (UNIT 16.11, Basic Protocol 1). Determine the expression level of the recombinant protein of interest and analyze its biological activity (UNIT 16.11, Support Protocols 1 and 2).

CHOOSING A BACULOVIRUS TRANSFER VECTOR

Overview of the Baculovirus Expression System

The majority of available baculovirus vectors are pUC-based and confer ampicillin resistance. Most contain the polyhedrin gene promoter and insertion site(s) for cloning a foreign gene of interest, flanked by viral sequences that lie 5′ to the promoter and 3′ to the foreign gene insert. These flanking sequences facilitate ho-

mologous recombination between the vector and baculovirus DNA. Other baculovirus vectors contain the p10 promoter, another strong, very late promoter, or the basic protein promotor expressed late in the infection process. Some vectors are designed to express more than one heterologous gene or to express genes as fusions to N-terminal signal sequences or leader peptides, which facilitate secretion and purification of the recombinant protein. For more information on these vectors, see Table 16.9.1, O’Reilly et al. (1992), and relevant catalogs from Pharmingen, Clontech, Invitrogen, Novagen, and Stratagene. A major consideration when choosing the appropriate baculovirus expression vector is whether to express the recombinant protein as a fusion or nonfusion protein in insect cells. Fusion proteins containing a specific tag have the advantage of easy purification and detection. For nonfusion proteins, there are several vectors available—notably pVL1392 and pVL1393 (Pharmingen and Invitrogen)—each of which contains a polylinker in the opposite orientation from the other. These vectors are derived from pAcYM1 (Matsuura et al., 1987) and pAcCL29 (Livingston and Jones, 1989) and differ only in the order of the cloning sites. Other nonfusion vectors include pBacPAK8 and pBacPAK9 (Clontech), pBac-1 (Novagen), pFAST Bac 1 (Invitrogen), and pAcSG2 (Pharmingen). To express the protein as a polyhistidine fusion protein, there are several available vectors: pAcHLT-A, -B, and -C (Pharmingen), pBlueBacHis-A, -B, and -C and pFAST BacHT -A, -B, and -C (Invitrogen), and pBac2cp (Novagen). These vectors allow easy purification of the recombinant protein using a nickel-chelating resin (UNIT 10.11B). To express the protein as a glutathione-S-transferase (GST) fusion protein, the pAcGHLT-A, -B, and -C vectors are commercially available (Pharmingen). These produce an N-terminal GST fusion protein. As with all fusion protein vectors, the gene of interest has to be cloned in the proper reading frame with respect to the tag. Several other affinity tags, such as FLAG, HA, and myc, have also been used successfully for purification using commercially available affinity resins. These tags can be fused to coding sequences using PCR. Secretion of recombinant proteins into insect cell medium simplifies purification and characterization of expressed recombinant proteins. The ease of purification is further enhanced by the use of serum-free medium. There are several vectors that contain signal se-



prepare insect cell (e.g., Sf 9) culture (UNIT 16.10, Basic Protocol 1)

prepare wild-type baculoviral DNA (UNIT 16.10, Alternate Protocol 1)

subclone gene of interest into baculoviral vector (UNIT 16.10)

cotransfect insect cell with wild-type viral DNA and recombinant vector (UNIT 16.10, Alternate Protocol 1)

buy linearized ORF 1629–negative baculoviral DNA

cotransfect insect cell with linearized viral DNA and recombinant vector (UNIT 16.10, Basic Protocol 2)

visually screen for recombinants by plaque assay (UNIT 16.10, Basic Protocol 4)

obtain recombinant virus by several rounds of plaque purification (UNIT 16.10, Basic Protocol 4)

purify recombinant virus by one round of plaque purification (optional; UNIT 16.10, Basic Protocol 4)

amplify recombinant virus by infecting new insect cells (UNIT 16.10, Basic Protocol 3)

analyze recombinant protein expression level (UNIT 16.11, Basic Protocol 1 and Support Protocols 1 and 2)

produce and purify recombinant protein on a large scale (UNIT 16.11, Basic Protocol 2)

Figure 16.9.2 Flow chart for the expression of proteins in insect cells using the baculovirus system. The flow chart describes the production of recombinants using circular wild-type baculovirus DNA and the more recently developed linearized ORF 1629–negative variants. The use of linearized baculovirus DNA, now widely accepted, simplifies the whole protocol and produces viral-infection stocks with little or no background of nonrecombinants, which may be used directly without plaque purification.

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Table 16.9.1

Baculoviral Expression Vectorsa

Promoter type

Fusion protein

Polyhedrin locus–based Single-baculovirus promoter vectors pVL1392/1393 (pair) P pAcSG2 P pAcMP2/3 (pair) BP

VL VL L

No —c No

pAcUW21 pBacPak 8/9 (pair) pBAC-1 pBacgus-1 pBlue Bac III pAcGHLT-A, B, C (set)

P P P P P P

VL VL VL VL VL VL

No No —c —c No Yes

pAcHLT-A, B, C (set) pBac-2cp pBACgus-2cp

P P P

VL VL VL

Yes Yes Yes

pBlue Bac His, A, B, C (set) P pAc 360 P

VL VL

Yes Yes

pAcG1 pAcG2T pAcG3X BioColors BV Control (set) BioColors His (set)

P P P P P

VL VL VL VL VL

Yes Yes Yes Yes Yes

Secretory-signal vectors pAcGP67, A, B, C (set) pAcSecG2T

P P

VL VL

Yes Yes

pPbac

P

VL

Yes

pMbac

P

VL

Yes

pBAc surf-1

P

VL

Yes

Ligation-independent cloning vectors pAcSG2-LIC P pAcGST-LIC-2T P

VL VL

No No

pAcGST1-LIC

VL

No

Vector

Promoter

P

Features

Sourceb

Standard transfer vectors Recommended for large inserts Facilitates post-translational modifications Polyhedrin gene, F1 origin Standard transfer vectors F1 origin gus reporter gene lacZ gene GST and 6×His tags, thrombin cleavage site 6×His tag, thrombin cleavage site 6×His and S tags, F1 origin 6×His and S tags, F1 origin, gus reporter gene lacZ gene, 6×His tag Translational fusion with polyhedrin gene GST tag GST tag, thrombin cleavage site GST tag, factor Xa cleavage site Fusion with GFP or its variants Fusion with GFP or its variants, 6×His tag, thrombin cleavage site

PG, IN PG PG

gp67 signal sequence for secretion gp67 signal sequence for secretion, GST tag Placental AKP signal sequence for secretion Melittin signal sequence for secretion gp67 signal sequence for secretion, F1 origin

PG PG

LIC site for fast PCR cloning GST tag, thrombin cleavage, and LIC sites GST tag, LIC site

PG PG

PG CT NG NG IN PG PG NG NG IN IN PG PG PG PG PG

SG SG NG

PG continued




Table 16.9.1

Baculoviral Expression Vectorsa, continued

Vector

Promoter

Promoter type

Fusion protein

Features

Sourceb

gus reporter gene, 6×His and S tags, thrombin cleavage site, F1 origin, LIC site 6×His and S tags, thrombin cleavage site, F1 origin, LIC site

NG

Expression of 2 foreign genes, F1 origin Expression of 2 foreign genes Expression of 3 foreign genes Expression of 4 foreign genes Expression of 2 foreign genes, M13 origin Expression of 4 foreign genes, F1 origin Expression of 4 foreign genes, gus reporter gene, F1 origin

PG

Ligation-independent cloning vectors, continued pBACgus-2cp LIC

P

VL

No

pBAC-2cp LIC

P

VL

No

Multiple baculovirus promoter vectors pAcUW51 P, p10

VL

No

p2Bac pAcAB3 pAcAB4 pAcUW31

P, p10 P, p10 P, p10 P, p10

VL VL VL VL

No No No No

pBAC4x-1

P, p10

VL

—c

pBACgus 4x-1

P, p10

VL

—c

p10 locus-based Single-baculovirus promoter plasmids pAcUW1 p10

VL

No

Standard p10 locus vector

PG

Multiple promoter vector pAcUW32/43 (pair)

VL

No

Expression of 2 foreign genes, F1 origin

PG

Bacmid expression vectors FastBac1 P FastBacHT A,B,C (set) P

VL VL

No Yes

FastBacDUAL

VL

Yes

Bacmid expression system IN Bacmid expression system, 6×His IN tag Bacmid expression system IN

P, p10

P

NG

IN PG PG CT NG NG

aAbbreviations: 6×His, six-histidine (tag); BP, basic protein (promoter); CT, Clontech; GFP, green fluorescent protein; GST, glutathione-S-

transferase (tag); gus, β-glucuronidase; IN, Invitrogen; L, late (promoter); LIC, ligation-independent cloning; NG, Novagen; P, polyhedrin; PG, Pharmingen; SG, Stratagene; S tag, peptide tag from Novagen; VL, very late (promoter). bSee APPENDIX 4 for source addresses and telephone numbers. cFusion protein optional.

quences under strong polyhedrin-promoter control that direct the nascent polypeptide chain toward the secretory pathway of the cell. The sequences to be expressed are inserted downstream with respect to the signal sequences to generate a fusion gene that is transcribed under strong polyhedrin-promoter control. The secretion of biologically active protein from insect cells is the final step in a complex pathway of posttranslational modifications performed in the endoplasmatic reticulum (ER) and the Golgi complex (GC). Proteins destined for secretion are first cotranslationally translo-

cated into the lumen of the ER, where initial steps of carbohydrate processing occur. Later, the protein is transported to the GC, where further modifications take place. During translocation, the amino-terminal leader peptide sequence is, in most cases, proteolytically removed. The major determinants for the final form of posttranslational modification of a protein are its primary structure and the conformation presented to successive processive steps. Each secreted protein will present its own characteristics and potential problems for efficient secretion in a biologically active form. In gen-

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eral, the protein produced by baculovirus-infected insect cells carries modifications that are very similar to those of the native protein. With no need for cell lysis, purification of the secreted recombinant proteins is extremely easy. For an example of a recombinant protein expressed and purified in this way, see Murphy et al. (1993). Several vectors have been developed that utilize the gp67 secretory sequence of the baculovirus envelope protein: pAcGP67-A, -B, or -C (Pharmingen) and pBac surf-1 (Novagen). Two additional vectors, pMbac and pPbac (Stratagene), have insertions, respectively, of the melittin and human placental alkaline phosphatase secretory signal sequences. Additionally, these two vectors contain a p10promoter-driven lacZ gene, allowing color selection of recombinants on Xgal plates. To improve screening of recombinant baculoviruses when using wild-type AcMNPV DNA, pBlueBacIII (Invitrogen), derived from pJVNheI (Vialard et al., 1990), was developed. pBlueBacIII has a multiple cloning site and contains two promoters—the polyhedrin promoter and the early-to-late (ETL) promoter— downstream with respect to which the lacZ gene has been inserted. As with the other baculovirus vectors, the gene of interest is cloned downstream of the polyhedrin promoter, which then controls synthesis of the recombinant protein. The recombinant virus is plaqued using agarose overlays containing 150 µg/ml Xgal; the plaques are visualized as described in Basic Protocol 4 of UNIT 16.10. Recombinant viruses generate plaques that are blue and lack occlusion bodies.

Cotransfection of the linearized baculoviral DNA with a complementing plasmid construct rescues the lethal deletion of the essential gene—ORF 1629—that lies downstream of the AcMNPV polyhedrin gene of the baculovirus DNA (see Fig. 16.9.3). Therefore, the baculovirus transfer vector must be polyhedrin-locus based to rescue this deficiency. This means that the flanking sequences of its promoter region must be derived from the polyhedrin locus of the AcMNPV wild-type virus, otherwise it will not recombine with the polyhedrin locus of linearized ORF 1629–deleted baculoviral DNA. Protocols for both methods are covered in UNIT 16.10, which also discusses a new baculovirus variant allowing the gene of interest to be cloned directly into the baculovirus genome, thus obviating the need for transfer vectors. Another method for avoiding the time-consuming purification by plaque assay is the Bacto-Bac system (Invitrogen). The gene of interest is cloned into a donor plasmid, pFastBac1, and transformed into competent E. coli cells containing a helper plasmid and a baculovirus shuttle vector (bacmid). pFastBac1 contains Tn7 sites, and is transposed into the bacmid using functions supplied by the helper plasmid in trans. The recombinant bacmid is isolated from the competent bacteria by miniprep and transfected into insect cells using a cationic lipid reagent (Cellfectin). Screening for recombinants is done in E. coli and can therefore be done much more quickly than via multiple rounds of plaque purification, as is done in generation of recombinant baculovirus using wild-type virus.

CHOOSING A BACULOVIRUS DNA


There are several methods for generating recombinant baculovirus. Initially it was required that researchers cotransfect the recombinant transfer vector with wild-type baculovirus DNA, generate a supernatant containing recombinant baculovirus, and screen out the nonrecombinant wild-type virus background through several rounds of plaque purification. This was a time-consuming process requiring technical expertise developed over a long period of time. The development of modified linearized baculovirus DNA allowed the generation of an initial viral stock containing little or no nonrecombinant virus, thus abolishing the need for plaque purification. The principle of this technique lies in the construction of a modified type of baculovirus DNA which, after linearization, contains a lethal deletion and no longer codes for any viable virus.

REAGENTS, SOLUTIONS, AND EQUIPMENT FOR THE BACULOVIRUS SYSTEM Reagents and solutions commonly used for expression of proteins using baculovirus vectors are summarized below. See APPENDIX 4 for supplier contact information. 1. Suitable insect cell lines. Sf 9 cells are derived from the ovaries of the fall armyworm (Spodoptera frugiperda) and are available from American Type Culture Collection, Pharmingen, or Invitrogen. A similar cell line, Sf21, is available from the same vendors. As an alternative to S. frugiperda cell lines, the Trichoplusia ni High Five line, derived from T. ni egg-cell homogenates, is available from Invitrogen. Several proteins have been reported to have significantly higher expression using this T. ni cell line. Additionally, High Five cells have a



AcMNPV wild-type DNA

polyhedrin gene

p10 gene

Bsu 36I Bsu 36I

Bsu 36I uncut BaculoGold DNA

ORF 603

lacZ

ORF 1629

polyhedrin promoter

∆ORF 603

∆lacZ

∆ORF 1629

cleaved BaculoGold DNA (ORF 1629 –deleted)

Figure 16.9.3 Generation and purification of recombinant baculovirus.

rapid doubling time as adherent cultures and adapt quickly to serum-free media. 2. Fully prepared TNM-FH insect medium. This will contain trace metals, lactalbumin hydrolysate, yeastolate, 10% fetal bovine serum (FBS), and gentamicin. The medium can be purchased from Pharmingen and several other vendors. It can also be prepared from Grace’s insect-cell culture medium (available at 1× and 2× concentration in powdered or liquid form from Invitrogen). For instructions on preparing media from individual components, see O’Reilly et al. (1992). FBS is available from many vendors. Obtain and test different lots of serum from a number of suppliers. The lot that promotes the best growth rate and cell viability should be purchased in bulk. See APPENDIX 3F for additional discussion of FBS. Alternatively, a serum-free insect cell culture medium can be purchased from several vendors (BaculoGold medium from Pharmingen, Sf-900 II from Invitrogen, HYQ-CCM3 and SFX-Insect from Hyclone, or ExCell 401 from JRH Biosciences). These synthetic, lowprotein media are recommended for secreted proteins and facilitate subsequent purification. 3. Incubator at 27°C ± 1°C. CO2 is not required and humidification is optional. The Biological Oxygen Demand (B.O.D.) low-temperature incubator (VWR Scientific) or the larger Isotemp (Fisher) are good examples. 4. Magnetic spinner flasks. These are available in a variety of sizes from Techne or Bellco.

5. Stir plate for multiple spinners. This is available from Techne or Bellco. 6. SeaKem ME agarose. (FMC Bioproducts). 7. 60-mm, 100-mm, and 150-mm tissue culture plates. (Falcon or Corning). 8. Antibiotics (optional). Gentamicin (available from numerous vendors) and amphotericin B (Fungizone from Flow Laboratories) are used. 9. Microscope. Either an inverted light microscope or a dissecting microscope is required. 10. Appropriate cloning vectors. These are available from many vendors (see Table 16.9.1). Several additional vectors, a manual of methods, and wild-type baculovirus DNA are also available upon request from Dr. Max D. Summers, Department of Entomology, Texas Agricultural Experiment Station, Texas A & M University, College Station, Texas 77843 (Phone: 409-845-9730). It is necessary to sign a licensing agreement before the material will be sent. Commercial kits are available from Pharmingen, Invitrogen, Clontech, Novagen and Stratagene. 11. Linearized ready-to-use baculovirus DNA. This can be purchased from many of the same vendors. 12. Shaking incubators (90 to 150 rpm) can also be used to culture Sf9 cells. In this case, disposable polycarbonate Erlenmeyer flasks (Corning) or Fernbach flasks with screw caps

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(Bellco) should be used instead of magnetic spinner flasks.

LITERATURE CITED Carbonell, L.F., Klowden, M.J., and Miller, L.K. 1985. Baculovirus-mediated expression of bacterial genes in dipteran and mammalian cells. J. Virol. 56:153-160. Davidson, D.J.C., Fraser, M.J., and Castellino, F.J. 1990. Oligosaccharide processing in the expression of human plasminogen cDNA by lepidopteran insect (Spodoptera frugiperda) cells. Biochemistry 29:5584-5590. Davis, T.R. and Wood, H.A. 1995. Intrinsic glycosylation potentials of insect-cell cultures and insect larvae. In Vitro Cell. Dev. Biol. Animal 31:659-663. Doerfler, W. and Bohm, P. 1986. The Molecular Biology of Baculoviruses. Springer-Verlag, New York. Grabenhorst, E., Hofer, B., Nimitz, M., Jager, V., and Contradt, H.S. 1993. Biosynthesis and secretion of human interleukin-2 glycoprotein variants from baculovirus-infected insect Sf21 cells: Characterization of polypeptides and post-translational modifications. Eur. J. Biochem. 215:189-197. Hellers, M., Gunne, H., and Steiner, H. 1991. Expression and post-translational processing of preprocecropin-A using a baculovirus vector. Eur. J. Biochem. 199:435-439. James, D.C., Freedman, R.B., Hoare, M., Ogonah, O.W., Rooney, B.C., and Larionov, O.A. 1995. N-glycosylation of recombinant human interferon-γ produced in different animal expression systems. Bio/Technology 13:592-596. Jarvis, D.L. and Finn, E.E. 1995. Biochemical analysis of the N-glycosylation pathway in baculovirus-infected lepidopteran insect cells. Virology 212:500511. Jarvis, D.L. and Summers, M.D. 1992. Baculovirus expression vectors. In Recombinant DNA Vaccines: Rationale and Strategies (R.E. Isaacson, ed.) pp. 265-291. Marcel Dekker, New York. Kitts, P.A. and Possee, R.D. 1993. A method for producing recombinant baculovirus expression vectors at high frequency. BioTechniques 14:810-817. Livingston, C. and Jones, I. 1989. Baculovirus expression vector with single-strand capability. Nucl. Acids Res. 17:2366. Luckow, V.A. 1991. Cloning and expression of heterologous genes in insect cells with baculovirus vectors. In Recombinant DNA Technology and Applications (A. Prokop, R.K. Bajpai, and C. Ho, eds.) pp. 97-152. McGraw-Hill, New York. Luckow, V.A. and Summers, M.D. 1988. Trends in the development of baculoviral expression vectors. Bio/Technology 6:47-55.


Matsuura, Y., Possee, R.D., and Bishop, D.H.L. 1987. Baculovirus expression vectors: The requirements for high level expression of proteins, including glycoproteins. J. Gen. Virol. 68:12331250.

Miller, L.K. 1988. Baculoviruses as gene expression vectors. Annu. Rev. Microbiol. 42:177-199. Miller, D.W., Safer, P., and Miller, L.K. 1986. An insect baculovirus host vector for high-level expression of foreign genes. In Genetic Engineering, Vol. 8 (J.K. Setlow and A. Hollaender, eds.) pp. 277-298. Plenum, New York. Murphy, C.I., McIntire, J.R., Davis, D.R., Hodgdon, H., Seals, J.R., and Young, E. 1993. Enhanced expression, secretion, and large-scale purification of recombinant HIV-1 gp120 in insect cells using the baculovirus egt and p67 signal peptides. Protein Expr. Purif. 4:349-357. Ogonah, O.W., Freedman, R.B., Jenkins, N., Patel, K., and Rooney, B.C. 1996. Isolation and characterization of an insect cell line able to perform complex N-linked glycosylation on recombinant proteins. Bio/Technology 14:197-202. O’Reilly, D.R., Miller, L.K., and Luckow, V.A. 1992. Baculovirus Expression Vectors. W.H. Freeman, New York. Piwnica-Worms, H., Williams, N.G., Cheng, S.H., and Roberts, T.M. 1990. Regulation of pp60 c-src and its association with polyoma virus middle T antigen in insect cells. J. Virol. 64:61-68. Summers, M.D. and Smith, G.E. 1987. A manual of methods for baculovirus vectors and insect cell culture procedures. Texas Agricultural Experiment Station Bulletin No.1555. College Station, Texas. Vialard, J., Lalumiere, M., Vernet, T., Briedis, D., Alkhatib, G., Henning, D., Levin, D., and Richardson, C. 1990. Synthesis of the membrane fusion and hemagglutinin proteins of measle virus, using a novel baculovirus vector containing the β-galactosidase gene. J. Virol. 64:37-50.

KEY REFERENCE O’Reilly et al., 1992. See above. A guide assembled to aid researchers using the baculoviral expression system, containing detailed protocols for using this system effectively.

Contributed by Cheryl Isaac Murphy Aquila Biopharmaceuticals Worcester, Massachusetts Helen Piwnica-Worms Washington University School of Medicine St. Louis, Missouri Stefan Grünwald Pharmingen San Diego, California William G. Romanow Corning Costar Portsmouth, New Hampshire Nicole Francis and Hua-Ying Fan Massachusetts General Hospital Boston, Massachusetts



Maintenance of Insect Cell Cultures and Generation of Recombinant Baculoviruses

UNIT 16.10

This unit describes the maintenance and care of insect cell cultures as well as the generation, purification, and storage of recombinant baculoviruses. Procedures are included for maintenance and subculturing of insect cells (see Basic Protocol 1) and cotransfection of insect cells with linearized baculovirus DNA and recombinant transfer plasmid containing the gene of interest (see Basic Protocol 2). Bacmid DNA may be prepared for this procedure (see Alternate Protocol 1). In the event that the linearized virus is not available, wild-type baculovirus (AcMNPV) DNA may be used to produce recombinant baculoviruses (see Alternate Protocol 2). A procedure is also included for the generation of recombinant baculoviruses using a novel method, direct cloning (see Alternate Protocol 3), which eliminates the need to first clone the gene of interest into a baculoviral transfer vector. Preparation of baculovirus infection stocks from both monolayer and suspension cultures is also described (see Basic Protocol 3). Finally, a protocol is given for a plaque assay to be used for determining the titer of baculoviral stocks as well as for selection of recombinants and plaque purification (see Basic Protocol 4). NOTE: All reagents and equipment coming into contact with live cells must be sterile and proper sterile technique should be used accordingly. MAINTENANCE AND CULTURE OF INSECT CELLS This protocol describes how to maintain and subculture Spodoptera frugiperda (Sf 9) cells in both monolayer and suspension (spinner) cultures, in either serum-containing or serum-free medium. A culture of insect cells is begun using frozen Sf 9 cells. Cultures are maintained by subculturing and their viability is checked periodically. Aliquots of these cultures can be frozen in a liquid nitrogen freezer for long-term storage.

BASIC PROTOCOL 1

Materials TNM-FH insect medium (see recipe) containing 10% fetal bovine serum (FBS), with and without 20% (v/v) DMSO Spodoptera frugiperda (Sf 9) cells (ATCC #CRL 1711) derived from fall armyworm ovaries (also see UNIT 16.9) 70% ethanol 0.4% trypan blue stain (Life Technologies) Serum-free insect cell culture medium (BaculoGold Protein-Free Insect Medium from Pharmingen; Sf-900 II from Life Technologies; HyQ-CCM3 and HyQSFX-Insect from Hyclone; or ExCell 401 from JRH Biosciences) 60-mm tissue culture plates or 25-cm2 flasks 27°C incubator (humidification optional) Spinner culture flasks (for suspension cultures) and stir plate for multiple spinner flasks (all available from Techne or Bellco) or disposable (plastic) shaker flasks (Corning) and shaking incubator set to 27°C, 90 to 150 rpm Beckman GPR centrifuge with GH-3.7 horizontal rotor (or equivalent) Screw-top cryostat freezing vials Liquid nitrogen freezer Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3F) Protein Expression Contributed by Cheryl Isaac Murphy, Helen Piwnica-Worms, Stefan Grünwald, William G. Romanow, Nicole Francis, and Hua-Ying Fan

16.10.1


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Begin culture of Sf 9 cells 1. Place 3 ml TNM-FH medium containing 10% FBS in a 60-mm tissue culture plate or 25-cm2 flask. 2. Thaw a frozen ampule of Sf 9 cells rapidly in a 37°C water bath by moving it back and forth by hand. When ampule contents are almost completely thawed, immerse ampule in 70% ethanol to sterilize the outside. 3. Break the neck of the ampule and transfer contents to 60-mm tissue culture plate or 25-cm2 flask from step 1. Rock plate gently by hand to distribute the cells evenly and incubate 2 to 3 hr at 27°C until cells have attached. 4. Remove old medium and replace with 3 ml fresh TNM-FH/10% FBS. Continue incubation, feeding culture every 3 days (by removing old medium and replacing with fresh) until cells reach confluency (form a packed monolayer). It is important to hold the plates at an angle and remove and add medium at one corner so as not to dislodge the cells from the monolayer.

Maintain and subculture monolayer cultures 5. Remove the old medium from a confluent plate or flask of Sf 9 cells and resuspend cells by gently spritzing them with medium from a pipet. 6. Count the cells using a hemacytometer designed for tissue culture cells (APPENDIX 3F). Each cell in a small square of the hemacytometer is equivalent to 104 cells/ml.

7. Seed 1–2 × 106 cells from step 5 in new 60-mm plates or 25-cm2 flasks and rock to evenly distribute the cells (or use a larger plate or flask with more medium if preparing a larger culture of cells). Add fresh TNM-FH/10% FBS to bring the volume to 3 ml. 8. Incubate at 27°C, feeding the culture every 3 days with TNM-FH/10% FBS, until the cells reach confluency. Maintain and subculture suspension cultures 9. Remove medium and resuspend cells from confluent monolayer culture as described in step 5. Count the cells using a hemacytometer (APPENDIX 3F). 10a. Seed cells in a spinner culture flask at ∼4–5 × 105 cells/ml. Incubate at 27°C with constant stirring on a stir plate set at 60 to 80 rpm. Leave the side-arm caps slightly loosened to ensure adequate aeration. 10b. Alternatively, seed cells in a shaker flask and incubate at 27°C in a shaking incubator. When initially adapting cells, shake at 80 to 90 rpm. Increase by 10 rpm each time cells are split until cultures are shaking at 150 rpm. The volume of cell suspension in the shaker flasks should not exceed 40% of the total capacity.

11. Count cells every 2 to 3 days using a hemacytometer (APPENDIX 3F). Subculture when cells reach a concentration of 2–2.5 × 106 cells/ml by transferring the appropriate number of cells to a new flask containing fresh TNM-FH/10% FBS to achieve a final density of 4–5 × 105 cells/ml. Alternatively, pour out the appropriate volume of cell suspension and replace it with fresh medium. Maintenance of Insect Cell Cultures and Generation of Recombinant Baculoviruses

12. Determine cell viability by adding 0.1 ml of 0.4% trypan blue to 1 ml log-phase cells and examining the cells under a microscope at low power. Count the number of cells that take up trypan blue (dead cells) and count the total number of cells, then calculate the percentages of dead cells and viable cells.



A healthy culture of cells should be >97% viable. To maintain a sufficient transfer of oxygen to the cells in suspension, a minimum ratio of surface area to volume of culture must be maintained (Maiorella et al., 1988). If this value decreases, the cells will not grow exponentially and will stop growing at a lower cell density. This ratio is adequate for a 100-ml culture grown in a 100-ml spinner flask, but decreases when larger spinner flasks are used at maximum volume. Thus, to maintain sufficient oxygen transfer, smaller volumes should be used in the larger flasks unless an outside source of air is introduced into the flask (UNIT 16.11).

Adapt cells to serum-free medium 13. Subculture monolayer cells (from step 5) or suspension cells (from step 11) into medium composed of one part complete TNM-FH/10% FBS and one part serum-free medium (BaculoGold, Sf-900 II, or ExCell 401). Allow cells to grow to confluency (monolayer cultures) or to a density of 2–3 × 106 cells/ml (suspension cultures). Other commercially available serum- or protein-free media besides BaculoGold, Sf-900 II, and ExCell 401 may be used. The final choice of serum-free medium should be based on a comparison of cell growth curves and production of recombinant protein in different media.

14. Repeat the subculture and growth procedure as in step 13 using a medium composed of one part FBS-containing complete medium and three parts serum-free medium. 15. Repeat the subculture and growth procedure as in step 13 using a medium composed of one part FBS-containing complete medium and between 7 and 9 parts serum-free medium. 16. Subculture the cells into serum-free medium. Cells may adapt slowly to serum-free medium and may require several passages before growth rates and viability return to normal.

Freeze cells 17. Count cells to be frozen from an exponentially growing culture using a hemacytometer (APPENDIX 3F). 18. Centrifuge cells 10 min at 1000 × g (2000 rpm in a GH-3.7 rotor), room temperature, and discard supernatant. 19. Resuspend cell pellet at 1–2 × 107 cells/ml in TNM-FH/10% FBS. Add an equal volume of TNM-FH/10% FBS containing 20% DMSO and place cells on ice. Dispense 1-ml aliquots of this cell suspension into screw-top cryostat freezing vials and incubate 1 hr at −20°C, then overnight at −70°C. Alternatively, cells can be frozen using serum-free medium (with and without DMSO) if the cells have been adapted to serum-free medium as described in steps 13 to 16.

20. Transfer frozen cells to a liquid nitrogen freezer for long-term storage. COTRANSFECTION OF INSECT CELLS USING LINEARIZED BACULOVIRAL DNA

BASIC PROTOCOL 2

There are several protocols available for preparing baculoviral supernatant, including transfection with bacmid DNA, transfection with linearized baculoviral DNA, and cotransfection with wild-type baculoviral DNA. Reagents and protocols for these methods are available commercially, and are summarized below. In general, the authors have found that preparation of bacmid DNA is the simplest means of generating baculovirus (Bac-to-Bac system, Invitrogen; see Alternate Protocol 1); however, cotransfection with linearized baculoviral DNA is still also used widely.

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One of the most commonly used methods of introducing baculovirus and transfer-plasmid DNA into susceptible insect cells is to coprecipitate the DNA with calcium phosphate and present the mixture to insect cells. For cotransfection, prepare ≥10 µg of purified plasmid DNA. Care must be taken that the plasmid be as clean as possible. With impure plasmids, cells may lyse shortly after transfection, resulting in a lower viral titer. At ∼24 hr post-transfection, Sf 9 cell viability should be greater than 97%. It should be noted that this protocol is optimized for use with Pharmingen linear DNA. Clontech and Invitrogen (see APPENDIX 4) have their own protocols for use with the linear DNA that they offer. Materials Spodoptera frugiperda (Sf 9) cells growing in tissue culture at 50% to 70% confluence or growing in suspension culture at 1–1.5 × 106 cells/ml (see Basic Protocol 1) TNM-FH insect medium (see recipe) containing 10% fetal bovine serum (FBS) Linearized ORF 1629–deleted AcMNPV DNA (e.g., BaculoGold from Pharmingen; see Fig. 16.9.3) Recombinant baculovirus transfer vector containing gene of interest (UNIT 16.9) Transfection buffer B (see recipe) Control transfer vector: pVL1392-XylE (Pharmingen) Transfection buffer A (see recipe) 500 mM catechol/50 mM sodium bisulfate 60-mm tissue culture plates 27°C incubator (humidification optional) Inverted microscope 15-ml conical centrifuge tubes Beckman GPR centrifuge with GH-3.7 horizontal rotor (or equivalent), 4°C Additional reagents and equipment for amplification of viral supernatants to produce baculoviral stocks (see Basic Protocol 3) and plaque assay of baculovirus (see Basic Protocol 4) Prepare cells and DNA 1. In each of three 60-mm tissue culture plates seed 2 × 106 Sf 9 cells in TNM-FH medium containing 10% FBS. Incubate in a 27°C incubator until cells attach. Cell attachment should be done on a flat and even surface, allowing the cells to attach firmly, which usually takes ∼5 min. If cells do not attach after that time, either they are not healthy or the wrong plates (e.g., petri dishes that have not been tissue culture treated) have been used. The quality of the insect cells is very important and only rapidly dividing cells should be used.

2. While cells are attaching, combine in a microcentrifuge tube 0.5 µg linearized ORF 1629–deleted AcMNPV DNA and 2 to 5 µg recombinant baculovirus transfer vector containing the gene of interest. Mix well by gently vortexing or flicking the tube. Let mixture sit 5 min, then add 1 ml transfection buffer B.


3. Prepare positive control by combining, in a microcentrifuge tube, 0.5 µg linearized ORF 1629–deleted AcMNPV DNA and 2 µg pVL1392-XylE control transfer vector DNA. Mix well by gently vortexing or flicking the tube. Let mixture sit 5 min, then add 1 ml transfection buffer B.



4. Label the first plate (from step 1) as the cotransfection plate. Aspirate old medium and replace with 1 ml transfection buffer A, making sure that the entire surface of the plate is covered to prevent the cells from drying out. 5. Label the second plate (from step 1) as the positive control. Aspirate old medium and replace with 1 ml of transfection buffer A, as in step 4. 6. Label the third plate (from step 1) as negative control. Aspirate old medium and replace with 3 ml fresh TNM-FH/10% FBS, without adding any DNA. Transfect cells 7. Add 1 ml of the solution prepared in step 2 (containing the vector with the gene of interest) drop-by-drop to the cotransfection plate. After every three to five drops, gently rock the plate back and forth to mix the drops with the medium. During this procedure, a fine calcium phosphate/DNA precipitate should form. The quality of the precipitate that is optimal for transfection can be assessed visually. It should be of a fine white milky appearance.

8. Add 1 ml of the solution prepared in step 3 (containing the positive control vector) drop-by-drop to the positive control plate, repeating the procedure in step 7. 9. Incubate all three plates 4 hr in a 27°C incubator. The time of exposure to the calcium precipitate is critical for optimal transfection results. If the incubation time is too long, cell viability will be dramatically reduced. For different cell lines, the optimal incubation time varies. For Sf 9 cells, the optimal time is 4 hr.

10. After 4 hr, remove the medium from the cotransfection plate and the positive control plate (but not the negative control plate). Add 3 ml fresh TNM-FH/10% FBS to each plate, rock the plate back and forth several times, then remove all the medium again. Add 3 ml of fresh TNM-FH/10% FBS to each plate and incubate all three plates 4 to 5 days at 27°C. It is not necessary to change the medium of the negative control plate.

Check for successful transfection 11. After 4 days, check the three plates for signs of infection using an inverted microscope. Compare the negative and positive controls to the cotransfection plate. Infected cells are much larger than uninfected cells and have enlarged nuclei. Because they stop dividing early in infection, their cell density will be much lower as compared to the uninfected population. Furthermore, infected cells do not attach well to the plate and a high percentage of them will float in the medium. Many of these infection signs may not be visible at this time because the virus titer during the cotransfection is usually low. A further amplification step (see below) may be needed to visualize these changes.

12. After 5 days, collect the supernatants of the cotransfection and positive control plates. Determine viral titer by plaque screening (see Basic Protocol 4). Alternatively, cotransfection efficiency may by assessed by endpoint dilution assay (see Basic Protocol 4, step 7 annotation).

13. Check the expression of the protein of interest by lysing the transfected cells (for recombinant proteins that are not secreted) or using an aliquot of the supernatant (for recombinant proteins that are secreted), and performing an appropriate assay. Unless a sensitive assay is available for the protein of interest, expression of the recombinant protein may not be detectable at this stage. Protein Expression


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14. Assay for cells expressing the XylE protein in the positive control plate by adding 100 µl of 500 mM catechol/50 mM sodium bisulfate. Infected cells expressing XylE protein will turn bright yellow in ∼5 min.

15. Transfer the transfection supernatants from each plate to sterile conical 15-ml centrifuge tubes and centrifuge 10 min at 1000 × g (2000 rpm in GH-3.7 rotor), 4°C. Transfer viral supernatant to new sterile tubes and store at 4°C in the dark. 16. Amplify viral supernatant to produce a high-titer virus stock for production of the recombinant protein by infection of insect cells (see Basic Protocol 3). Alternatively, a single recombinant virus, obtained by plaque purification (see Basic Protocol 4), may be used for virus amplification. ALTERNATE PROTOCOL 1

GENERATION OF RECOMBINANT BACULOVIRUS BY PREPARATION AND TRANSFECTION OF BACMID DNA USING THE BAC-TO-BAC SYSTEM The Bac-to-Bac manual from Invitrogen provides detailed protocols for preparation and transfection of bacmid DNA. The following is a brief overview of the protocol starting from the cDNA being cloned into a pFastBac vector. All protocols and reagents are available from Invitrogen (http://www.invitrogen.com). Materials cDNA of interest (e.g., UNIT 5.5) Bac-to-Bac Baculovirus Expression System (Invitrogen) including: pFastBac plasmid vector and control plasmid DH10Bac competent E. coli cells Bac-to-Bac manual (available online at http://invitrogen.com/content/sfs/ manuals/bactobac_man.pdf) Cellfectin lipid transfection reagent Selection plates for transposition (see recipe) Bacmid selection medium (see recipe) Sf9 cells in serum-free medium Serum-free medium 27°C incubator 6-well tissue culture plates Additional reagents and equipment for cloning DNA into plasmids (UNIT 3.16), introduction of plasmid DNA into E. coli cells (UNIT 1.8), growing E. coli on solid (UNIT 1.3) and in liquid (UNIT 1.2) medium, alkaline lysis miniprep (UNIT 1.6), and preparation of insect cells for transfection (see Basic Protocol 2, steps 1 to 6, in this unit), and amplification of recombinant baculovirus to produce stocks (see Basic Protocol 3 in this unit) 1. Transpose cDNA into bacmid to create recombinant bacmid as follows (also see Bac-to-Bac manual). a. Clone the cDNA of interest into the pFastBac plasmid vector (also see UNIT 3.16). b. Transform plasmid DNA into DH10Bac competent cells (also see UNIT 1.8).


The DH10Bac competent cells carry the bacmid DNA and helper plasmid (encoding proteins required for transposition).

c. Incubate 4 hr at 37°C to allow transposition of the gene of interest into the bacmid.



2. Grow colonies (also see UNIT 1.2) for 24 hr on selection plates for transposition. Identify recombinant bacmid-containing clones by blue-white selection. Transposition disrupts the bacmid lacZα gene, enabling selection via Bluo-Gal, which is included in the selection plates. Also see UNIT 1.4.

3. Pick colonies and culture 24 hr in bacmid selection medium. 4. Prepare bacmid DNA by alkaline lysis miniprep (UNIT 1.6). 5. Prepare Sf9 cells (see Basic Protocol 2, steps 1 to 6, in this unit). Use Cellfectin lipid transfection reagent to transfect cells with the bacmid DNA (see Bac-to-Bac manual), then incubate for 3 days at 27°C. 6. Collect transfection supernatant (containing recombinant baculovirus) and amplify to produce baculovirus stock (see Basic Protocol 3). GENERATION OF RECOMBINANT BACULOVIRUS USING WILD-TYPE BACULOVIRAL DNA


As an alternative to linearized ORF 1629–deleted baculoviral DNA, circular wild-type AcMNPV DNA can be used for contransfection of insect cells with baculoviral transfer plasmids (also see Basic Protocol 2). However, recombination efficiency is dramatically lower (usually around 0.1% to 0.2%) as compared to that obtained with linearized DNA. The technique described here thus requires the identification and purification of recombinants by multiple rounds of plaque assay. The following protocol describes how to isolate and purify AcMNPV wild-type baculoviral DNA, which can then be used to cotransfect susceptible insect cells (e.g., Sf9) with an appropriate plasmid vector to generate recombinant baculoviruses. The cotransfection of wild-type baculovirus DNA and recombinant transfer plasmid should be performed as described in Basic Protocol 2 (using wild-type baculoviral DNA instead of linearized ORF 1629–negative AcMNPV DNA). To facilitate screening between wild-type and recombinant viral plaques, pBlueBacIII (Invitrogen) or pVL1393-XylE (Pharmingen) can be used as a positive control, making it possible to visualize differences between nonrecombinant wild-type viral plaques and recombinant viral plaques. Wild-type baculovirus available upon request from Dr. Max D. Summers, Department of Entomology, Texas Agricultural Experiment Station, Texas A & M University, College Station, Texas 77843 (Phone: 409-845-9730). Additional Materials (also see Basic Protocol 2) Wild-type baculovirus (Dr. Max D. Summers, see above, or Pharmingen) Sucrose cushion solution (see recipe) 0.1× and 1× TE buffer, pH 7.4 (APPENDIX 2) 25% and 56% (w/v) sucrose (ultrapure) in 0.1× TE buffer (filter sterilize and store up to 1 month at 4°C) Extraction buffer (see recipe) 10 mg/ml proteinase K (prepare fresh) 10% N-lauroylsarcosine (sodium salt; filter sterilize and store up to 1 year at 4°C) 25:24:1 phenol/chloroform/isoamyl alcohol (UNIT 2.1) 100% (ice-cold) and 70% (room temperature) ethanol 3 M sodium acetate, pH 5.2 (APPENDIX 2) Protein Expression


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150-mm tissue culture dishes 50-ml conical centrifuge tubes Beckman GPR centrifuge with GH-3.7 horizontal rotor (or equivalent), 4°C Beckman ultracentrifuge with SW-27 or SW-28 rotor and SW-41 rotor (or other ultracentrifuge with equivalent rotors), 4°C, and appropriate tubes Sucrose gradient maker 15-ml polypropylene centrifuge tubes 50°C water bath 5- or 10-ml wide-mouth pipets Additional reagents and equipment for phenol/chloroform extraction and ethanol precipitation of DNA (UNIT 2.1) and quantitating DNA by absorbance spectrometry (APPENDIX 3D) Prepare viral supernatant and collect virus 1. Seed at least ten 150-mm plates with 2.0 × 107 Sf 9 cells/plate in 30 ml TNM-FH medium containing 10% FBS. Incubate 1 hr at 27°C to allow the cells to attach firmly, then infect them with AcMNPV wild-type virus at a multiplicity of infection (MOI) of 0.1. Incubate 3 to 5 days at 27°C, examining plates periodically with an inverted microscope for the presence of occlusion bodies. Dr. Max D. Summers provides a manual with his materials that includes details for these procedures. Pharmingen provides similar documentation with its wild-type baculovirus. Also see O’Reilly et al. (1992). MOI is equal to plaque-forming units (pfu; see Basic Protocol 4) divided by the number of cells (pfu/cell). If the supplier of the virus does not provide adequate information regarding pfu, this can be determined by plaque assay (see Basic Protocol 4). Occlusion bodies are highly refractile, giving them a yellowish-green crystalline appearance that is readily detected under a light microscope.

2. When occlusion bodies are observed in most cells, pool the viral supernatant (∼30 ml per plate) in six 50-ml conical tubes. Centrifuge 10 min at 1000 × g (2000 rpm in GH-3.7), 4°C, then pour viral supernatant into six new tubes. Repeat centrifugation to completely remove any remaining cells. Sterile technique is not required in this step nor for any of the remaining steps of this protocol.

3. Place 6 to 35 ml of viral supernatant in each of an appropriate number of ultracentrifuge tubes for the SW-27 or SW-28 rotor and balance the tubes. Underlay with 3 ml sucrose cushion solution. Precool the ultracentrifuge and rotor to 4°C. The amount of sucrose cushion may have to be increased for larger volumes of viral supernatant.

4. Centrifuge 60 min at 100,000 × g (24,000 rpm in an SW-27 or -28 rotor), 4°C, to pellet the virus. Pour off supernatants and invert tubes on a Kimwipe to drain as much liquid as possible.


Separate virus from cellular contaminants (if necessary) 5. Examine the viral pellets carefully. If the viral pellet is pure (i.e., has an light bluish appearance), proceed to DNA isolation (step 11). If the pellet appears yellowish, separate the virus from cellular contaminants by steps 6a to 10a or 6b to 10b.



To purify viral pellet by sucrose-gradient fractionation 6a. Add 2 ml of 0.1 × TE buffer to one of the viral pellets and repeatedly pipet up and down with a Pasteur pipet to resuspend. Transfer the buffer with the resuspended virus to a second tube containing a pellet and repeat the resuspension, then repeat in turn for each pellet until all pellets are pooled in the same 2 ml. If pellet is difficult to resuspend, incubate preparation overnight at 4°C.

7a. Place 25% and 56% sucrose solutions in 0.1× TE buffer in the reservoirs of a gradient maker and prepare two linear 25% to 56% sucrose gradients in SW-41 ultracentrifuge tubes. If a gradient maker is unavailable, simply layer the 25% sucrose carefully atop the 56% sucrose to form a step gradient. Some investigators report that a step gradient gives a sharper band.

8a. Carefully layer 1 ml of the pooled viral suspension on top of each sucrose gradient. Centrifuge 90 min at 100,000 × g (28,000 rpm in an SW-41 rotor), 4°C. After centrifugation, virus should be visible as a broad bluish-white band inside the gradient.

9a. Using a Pasteur pipet, transfer the viral bands to a new SW-41 ultracentrifuge tube. Add enough 0.1× TE buffer to fill the tube (∼35 ml), then centrifuge 30 min at 100,000 × g (28,000 rpm in an SW-41 rotor), 4°C, to pellet the virus. Decant supernatant and invert tube on a Kimwipe to drain as much liquid as possible. 10a. Resuspend the virus pellet in 9 ml extraction buffer and transfer 4.5-ml aliquots to two 15-ml polypropylene centrifuge tubes. Proceed to step 11. To purify viral pellet by microcentrifugation 6b. Add 3 ml extraction buffer to one of the viral pellets and repeatedly pipet up and down with a Pasteur pipet to resuspend. Transfer the buffer with the resuspended virus to a second tube containing a pellet and repeat the suspension, then repeat in turn for each pellet until all pellets are pooled in the same 3 ml. If pellet is difficult to resuspend, incubate preparation overnight at 4°C.

7b. Transfer 1.5 ml viral suspension into each of two 1.5-ml microcentrifuge tubes. Microcentrifuge 5 min at maximum speed and pool supernatants in one 15-ml polypropylene centrifuge tube. 8b. Resuspend each pellet in 1 ml extraction buffer. 9b. Microcentrifuge the pellets 5 min at maximum speed and combine the two supernatants with the pooled supernatants in the 15-ml tube. 10b. Bring volume in the 15-ml tube to 9 ml with extraction buffer and transfer 4.5-ml aliquots to two new 15-ml polypropylene centrifuge tubes. Proceed to step 11. Isolate DNA from purified virions 11. Add 200 µl of 10 mg/ml proteinase K to each tube and incubate 1 to 2 hr at 50°C. 12. Add 0.5 ml of 10% N-lauroylsarcosine to each tube and incubate 2 hr or overnight at 50°C. 13. Extract DNA twice with an equal volume of 25:24:1 phenol/chloroform/isoamyl alcohol (see UNIT 2.1 for additional details on extraction of DNA). Extreme care should be taken to be as gentle as possible to avoid shearing the DNA at this point. Use a wide-bore Pasteur pipet and mix DNA solutions by inverting the tubes rather than by vortexing.

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14. Transfer the aqueous phase containing the DNA to another 15-ml tube using a wide-mouth 5- to 10-ml pipet. Add 10 ml of ice-cold 100% ethanol to each tube and mix gently by inverting the tubes several times. Incubate 10 min at −80°C (see UNIT 2.1 for additional details on ethanol precipitation of DNA). 15. Centrifuge 20 min at 1500 × g (2500 rpm in GH-3.7 rotor), 4°C, and discard supernatant. Rinse the DNA pellet with 70% ethanol and air dry pellet for 30 to 60 min. Resuspend pellet in 800 µl of 1× TE buffer. 16. Transfer 400 µl of the resuspended DNA to each of two microcentrifuge tubes and reprecipitate the DNA by adding 40 µl of 3 M sodium acetate and 2 vol of ice-cold 100% ethanol to each tube. Incubate 10 min at −80°C. 17. Microcentrifuge 10 min and discard supernatant. Rinse DNA pellet with 70% ethanol and air dry pellet. Resuspend DNA in 0.3 to 1.0 ml of 1× TE buffer. 18. Quantitate DNA by measuring A260 (APPENDIX 3D) and calculate yield. Store the circular wild-type baculoviral DNA at 4°C (stable for several months). This method should yield 50 to 100 ìg viral DNA per ten 150-mm dishes. If difficulty is encountered resuspending the DNA, heat mixture for ∼15 min at 65°C.

19. Cotransfect Sf 9 cells with the circular wild-type baculoviral DNA and a baculoviral transfer plasmid containing a gene of interest as in Basic Protocol 2, substituting the wild-type DNA for the linearized ORF 1629–deleted DNA in step 2 of that protocol. Recombination efficiency is dramatically lower with wild-type DNA as compared to that obtained with linearized DNA. The technique described here thus requires the identification and purification of recombinants by multiple rounds of plaque assay (see Basic Protocol 4). ALTERNATE PROTOCOL 3


GENERATION OF RECOMBINANT BACULOVIRUSES BY DIRECT CLONING This protocol describes the generation of recombinant virus by direct cloning methods (see Fig. 16.10.1), which may be applicable to the production of high-diversity expression libraries in baculovirus. Two modified AcMNPV baculoviruses—vEHuni and vECuni— have been constructed that have two Bsu36I sites downstream, respectively, of the hsp70 promoter and the synthetic promoter PcapminXIV (Lu and Miller, 1996). Cleavage of the Bsu36I sites produces overhanging TTA ends, which are filled in by incubation with the Klenow fragment of DNA polymerase I in presence of dTTP, thus leaving a TT overhang. Ready-to-use vEHuni and vECuni baculoviral DNA can be purchased from Pharmingen. The gene to be cloned must be flanked by EcoRI sites and the EcoRI ends produced by cleavage at these sites have to be partially filled in with Klenow fragment in the presence of dATP to leave AA overhangs. The gene can then be cloned directly into the vEHuni or vECuni genome. Additional Materials (also see Basic Protocol 2) Purified vector containing gene of interest, flanked by EcoRI sites Klenow fragment of DNA polymerase I (UNIT 3.5) Reaction buffer for Klenow fragment (see recipe) 25:24:1 phenol/chloroform/isoamyl alcohol (UNIT 2.1) 1× TE buffer (APPENDIX 2) vEHuni or vECuni (Pharmingen), linearized, partially filled in by Klenow fragment treatment, and containing TTA ends T4 DNA ligase (UNIT 3.14) 15°C water bath Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5A) and phenol/chloroform extraction and ethanol precipitation of DNA (UNIT 2.1)



Bsu36I promoter

Bsu36I

CCTAAGG CCTTAGG MCS GGATTCC GGAATCC

vector GAATTC CTTAAG

AcMNPV genome

gene

GAATTC CTTAAG

+ Bsu36I EcoRI promoter

TTAGG CC

CC GGATT

+ EcoRI

AcMNPV genome

+ dTTP + Klenow fragment promoter

AATTC G

AcMNPV genome

gene

G CTTAA

+ dATP + Klenow fragment

TTAGG TCC

CCT GGATT

EcoRI

AATTC AAG

gene

GAA CTTAA

ligate promoter

CCTAAT TC GGATTAAG

gene

GAATTAGG CTTAATCC

AcMNPV genome

Figure 16.10.1 Direct cloning of a gene of interest into baculovirus DNA. At the left is a diagram representing an AcMNPV recombinant containing two different Bsu36I sites. Digestion of this viral DNA with Bsu36I followed by a partial fill-in reaction with dTTP and Klenow fragment of DNA polymerase I generates a linear viral DNA with TT overhanging ends. At the right, a foreign gene with flanking EcoRI sites is digested with EcoRI to generate overhanging ends, which are then partially filled in using dATP. The resulting AA overhanging ends are then compatible with the TT overhanging ends of the viral DNA. The viral DNA and foreign gene DNAs are then combined, ligated, and transfected into insect cells.

1. Digest vector containing gene of interest with EcoRI in a total volume of 50 µl for 16 hr at 37°C and isolate gene of interest that contains flanking EcoRI sites by agarose gel electrophoresis (UNIT 2.5A). If gene of interest does not contain flanking EcoRI sites, a polymerase chain reaction with specific primers containing EcoRI sites can be used to insert flanking EcoRI sites. Alternatively, EcoRI linkers can be ligated to the purified gene.

2. In a total volume of 50 µl, incubate up to 5 µg of EcoRI-digested and purified gene of interest with 5 U Klenow fragment in reaction buffer for Klenow fragment for 20 min at 37°C. 3. Extract reaction mix with 1 vol of 25:24:1 phenol/chloroform/isoamyl alcohol, then ethanol precipitate DNA and resuspend in 10 µl of 1× TE buffer (UNIT 2.1). 4. In a total volume of 50 µl, mix 0.5 µg vEHuni or vECuni DNA with 0.1 to 1 µg of the treated gene fragment from step 3 (to obtain an ∼1:60 molar ratio of baculoviral DNA to gene fragment), and add 2 U T4 DNA ligase. Incubate overnight at 15°C. The ligated AcNPV DNA is now ready to be transfected into susceptible insect cells. Protein Expression


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5. Transfect the entire ligation mixture from step 4 into 2 × 106 Sf 9 cells as described in Basic Protocol 2. The ligation mixture from this protocol replaces the reaction mixture composed of ORF 1629–deleted AcMNPV DNA and recombinant baculovirus transfer vector used in step 2 of Basic Protocol 2. As in Basic Protocol 2, use pVL1392-XylE as a control. To purify recombinant plaques obtained by direct cloning, see Basic Protocol 4. BASIC PROTOCOL 3

PREPARATION OF BACULOVIRUS STOCKS This protocol describes how to prepare a large-scale stock of wild-type or recombinant AcMNPV virus from either monolayer or suspension culture. Growing insect cells (Sf 9) are infected with virus at a low MOI (100 ml), an MOI of 1 or 2 is sufficient. The virus is stable for 6 months at 4°C but must be shielded from light to maintain the titer (Jarvis and Garcia, 1994). Placement of 1 ml of each stock in liquid

nitrogen is recommended for long-term storage. It is highly recommended that linear wildtype DNA be used for cotransfection or that bacmid DNA be prepared in E. coli and used for transfection. This greatly simplifies the screening procedure and saves time in obtaining a pure recombinant viral stock. If circular wild-type DNA is used, it is important to purify the recombinant viral DNA through several rounds of plaque assay. Degradation by mechanical shearing and by nuclease contamination are the major problems encountered when trying to purify viral DNA from intact virions. Use of wide-mouth 5- or 10-ml pipets or Pasteur pipets with the necks broken off to handle the viral DNA helps to avoid shearing. In addition, tubes containing viral DNA should be mixed by inversion rather than by vortexing when performing phenol extractions and ethanol precipitations, and the purified DNA should be stored at 4°C rather than frozen and thawed. It is very useful to practice visualizing recombinant plaques using the β-Gal recombinant virus as described in Basic Protocol 4. As mentioned, screening for recombinants is the most time-consuming and variable part of the entire expression system. The cell density at the time of plaquing seems to be the most critical parameter for achieving good plaques in the shortest period of time. Thus, it is helpful to seed duplicate plates at a couple of different cell densities. A perfect plaque assay will yield easily visualized plaques within 6 to 8 days.

Anticipated Results Maintenance and culture of insect cells When grown at 27°C, healthy insect cells should double every 18 to 24 hr. Healthy, logarithmically growing cells will be maintained in suspension cultures most readily when they are cultured at densities between 5 × 105 and 2 × 106 cells/ml. Cell viability should be ∼97%. Cotransfection After 5 days, 20% to 100% of the cells should be showing signs of infection. Infected cells are of increased size with enlarged nuclei. They stop dividing and do not attach well to the plate. Many of the signs of infection may not be obvious if the transfection efficiency results in a low viral titer. In this case, it is necessary to amplify the virus in order to verify the success of the cotransfection. When generating recombinant baculovirus using wild-type bacu-



loviral DNA (see Alternate Protocol 2), the recombination frequency will be 0.1% to 0.2%. This method requires selection based upon identification of viral occlusion bodies. The direct cloning method (see Alternate Protocol 3) results in ∼99% recombinant virus with ∼50% containing the foreign gene in the correct orientation. Plaques can generally be identified within 5 to 10 days post-infection and counted to determine viral titers. When individual plaques are picked to isolate recombinant virus, the resulting viral titer is generally low (5 × 105 pfu/ml) and will need to be amplified several times to produce a high-titer stock.

Time Considerations Purification of wild-type viral DNA takes ∼7 days to complete. Subcloning and largescale production of recombinant DNA should take 1 to 2 weeks. The transfection procedure takes 1 day, and the transfection supernatant is harvested after 4 to 5 days. Plaquing takes l day, and it generally takes 1 week before the plaques are sufficiently formed to begin screening. Screening takes an additional day. It takes 4 to 5 days from the time of infecting Sf 9 cells with a plaque to the time of harvesting the culture supernatant for an expanded virus stock. Purifying recombinants takes from 1000 Ci/mmol; Du Pont NEN) Additional reagents and equipment for one-dimensional SDS-PAGE (UNIT 10.2) and autoradiography (APPENDIX 3A) 1. Seed 2.5 × 106 cells into 60-mm tissue culture plates containing 3 ml TNM-FH medium with 10% FBS. Prepare one plate to be infected with each putative recombinant virus and one control plate to be infected with wild-type baculovirus. 2. Incubate 1 hr at 27°C to allow cells to attach. Aspirate medium, then add 1 ml recombinant virus or 1 ml medium containing wild-type virus at an MOI of 5 to 10. Incubate 1 hr at room temperature. 3. Remove the medium from each plate by aspiration. Add 3 ml TNM-FH/10% FBS medium to the cells and incubate 24 to 48 hr at 27°C. 4. Carefully remove medium from each plate, then rinse cells once with methionine-free or methionine-free/cysteine-free medium. Add 1 ml methione-free or cysteine-free medium to each plate. Incubate cells 30 min at 27°C, then add 0.25 to 0.5 µCi EXPE35S35S per plate and incubate 3 to 4 hr at 27°C. It is important to hold the plates at an angle and remove and add medium at one corner so as not to dislodge the cells from the monolayer.

5. Transfer cells and culture supernatant from each plate to a separate 15-ml polypropylene centrifuge tube and centrifuge 10 min at 1000 × g (2000 rpm in a GH-3.7 rotor), 4°C. 6. Process and analyze the cells (if the recombinant protein is not secreted) or the supernatant (if the recombinant protein is secreted) according to the appropriate procedures in Basic Protocol 1 (see Basic Protocol 1, steps 5a or 5b, 6a or 6b, and 7). If an antibody is available, it is recommended that the labeled lysates be immunoprecipitated (UNIT 10.16) prior to boiling in SDS sample buffer and resolution by SDS-PAGE. Immunoprecipitation will help detect the recombinant protein if it comigrates with a labeled host or late-viral-specific protein. Purification of Proteins Using Baculovirus

Polyhedra (UNIT 16.9) are solubilized only under very alkaline conditions (0.1 M final NaOH concentration). Without prior disruption under alkaline conditions, 97% viability can be achieved. Suspension cultures are also preferred when growing large quantities of cells (e.g., to prepare large viral stocks or large

quantities of recombinant protein), because the entire process can be carried out in one flask. For small-scale procedures, by contrast, monolayer cultures are preferred because they seem less susceptible to contamination and can more easily be assessed to monitor the progress of viral infections. Monolayer cultures are absolutely required for transfections as well as for plaquing viral supernatants, and they are also preferred for preparing viral stocks or producing recombinant protein.



Most cell lines are readily adaptable to serum-free medium when in suspension. The use of serum-free medium has several advantages. First, cells can be grown to higher cell densities in serum-free medium. Second, serum-free medium is more consistent and less expensive than serum-supplemented medium. In addition, serum-free medium has a low protein content, which aids in the purification of secreted recombinant proteins. It should be determined empirically which cell line and medium formulation is most productive for the particular recombinant protein of interest. Spodoptera frugiperda (Sf 9, Sf 21) cells grown in serumfree medium are more sensitive to centrifugation, stick more tightly to plasticware, undergo a growth lag when seeded at too low a density, and cannot be passaged more than ∼50 times. In regard to purification, secreted recombinant proteins are much easier to purify than nonsecreted proteins, because the ratio between recombinant protein and host proteins in the medium is much higher than in lysates, especially when protein-free medium has been used. The general strategy for purifying protein from the medium depends on the nature of the recombinant protein. If an antibody against the desired protein is available in large quantities, it can be used for affinity purification. Otherwise, conventional ion-exchange chromatography may perform equally well.

Critical Parameters The efficiency of heterologous gene expression in the baculovirus system can differ 1000fold as a result of the intrinsic nature of the gene and the encoded protein. Modifying the heterologous gene will generally influence gene expression by only 2- to 5-fold. Researchers should not feel compelled to modify their gene excessively. For some general rules regarding the improvement of gene expression, see O’Reilly et al. (1992). When expressing protein, a time-course experiment should be conducted to determine peak protein production. To optimize for protein production, it is recommended that log-phase Sf9 cells be infected at high multiplicity of infections (MOIs)—e.g., 5 to 10 for monolayer culture and 1 to 2 for suspension culture. Cells to be infected should be >97% viable. Use of highquality media and FBS also seems to contribute to high protein yields. The most common method for radiolabeling protein involves incorporating [35S]methionine or [35S]cysteine. To improve results, the intra-

cellular pool of these essential amino acids should be depleted prior to radiolabeling by incubating the cells in methionine/cysteinefree medium. If an antibody is available, the labeled lysate should be immunoprecipitated to help detect the recombinant protein if it comigrates with a labeled host protein. To maximize large-scale production of recombinant proteins, Sf 9 cells should be grown in suspension in serum-free medium. Flasks should remain 1, it is expected that most cells in the culture will become infected at the same time and thus produce protein within the same time frame. A time course can be run to test for peak protein production. Cells infected during logphase growth with a high surface-to-volume ratio to allow for proper gas exchange will produce protein optimally. Approximately 1 to 2 mg of 6×His and GST fusion protein are routinely obtained per liter of insect-cell culture. Affinity purifications to

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>90% homogeneity are easily achieved using single-step affinity purification with Ni-NTA, glutathione agarose, or other epitope tags.

Time Considerations It takes 1.5 to 2 weeks to obtain a spinner culture from a frozen vial of Sf9 cells. A viral stock can be obtained in ∼4 days and plaqued in 1 day. It takes ∼1 week before the plaques are formed and ready to be counted. It takes an additional 2 days to infect Sf9 cells to obtain cells or medium for protein analysis. Analyzing for the recombinant protein after harvesting should take l to 3 days. Finally, it will take ∼4 days to complete the time course and another 2 to 3 days to harvest and analyze the infected cell pellets or culture supernatants.

Literature Cited Coligan, J.E., Dunn, B.M., Ploegh, H.L., Speicher, D.W., and Wingfield, P.T. (eds.) 1997. Current Protocols in Protein Science. John Wiley & Sons, New York. Guan, K. and Dixon, J.E. 1991. Eukaryotic proteins expressed in Escherichia coli: An improved thrombin cleavage and purification procedure of fusion proteins with glutathione S-transferase. Anal. Biochem. 192:262-267. Haun, R.S. and Moss, J. 1993. Ligation-independent cloning of glutathione S-transferase fusion genes for expression in Escherichia coli. Gene 112:3743. O’Reilly, D.R., Miller, L.K., and Luckow, V.A. 1992. Baculovirus Expression Vectors. W.H. Freeman, New York. Phelan, M.L., Sif, S., Narlikar, G.J., and Kingston, R.E. 1999. Reconstitution of a core chromatin remodeling complex from SWI/SNF subunits. Mol. Cell 3:247-53.

Summers, M.D. and Smith, G.E. 1987. A manual of methods for baculovirus vectors and insect cell culture procedures. Texas Agricultural Experiment Station Bulletin No. 1555. College Station, Texas. Wu, H., White, G.C., Workman, E.F., Jenzano, J.W., and Lundblad, R.L. 1992. Affinity chromatography of platelets on immobilized thrombin: retention of catalytic activity by platelet-bound thrombin. Thrombosis Res. 67:419-427. Zhang, Y., Ng, H.-H., Erdjument-Bromage, H., Tempst, P., Bird, A., and Reinberg, D. 1999. Analysis of the NuRD subunits reveals a histone deacetylase core complex and a connection with DNA methylation. Genes Dev. 13:1924-1935.

Key Reference O’Reilly et al., 1992. See above. A laboratory manual that will aid researchers in the expression and purification of recombinant proteins using the baculovirus system.

Contributed by Cheryl Isaac Murphy Aquila Biopharmaceuticals Worcester, Massachusetts Helen Piwnica-Worms Washington University School of Medicine St. Louis, Missouri Stefan Grünwald Pharmingen San Diego, California William G. Romanow Corning Costar Portsmouth, New Hampshire Nicole Francis and Hua-Ying Fan Massachusetts General Hospital Boston, Massachusetts

Purification of Proteins Using Baculovirus



EXPRESSION OF PROTEINS IN MAMMALIAN CELLS

SECTION III

Transient Expression of Proteins Using COS Cells

UNIT 16.12

This unit describes the use of COS cells to efficiently produce a desired protein in a short period of time. These cells express high levels of the SV40 large tumor (T) antigen, which is necessary to initiate viral DNA replication at the SV40 origin. Three factors contribute to make COS cell expression systems appropriate for the high-level, short-term expression of proteins: (1) the high copy number achieved by SV40 origin–containing plasmids in COS cells 48 hr post-transfection, (2) the availability of good COS cell expression/shuttle vectors, and (3) the availability of simple methods for the efficient transfection of COS cells. Each COS cell transfected with DNA encoding a cell-surface antigen (in the appropriate vector) or cytoplasmic protein will express several thousand to several hundred thousand copies of the protein 72 hr post-transfection. If the transfected DNA encodes a secreted protein, up to 10 µg of protein can be recovered from the supernatant of the transfected COS cells 1 week post-transfection. COS cell transient expression systems have also been used to screen cDNA libraries, to isolate cDNAs encoding cell-surface proteins, secreted proteins, and DNA binding proteins, and to test protein expression vectors rapidly prior to the preparation of stable cell lines (UNIT 9.5).

BASIC PROTOCOL

This transfection protocol is a modification of that presented in UNIT 9.2 and gives conditions for optimal transfection of COS cells. The main difference between this procedure and that in UNIT 9.2 is the composition of the DEAE-dextran/chloroquine solution, which is prepared here in PBS, not TBS, and contains chloroquine to prevent the acidification of endosomes presumed to carry the DEAE-dextran/DNA into the cell. (This acidification results in acid hydrolysis of the DNA, giving rise to mutations and destruction of the DNA.) With this protocol, 40% to 70% of the cells can be routinely transfected. Materials Appropriate vector (see Background Information) COS-7 cells to be transfected (see Critical Parameters), adapted to growth in 2% serum (see annotation to step 2) Dulbecco’s minimum essential medium (DMEM) with 2% calf serum (DMEM-2 CS) DMEM with 2% NuSerum (Collaborative Research) (DMEM-2 NS), 37°C Phosphate-buffered saline (PBS; APPENDIX 2) DEAE-dextran/chloroquine solution: PBS containing 10 mg/ml DEAE-dextran (Sigma) + 2.5 mM chloroquine (Sigma) 10% dimethyl sulfoxide (DMSO; Sigma) in PBS (see APPENDIX 2 for PBS) 0.5 mM EDTA in PBS 100-mm tissue culture dishes Humidified 37°C, 6% CO2 incubator Phase-contrast microscope Sorvall RT-6000B rotor (or equivalent) Protein Expression Contributed by Alejandro Aruffo Current Protocols in Molecular Biology (1998) 16.12.1-16.12.7 Copyright © 2002 by John Wiley & Sons, Inc.


Additional reagents and equipment for subcloning of DNA (UNIT 3.16), preparation of miniprep DNA (UNIT 1.6), purification of DNA by CsCl/ethidium bromide equilibrium centrifugation (UNIT 1.7), and flow cytometric analysis (Otten et al., 1995) NOTE: All cell culture incubations should be carried out in a humidified 37°C, 6% CO2 incubator unless otherwise stated. 1. Subclone the gene of interest into the appropriate vector to obtain the desired recombinant DNA (UNIT 3.16). Purify the recombinant DNA by a miniprep procedure (5-ml culture UNIT 1.6), or by CsCl/ethidium bromide centrifugation (UNIT 1.7). 2. Seed COS-7 cells in DMEM-2 CS at ∼20% confluence per 100-mm dish the day prior to transfection (so they will be ∼50% confluent the next day). Grow cells overnight to ∼50% confluence. The COS-7 cells should be adapted to 2% serum (from the usual 10% serum) by passaging them through decreasing concentrations of serum, in 2% increments, over ∼1 month. The procedure can be carried out in 10% serum, but at much greater expense. A confluent dish of COS-7 cells (∼106 cells) is usually split 1:5 on the day prior to transfection to give 2 × 105 cells/100-mm dish in 10 ml of DMEM-2 CS. In some cases cells may benefit from an extra day of growth prior to transfection (see step 6 annotation).

3. Just before use (for each 100-mm dish of COS cells to be transfected), thoroughly mix 5 ml of 37°C DMEM-2 NS with 0.2 ml of DEAE-dextran/chloroquine solution. Add 5 to 10 µg recombinant DNA and mix. It is important that the DEAE-dextran be well mixed with the medium before the DNA is added—otherwise, the DNA, a negatively charged polymer, will form large precipitates with DEAE-dextran, a positively charged polymer. These large precipitates cannot be taken up by the cell, resulting in a reduced transfection efficiency. When larger dishes are used, the amount of medium/DEAE/DNA should be sufficient to easily cover the cells and should include 400 ìg/ml DEAE-dextran, 100 ìM chloroquine, and 1 to 2 ìg/ml DNA in DMEM-2 NS. Either CsCl-purified (UNIT 1.7) or miniprep (UNIT 1.6) plasmid DNA can be used for the transfections. If miniprep DNA is used, use one-fifth of the miniprep per transfection.

4. Aspirate medium from COS cells and add the DMEM-2 NS/DEAE-dextran/DNA prepared in step 3. Incubate cells 3 to 4 hr. Observe the cells using a phase-contrast microscope. The DEAE-dextran will cause cells to retract and become vacuolated. Efficiency of transfection increases with longer incubation periods; on the other hand, so does cell death. The 3- to 4-hr incubation suggested here is a good starting point. However, several time points should be tried to optimize transfection of the particular population of cells used.

5. Aspirate DMEM/DEAE-dextran/DNA and add 5 ml of 10% DMSO (prepared in PBS). Incubate cells 2 min at room temperature. The DMSO shock results in increased transfection efficiencies. Without this step, transfection efficiencies might be lower by a factor of two or more.

6. Aspirate DMSO and add 10 ml DMEM-2 CS. Grow cells overnight (12 to 20 hr). If a significant number of the cells have lifted from the plate after overnight growth, the procedure will need to be repeated, but cells should be allowed to grow an extra day at step 2. Transient Epression of Proteins Using COS Cells

7. Passage (split and replate) each 100-mm dish of transfected COS cells into two new 100-mm dishes. After transfection, the COS cells will look unhealthy. Passaging them the day after transfection facilitates recovery, resulting in better levels of protein expression. In addition,



DEAE-dextran treatment makes the cells sticky, and passaging the cells the morning after transfection restores their adhesion characteristics so that they may be once again lifted by a gentle treatment with PBS and EDTA (see step 8b).

8a. When expressing secreted proteins: 96 hr (4 days) after completing step 7, add 5 ml DMEM-2 CS and incubate 4 days. Harvest the medium, remove dead cells and debris by centrifuging 10 min in a Sorvall RT-6000B rotor at ∼2000 rpm (∼1000 × g), room temperature, and save the supernatant (see Anticipated Results). Detect secreted proteins by bioassay (UNIT 9.5) or by metabolic labeling and immunoprecipitation (UNITS 10.18 & 10.16), immunoaffinity chromatography (UNIT 10.11), radioimmunoassay (UNIT 11.17), or western blotting (UNIT 10.8). Do not aspirate the old medium prior to addition of 5 ml DMEM-2 CS because this medium contains the secreted protein. Addition of extra medium 96 hr post-transfection results in better yield of expressed protein; however, it also increases the level of total protein (since the medium contains 2% serum), which could complicate protein purification. To eliminate this problem, COS cells can be placed in serum-free medium 10 to 12 hr after they have been replated, although (in our hands) this results in a 10-fold lower yield of expressed protein than in the presence of serum. Thus, unless it is absolutely necessary to remove additional contaminating protein, serum should be present in the medium even at reduced levels (1%).

8b. When expressing cell-surface or intracellular proteins: Aspirate medium from cells 72 hr (3 days) after transfection in step 6. Add 5 ml PBS, swirl, and aspirate PBS. Add 5 ml of 0.5 mM EDTA in PBS and incubate 15 min. Lift cells from the dish by gently dislodging them with a Pasteur pipet. Stain cell-surface proteins with the appropriate fluorescent antibody and detect by microscopy or flow cytometry (Otten et al., 1995). Transfected COS cells will tend to clump when lifted from the dish. Pipetting the cells up and down will tend to disrupt these clumps. More effective dispersion of the clumps can be obtained by forcing the cells through a 100-ìM nylon mesh.

COMMENTARY Background Information COS cells COS cells are African green monkey kidney cells (CV-1) that have been transformed with an origin-defective SV40 virus, which has integrated into COS cell chromosomal DNA. Therefore, COS cells produce wild-type SV40 large T antigen but no viral particles. Since SV40 large T antigen is the only viral protein required in trans for viral replication (i.e., its coding sequence need not be located on the DNA molecule on which it acts), SV40 origin– containing plasmids replicate in these cells to a high copy number (10,000 to 100,000 copies/cell) 48 hr post-transfection. If the plasmid carries a cDNA or genomic insert encoding a desired protein (under the control of the appropriate promoter), COS cells will express the protein at relatively high levels over a short period of time. Transfected COS cells produce protein in a burst that starts ∼24 hr post-trans-

fection and can last for up to a week. However, due to the excessive burden placed on the transfected cell by the replicating plasmid and the high levels of protein production, the transfected cells typically either die or lose the plasmid a week after transfection. COS cells were developed by Yakov Gluzman (1981) as a host for the propagation of SV40 virus early-region mutants. The first SV40 origin–containing plasmids to be used in conjunction with COS cells were made by Lusky and Botchan (1981). Short-term expression systems using both COS cells and SV40 origin–containing plasmids were initially used to identify DNA sequences required for transcription of the human α1-globin gene (Mellon et al., 1981). COS cells were first used to produce cellsurface and secreted proteins by Rose and Bergmann (1982), who looked at the expression of wild-type and mutant vesicular stomatitis virus glycoprotein in transfected cells. This

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Transient Epression of Proteins Using COS Cells

technique was subsequently used to study the expression of insulin (Laub and Rutter, 1983), somatostatin (Warren and Shields, 1984), and acetylcholine receptors (Mishina et al., 1984). These experiments demonstrated that COS cells could be used to express biologically active cell-surface and secreted proteins. Furthermore, these proteins were correctly processed although they are normally not produced by COS cells. COS cell expression was initially used to screen a cDNA library to isolate a cDNA encoding human granulocyte/macrophage colony-stimulating factor (Wong et al., 1985). This was subsequently extended to isolation of cDNAs encoding cell-surface proteins (Aruffo and Seed, 1987a; Seed and Aruffo, 1987) and DNA-binding proteins (Tsai et al., 1989). The expressed protein produced in COS cells is in most cases biologically active. However, although COS cells are able to carry out some post-translational modifications, they may not modify the expressed protein in exactly the same way as the cell that would normally produce it. For example, COS cells do not express the α-(1,3)fucosyltransferase, which is capable of transferring fucose to either sialyl or asialyl precursors (Goelz et al., 1990). In addition, insufficient post-translational modification occurs in the case of lymphocyte cell-surface proteins, which tend to be underglycosylated in COS cells (Aruffo and Seed, 1987b). This might be due to an overburdening of the COS cell glycosylation machinery by the high levels of protein expression and/or by the lack of enzymes required to carry out the full posttranslational modifications. An alternative to COS cells is provided by WOP cells (Dailey and Basilico, 1985), which are mouse 3T3 cells transformed by an origindefective polyoma virus. Like COS cells, they produce no viral particles. However, they produce polyoma large T antigen and are therefore capable of replicating a plasmid containing a polyoma origin of replication to a copy number that is typically ten times lower than that obtained in COS cells. WOP cells are also more delicate than COS cells, making them harder to transfect. For these two reasons, COS cells should be used whenever possible. However, WOP cells should be used in those cases where a monoclonal antibody that is used to identify and/or purify the protein being expressed crossreacts with COS cell proteins. This situation has arisen when transient expression in COS cells was carried out in order to clone a human

protein with a mouse anti-human monoclonal antibody that also recognizes the equivalent monkey protein (Seed and Aruffo, 1987). In this situation, the mouse cell line presented a useful alternative to COS cells by avoiding monoclonal antibody cross-reactivity. Vectors The main requirements of any COS cell expression/shuttle vector are: (1) an SV40derived origin of replication, (2) appropriate eukaryotic transcription regulatory elements (i.e., enhancer, promoter, and polyadenylation signal sequences), (3) a prokaryotic origin of replication, and (4) a prokaryotic genetic marker for selection in Escherichia coli. A particularly useful example of such a vector is CDM8 (Fig. 16.12.1; Seed, 1987). The eukaryotic transcription element of CDM8 is composed of the cytomegalovirus (CMV) enhancer-promoter, with an SV40 virus–derived intron and polyadenylation signal; the CMV promoter is a cis element (i.e., one that must be located adjacent to the DNA it acts on) that directs transcription of the DNA subcloned downstream from it. The prokaryotic genetic marker in CDM8 is provided by the supF (amber suppressor) gene. CDM8 is propagated in host bacterial cells containing helper plasmid P3, which contains amber mutations in the genes responsible for tetracycline and ampicillin resistance. (P3 has been introduced into many E. coli strains.) When CDM8 is transformed into an E. coli strain containing P3, the amber mutations are suppressed, rendering the host resistant to tetracycline and ampicillin. In addition to these elements, the CDM8 expression/shuttle vector contains an M13 origin of replication so that it can be used for the production of single-stranded DNA, a T7 RNA polymerase promoter for preparation of mRNA in vitro, and a polyoma virus–derived origin of replication which permits plasmid replication in WOP cells. Other vectors commonly used for COS cell transient expression include pXM (Yang et al., 1986) and pDC201 (Sims et al., 1988). These two plasmids contain the adenovirus-2 major late promoter and tripartite mRNA leader. This element acts in conjunction with the adenovirus VA RNA (also produced by the vectors) to increase the translatability of the mRNA encoding the desired protein. It is thought that adenovirus VA RNAs increase translation efficiency of mRNAs containing the major late promoter tripartite leader by facilitating the



enhancer- pcmv/ T7

supF

Hin dIII XbaI Xho I Bst XI

M13 ori vx ori

stuffer CDM8 ~ 4800 bp

SV40 ori

splice and poly(A)+

Bst XI XbaI Pst I Not I

Py ori

Figure 16.12.1 CDM8. CDM8 (Seed, 1987) is a COS cell expression/shuttle vector that contains an SV40-derived origin of replication (SV40 ori), eukaryotic transcription regulatory elements [splice and poly(A)+], a prokaryotic origin of replication (πvx; derived from the pBR322 ori), and a prokaryotic genetic marker (supF). In addition, CDM8 contains an M13 origin of replication (M13 ori), a T7 RNA polymerase promoter (pCMV/T7), and a polyoma-derived origin of replication (Py ori). Any restriction endonuclease sites shown can be used for cloning, but the inserted fragment must have its 5′ end nearest the enhancer pCMV/T7. The stuffer sequence is used to detect a size difference after restriction digestion.

interaction between mRNA and a 43S ribosomal protein translation preinitiation complex (Kaufman, 1985).

Critical Parameters Efficiency of transfection depends critically on the length of time that the cells (COS or other cells) are incubated in the presence of DEAEdextran/DNA. Longer periods of time result in higher transfection efficiencies. However, the DEAE-dextran/chloroquine solution is quite toxic to cells and in general, cells should not be in its presence for >4 hr. In the past, DEAEdextran transfections were carried out in the absence of serum, because a precipitate of unknown composition that seemed to be very toxic formed in DEAE-dextran/calf serum mixtures. Medium containing NuSerum, on the other hand, does not form this precipitate and tends to enhance the ability of the cells to tolerate DEAE-dextran; thus NuSerum should always be included in transfection medium. Efficiency of transfection can also be affected by the quality of the DNA and the age of the DEAE-dextran/chloroquine solution. It is preferable to use CsCl-purified or other highly purified DNA whenever possible (UNIT

1.7). However, miniprep DNA (UNIT 1.6) or DNA purified using a pZ523 column (5′→3′) or by other methods can also be used. The DEAEdextran chloroquine solution can be kept at 4°C for several months but it is wise to prepare it fresh about every 3 months. COS cells can be obtained from the American Type Culture Collection; several sublines exist, including COS-1 and COS-7. The COS-7 subline is recommended because it produces a higher plasmid copy number. These cells grow as a monolayer in DMEM-2 CS in a humidified 37°C, 5% CO2 incubator; however, the ATCC grows its COS cells in DMEM-10 fetal bovine serum (FBS). Since serum is expensive, changing the growth medium is worthwhile. The change from FBS to CS should be done slowly over 1 to 2 weeks; the concentration of CS should then be reduced to 2% in 2% increments over ∼1 month. Because the growth characteristics, transfectability, and protein expression properties of COS cells change with time and with repeated subculturing, and because these changes tend not to favor the production of high levels of proteins, it is prudent to freeze aliquots of the original COS cell stock in DMEM-10 CS/10%

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DMSO for later use in a −70°C freezer for 24 hr and then transfer them to a −150°C (liquid nitrogen) freezer. COS cells grow rapidly requiring passage every 4 to 5 days; typically, a confluent plate of cells is split 1 to 10. To obtain good levels of transient protein production from transfected COS cells, it is very important to replate the transfected cells onto new dishes with fresh medium the morning after tranfection. In addition to enhancing protein production, replating the transfected cells allows lifting of the cells from the dish using only PBS/EDTA (without trypsin). This is very important when transient expression by COS cells is used to produce cell-surface proteins.

Anticipated Results The Basic Protocol should yield transfection efficiencies of 40% to 70%. When COS cells are being used to produce cell-surface or intracellular proteins, it can be expected that each transfected cell will express several thousand copies of this protein (10,000 to 100,000 copies/cell) 72 hr post-transfection. If COS cells are used to produce secreted proteins, up to 1 µg/ml of protein can be recovered from the supernatant of a 100-mm dish of transfected cells 1 week post-transfection. However, the amount of protein produced by COS cells can vary dramatically depending on the protein being produced. This was the case when COS cells were used to produce soluble immunoglobulin fusion forms of cell-surface proteins (Aruffo et al., 1990). In this case, one of the fusion proteins, CD8 immunoglobulin, was secreted from COS cells at high levels (1 µg/ml) while the other, CD44 immunoglobulin, was secreted very poorly if at all. It was found that the CD44 fusion protein was sequestered inside the cell. To obtain efficient secretion of the CD44 fusion protein, it was necessary to change the amino-terminal signal sequence of the CD44 fusion protein. Interestingly, the native cell-surface forms of both CD8 and CD44 are expressed equally efficiently on the surface of transfected COS cells. For some of these immunoglobulin fusion proteins, 0.5 ml of medium contained plenty of protein (∼500 ng) after concentration using a protein A affinity matrix. In some cases, it is possible to use such COS cell supernatants directly without further purification (Aruffo et al., 1990). Transient Epression of Proteins Using COS Cells

Time Considerations It is important not to transfect the cells too soon after replating; >8 to 12 hr should pass

between the time the cells are seeded on the plate and the time of transfection. Once the transfection has started, the DEAE-dextran/DNA mixture should be left on the cells for a minimum of 2 hr and a maximum of 4 hr; because the mixture increases transfection efficiency it should remain in contact with the cells as long as they appear viable. After transfection, the cells look quite unhealthy, and 12 to 24 hr post-transfection they should be replated in new dishes with fresh medium. The peak of plasmid replication in transfected COS cells occurs 48 to 72 hr post-transfection. Protein production starts 24 hr posttransfection but peaks 72 to 96 hr post-transfection. Thus, when expressing cell-surface or cytoplasmic proteins, the cells should be harvested 72 to 96 hr post-transfection. However, transfected cells continue to produce protein for up to a week post-transfection and when expressing secreted proteins, the supernatants should be harvested a week post-transfection.

Literature Cited Aruffo, A. and Seed, B. 1987a. Molecular cloning of a CD28 cDNA by a high-efficiency COS cell expression system. Proc. Natl. Acad. Sci. U.S.A. 84:8573-8577. Aruffo, A. and Seed, B. 1987b. Molecular cloning of two CD7 (T cell leukemia antigen) cDNAs by a COS cell expression system. EMBO J. 6:33133316. Aruffo, A., Stamenkovic, I., Melnick, M., Underhill, C.B., and Seed, B. 1990. CD44 is the principal cell-surface receptor for hyaluronate. Cell 61:1303-1313. Dailey, L. and Basilico, C.J. 1985. Sequences in the polyoma virus DNA regulatory region involved in viral DNA replication and early gene expression. J. Virol. 54:739-749. Gluzman, Y. 1981. SV-40 transformed simian cells support the replication of early SV40 mutants. Cell 23:175-182. Goelz, S.E., Hession, C., Goff, D., Griffiths, B., Tizard, R., Newman, B., Chi-Rosso, G., and Lobb, R. 1990. ELFT: A gene that directs the expression of an ELAM-1 ligand. Cell 63:13491356. Kaufman, R.J. 1985. Identification of the components necessary for adenovirus translational control and their utilization in cDNA expression vectors. Proc. Natl. Acad. Sci. U.S.A. 82:689693. Laub, O. and Rutter, W.J. 1983. Expression of the human insulin gene and cDNA in a heterologous mammalian system. J. Biol. Chem. 258:60436050. Lusky, M. and Botchan, M. 1981. Inhibition of SV40 replication in simian cells by specific pBR322 DNA sequences. Nature 293:79-81.



Mellon, P., Parker, V., Gluzman, Y., and Maniatis, T. 1981. Identification of DNA sequences required for transcription of the human α1-globin gene in a new SV40 host-vector system. Cell 27:279288.

Tsai, F., Martin, D.I.K., Zon, L.I., D’Andrea, D., Wong, G.G., and Orkin, S.H. 1989. Cloning of cDNA for the major DNA-binding protein of the erythroid lineage through expression in mammalian cells. Nature 339:446-451.

Mishina, M., Kurosaki, T., Tobimatsu, T., Morimoto, Y., Noda, M., Yamamoto, T., Terao, M., Lindstrom, J., Takahashi, T., Kuno, M., and Numa, S. 1984. Expression of functional acetylcholine receptor from cloned cDNAs. Nature 307:604-608.

Warren, T.G. and Shields, D. 1984. Expression of preprosomatostatin in heterologous cells: Biosynthesis, posttranslational processing, and secretion of mature somatostatin. Cell 39:547-555.

Otten G., Yokoyama, W.M. and Holmes, K.H. 1995. Flow cytometry analysis using the Becton Dickinson FACScan. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 5.4.1-5.4.19. John Wiley & Sons, New York. Rose, J.K. and Bergmann, J.E. 1982. Expression from cloned cDNA of cell-surface, secreted forms of the glycoprotein of vesicular stomatitis virus in eucaryotic cells. Cell 30:753-762. Seed, B. 1987. An LFA-3 cDNA encodes a phospholipid-linked membrane protein homologous to its receptor CD2. Nature 329:840-842. Seed, B. and Aruffo, A. 1987. Molecular cloning of the CD2 antigen, the T cell erythrocyte receptor, by a rapid immunoselection procedure. Proc. Natl. Acad. Sci. U.S.A. 84:3365-3369. Sims, J.E., March, C.J., Cosman, D., Widmer, M.B., MacDonald, H.R., McMahan, C.J., Grubin, C.E., Wignall, J.M., Jackson, J.L., Call, S.M., Friend, D., Alpert, A.R., Gillis, S., Urdal, D.L., and Dower, S.K. 1988. cDNA expression cloning of the IL-1 receptor, a member of the immunoglobulin superfamily. Science 241:585-589.

Wong, G.G., Witek, J.S., Temple, P.A., Wilkens, K.M., Leary, A.C., Luxenber, D.P., Jones, S.S., Brown, E.L., Kay, R.M., Orr, E.C., Shoemaker, C., Golde, D.W., Kaufman, R.J., Hewick, R.M., Wang, E.A., and Clark, S.C. 1985. Human GMCSF: Molecular cloning of the complementary DNA and purification of the natural and recombinant proteins. Science 228:810-815. Yang, Y., Ciarletta, A.B., Temple, P.A., Chung, M.P., Kovacic, S., Witek-Giannotti, J.S., Leary, A.C., Kriz, R., Donahue, R.E., Wong, G.G., and Clark, S.C. 1986. Human IL-3 (multi-CSF): Identification by expression cloning of a novel hematopoietic growth factor related to murine IL3. Cell 47:3-10.

Key Reference Warren and Shields, 1984. See above. This article shows that COS cells can be used as an efficient, short-term, mammalian epression system for the production of proteins.

Contributed by Alejandro Aruffo Bristol-Myers Squibb Princeton, New Jersey

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Expression and Purification of EpitopeTagged Multisubunit Protein Complexes from Mammalian Cells

UNIT 16.13

Biochemical characterization and functional studies of mammalian proteins are often hampered by the availability of the purified protein, in particular, when the functional entity is present as a multisubunit protein complex in the cell. To overcome the difficulties in the purification of multisubunit protein complexes from mammalian cells, one may create stable cell lines containing epitope-tagged protein. In general, a stable cell line that expresses an epitope-tagged subunit of a protein complex is first established either by retrovirus-mediated gene transfer (for constitutive expression) or by a tetracycline-regulated system (for inducible expression). Immunoaffinity purification using epitope-specific monoclonal antibody-conjugated beads is then performed to pull down the epitope-tagged multisubunit protein complex, which is finally recovered by peptide elution under neutral pH or physiological conditions and is readily available for functional assays. Several commonly used epitopes are listed in Table 10.15.1 (UNIT 10.15). For simplicity, only protocols using the FLAG epitope (Asp-Tyr-Lys-Asp-Asp-Asp-Asp-Lys, or DYKDDDDK) are described here. The authors normally use human HeLa cells, which can grow in monolayer as well as in suspension, as parental cells for establishing stable cell lines, although other cells, such as human 293, can also be used for the experiments. By and large, HeLa-derived cell lines expressing an epitope-tagged protein are initially isolated in monolayer culture and then adapted to suspension culture for the purpose of collecting large quantities of cells for protein purification. The first protocol in this unit (see Basic Protocol 1) describes the procedures involved in the establishment of a stable cell line constitutively expressing the FLAG-tagged protein by retrovirus-mediated gene transfer and immunoaffinity purification of the epitope-tagged multisubunit protein complex. The next protocol (see Basic Protocol 2) outlines the steps involved in the establishment of an inducible cell line conditionally expressing the FLAG-tagged protein by a tetracycline-regulated system, and the one-step immunoaffinity purification of the multisubunit protein complex. Since an epitope-tagged protein may exist as different complexes in the cell, it is also important to isolate various forms of multisubunit protein complexes. This can sometimes be achieved simply by careful selection of the starting material (e.g., using nuclear extract, cytoplasmic S100 fraction, or the chromosomal fraction) for immunoaffinity purification or by manipulating the wash conditions with different concentrations of salt, detergent, and chaotropic agents during immunoaffinity purification. The Alternate Protocol provides an excellent example for the purification of different forms of human RNA polymerase II complexes, achieved simply by choosing the appropriate starting material and by varying wash conditions. Frequently, multiple forms of the epitope-tagged protein complexes coexist in the same starting material. In that case, a simple chromatographic column, such as the P11 ion-exchange resin, can be used to separate multiple forms of protein complexes into different fractions prior to immunoaffinity purification. The isolation of various human TATA-binding protein (TBP)–containing complexes, described in the Support Protocol, is a good example of combining the P11 column and immunoaffinity purification. These protocols, collectively, illustrate a powerful methodology in applying epitope tagging and stable cell line approaches for the purification of multisubunit protein complexes from mammalian cells. Protein Expression Contributed by Shwu-Yuan Wu and Cheng-Ming Chiang Current Protocols in Molecular Biology (2001) 16.13.1-16.13.17 Copyright © 2002 by John Wiley & Sons, Inc.


BASIC PROTOCOL 1

PURIFICATION OF MULTISUBUNIT PROTEIN COMPLEXES FROM CLONAL CELL LINES CONSTITUTIVELY EXPRESSING A FLAG-TAGGED PROTEIN To purify a multisubunit protein complex, the authors usually begin with a constitutive expression system using retrovirus-mediated gene transfer for the establishment of clonal cell lines expressing the FLAG-tagged subunit of a protein complex. Retrovirus-mediated gene transfer (UNITS 9.9-9.14) provides a gene delivery system with few copies of the viral integrant to ensure the expressed FLAG-tagged protein can be efficiently assembled into a large protein complex, in a stoichiometric ratio, with endogenous cellular proteins. A retroviral vector containing a drug-selection marker and the FLAG sequence introduced either at the N-terminus or at the C-terminus of the protein-coding region is first created and delivered into a packaging cell line that provides all the viral proteins essential for virion assembly. Viral particles are then collected from the cell culture medium and used to infect HeLa or other chosen cells. Cellular clones potentially expressing the tagged protein are initially selected by drug resistance and further identified by immunoblotting from individually expanded colonies. Clonal cell lines expressing the FLAG-tagged protein are then adapted to suspension culture and used for immunoaffinity purification of the epitope-tagged protein complexes. NOTE:All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Materials FLAG-tagged protein-coding sequence Retroviral vector (e.g., pBabe neo, Morgenstern and Land, 1990; also see UNITS 9.9-9.14) 2× HEPES-buffered saline (HBS), pH 7.12 (UNIT 16.14) 0.25 M CaCl2 Retrovirus-packaging cells (e.g., ψCRIP, Danos and Mulligan, 1988) Dulbecco’s modified Eagle’s medium (DMEM) with and without 10% calf serum 15% (v/v) glycerol/DMEM DMEM with 10% fetal bovine serum (FBS; APPENDIX 3F) 2.5 mg/ml polybrene stock solution 50% confluent HeLa cells Drug-containing selection medium (e.g., 1 mg/ml of G418 (total weight), 0.2 mg/ml of hygromycin, or 0.5 µg/ml of puromycin; see UNIT 9.5) Trypsin-EDTA (APPENDIX 3F) 1× PBS (APPENDIX 2) 1× SDS-PAGE protein sample buffer (UNIT 10.2A) Anti-FLAG M2 monoclonal antibody (and/or anti-protein antibodies; Sigma) Joklik’s medium (Sigma) with 10% calf serum and 5% calf serum Anti-FLAG M2 monoclonal antibody-conjugated beads (Sigma) BC100 (see recipe) BC300 (see recipe)/0.1% Nonidet P-40 FLAG peptide (100 mg/ml in water) Liquid nitrogen

Purification of Epitope-Tagged Multisubunit Protein Complexes

5-ml round-bottom Falcon tube 60-mm and 100-mm tissue culture plates 15-ml sterile tube 0.45-µm cellulose acetate syringe filter 24-well tissue culture plate 1.5-ml and 0.5-ml microcentrifuge tubes



250-ml, 500-ml, 1-liter, 3-liter, and 12-liter spinner flasks End-over-end tube rotator Rotor (e.g., Sorvall H-6000A or equivalent) Microcentrifuge spin column Additional reagents and equipment for use of retroviral vectors (UNITS 9.9-9.14), preparation of nuclear extracts (UNIT 12.1), and immunoblotting (UNIT 10.8) Establish stable cell lines by retrovirus-mediated gene transfer 1. Clone the FLAG-tagged protein-coding sequence into a retroviral vector containing a drug-selection marker (e.g., neomycin). UNITS 9.9-9.14 describe

the use of retroviral vectors.

2. Transfer 20 µg plasmid DNA into a 5-ml round-bottom Falcon tube and mix with 0.5 ml of 2× HBS, pH 7.12. The following transfection procedure works well for ψCRIP and many other packaging cell lines.

3. Add 0.5 ml of 0.25 M CaCl2 dropwise while vortexing. Leave for 20 to 30 min at room temperature. 4. Vortex the calcium phosphate–DNA coprecipitates vigorously and then add the solution to 50% confluent ψCRIP cells, which are maintained in DMEM plus 10% calf serum in a 100-mm tissue culture plate. Leave cells for 3 to 4 hr in a 37°C, 5% CO2 incubator. 5. Remove medium and add 3 ml of 15% (v/v) glycerol/DMEM for 3 min at room temperature. 6. Add 8 ml DMEM to quickly dilute glycerol, mix well, and remove solution. 7. Repeat washes three additional times with 3 ml DMEM for each wash. 8. Add 10 ml DMEM with 10% calf serum to transfected cells and leave plates for 18 hr in a 37°C, 5% CO2 incubator. 9. Collect viral particles in a 15-ml sterile tube after filtering cell culture supernatant (18-hr post glycerol shock) through a 0.45-µm cellulose acetate syringe filter to remove floating cells and large cellular debris. 10. Mix 1 ml of filtered viral stock with 1 ml of DMEM plus 10% FBS and polybrene (final concentration 8 µg/ml), and incubate mixture with 50% confluent HeLa cells, which are grown in DMEM with 10% FBS in 100-mm cell culture plates and have been prewashed with medium two times, for 2.5 hr at 37°C with intermittent mixing. 11. Add an additional 8 ml of DMEM with 10% FBS, without removing the original infection medium, and leave cells for 1 to 1.5 days in a 37°C, 5% CO2 incubator. 12. Split cells 1:5 in drug-containing medium and change the selection medium every 3 to 4 days. The drug concentrations normally used for HeLa cells, depending on individual drug-selection markers, are: 1 mg/ml of G418 (total weight), 0.2 mg/ml of hygromycin, or 0.5 ìg/ml of puromycin. See UNIT 9.5 for additional details. Drug-resistant colonies are usually visible after 2 to 3 weeks. Protein Expression


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Identify clonal cell lines expressing FLAG-tagged protein 13. Prepare a 24-well plate and add 2 drops (∼100 µl) of trypsin-EDTA to each well at alternate positions to avoid cross contamination between neighboring wells. 14. Circle the drug-resistant colonies (∼2- to 3-mm diameter), to be picked up, on the bottom of the plate with a black (or blue) marker pen, remove medium, wash cells once with 1× PBS, and then remove solution. 15. Take ∼50 µl of trypsin-EDTA from the first well using a 200-µl pipet tip and transfer a circled colony back to the first well after pipetting up and down several times. 16. Repeat the transfer process for a total of 12 colonies. 17. Add 1 to 2 ml of selection medium to each well and put the 24-well cell culture plate back to a 37°C, 5% CO2 incubator for cells to attach and grow. Depending on the number of viable cells transferred, it may take 3 to 14 days for cells to become confluent in the wells.

18. Transfer cells to a 60-mm cell culture plate when they become confluent in the 24-well plate and continue the expansion for individual cellular clones. 19. When cells are nearly confluent on the 60-mm cell culture plate, split cells again to one 60-mm cell culture plate and two 100-mm cell culture plates. The cells grown on the 100-mm plates will be frozen down as individual cloned cell lines.

20. Prepare whole-cell lysates from 80% to 90% confluent cells still in log phase grown on the 60-mm cell culture plate by adding 300 µl of 1× SDS-PAGE protein sample buffer to plate, after washing cells two times with 1× PBS. Pipet lysate up and down many times until the sample is no longer viscous, and then transfer the lysate to a 1.5-ml microcentrifuge tube. 21. Perform immunoblotting (UNIT 10.8) on individually collected whole-cell lysates with anti-FLAG M2 monoclonal antibody and/or anti-protein antibodies to identify the cellular clones that express FLAG-tagged protein. Purify FLAG-tagged protein complexes from established cell lines 22. Select a positive clone expressing FLAG-tagged protein for further expansion by adapting it to suspension culture. Combine cells from twelve 100-mm cell culture plates into a 250-ml spinner flask and adjust the cell density to 0.5 × 106 cells/ml using Joklik’s medium with 10% FBS. Maintain cells at the same density using Joklik’s medium with 10% FBS for ≥3 days. If cells are healthy and nearly grow exponentially, start to feed cells using Joklik’s medium with 10% calf serum. Continue feeding cells using Joklik’s medium with 10% calf serum for at least three days. If cells still look healthy and grow well, begin to feed cells using Joklik’s medium with 7.5% (or 5%) calf serum. During the expansion and serum-switching/reducing process, if cells look unhealthy or cell density drops, centrifuge cells for 5 min at 300 × g and resuspend cells in fresh Joklik’s medium with the same concentration of calf serum, this helps to remove cell debris. Readjust the cell density to 0.5 × 106 cells/ml. Once cells are healthy and continue to grow, begin to lower the concentration of serum as previously described. Repeat the process until cells are maintained in Joklik’s medium with 5% calf serum. Purification of Epitope-Tagged Multisubunit Protein Complexes

23. Grow and expand the cells growing in Joklik’s medium with 5% calf serum continually to 20 liters or any other desired volume.



24. Prepare nuclear extract, cytoplasmic S100, and nuclear pellet (or chromosomal fraction) according to UNIT 12.1 (Dignam et al., 1983). 25. Perform immunoblotting (UNIT FLAG-tagged protein.

10.8)

to identify the cellular fractions containing

26. Add 0.4 ml of anti-FLAG M2 monoclonal antibody-conjugated beads, which have been prewashed several times with BC100, to a 15-ml tube containing 10 to 14 ml of nuclear extract (or S100, or solubilized nuclear pellet). Incubate the sample for 6 to 12 hr at 4°C on an end-over-end tube rotator. 27. Centrifuge for 2 min at 300 × g (Sorvall H-6000A rotor 1000 rpm), 4°C, to separate supernatant from beads. 28. Remove the supernatant and wash protein-bound beads five times with BC300/0.1% Nonidet P-40. Inclusion of Nonidet P-40 prevents protein/bead aggregation and also helps the immunoaffinity beads rotate better during the elution step.

29. Transfer the M2-conjugated beads with bound proteins to a microcentrifuge spin column. 30. Microcentrifuge for 10 sec at maximum speed (10,000 rpm) to remove the residual liquid. 31. Add 0.5 ml of BC300/0.1% Nonidet P-40 plus 0.2 mg/ml of FLAG peptide to the dried beads. Mix well and incubate for 20 to 60 min at 4°C with continuous rotation. 32. Collect the eluate (i.e., the first elution) by centrifuging for 10 sec at 4°C. 33. Repeat the elutions (steps 31 and 32) for a total of four times to recover the majority of FLAG-tagged protein complexes. 34. Dispense samples into small aliquots (e.g., 20 µl) in 0.5-ml microcentrifuge tubes and freeze in liquid nitrogen. Store aliquots of the purified complexes at −80°C until use. PURIFICATION OF MULTISUBUNIT PROTEIN COMPLEXES FROM CLONAL CELL LINES CONDITIONALLY EXPRESSING THE FLAG-TAGGED PROTEIN

BASIC PROTOCOL 2

Sometimes, the introduced FLAG-tagged protein-coding sequence is not expressed in the established clonal cell lines following retrovirus-mediated gene transfer and drug selection. This is likely due to cytotoxicity resulting from unregulated or constitutive expression of the ectopic gene product. To overcome this problem, an inducible mammalian expression system can be used to turn “on” or “off” expression of the FLAG-tagged protein based on the use of tetracycline. In the process of establishing FLAG-tagged protein-expressing cell lines, tetracycline is continuously added to silence ectopic gene expression. The inducibility of individual cellular clones is then screened by immunoblotting using whole-cell lysates collected in the presence and absence of tetracycline. Once identified, the inducible cell line is adapted to suspension culture and used for immunoaffinity purification of FLAG-tagged multisubunit protein complexes after removing tetracycline.

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Materials Protein-coding sequence Tetracycline-regulated expression plasmid (e.g., pTetCMV-F°(S), Wu and Chiang, 1996; UNIT 16.14) Sheared calf thymus DNA SacI-linearized plasmid containing a drug-selection marker (e.g., pREP4, Invitrogen) Cell line expressing a tetracycline-controlled transactivator (e.g., HtTA-1, Gossen and Bujard, 1992) DMEM with 10% FBS (APPENDIX 3F) G418 sulfate Trypsin-EDTA (APPENDIX 3F) 1 M BES buffer, pH 7.2 (see recipe) Hygromycin B Tetracycline 1× SDS-PAGE protein sample buffer (UNIT 10.2A) 1× PBS (APPENDIX 2) Anti-FLAG M2 monoclonal antibody and/or anti-protein antibodies Joklik’s medium (Sigma) with 10% calf serum Joklik’s medium with 5% calf serum HeLa cells Anti-FLAG M2-conjugated beads BC100 and BC300 (see recipe) 10% Nonidet P-40 FLAG peptide Liquid nitrogen 0.4-cm cuvettes for electroporation 24-well, 60-mm, and 100-mm tissue culture plates 15-ml and 50-ml Falcon tubes Rotor (e.g., Sorvall H-6000A or equivalent) Electroporator (e.g., Bio-Rad Gene Pulser II) Disposable glass pipet 1.5-ml and 0.5-ml microcentrifuge tubes 250-ml, 500-ml, 1-liter, 3-liter, and 12-liter spinner flasks 250-ml sterile conical centrifuge tubes Microcentrifuge spin column Additional reagents and equipment for tissue culture (APPENDIX 3F) and immunoblotting (UNIT 10.8) Establish tetracycline-regulated cell lines by electroporation 1. Clone the protein-coding sequence into a tetracycline-regulated expression plasmid containing the FLAG epitope sequence, such as pTetCMV-F°(S). 2. Mix 10 µg of linearized tetracycline-regulated FLAG-tagged protein-expressing plasmid (e.g., PvuI used to linearize most pTetCMV-F°(S)-derived constructs), 50 µg of sheared calf thymus DNA, and 0.5 µg of SacI-linearized pREP4, which contains a hygromycin-selection marker, into an electroporation cuvette.


3. Combine HtTA-1 cells (or other cells expressing a tetracycline-controlled transactivator), which are grown in DMEM with 10% FBS containing 0.6 mg/ml of G418, from several 100-mm cell culture plates into a 50-ml Falcon tube after trypsin-EDTA treatment, centrifuge for 5 min at 300 × g (Sorvall H-6000A rotor 1000 rpm), and



resuspend cell pellets in DMEM with 10% FBS and 5 mM BES buffer, pH 7.2, at a density of 2 × 107 cells/ml. 4. Add 250 µl of resuspended cells, using a 200-µl pipet tip to transfer 125 µl of cells two times, to the electroporation cuvette containing plasmids and carrier DNA (see step 2). Mix well by pipetting up and down several times. 5. Electroporate DNAs into cells using the Bio-Rad Gene Pulser II with setting at 960 µF and 200 V. Leave cuvette for 10 min at room temperature and then transfer cells, using a disposable glass pipet, to a 15-ml centrifuge tube containing 10 ml DMEM with 10% FBS. 6. Centrifuge for 5 min at 300 × g. Aspirate and discard the supernatant, which contains untransfected DNAs and cell debris generated by electroporation. 7. Resuspend cells in 5 ml DMEM with 10% FBS and distribute cells onto five 100-mm cell culture plates, each containing 9 ml DMEM with 10% FBS, and leave for 2 days in a 37°C, 5% CO2 incubator. 8. Initiate drug selection by changing medium to DMEM with 10% FBS containing 400 µg/ml of G418, 200 µg/ml of hygromycin B, and 2 µg/ml of tetracycline. 9. Change medium every 3 or 4 days and continue drug selection for ∼3 weeks until drug-resistant colonies are clearly visible. Identify clonal cell lines conditionally expressing FLAG-tagged protein 10. Pick up 12 to 24 drug-resistant colonies and expand them into cell lines following the procedures described in Basic Protocol 1, steps 13 to 18. 11. When cells are nearly confluent on the 60-mm cell culture plate during expansion, split cells to one 100-mm cell culture plate and two 60-mm cell culture plates. The cells grown on the 100-mm cell culture plate will be frozen down as individual cloned cell lines. The cells grown on the two 60-mm cell culture plates are plated in selection medium, one with and the other without tetracycline, for 48 to 72 hr.

12. Prepare whole-cell lysates from cells grown on the two 60-mm cell culture plates by adding 300 µl of 1× SDS-PAGE protein sample buffer to each plate, after washing the cells two times with 1× PBS. Pipet the lysate up and down many times until the sample is no longer viscous and then transfer lysate to a 1.5-ml microcentrifuge tube. 13. Perform immunoblotting (UNIT 10.8) on an aliquot of the individually collected whole cell lysates with anti-FLAG M2 monoclonal antibody and/or anti-protein antibodies to identify the cellular clones that express FLAG-tagged protein only in the absence of tetracycline. Purify FLAG-tagged protein complexes from established cell lines 14. Select a positive clone expressing FLAG-tagged protein for further expansion by adapting it to suspension culture. Combine cells from twelve 100-mm cell culture plates into a 250-ml spinner flask (at a density of 0.5 × 106 cells/ml) and gradually replace medium from DMEM with 10% FBS to Joklik’s medium with 10% calf serum, and eventually to Joklik’s medium with 5% calf serum, but include 1 µg/ml of tetracycline and 0.6 mg/ml of G418 to the medium throughout the expansion process until culture volume reaches 250 ml. 15. Continue to expand cells in tetracycline-containing medium, but omit G418 after 250 ml culture.

Protein Expression


Supplement 60

16. After expansion of cell culture to 12 liters, remove tetracycline by centrifuging cells for 5 min at 300 × g (Sorvall H-6000A rotor 1000 rpm) using 250-ml sterile conical tubes. Repeat the process until all of the cells are consolidated into two 250-ml conical tubes. 17. Wash cell pellets with 250 ml of 1× PBS in each tube. Centrifuge for 5 min at 300 × g (Sorvall H-6000A rotor 1000 rpm). Repeat washes for a total of 4 to 6 times. 18. Resuspend cell pellets in ∼120 ml of Joklik’s medium with 5% calf serum without tetracycline, and dispense cells into six 12-liter flasks, each containing 1.5 liters of Joklik’s medium with 5% calf serum. 19. Four days after protein induction with medium doubling every day, prepare nuclear extract, cytoplasmic S100, and nuclear pellet (or chromosomal fraction) according to UNIT 12.1. Protein induction in suspension culture usually takes longer than induction in monolayer culture. From the time course study, protein induction after 4 days maximizes the quantity of the induced FLAG-tagged protein, in part due to a reduced doubling time in a larger volume of suspension culture using less expensive and lower concentrations of calf serum. Induction also appears to work better under a slightly dilute condition with medium doubling every day.

20. Perform immunoblotting (UNIT FLAG-tagged protein.

10.8)

to identify the cellular fractions containing

21. For immunoaffinity purification of FLAG-tagged protein complexes, follow the same procedures as described in Basic Protocol 1, steps 26 to 34. ALTERNATE PROTOCOL

PURIFICATION OF MULTIPLE FORMS OF EPITOPE-TAGGED PROTEIN COMPLEXES BY VARYING THE STARTING MATERIAL AND WASH CONDITIONS Using human RNA polymerase II (pol II) as an example, this protocol describes the purification of epitope-tagged protein complexes. RNA pol II is a multisubunit protein complex containing 12 polypeptides (RPB1-12). The largest subunit (RPB1) of pol II has a unique C-terminal domain (CTD) containing heptapeptide sequence, Tyr-Ser-Pro-ThrSer-Pro-Ser (YSPTSPS), which occurs 26 times in yeast and 52 times in humans. This CTD can be phosphorylated by many protein kinases, resulting in three forms of pol II commonly found in eukaryotes. The IIO form of pol II contains a highly or hyper-phosphorylated CTD, whereas the IIA form of pol II has a non- or hypo-phosphorylated CTD. The IIB form of pol II does not have the CTD, likely generated due to protease cleavage. In addition, pol II can form an extremely large protein complex, so-called pol II holoenzyme, by association with a subset of general transcription factors and other cellular proteins involved in chromatin remodeling, mRNA processing, and DNA repair. This Alternate Protocol exemplifies an application of the epitope-tagging and stable cell line approaches for the simultaneous purification of the IIO, IIA, and IIB forms of pol II as well as pol II holoenzyme, which contains only the nonphosphorylated form of RPB1. This is achieved simply by careful selection of the starting material and by modifying the composition of protein buffer used for immunoaffinity purification (see Fig. 16.13.1). Please note that the conditions outlined herein will vary with other tagged protein systems.


Additional Materials (also see Basic Protocols 1 and 2) 10 M urea BC850 (see recipe) Buffer B (see recipe)



Buffer D (see recipe) FLAG peptide elution buffer (see recipe) Ammonium sulfate Sonicator Rotor (e.g., Bechman 45-Ti or equivalent) Purify FLAG-tagged human pol II holoenzyme 1a. Establish a tetracycline-regulated HeLa-derived cell line conditionally expressing a FLAG-tagged subunit of human pol II, and prepare nuclear extract, cytoplasmic S100, and nuclear pellet according to the procedures described in Basic Protocol 2, steps 1 to 20. 2a. For immunoaffinity purification of FLAG-tagged human pol II holoenzyme from nuclear extract or S100, follow Basic Protocol 1, steps 26 to 34, except use BC100/0.1% Nonidet P-40 as wash and elution buffer. Purify IIA form of FLAG-tagged human pol II 1b. For immunoaffinity purification of FLAG-tagged human pol II from nuclear extract or S100, follow Basic Protocol 1, steps 26 to 34, except wash the protein-bound beads sequentially five times each with BC850/0.1% Nonidet P-40 containing 1.0 M urea,

cell line expressing FLAG-tagged human RPB9 (hRPB9-3)

cytoplasmic S100

nuclear extract

nuclear pellet

p low salt

GCN5

pol II

TFIIB

TFIIF TFIIE TFIIH

pol II

p p

p

high salt pol II

SWI/SNF SRBs

p

IIO

pol II

IIA

pol II

IIB

pol II holoenzyme

Figure 16.13.1 Purification of human RNA polymerase II complexes. The hRPB9-3 cells (Wu and Chiang, 1998), derived from human HeLa cells that conditionally express the FLAG-tagged RPB9 subunit of human RNA polymerase II, are first separated into cytoplasmic S100, nuclear extract, and nuclear pellet. Immunoaffinity purification is then performed with S100 or nuclear extract under either low salt (100 mM KCl-containing buffer) or high salt (850 mM KCl and 1.0 M urea-containing buffer) wash conditions, which results in the purification of human RNA polymerase II (pol II) holoenzyme and pol II, respectively. The pol II holoenzyme complex purified from S100 or nuclear extract contains pol II, a subset of general transcription factors (TFIIB, TFIIE, TFIIF, and TFIIH), SRBs, histone acetyltransferase GCN5, and chromatin remodeling factor SWI/SNF (Wu and Chiang, 1998; Wu et al., 1999). Additional forms of pol II complexes with different phosphorylation status on the largest subunit (RPB1) of pol II can also be purified from the nuclear pellet under the high salt wash condition. The IIO form of pol II contains the highly phosphorylated carboxy-terminal domain (CTD), indicated by a tail, in RPB1, whereas the IIA and IIB forms of pol II have non-phosphorylated or truncated CTD in RPB1, respectively.

Protein Expression


Supplement 60

and BC100, then finally elute the bound proteins with BC100 containing 0.2 mg/ml of FLAG peptide (FLAG peptide elution buffer). Inclusion of high salt and urea during the washes disrupts the interactions between pol II and other weakly associated polypeptides found in the pol II holoenzyme, thereby allowing the recovery of only the IIA form of the core pol II, which forms a tight complex resistant to high salt and urea washes.

Purify FLAG-tagged human pol II containing a mixture of IIA, IIB, and IIO forms 1c. Slowly agitate 35 ml of nuclear pellet isolated from hRPB9-3, which is a HeLa-derived cell line conditionally expressing the FLAG-tagged RPB9 subunit of human pol II, with 70 ml of buffer B at 4°C (or on ice) and then with 11 ml of 3 M (NH4)2SO4 to adjust the ammonium sulfate concentration to 0.3 M. Stir for an additional 30 min. The use of buffer B together with ammonium sulfate provides both a magnesium ion and high salt concentration to help dissociate pol II complexes from the chromosomal fraction and at the same time preserve pol II activity.

2c. Sonicate five times, for 1 min each, on an ice water slurry with 20-sec intervals between bursts. 3c. Centrifuge the mixture for 90 min at 185,000 × g (Beckman 45-Ti rotor 40,000 rpm), 4°C, and then pour the supernatant into a 500-ml beaker. 4c. Add 2 volumes of buffer B dropwise with a syringe to gradually adjust the ammonium sulfate concentration to 0.1 M. 5c. Remove precipitated material by centrifuging for 60 min at 185,000 × g (Beckman 45-Ti rotor 40,000 rpm) and pour supernatant to another 500-ml beaker. 6c. Precipitate pol II by slowly adding solid ammonium sulfate to 65% saturation (i.e., 0.42 g/ml of suspension) and stir for an additional 30 min. 7c. Centrifuge for 60 min at 142,000 × g (Beckman 45-Ti rotor 35,000 rpm) and resuspend the pellet in 40 ml of buffer D. 8c. Dialyze the sample against 4 liters of BC100 for 6 hr with at least one change of buffer. The final recovered sample is ∼46 ml.

9c. For immunoaffinity purification of FLAG-tagged human pol II complexes containing a mixture of IIA, IIB, and IIO forms, follow Basic Protocol 1, steps 26 to 34, except sequentially wash the protein-bound beads with five times each of BC850/0.1% Nonidet P-40 with 1.0 M urea, and BC100, before final elutions with BC100 containing 0.2 mg/ml of FLAG peptide (FLAG peptide elution buffer). The inclusion of high salt and urea during the initial washes disrupts the interactions between pol II and other weakly associated polypeptides found in the pol II holoenzyme, thereby allowing the recovery of core pol II complexes with different phosphorylation states. Equilibration of the immobilized complexes to BC100 is necessary before final elutions with FLAG peptide BC100 elution buffer.




PURIFICATION OF MULTIPLE FORMS OF EPITOPE-TAGGED PROTEIN COMPLEXES FOLLOWING A P11 ION-EXCHANGE CHROMATOGRAPHIC COLUMN

SUPPORT PROTOCOL

Human TATA-binding protein (TBP) can associate with different classes of cellular proteins to form distinct protein complexes, such as SL1, TFIID, and TFIIIB, which are required for transcription by RNA polymerase I, II, and III, respectively. These distinct protein complexes are all found in nuclear extract. A simple chromatographic step, such as a P11 ion-exchange column, is then used to separate various TBP-containing complexes into different fractions prior to immunoaffinity purification (see Fig. 16.13.2). This Support Protocol illustrates an application of conventional chromatography together with immunoaffinity purification for the isolation of multiple protein complexes containing a common subunit. Additional Materials (also see Basic Protocol 1) P11 ion-exchange resin Various BC buffers (BC100, BC300, BC500, BC850, and BC1200; see recipe) Chromatographic column (∼100 ml capacity) Flow adapter Chart recorder Fraction collector Dialysis tubing (MWCO 12,000; Sigma) Additional reagents and equipment for dialysis (APPENDIX 3C) Purify TFIID and TFIIIB from a cell line expressing FLAG-tagged human TBP 1. Establish a HeLa-derived cell line expressing FLAG-tagged human TBP by retrovirus-mediated gene transfer (UNITS 9.9-9.14) and prepare nuclear extract, S100, and nuclear pellet from the established cell line using the procedures described in Basic Protocol 1, steps 1 to 24. 2. Pack a P11 ion-exchange column containing ∼60 ml of resin in a chromatographic column. 3. Connect the column to a flow adapter, a chart recorder, and a fraction collector, and equilibrate the whole system with BC100 overnight. Set the flow rate at ∼1 column volume (CV) per hr (i.e., 1 ml/min).

cell line expressing FLAG-tagged human TBP (3-10) nuclear extract

P11 0.1

0.3

0.5

TBP complex 1

M2-agarose

TFIIIB

0.85 M KCl TBP complex 2

M2-agarose

TFIID

Figure 16.13.2 Purification of different FLAG-tagged TBP complexes. Nuclear extract prepared from 3-10 (Chiang et al., 1993), which is a HeLa-derived cell line constitutively expressing FLAG-tagged TBP, is loaded onto a P11 ion-exchange column. Proteins bound to the P11 resin are sequentially eluted off the column by step elutions with 0.1, 0.3, 0.5, and 0.85 M KCl-containing buffer. The FLAG-tagged TBP complexes are mainly detected at the 0.3 and 0.85 M KCl fractions, which are then used for immunoaffinity purification by anti-FLAG M2 monoclonal antibody-conjugated beads to isolate TFIIIB and TFIID complexes, respectively.

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4. Thaw ∼100 ml of nuclear extract in an ice bucket overnight at 4°C. 5. Centrifuge nuclear extract for 20 min at 105,000 × g (Beckman 45-Ti rotor 30,000 rpm), 4°C. 6. Pour supernatant into 50-ml tubes and load the sample onto an equilibrated P11 column at a flow rate of 1 CV/hr. Collect the eluate, ∼12 ml/tube, with a fraction collector. Remember to save a small aliquot (∼100 ìl) for immunoblotting analysis.

7. Wash the column with ∼2 CV of BC100 or until the absorption profile, monitored at 280 nm, almost reaches the baseline. 8. Step elute the bound proteins by sequentially switching to BC300, BC500, BC850, and BC1200, when the protein curve of each elution buffer nearly reaches the baseline. 9. Combine the fractions representing each of the BC100, BC300, BC500, and BC850 protein peaks. The BC1200 fractions, containing a minor amount of SL1, are not collected for further purification.

10. Dialyze BC500 and BC850 samples against 4 liters of BC100 for 5 hr at 4°C. 11. Centrifuge samples for 15 min at 46,000 × g (Beckman 45-Ti rotor 20,000 rpm), 4°C. 12. Aliquot the supernatant of BC500 and BC850, as well as the BC100 and BC300 fractions, individually, into several 15-ml tubes (for immunoaffinity purification) and 1.5-ml microcentrifuge tubes (for immunoblotting and protein analysis). 13. Freeze the aliquots in liquid nitrogen and store samples at −80°C until use. 14. Perform immunoblotting (UNIT 10.8) to locate the P11 fractions containing FLAGtagged TBP. 15. Purify TFIIIB from the P11 0.3 M KCl fraction and TFIID from the P11 0.85 M KCl fraction according to the procedures described in Basic Protocol 1, steps 26 to 34. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

BES buffer, 1 M (pH 7.2) 21.32 g of BES (N, N-bis[2-hydroxyethyl]-2-aminoethanesulfonic acid) (Sigma) 60 ml H2O Adjust pH to 7.12 with 1 M NaOH and bring up total volume to 100 ml Autoclave and store up to 6 months at room temperature


BC100 20 mM Tris⋅Cl, pH 7.9 at 4°C (APPENDIX 2) 20% glycerol 0.2 mM EDTA, pH 8.0 (APPENDIX 2) 100 mM KCl 1 mM DTT (added prior to use) 0.5 mM PMSF (added prior to use) Store up to 6 months at 4°C



BC300, BC500, BC850, and BC1200 Prepare as for BC100 (see recipe) except the KCl concentration is adjusted to 300, 500, 850, and 1200 mM, respectively. Buffer B 50 mM Tris⋅Cl, pH 7.9 at 4°C (APPENDIX 2) 25% glycerol 5 mM MgCl2 5 mM EDTA, pH 8.0 (APPENDIX 2) 5 mM EGTA, pH 8.0 5 mM DTT (added prior to use) 0.5 mM PMSF (added prior to use) Store up to 6 months at 4°C Buffer D 50 mM Tris⋅Cl, pH 7.9 at 4°C (APPENDIX 2) 25% glycerol 5 mM EDTA, pH 8.0 (APPENDIX 2) 5 mM EGTA, pH 8.0 2 mM DTT (added prior to use) 0.5 mM PMSF (added prior to use) Store up to 6 months at 4°C FLAG peptide elution buffer Diluting FLAG peptide from 100 mg/ml of FLAG peptide (in water) in BC100 or BC300 (see recipe) prior to use. COMMENTARY Background Information The FLAG epitope consists of 8 amino acids that are mostly charged residues (Hopp et al., 1988). The popularity of this epitope lies in the following facts. 1. It is synthetic, so that very few, if any, cross-reacting species are detected in bacterial, yeast, insect, or mammalian cells. 2. It is very hydrophilic and tends to help proteins stay soluble in aqueous solution. 3. It is small and highly antigenic. The introduction of the FLAG sequence rarely changes protein conformation and has a tendency to stimulate the immune response when injected into animals for antibody production (Chiang and Roeder, 1995). 4. A heart muscle kinase site linked to the FLAG sequence offers a unique way to label proteins with 32P after purification (Blanar and Rutter, 1992). The labeled proteins can be used for interaction studies, expression library screening and protein tracking (Chiang and Roeder, 1995). 5. A peptide elution method has been developed to elute the FLAG-tagged protein (Chiang and Roeder, 1993) or protein complexes (Chiang et al., 1993) off the anti-FLAG M2

monoclonal antibody-conjugated beads under neutral pH or physiological conditions, thereby allowing the recovery of fully active proteins ready for functional assays. 6. The fold purification achieved by M2agarose beads, which are commercially available, is exceedingly high, making it possible to purify FLAG-tagged multisubunit protein complexes to near homogeneity via one-step immunoaffinity purification (Chiang et al., 1993; Fondell et al., 1996; Kershnar et al., 1998; Ogryzko et al., 1998; Sif et al., 1998; Wu et al., 1998; Gu et al., 1999; Ikura et al., 2000). Although purification of recombinant FLAG-tagged protein can be easily achieved by using bacterial and baculovirus expression systems (Chiang and Roeder, 1993; Wu et al., 1999), the reconstitution of a fully functional protein complex from individually purified subunits or from insect cells coinfected with several recombinant baculoviruses expressing different subunits remains challenging (Bell et al., 1995; Chen and Tjian, 1996; Guermah et al., 1998; Phelan et al., 1999). Therefore, we and others have applied retrovirus-mediated gene transfer to deliver an epitope-tagged subunit of a large protein complex into human

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HeLa cells for the natural assembly of multisubunit protein complexes in a mammalian cell. Isolation of the in vivo-assembled large protein complexes is then facilitated by immunoaffinity purification using epitope-specific monoclonal antibody-conjugated beads and peptide elution (Zhou et al., 1992; Chiang et al., 1993). This methodology, combining epitope-tagging and stable cell line approaches, has made it possible to purify many multisubunit protein complexes, as long as a clonal cell line expressing the tagged subunit of a protein complex is available. Sometimes, cytotoxicity caused by constitutive expression of the tagged protein prevents the establishment of a stable cell line by retrovirus-mediated gene transfer. The application of an inducible mammalian expression system, in combination with FLAG epitopetagging and peptide elution methods, has further expanded our ability to establish stable cell lines expressing essentially any epitope-tagged protein (Wu and Chiang, 1996). The purification of human pol II and TFIIH from tetracycline-regulated clonal cell lines conditionally expressing FLAG-tagged RPB9 and the FLAG-tagged p62 subunit of human TFIIH, respectively, was the first documentation detailing the procedures for biochemical purification of multisubunit protein complexes from an inducible mammalian expression system (Kershnar et al., 1998). The protocols described here not only greatly simplify the purification steps, but also dramatically reduce the cost of preparation.



Location of the epitope tag. In general, the FLAG epitope sequence is introduced at either the N-terminus or the C-terminus of the protein-coding region. A critical parameter is to keep the tag away from essential regions of the protein. In many cases, however, it is hard to predict where the best location is in a protein and which subunit may be exposed on the surface of a protein complex. The suggestion is to tag different subunits in parallel to enhance the successful rate. Isolation of drug-resistant colonies. The cellular colonies are ready to be picked up when their diameter is between 2-3 mm. However, some slow-growing colonies should not be ignored, as expression of potentially toxic proteins may reduce the growth rate of the cells. The suggestion is to pick up colonies of various

sizes, especially if there are clear differences in the diameter of the colonies. Identification of positive clones. It is very important to use anti-FLAG M2, not M1, monoclonal antibody for immunoblotting, as anti-FLAG M2 monoclonal antibody can recognize the FLAG sequence at any location on the protein, whereas the M1 monoclonal antibody only recognizes the FLAG sequence at the very N-terminal region of the protein. In addition, a negative result with the M2 monoclonal antibody does not necessarily indicate a failure in establishing clonal cell lines. The FLAG epitope may sometimes be masked by the fusion partner, thereby preventing its detection by anti-epitope antibodies. In this case, it is essential to further confirm the results by anti-protein antibodies. Induction in suspension culture. It often happens that the tagged protein is not successfully induced after continued expansion in suspension culture. This is probably caused by residual tetracycline, which prevents tetracycline-dependent transactivator (tTA) from binding to the regulated gene, or by a reduced expression of tTA in the clonal cell line. To overcome these problems and to ensure a high level of protein induction, we usually follow these four guidelines. First, preselect cells with antibiotic (G418) in a small culture (∼100 ml) in the presence of tetracycline for at least three days before further expansion in the absence of antibiotic (G418). Presumably, this will boost the expression of tTA in the cell, thereby enhancing the level of transactivation from the tetracycline-regulated promoter. Second, wash cells at least 4 to 6 times with 1× PBS to remove tetracycline prior to induction. Third, use 250ml conical centrifuge tubes instead of 1-liter flat bottom bottles to spin down cells when washing with 1× PBS. Conical centrifuge tubes allow for better cell adhesion; thus medium and PBS washes can be completely removed after each successive centrifugation without losing cell volume. The 1-liter flat bottom centrifuge bottles, although convenient to use, do not permit complete removal of the original medium without disturbing the cell pellet; a minor residual tetracycline contaminant is sufficient to prohibit the induction. Fourth, use tetracyclinefree serum as a growth supplement when protein induction is desired. The authors have discovered that some batches of calf serum contain endogenous tetracycline, likely resulting from the use of tetracycline in cattle feed to protect animals from bacterial infection.



Removal of FLAG peptide. The presence of the free FLAG peptide in the purified protein complexes, following elution from the M2conjugated beads, is unlikely to cause problems in functional assays. In fact, the free peptide can enhance the stability of the purified protein complexes, which are often present in dilute concentrations after the final step of purification. However, if the presence of FLAG peptide presents a problem, e.g., by masking the surfaces of protein complexes or by competing for resin binding to an ion-exchange chromatographic column, a dialysis step can be included to remove the free peptide. Thus, depending on the purpose, FLAG peptide may or may not be removed after immunoaffinity purification. Specificity of purified protein complexes. Normally, a control purification with extract prepared from the parental cells is conducted in parallel to identify polypeptides binding non-specifically to the M2-conjugated beads. This is critical, especially for the characterization of the protein composition in a purified protein complex. It is also a good idea to perform a small-scale purification, e.g., by using 200 µl of protein sample and 10 µl of M2-conjugated beads in each tube (Chiang et al., 1993), in order to compare different washing conditions. Once the condition is optimized, a large-scale purification is then conducted. In this way, the specificity of the protein complexes is enhanced, while the nonspecific background is minimized. Consideration of elution buffer. The elution conditions also play an important role in immunoaffinity purification. Ideally, proteins are stored in low salt-containing buffers which allow for better manipulation of the salt conditions in later functional assays. But when comparing the elution conditions, we find that some proteins, such as the FLAG-tagged human papillomavirus E2 protein, elute better in 300 mM KCl–containing buffer (Hou et al., 2000), while the majority of other proteins elute well in 100 mM KCl–containing buffer. Inclusion of a minor amount (0.1% to 0.01%) of detergents, such as Nonidet P-40 or Triton X-100, in the elution buffer prevents protein/bead aggregation, allowing the immunoaffinity beads to rotate better during the elution step. A caveat in using detergent, however, is that the activity of some proteins (or complexes) may be sensitive to different concentrations of detergent. Luckily, protein complexes purified with these procedures, including human TFIID, TFIIH,

RNA polymerase II, and RNA polymerase II holoenzyme, are fully functional in both basal and activated transcription (Chiang et al., 1993; Chiang and Roeder, 1995; Kershnar et al., 1998; Wu and Chiang, 1998; Wu et al., 1998, 1999). Combination of immunoaffinity purification and conventional column chromatography. Additional steps of conventional column chromatography, such as ion-exchange and gel filtration, can be included prior to or following immunoaffinity purification to further enhance the purity of the target protein complex.

Anticipated Results The percentage of drug-resistant colonies exp ressing FLAG-tagg ed pr otein via retrovirus-mediated gene transfer is theoretically 100%. However, integration of the coding cassette into a transcriptionally silenced region of the cellular chromosomes may prevent the expression of FLAG-tagged protein, thereby reducing the number of positive clones. Even so, there are usually several clonal cell lines expressing FLAG-tagged protein from a dozen cellular isolates. In contrast, only 5 to 10% of clonal cell lines expressing FLAG-tagged protein were identified when the tetracycline-regulated expression system was used to establish stable cell lines. This is in part due to random integration of the tetracycline-regulated expression plasmid into cellular chromosomes. Thus, more drug-resistant colonies (>12) should be expanded when using the inducible expression system. The level of induced FLAG-tagged protein, which can vary over three orders of magnitude, is modulated by the amount of tetracycline included in the growth medium (Wu and Chiang, 1996). A maximal induction is usually achieved by complete removal of tetracycline. Normally the concentration of tetracycline is kept at either zero or 1 µg/ml to turn “on” or “off” the expression of the tagged protein throughout the entire experiment. The purified protein complexes are finally analyzed, in comparison with a control sample purified in parallel, by silver staining and immunoblotting. A purification table listing variables, including protein amount of the starting material, units, specific activity, fold purification, and yield, is typically generated. This is helpful for estimating the efficiency of immunoaffinity purification and other chromatographic steps (Kershnar et al., 1998). Protein Expression


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Time Considerations It usually takes three to four weeks to see drug-resistant colonies and an additional three weeks to identify clonal cell lines expressing FLAG-tagged protein by either retrovirus-mediated gene transfer or the tetracycline-regulated expression system. Expansion of a clonal cell line in suspension culture ready for extract preparation may take another 3 to 4 weeks. Once cells are ready for harvesting, it takes only one day to prepare nuclear extract, S100, and nuclear pellet, and another two days for immunoaffinity purification. If a chromatography step in involved, only one additional day is needed. Taken together, it may take 2 to 3 months to purify a multisubunit protein complex when beginning with an expression plasmid containing the FLAG-tagged protein-coding sequence.

Literature Cited Bell, S.P., Mitchell, J., Leber, J., Kobayashi, R., and Stillman, B. 1995. The multidomain structure of Orc1p reveals similarity to regulators of DNA replication and transcriptional silencing. Cell 83:563–568. Blanar, M.A. and Rutter, W.J. 1992. Interaction cloning: identification of a helix-loop-helix zipper protein that interacts with c-Fos. Science 256:1014-1018. Chen, J.L. and Tjian, R. 1996. Reconstitution of TATA-binding protein-associated factor/TATAbinding protein complexes for in vitro transcription. Methods Enzymol. 273:208-217. Chiang, C.-M. and Roeder, R.G. 1993. Expression and purification of general transcription factors by FLAG epitope-tagging and peptide elution. Peptide Res. 6:62-64. Chiang, C.-M. and Roeder, R.G. 1995. Cloning of an intrinsic human TFIID subunit that interacts with multiple transcriptional activators. Science 267:531-536. Chiang, C.-M., Ge, H., Wang, Z., Hoffmann, A., and Roeder, R.G. 1993. Unique TATA-binding protein-containing complexes and cofactors involved in transcription by RNA polymerases II and III. EMBO J. 12:2749-2762. Danos, O. and Mulligan, R.C. 1988. Safe and efficient generation of recombinant retroviruses with amphotropic and ecotropic host ranges. Proc. Natl. Acad. Sci. U.S.A. 85:6460-6464. Dignam J.D., Lebovitz, R.M., and Roeder, R.G. 1983. Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11:14751489. Purification of Epitope-Tagged Multisubunit Protein Complexes

Fondell, J.D., Ge, H., and Roeder, R.G. 1996. Ligand induction of a transcriptionally active thyroid hormone receptor coactivator complex. Proc. Natl. Acad. Sci. U.S.A. 93:8329-8333.

Gossen, M. and Bujard, H. 1992. Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc. Natl. Acad. Sci. U.S.A. 89:5547-5551. Gu, W., Malik, S., Ito, M., Yuan, C.X., Fondell, J.D., Zhang, X., Martinez, E., Qin, J., and Roeder, R.G. 1999. A novel human SRB/MED-containing cofactor complex, SMCC, involved in transcription regulation. Mol. Cell 3: 97-108. Guermah, M., Malik, S., and Roeder, R.G. 1998. Involvement of TFIID and USA components in transcriptional activation of the human immunodeficiency virus promoter by NF-κB and Sp1. Mol. Cell. Biol. 18:3234-3244. Hopp, T.P., Prickett, K.S., Price, V.L., Libby, R.T., March, C.J., Cerretti, D.P., Urdal, D.L., and Conlon, P.J. 1988. A short polypeptide marker sequence useful for recombinant protein identification and purification. Bio-Technology 6:12041210. Hou, S.Y., Wu, S.-Y., Zhou, T., Thomas, M.C., and Chiang, C.-M. 2000. Alleviation of human papillomavirus E2-mediated transcriptional repression via formation of a TATA binding protein (or TFIID)-TFIIB-RNA polymerase II-TFIIF preinitiation complex. Mol. Cell. Biol. 20:113-125. Ikura, T., Ogryzko, V.V., Grigoriev, M., Groisman, R., Wang, J., Horikoshi, M., Scully, R., Qin, J., and Nakatani, Y. 2000. Involvement of the TIP60 histone acetylase complex in DNA repair and apoptosis. Cell 102:463-473. Kershnar, E., Wu, S.-Y., and Chiang, C.-M. 1998. Immunoaffinity purification and functional characterization of human transcription factor IIH and RNA polymerase II from clonal cell lines that conditionally express epitope-tagged subunits of the multiprotein complexes. J. Biol. Chem. 273:34444-34453. Morgenstern, J.P. and Land, H. 1990. Advanced mammalian gene transfer: High titre retroviral vectors with multiple drug selection markers and a complementary helper-free packaging cell line. Nucleic Acids Res. 18:3587-3596. Ogryzko, V.V., Kotani, T., Zhang, X., Schlitz, R.L., Howard, T., Yang, X.J., Howard, B.H., Qin, J., and Nakatani, Y. 1998. Histone-like TAFs within the PCAF histone acetylase complex. Cell 94:35-44. Phelan, M.L., Sif, S., Narlikar, G.J., and Kingston, R.E. 1999. Reconstitution of a core chromatin remodeling complex from SWI/SNF subunits. Mol. Cell 3:247-253. Sif, S., Stukenberg, P.T., Kirschner, M.W., and Kingston, R.E. 1998. Mitotic inactivation of a human SWI/SNF chromatin remodeling complex. Genes Dev. 12:2842-2851. Wu, S.-Y. and Chiang, C.-M. 1996. Establishment of stable cell lines expressing potentially toxic proteins by tetracycline-regulated and epitopetagging methods. BioTechniques 21:718-725. Wu, S.-Y. and Chiang, C.-M. 1998. Properties of PC4 and an RNA polymerase II complex in



directing activated and basal transcription in vitro. J. Biol. Chem. 273:12492-12498. Wu, S.-Y., Kershnar, E., and Chiang, C.-M. 1998. TAFII-independent activation mediated by human TBP in the presence of the positive cofactor PC4. EMBO J. 17:4478-4490.

Chiang and Roeder, 1993. See above. This paper documents the development of the FLAG peptide elution method for purification of recombinant FLAG-tagged protein from bacteria. Kershnar, et al., 1998. See above.

Wu, S.-Y., Thomas, M.C., Hou, S.Y., Likhite, V., and Chiang, C.-M. 1999. Isolation of mouse TFIID and functional characterization of TBP and TFIID in mediating estrogen receptor and chromatin transcription. J. Biol. Chem. 274:2348023490.

This paper is the first documentation detailing the procedures for biochemical purification of multisubunit protein complexes from an inducible mammalian expression system.

Zhou, Q., Lieberman, P.M., Boyer, T.G., and Berk, A.J. 1992. Holo-TFIID supports transcriptional stimulation by diverse activators and from a TATA-less promoter. Genes Dev. 6:1964-1974.

This paper describes the construction of tetracycline-regulated FLAG-tagged expression plasmids, and the isolation and characterization of the inducible cell lines.

Wu and Chiang, 1996. See above.

Key References Chiang et al., 1993. See above. This paper describes the purification of human TFIID and TFIIIB complexes from a stable cell line constitutively expressing FLAG-tagged TBP.

Contributed by Shwu-Yuan Wu and Cheng-Ming Chiang Case Western Reserve University School of Medicine Cleveland, Ohio

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Inducible Gene Expression Using an Autoregulatory, Tetracycline-Controlled System

UNIT 16.14

Tetracycline-regulated gene expression systems have been developed to overcome some of the obstacles encountered using other strategies for inducible gene expression in mammalian cells. These difficulties include pleiotropic, nonspecific effects or toxicity of inducing agents or treatments, and high uninduced background levels of expression. This unit describes protocols for using a modified tetracycline-regulated system in which a transcriptional transactivator drives expression of itself and a target gene in cultured cells and, to some extent, in transgenic mice. This transactivator (tTA) is a fusion protein consisting of the tetracycline-repressor of E. coli and the transcriptional activation domain of the VP16 protein of herpes simplex virus. In the absence of tetracycline, tTA binds to and activates genes preceded by a heptamerized version of the tetracycline-resistance operator of Tn10 plus a minimal CMV promoter (here collectively referred to as Tet P). Binding of tTA to Tet P and subsequent gene activation are blocked in the presence of tetracycline. The plasmid pTet-Splice (Fig. 16.14.1A) contains Tet P upstream, and SV40 splice and polyadenylation signals downstream, of a multiple cloning site into which sequences encoding the open reading frame (ORF) of a target gene of choice is easily inserted. Autoregulatory tTA expression is driven from the plasmid pTet-tTAk (Fig. 16.14.1B), in which the tTA ORF (including an optimal sequence for initiation of translation according to Kozak) has been inserted into pTet-Splice. The protocols in this unit describe the transfection of adherent cells and the testing of resultant clones for inducible transactivator or target gene protein expression. Stably transfected fibroblast cell lines expressing transactivator and target gene(s) can be derived by first cotransfecting pTet-tTAk and a plasmid encoding a selectable marker and obtaining stable lines with inducible transactivator expression (see Basic Protocol). These lines are subsequently stably cotransfected with plasmids encoding the target gene(s) and a second selectable marker. The procedure may also be used to cotransfect pTet-tTAk with the target gene–encoding plasmid(s) and a single selectable marker plasmid. The choice of method depends upon the feasibility of screening for the protein products of the target genes. While the consecutive method is more systematic, cotransfection may be faster given a relatively straightforward screening method for expression of the target gene (see Critical Parameters). A Support Protocol also describes methods to test stably transfected cell lines for inducible gene expression, for transient transfection and induction of tet-regulated plasmids, and for detection of the tTAk gene in cells (or transgenic mice). CALCIUM PHOSPHATE-MEDIATED STABLE TRANSFECTION OF NIH3T3 CELLS WITH pTet-tTAk AND TETRACYLINE-REGULATED TARGET PLASMIDS This protocol describes the stable transfection of adherent cells with pTet-tTAk for the derivation of cell lines expressing inducible tTA. In the first round of transfection stable cell lines expressing inducible tTA alone are produced. The single transfection procedure may also be used for stable cotransfection of pTet-tTAk and plasmids expressing the target gene(s). In the second round of transfection tTA expressing lines are transfected with plasmids expressing the target gene(s).

BASIC PROTOCOL

Protein Expression Contributed by Penny Shockett and David Schatz Current Protocols in Molecular Biology (1997) 16.14.1-16.14.9 Copyright © 2002 by John Wiley & Sons, Inc.


A PvuI

ScaI

SspI SspI XmnI

BssH II PvuI PvuII

ampr

SstI Bst XI NotI Xbal PstI

poly(A) pTet–Splice (5178 bp)

BamHI PvuII

TetP

BssH II

+

splice

mcs XhoI PstI

EcoRI

Xbal SaII

BamHI

Clal EcoRV HindIII

Xbal SpeI BamHI

EcoRI

B ampr

poly(A) pTet–tTAk (6206 bp)

splice TetP +

XbaI

tTAk SalI

Figure 16.14.1 (A) The plasmid pTet-Splice (Shockett et al., 1995) is designed to drive tetracycline-regulated expression of a target gene inserted into the multiple cloning site (mcs). The tetracycline-regulated promoter (TetP) consists of a heptamerized tetracycline operator (doubleended arrow) upstream of a minimal human CMV promoter that includes bases −53 (triangle) to +75. The transcriptional start site (+) and TATAA box (small rectangle) are also indicated. This TetP fragment is an Xhol-SaII fragment derived from pUHC13-3 (Gossen and Bujard, 1992). SV40-derived sequences downstream of the MCS drive mRNA splicing and polyadenylation. The backbone of the plasmid is from Bluescript II KS+ (Stratagene) and carries the ampicillin resistance gene (ampr). (B) pTet-tTAk (Shockett et al., 1995) consists of the tTAk open reading frame inserted into the HindIII-EcoRV sites of pTet-Splice.

Inducible Gene Expression



Materials NIH3T3 cells Complete DMEM-10 medium (see recipe) Complete DMEM/tet: complete DMEM-10 medium (see recipe) containing 0.5 µg/ml tetracycline hydrochloride (Sigma; dilute 10 mg/ml stock in 70% ethanol and store protected from light at −20°C) Selection medium (see recipe) containing 125 µM, 250 µM, or 500 µM L-histidinol Plasmids for first-round or cotransfection procedure: pTet-tTAk (Life Technologies) and plasmids containing target gene ORF(s) cloned into pTet-Splice (Life Technologies), pSV2-His, or another selectable marker plasmid; purified by CsCl banding (UNIT 1.7) or anion-exchange chromatography (UNIT 2.1B) Plasmids for second round transfection procedure: plasmids containing target gene ORF(s) cloned into pTet-Splice, pPGKPuro, or another selectable marker plasmid; purified by CsCl banding (UNIT 1.7) or anion-exchange chromatography (UNIT 2.1B) 2 M CaCl2 HEPES-buffered saline (HeBS; see recipe) 10 mg/ml chloroquine (19 mM; optional; Sigma); dilute in water and store at −20°C 85% HeBS/15% glycerol, prewarmed to 37°C 3 mg/ml puromycin (Sigma) diluted in PBS (APPENDIX 2) Phosphate-buffered saline (PBS; APPENDIX 2) 1× trypsin/EDTA (Life Technologies) 10-cm and 6-cm tissue culture plates 4-ml polystyrene tubes (Falcon) 24-well and 6-well tissue culture plates NOTE: All tissue culture incubations are performed in a humidified 37°C, 5% CO2 incubator. Grow the cells 1. First round only: Grow cells in complete DMEM-10 medium. The day before transfection split cells into 10-cm tissue culture plates in complete DMEM/tet to achieve one-third confluence on the day of the transfection. From this point on cells are kept in the presence of 0.5 ìg/ml tet. One plate per transfection is needed at this stage. A typical experiment might include one plate for tTA only, one for tTA plus target gene, and one to serve as the untransfected control plate.

Second round only: Grow stable cell lines that inducibly express autoregulatory tTA in selection medium/500 µM L-histidinol. The day before transfection split into 10-cm plates in this same medium to achieve one-third confluence on the day of transfection. Transfect the cells 2. Linearize plasmids prior to transfection and adjust concentration to ≥0.5 mg/ml. See Damke et al. (1995) for discussion of other selectable markers. All plasmids should be purified by CsCl banding (UNIT 1.7) or on a Qiagen column (UNIT 2.1B).

3. First round only: Mix 10 to 20 µg of pTet-tTAk (in the presence or absence of an equimolar amount of target gene plasmids) plus 1 to 2 µg pSV2-His (a molar ratio

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of ∼10:1 of each tet plasmid to selectable marker plasmid) with 500 µl HeBS in a clear 4-ml polystyrene tube. A control mock transfection should be performed with no DNA added to the transfection. All of these cells should die in the selection medium/125 ìM L-histidinol introduced in step 14.

Second round only: Mix 10 to 20 µg each of target gene plasmid(s) plus 1 to 2 µg pPGKPuro (a molar ratio of ∼10:1 of each tet plasmid to selectable marker plasmid) with 500 µl HeBS in a clear 4-ml polystyrene tube. A control mock transfection should be performed with no DNA added to the transfection. All of these cells should die in the presence of the puromycin introduced in step 14. The optimal killing concentration for puromycin (lowest dose between 0.1 ìg/ml to 10 ìg/ml that kills all untransfected cells within a few days) should be determined empirically prior to the transfection and varies with the cell type.

4. Add 32.5 µl of 2 M CaCl2 to plasmid DNA and mix immediately by gentle vortexing. With occasional gentle mixing, allow precipitate to form for 15 to 30 min at room temperature or until solution is visibly cloudy when compared to a tube containing water. 5. Aspirate all of the medium from cells, doing one plate at a time. Mix precipitate a few times by pipetting with a Pasteur pipet, and apply dropwise and evenly over cells. 6. Incubate 30 min, gently rocking the plate after 15 min to ensure even coverage over entire plate. 7. First round only: Add 10 ml complete DMEM/tet, with or without 25 µM chloroquine (final), to each plate. Although the use of chloroquine may further reduce cell integrity during the glycerol shock (step 9), it can improve transfection efficiency.

Second round only: Add 10 ml selection medium/500 µM L-histidinol, with or without 25 µM chloroquine (final), to each plate. 8. Incubate 4 to 5 hr. The optimal length of incubation may vary for different cell types.

9. Gently aspirate medium from cells with minimal disruption of the precipitate that has settled onto the cells. Shock cells by adding dropwise 2.5 ml of prewarmed 85% HeBS/15% glycerol. It is normal for the cells to look somewhat ragged before and especially after glycerol shock. Two to four plates may be shocked at one time, depending on the speed of the researcher.

10. Aspirate HeBS/glycerol after exactly 2.5 min. Work quickly, as glycerol can be very toxic to the cells. The length of time cells are exposed to glycerol solution can be varied and increased up to 4 to 5 min to optimize transfection efficiency for different cell types. Cells should be shocked the maximal length of time which results in the least cell death.

11. First round only: Immediately, gently, and quickly wash cells twice by adding 10 ml complete DMEM/tet and immediately aspirating. Because cells tend to come loose from the plate after glycerol addition, add all medium to a single spot on the plate.


Second round only: Immediately, gently, and quickly wash cells twice by adding 10 ml selection medium/500 µM L-histidinol and immediately aspirating. Again, add medium to a single spot on the plate to avoid loosening the cells.



12. First round only: Add 10 ml complete DMEM/tet. Incubate cells overnight. Second round only: Add 10 ml selection medium/500 µM L-histidinol. Incubate cells overnight. 13. First round only: The morning after the transfection, aspirate the medium and replace with 10 ml complete DMEM/tet. Continue incubation. Second round only: The morning after the transfection, aspirate the medium and replace with 10 ml selection medium/500 µM L-histidinol. Continue incubation. Select and clone transfected cells 14. First round only: At 48 hr posttransfection, split cells into selection medium/125 µM 4 6 L-histidinol at several dilutions ranging from 3 × 10 to 1 × 10 cells per 10-cm plate. Make more than one plate in the mid-range that corresponds to an approximate split from one confluent plate of 1:16 to 1:32. Second round only: At 48 hr posttransfection, split cells as above, using selection medium/500 µM L-histidinol containing 3 µg/ml puromycin (final). The optimal killing concentration for puromycin (lowest dose between 0.1 ìg/ml to 10 ìg/ml that kills all untransfected cells within a few days) should be determined empirically prior to the transfection and varies with the cell type. The concentration of 3 ìg/ml puromycin is sufficient for selection of transfected NIH3T3 cells.

15. First round only: Refeed cells 4 days later with selection medium/125 µM L-histidinol. When colonies have formed, increase the concentration of L-histidinol in the selection medium to 250 µM. L-histidinol

is normally toxic to cells. The concentration of L-histidinol in the selection medium is therefore kept low initially and is raised as the number of cells expressing pSV2-His at high levels reaches a critical mass.

Second round only: Refeed cells 4 days later with selection medium/500 µM L-histidinol/puromycin. 16. When colonies are well established (at about day 12 to 14 of selection), circle their borders with a marker. Aspirate medium from plate and place a plastic cloning ring (autoclaved upright in vacuum grease) on the plate to surround an individual clone. Wash clones quickly with ∼100 µl PBS and add 2 drops of trypsin (∼100 µl) for 30 sec to 1 min. Pick cells from plates on which individual colonies are moderately spaced and can easily be distinguished.

17. First round only: Loosen cells by pipetting up and down with a Pasteur pipet and transfer colonies to wells of a 24-well plate into 1 ml selection medium/250 µM L-histidinol. Second round only: Loosen cells as for first round, transferring them into 1 ml selection medium/500 µM L-histidinol/puromycin. 18. First round only: When cells are heavy in wells, split into 6-cm dishes in selection medium/500 µM L-histidinol. Second-round only: When cells are heavy in wells, split into 6-cm dishes in selection medium/500 µM L-histidinol/puromycin. All trypsinization is performed by standard methods (APPENDIX 3F), involving a quick PBS wash, a 1 to 3 min trypsin/EDTA incubation (2 ml per confluent 10-cm plate), and using 3rd selection medium/500 ìM L-histidinol (± puromycin) and containing 10% calf serum to dilute and stop the trypsin.

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19. First round only: Expand cells for testing in selection medium/500 µM L-histidinol. Freeze aliquots of cells for storage in liquid nitrogen and grow in selection medium/500 µM L-histidinol from this point on. Test for tTA or target gene expression (if applicable; see Support Protocol for methods that may be used). Or, if applicable, repeat transfection procedure with target gene plasmid(s), following steps 1 to 18 and using the options listed for second-round transfection. Second round only: Test for target gene expression by northern or immunoblotting after induction (see Support Protocol). Freeze aliquots for storage in liquid nitrogen and grow in selection medium/500 µM L-histidinol/puromycin from this point on. SUPPORT PROTOCOL

ANALYSIS OF TARGET GENE PROTEIN EXPRESSION This protocol outlines methods for the analysis of target gene expression and inducibility. Instructions for inducing stable cell lines, for examining transient target gene expression with and without induction, and for PCR amplification of the tTA gene are included, with references to detection procedures such as Southern, northern, and immunoblotting techniques. Induction of Stable Cell Lines Stable cell lines can be tested for tTA or target gene expression by comparing induced to uninduced cells for tTA mRNA or target gene mRNA (see Detection of tTA Transgene in Cellular or Tail DNA by Southern Blotting), or protein expression or protein activity. Multiple lines may be screened at a time. The night before induction, the cells are plated in selection medium/500 µM L-histidinol (see recipe) containing 3 µg/ml puromycin at an appropriate density such that cells will be subconfluent to confluent at the time of harvest. Cells are washed three times with PBS (APPENDIX 2), with gentle swirling. Immediately, the medium is replaced with selection medium without 0.5 µg/ml tetracycline hydrochloride (tet). (For tet+ controls, simply aspirate medium and replace with fresh selection medium containing tet.) Cells are incubated 6 to 48 hr in a humidified 37°C, 5% CO2 incubator, then trypsinized (APPENDIX 6 3F) and harvested at 4°C, and an aliquot of 0.15–0.4 × 10 cells is analyzed by immunoblotting (see UNIT 10.8). Alternatively, cells may be grown in selection medium in the presence of tet, transferred to tubes [with a quick wash with cold PBS followed by trypsinization (APPENDIX 3F) and stopping of the trypsin by addition of selection medium containing tet], washed three times with PBS (or just pelleted, for tet+ controls), and replated into selection medium with and without tet at an appropriate density such that the cells will be subconfluent to confluent at the time of harvest. Induction of Gene Expression in Transiently Transfected Cells Transient transfection of tet-regulated plasmids is useful in several situations, including the initial testing of the autoregulatory system in a given cell line, screening stable tTA expressors for inducible expression, and biological applications where transient expression is specifically desired.


The night before the transfection, cells are split into medium containing 0.5 µg/ml tetracycline hydrochloride; the following day they are then transfected by methods appropriate for the cells being used (UNITS 9.1-9.4). Cells are induced by washing them three times in medium without tet. For CaPO4 transfection, washes are incorporated into those normally performed after glycerol shock (see Basic Protocol, step 11). Uninduced cell



controls are washed with medium containing tet. Medium with and without tet is added to the appropriate plates, then the cells are incubated for 12 to 48 hr in a humidified 37°C, 5% CO2 incubator. The cells are harvested at 4°C and, if trypsinized (APPENDIX 3F), cold medium containing 10% FBS (with and without tet, as appropriate) is used to stop the action of the trypsin. Cells are pelleted for freezing or lysis, and tTA or target gene (experimental or reporter) expression can be analyzed by northern blotting (UNIT 4.9), immunoblotting (UNIT 10.8), or by an appropriate activity assay (see Commentary). Detection of tTA Transgene in Cellular or Tail DNA by PCR PCR is routinely used to detect the Tet-tTAk transgene in candidate transgenic mouse tail DNA. The forward primer derives from the minimal human CMV promoter, CMV-F1: 5′-TGACCTCCATAGAAGACACC-3′ The reverse primer, TTA-REV1, is specific for the tTA ORF: 5′-ATCTCAATGGCTAAGGCGTC-3′ Hot-start PCR (UNIT 15.1) is performed on 150 ng of each tail DNA to be analyzed in a reaction mix containing 1.5 mM MgCl2, 0.5 µM each primer, and 0.2 mM each dNTP. PCR cycling conditions are as follows: 1 cycle: 30 cycles:

1 cycle

3 min 45 sec 45 sec 90 sec 10 min

94°C 80°C 94°C 58°C 72°C 72°C 8°C

(pause) add Taq polymerase (denaturation) (annealing) (extension) (extension) (end).

Products are analyzed on a 1% to 1.3% agarose gel; the main product of interest is visible as a 290-bp band after ethidium bromide staining. Detection of tTA Transgene in Cellular or Tail DNA by Southern Blotting The tTA transgene may also be detected by Southern blot analysis (UNIT 2.9). Tail DNA is digested with EcoRI and blots are probed with a 761-bp XbaI-SalI tTA insert from pTet-tTA. This fragment detects a 1094-bp tTA fragment of the transgene. This probe may also be used to detect tTA mRNA by northern blotting (UNIT 4.9). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Complete DMEM-10 Dulbecco’s minimal essential medium containing: 10% donor bovine calf serum (JRH Biosciences) 100 U/ml penicillin/100 µg/ml streptomycin (Life Technologies) 2 mM glutamine (Life Technologies) All DMEM complete medium used in this unit (with or without selection reagents or 0.5 ìg/ml tetracycline hydrochloride) may be stored protected from light ∼1 month at 4°C). Fetal bovine serum (FBS) may also be used in place of donor bovine calf serum, but the latter is less expensive. Protein Expression


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HEPES-buffered saline (HBS) 6 mM dextrose 137 mM NaCl 5 mM KCl 0.7 mM Na2HPO4⋅7H2O 21 mM HEPES (free acid) Adjust final pH to 7.05 with NaOH Filter sterilize and store in aliquots at −20°C Selection medium Complete histidine-free DMEM (Irvine Scientific, purchased without glutamine), containing: 10% donor bovine calf serum (JRH Biosciences) 100 U/ml penicillin/100 µg/ml streptomycin (Life Technologies) 2 mM glutamine (Life Technologies) 0.5 µg/ml tetracycline⋅HCl (Sigma; dilute 10 mg/ml stock in 70% ethanol and store protected from light at −20°C) 125 µM, 250 µM, or 500 µM L-histidinol (Sigma, dilute in water as a 125 mM stock and store at −20°C) COMMENTARY Background Information


Inducible, tetracycline-regulated gene expression systems were initially developed to allow the controlled expression in eukaryotic cells of foreign genes not tolerated constitutively in cultured cells or during the development of transgenic animals. The general features of tetracycline-regulated gene expression strategies and their improvements over previous inducible expression systems have been addressed in current review articles (Gossen et al., 1993; Barinaga, 1994; Damke et al., 1995; Shockett and Schatz, 1996). The autoregulatory tTA system used in this protocol derives directly from a constitutive tTA system described by Gossen and Bujard (1992). Although tight regulatory control and high inducibility was achieved with the original system in HeLa cells, the inability to detect clones expressing moderate to high levels of tTA by immunoblotting suggested that the tTA was toxic when expressed constitutively. The autoregulatory tTA system was designed to overcome possible toxic effects of constitutive tTA expression by making tTA expression itself tetracycline regulated. Autoregulated tTA expression theoretically allows for the selection of clones expressing higher levels of tTA via an autoregulatory feed-forward mechanism that is activated only in the absence of tetracycline. In the presence of tetracycline, low-level tTA and target gene expression are driven from the minimal human CMV promoter. However, any tTA produced is

unable to bind to tet operators upstream of the tTA or target gene. Conversely, when tetracycline is removed from the system, the small amounts of tTA protein expressed from the minimal promoter can bind the tet operators upstream of the tTA gene, driving higher levels of tTA (for controlled periods of time) and, subsequently, target gene expression. The theoretical benefits of the autoregulatory tTA system have been confirmed by experiments in stably transfected NIH3T3 cell lines (Shockett et al., 1995). In these experiments, expression of the recombination activating genes RAG-1 and RAG-2, and subsequent DNA recombination activated by these proteins, was higher and more frequently detected among stable transfectants expressing autoregulatory tTA than in constitutive tTA expressors. In transgenic mice expressing a luciferase reporter target transgene, the levels of expression appear to be 1 to 2 orders of magnitude greater with the autoregulatory system, although the uninduced levels also appear to be higher. Since the description of the early tTA systems, several laboratories have created modified vectors, including streamlined versions containing both tTA and the target gene, viral vectors, and vectors in which expression of two different target genes may be differentially or co-regulated. Some of these systems and their applications have recently been reviewed (Shockett and Schatz, 1996).



Critical Parameters and Troubleshooting Cell lines stably expressing both autoregulatory tTA and target genes have been derived at fairly high efficiencies by simultaneous transfection of all plasmids. This method may be faster, but it may require the screening of more clones than if stable lines with low basal and high induced levels of tTA are first derived and subsequently transfected with plasmids encoding the target genes. For the derivation of these clones, any selectable marker combination should theoretically work for consecutive cotransfection. Additionally, although the Basic Protocol describes calcium phosphate–mediated transfection of adherent fibroblast cell lines, the procedure can be adapted for other cell types using their optimal methods of transfection and selection. The protocol can also be scaled down to require fewer cells by using smaller dishes or wells and reducing all components proportionately. Using the autoregulatory tTA system, tTA mRNA induction appears to be a good indicator of induced tTA expression (see Support Protocol). Alternatively, the vector pUHC13-3 (Life Technologies) encoding luciferase under tet control may be transiently transfected into putative stable tTA expressors as previously described (see Support Protocol and Damke et al., 1995). Cells are then cultured for 12 to 48 hr in the presence and absence of tetracycline. Luciferase activity is easily measured in cell lysates using a kit (Luciferase Assay System and Dual-Luciferase Reporter Assay System; Promega) in which luciferase activity in cell lysates is normalized either to total protein determined using a Bradford protein assay (UNIT 10.1), or to a transfection control, respectively. Although basal expression of target plasmids tends to be higher when transiently transfected and luciferase detection is extremely sensitive, this method can be useful for the initial testing of the system in a given cell type (Damke et al., 1995). It is imperative after stable transfection with pTet-tTAk that cells be maintained in medium containing 0.5 µg/ml tetracycline to prevent any toxic effects of tTA expression and subsequent selection against clones expressing high levels of tTA.

Anticipated Results In the authors’ experience with stably transfected NIH3T3 cells, expression of induced

tTA and target gene has been observed by 6 hr and peaks at ∼12 hr after induction. In cells that stably express tTA, transient target gene expression has been observed by 12 hr. In cells transiently expressing tTA and a tet-sensitive luciferase reporter (pUHC13-3), luciferase activity induced by 2 orders of magnitude has been observed by 20 hr.

Time Considerations Starting with the plasmid vectors and following the transfection protocols above, stable clones expressing tTA (or tTA + target gene(s) if cotransfecting) are obtained in ∼12 to 14 days. Approximately 2 additional weeks are required for expansion and testing of candidate clones. Subsequent transfection of a stable inducible tTA clone with vectors expressing target genes will require the same amount of time. Transient transfection and inducible gene expression may be achieved within 48 hr.

Literature Cited Barinaga, M. 1994. Researchers devise a master gene control switch. Science 265:26-28. Damke, H., Gossen, M., Freundlieb, S., Bujard, H., and Schmid, S.L. 1995. Tightly regulated and inducible expression of dominant interfering dynamin mutant in stably transformed HeLa cells. Methods Enzymol. 257:209-220. Gossen, M. and Bujard, H. 1992. Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc. Natl. Acad. Sci. U.S.A. 89:5547-5551. Gossen, M., Bonin, A.L., and Bujard, H. 1993. Control of gene activity in higher eukaryotic cells by prokaryotic regulatory elements. Trends Biochem. Sci. 18:471-475. Shockett, P.E. and Schatz, D.G. 1996. Commentary: Diverse strategies for tetracycline-regulated inducible gene expression. Proc. Natl. Acad. Sci. U.S.A. 93:5173-5176. Shockett, P., Difilippantonio, M., Hellman, N., and Schatz, D. 1995. A modified tetracycline-regulated system provides autoregulatory, inducible gene expression in cultured cells and transgenic mice. Proc. Natl. Acad. Sci. U.S.A. 92:65226526.

Contributed by Penny Shockett Yale University School of Medicine New Haven, Connecticut David Schatz Howard Hughes Medical Institute and Yale University School of Medicine New Haven, Connecticut Protein Expression


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Overview of the Vaccinia Virus Expression System Vaccinia virus was introduced in 1982 as a vector for transient expression of genes in mammalian cells (Mackett et al., 1982; Panicali and Paoletti, 1982). This expression system differs from others in that transcription occurs in the cytoplasm of the cell rather than in the nucleus. As a vector, vaccinia virus has a number of useful characteristics, including a capacity that permits cloning large fragments of foreign DNA (>20 kbp) with retention of infectivity, a wide host range, a relatively high level of protein synthesis, and “appropriate” transport, secretion, processing, and posttranslational modifications as dictated by the primary structure of the expressed protein and the cell type used. For example, N- and O-glycosylation, phosphorylation, myristylation, and cleavage, as well as assembly of expressed proteins, occur in an apparently faithful manner. Laboratory applications of vaccinia virus vectors include production of biologically active proteins in tissue culture, analysis of mutant forms of proteins, and determination of transport and processing signals. In addition, recombinant vaccinia viruses have been important for immunological studies (Bennink and Yewdell, 1990). Infected cells can serve as targets to analyze the antigenic specificity of cytotoxic T cells. Recombinant viruses can be used to infect animals in order to determine cell-mediated and humoral responses to specific proteins, and are being evaluated as candidate vaccines for human and veterinary uses. Several variations of the vaccinia vector system have been developed and either standard or more attenuated, host-restricted strains may be used. After obtaining the virus stock (UNIT 16.16), the gene of interest is placed under control of a vaccinia virus promoter and integrated into the genome of vaccinia so as to retain infectivity (UNIT 16.17). Alternatively, expression can be achieved by transfecting a plasmid containing the vaccinia promoter– controlled gene into a cell that has been infected with vaccinia virus. These recombinant viruses are then characterized using various methods (UNIT 16.18). In still another variation, the bacteriophage T7 RNA polymerase gene can be integrated into the genome

UNIT 16.15

of vaccinia so that a gene controlled by a T7 promoter, either in a transfected plasmid or a recombinant vaccinia virus, will be expressed.

VACCINIA REPLICATION CYCLE Vaccinia is the prototypic member of the Orthopoxvirus genus of the Poxviridae family. Poxviruses differ from other eukaryotic DNA viruses in that they replicate in the cytoplasm rather than in the nucleus. Vaccinia virus has a linear, double-stranded DNA genome of nearly 200,000 bp that encodes most of the proteins needed for replication and transcription in the cytoplasm. The replication cycle of poxviruses is represented in Figure 16.15.1. The virus particle or virion consists of a complex core structure surrounded by a lipoprotein envelope. Remarkably, all proteins necessary for transcription of the early class of genes are packaged with the genome in the core. These include the following virus-encoded proteins: a multisubunit, DNA-dependent RNA polymerase, an early transcription factor, capping and methylating enzymes, and a poly(A) polymerase. The transcription system is activated upon infection, and early mRNAs and proteins can be detected within the first hour. The early mRNAs closely resemble their eukaryotic counterparts—they are capped, methylated, polyadenylated, and of discrete size. Termination of transcription o c c u r s ∼5 0 b a s e s a f te r t h e se q u e n c e TTTTTNT (where N can be any nucleotide; the termination signal is actually recognized in the nascent RNA as UUUUUNU). There is no evidence for splicing or other kinds of processing involving RNA cleavage. DNA replication begins within a few hours postinfection and leads successively to the intermediate and late phases of gene expression. The promoters associated with intermediate and late genes differ in sequence from early promoters and have different transcription factor requirements. The intermediate factors are synthesized early in order to facilitate transcription of newly replicated viral DNA. Some of the late factors are products of intermediate genes and hence are made and used after viral DNA replication. Protein Expression

Contributed by Bernard Moss and Patricia L. Earl Current Protocols in Molecular Biology (1998) 16.15.1-16.15.5 Copyright © 2002 by John Wiley & Sons, Inc.


uncoating

envelope RNA polymerase transcription factor poly(A) polymerase capping enzyme methylating enzymes

DNA replication (2-12 hr)

e tor as tiva c er a ns early mRNA lym tra po e t A dia DN erme t in early enzymes

growth factor

nucleus

mature virion

morphogenesis (4-20 hr)

resolution

core

concatem er

DNA

intermediate mRNA

late transactivators

late mRNA late enzymes early transcription factor structural proteins cleavage glycosylation acylation phosphorylation

cell

Figure 16.15.1 Replication cycle of vaccinia virus. After entry of vaccinia virus into the cells, early genes are expressed, leading to secretion of several proteins (including a growth factor), uncoating of the virus core, and the synthesis of DNA polymerase (and other replication proteins), RNA polymerase subunits, and transcriptional transactivators of the intermediate class of genes. After DNA replication, intermediate mRNAs are made, some of which encode late transcriptional transactivators. This in turn leads to expression of late genes, which encode structural proteins, enzymes, and early transcription factors that are packaged in the assembling virus particles. Some of the mature virions are wrapped in Golgi-derived membranes and are released from the cell. The bold arrows indicate products that exit the cell. Reprinted with permission from Raven Press (Moss, 1996a).

Overview of the Vaccinia Virus Expression System

The mRNAs transcribed from intermediate and late genes differ from typical early mRNAs as follows. First, the termination signal UUUUUNU is not recognized; consequently the mRNAs are long and heterogeneous in length, making procedures like northern blotting virtually useless. Second, the 5′ ends of intermediate and late mRNAs contain a capped poly(A) leader of ~35 nucleotides that is probably the result of an RNA polymerase slippage mechanism within the conserved AAA at the initiation site. Many early proteins are not synthesized beyond ∼6 hr postinfection (unless an inhibitor of DNA replication such as cytosine arabinoside is added) because of the cessation of early gene transcription and the relatively short half-life of all mRNAs at late times. Some genes, however, have tandem early and late promoters so that they are expressed throughout the growth cycle. Intermediate

genes are expressed for a relatively short period after DNA replication. Either because of intrinsic promoter strength, DNA copy number, or prolonged expression (>20 hr), much more protein is made from the strongest late promoters than from early or intermediate promoters.

EFFECTS OF VACCINIA INFECTION Standard vaccinia virus strains can productively infect most mammalian and avian cell lines, with a few exceptions such as Chinese hamster ovary (CHO) cells, but may not complete the replication cycle in primary lymphocytes or macrophages. The host-restricted modified vaccinia virus Ankara (MVA) strain has been shown to replicate efficiently only in chick embryo fibroblasts and BHK-21 cells, but viral and recombinant protein expression occurs in all cell lines infectable by standard strains. Vaccinia infection generally results in



rapid inhibition of host nucleic acid and protein synthesis. Inhibition of host protein synthesis is dramatic and probably results from several factors whose identities are uncertain; the relative contribution of each factor may depend on the virus multiplicity, cell type, and time of analysis. At the time of maximal late gene expression, host protein synthesis has been largely suppressed, facilitating the identification of viral or recombinant proteins by pulselabeling with radioactive amino acids (UNIT 10.18). In fibroblasts, the initial cytopathic effect is cell rounding, and is obvious by several hours postinfection. Nevertheless, the majority of cells remain intact for ≥48 hr. Some virus strains may cause less cytopathic effects than others, but mutants that efficiently express recombinant proteins but do not inhibit host gene expression have not been identified. Approximately 100 to 200 plaque-forming units (pfu), equivalent to ∼2500 to 5000 particles, are made per cell within a 20- to 40-hr period. With the commonly used vaccinia virus Western Reserve (WR) strain, >95% of the infectious virus remains cell-associated. With some other vaccinia virus strains, notably IHD-J, larger amounts of extracellular virus are produced.

VACCINIA VECTOR EXPRESSION SYSTEM Genes or cDNAs containing open reading frames derived from prokaryotic, eukaryotic, or viral sources have been expressed using vaccinia virus vectors. The gene of interest is usually placed next to a vaccinia promoter and this expression cassette is then inserted into the virus genome by homologous recombination or direct ligation (UNIT 16.17). Use of poxvirus promoters is essential because cellular and other viral promoters are not recognized by the vaccinia transcriptional apparatus. Strong late promoters are preferable when high levels of expression are desired. An early promoter, however, may be of use if it is desirable to express proteins prior to the occurrence of major cytopathic effects, or when the purpose is to make cells that express antigens in association with major histocompatibility class I molecules so they form cytotoxic T cell targets or prime animals for a cytotoxic T cell response. (The ability of vaccinia viral vectors to direct this type of antigen presentation seems to diminish late in infection.) The most versatile and widely used promoters contain early and late promoter elements. Transcripts originating early will terminate after the sequence

TTTTTNT; thus, any cryptic TTTTTNT termination motifs within the coding sequence of the gene should be altered by mutagenesis if an early poxvirus promoter is used (Earl et al., 1990). To mimic vaccinia virus mRNAs, untranslated leader and 3′-terminal sequences are usually kept short. A number of plasmids have been designed with restriction endonuclease sites for insertion of foreign genes downstream of vaccinia promoters (UNIT 16.17). The expression cassette is flanked by vaccinia DNA to permit homologous recombination when the plasmid is transfected into cells that have previously been infected with a vaccinia virus. The flanking vaccinia virus DNA is chosen so that recombination will not interrupt an essential viral gene. Without selection, the ratio of recombinant to parental vaccinia virus is usually ∼1:1000. Although this frequency is high enough to permit the use of plaque hybridization (UNITS 6.3 & 6.4) or immunoscreening (UNIT 6.7) to pick recombinant viruses, a variety of methods have been employed to facilitate identification of recombinant viruses. Some widely used selection or screening techniques are described in UNIT 16.17. Commonly, the expression cassette is flanked by segments of the vaccinia thymidine kinase (TK) gene so that recombination results in inactivation of TK. Virus with a TK− phenotype can then be distinguished from those with a TK+ phenotype by infecting a TK− cell line in the presence of 5-bromodeoxyuridine (BrdU), which must be phosphorylated by TK to be lethally incorporated into the virus genome. Alternatively, recombinant viruses can be selected by the co-expression of a bacterial antibiotic resistance gene such as guanine phosphoribosyltransferase (gpt). Co-expression of the Escherichia coli lacZ gene or other color markers allows rapid screening of recombinant virus plaques. Complementation of host-range and plaque-forming defects are also useful for isolation of recombinant viruses (UNIT 16.17).

STEPS FOR EXPRESSION OF GENES USING VACCINIA VECTORS The expression of genes using the vaccinia expression system is presented in detail in UNITS 16.16-16.18 and is outlined in the flowchart in Figure 16.15.2. A brief overview is presented below. 1. Prepare a stock of standard, host-rangerestricted, or bacteriophage T7 RNA polym-

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culture cell lines (UNIT 16.16, Basic Protocols 1 & 2)

infect cell line and prepare wild-type vaccinia stock (UNIT 16.16, Basic Protocol 3)

purify vaccinia virus (UNIT 16.17, Support Protocol 1)

titer vaccinia virus stock using plaque assay (UNIT 16.16, Support Protocol 1)

insert gene into transfer vector (UNIT 16.17, Basic Protocol 1)

isolate vaccinia virus DNA (UNIT 16.17, Support Protocol 2)

infect cells with vaccinia virus and transfect with vector (UNIT 16.17, Basic Protocol 1)

amplify a plaque (UNIT 16.17, B asic Protocol 3)

select and screen recombinant virus plaques (UNIT 16.17, Basic Protocol 2)

titer the amplified plaque (UNIT 16.16, Support Protocol 1)

prepare recombinant virus stock (UNIT 16.16, Basic Protocol 3)

express gene and analyze protein by • polyacrylamide gel electrophoresis • immunoblotting • immunoprecipitation

analyze recombinant virus by • polymerase chain reaction (UNIT 16.18, Basic Protocol 1) • Southern blot hybridization (UNIT 16.18, Basic Protocol 2) • DNA dot-blot hybridization (UNIT 16.18, Basic Protocol 3) • immunoblotting (UNIT 16.18, Alternate Protocol) • immunostaining (UNIT 16.16 )

Figure 16.15.2 Flowchart showing protocols for gene expression using the recombinant vaccinia virus system.

Overview of the Vaccinia Virus Expression System



erase–expressing vaccinia virus. At the same time, subclone the gene of interest into a plasmid transfer vector (UNITS 16.16 & 16.17). 2. Infect cells with vaccinia virus and transfect with the recombinant plasmid (UNIT 16.17). 3. Lyse the cells and plaque the virus under suitable selection or screening conditions (UNIT 16.17). 4. Pick plaques and confirm the presence and/or expression of the foreign gene (UNIT 16.18). 5. Amplify the plaque and prepare recombinant virus stock (UNIT 16.17). 6. Infect cells and analyze proteins synthesized (UNIT 16.18).

SAFETY PRECAUTIONS FOR USING VACCINIA Vaccinia virus is not to be confused either with variola virus, another member of the Orthopoxvirus genus that caused smallpox prior to its eradication, or with varicella virus, a herpes virus that causes chicken pox. Until 1972, vaccinia virus was routinely used in the United States as a live vaccine to prevent smallpox, and a residual scar, commonly on the upper arm, is evidence of that vaccination. To prevent laboratory infections, the Centers for Disease Control (CDC) and the National Institutes of Health (NIH) recommend that individuals who come into contact with vaccinia virus receive vaccinations at 10-year intervals (Richmond and McKinney, 1993). The CDC has supplied vaccine for such purposes when requested by qualified health workers. Eczema or an immunodeficiency disorder in the laboratory worker or a close contact, however, may be a contraindication to vaccination, which should only be given under medical supervision. The benefits of routine vaccination for healthy investigators have also been questioned (Baxby, 1989; Wenzel and Nettelman, 1989). Vaccinia virus is very stable and parenteral inoculation, ingestion, and droplet or aerosol exposure of mucous membranes are the primary hazards to laboratory or animal care personnel. Standard biosafety level 2 (BL-2) practices and class I or II biological safety cabinets should be employed (Richmond and McKinney, 1993). The NIH intramural program has lowered the safety requirements for highly attenuated vaccinia virus strains such as modified vaccinia virus Ankara (MVA; UNIT 16.6) and NYVAC (Tartaglia et al., 1992). However, local institutional biosafety offices should be contacted to determine current policy regarding vaccination and physical containment.

Additional precautions may be necessary for expression of certain genes such as toxins or large segments of other viral genomes, and guidelines for recombinant DNA work should be consulted. Approval of local biosafety committees may be necessary.

LITERATURE CITED Baxby, D. 1989. Smallpox vaccination for investigators. Lancet 2:919. Bennink, J.R. and Yewdell, J.W. 1990. Recombinant vaccinia viruses as vectors for studying T lymphocyte specificity and function. Curr. Topics Microbiol. Immunol. 163:153-184. Earl, P.L., Hügin, A.W., and Moss, B. 1990. Removal of cryptic poxvirus transcription termination signals from the human immunodeficiency virus type 1 envelope gene enhances expression and immunogenicity of a recombinant vaccinia virus. J. Virol. 64:2448-2451. Mackett, M., Smith, G.L., and Moss, B. 1982. Vaccinia virus: A selectable eukaryotic cloning and expression vector. Proc. Natl. Acad. Sci. U.S.A. 79:7415-7419. Moss, B. 1996a. Poxviridae: The viruses and their replication. In Virology (B.N. Fields, D.M. Knipe, and P. M. Howley, eds.) pp. 2637-2671. Raven Press, New York. Panicali, D. and Paoletti, E. 1982. Construction of poxviruses as cloning vectors: Insertion of the thymidine kinase gene from herpes simplex virus into the DNA of infectious vaccinia virus. Proc. Natl. Acad. Sci. U.S.A. 79:4927-4931. Richmond, J.Y. and McKinney, R.W. 1993. Biosafety in Microbiological and Biomedical Laboratories, 3rd ed. U.S. Government Printing Office, Washington, D.C. Tartaglia, J., Perkus, M.E., Taylor, J., Norton, E.K., Audonnet, J.C., Cox, W.I., Davis, S.W., Vanderhoeven, J., Meignier, B., Riviere, M., Languet, B., and Paoletti, E. 1992. NYVAC–A highly attenuated strain of vaccinia virus. Virology 188:217-232. Wenzel, R.P. and Nettelman, M.D. 1989. Smallpox vaccination for investigators using vaccinia recombinants. Lancet 2:630-631.

KEY REFERENCES Johnson, G.P., Goebel, S.J., and Paoletti, E. 1993. An update on the vaccinia virus genome. Virology 196: 381-401. Moss, B. 1996b. Genetically engineered poxviruses for recombinant gene expression, vaccination, and safety. Proc. Natl. Acad. Sci. U.S.A. 93:11341-11348.

Contributed by Bernard Moss and Patricia L. Earl National Institute of Allergy and Infectious Diseases Bethesda, Maryland

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Preparation of Cell Cultures and Vaccinia Virus Stocks

UNIT 16.16

This unit describes the maintenance of cell lines used with vaccinia virus, both in monolayer cultures (see Basic Protocol 1) and in suspension (see Basic Protocol 2). The suspended cell culture is then used in the preparation of vaccinia virus stocks (see Basic Protocol 3). The preparation of chick embryo fibroblasts (CEF) is also presented (see Basic Protocol 4), for use in the production of the highly attenuated and host range–restricted modified vaccinia virus Ankara (MVA) strain of vaccinia virus (see Basic Protocol 5). Additionally, support protocols are presented for the titration of standard and MVA vaccinia virus stocks (see Support Protocols 1 and 2, respectively). Because standard vaccinia virus strains have a broad host range, there is considerable latitude in the selection of cell lines; those described below (see Basic Protocols 1 and 2) have been found to give good results. BS-C-1 cells give the best results for a plaque assay, whereas HeLa cells are preferred for preparation of virus stocks. CV-1 cells can be used for both procedures, but they are generally used for transfection (UNIT 16.17). Human thymidine kinase–negative (TK−) 143B cells are used when TK selection is employed (UNIT 16.17), but they can be used for transfection as well as for a plaque assay. Either CEF or BHK-21 cells are used to propagate MVA and prepare recombinant MVA. Table 16.16.1 presents a summary of the uses for specific cell lines. NOTE: Carry out all procedures in this unit using sterile technique, preferably in a biosafety cabinet.

Table 16.16.1

Cell Lines Used in Specific Vaccinia Protocols

Cell line

Usea

Procedure

HeLa S3

Virus stock preparation Virus purification Plaque amplification

UNIT 16.16 Basic

Plaque assay Transfection (optional) XGPRT selection Plaque amplification (optional)

UNIT 16.16

Transfection Virus stock preparation (optional) Plaque assay (optional)

UNIT 16.17 Basic

TK selection Plaque assay (optional) Transfection (optional)

UNIT 16.17 Basic

MVA procedures

UNIT 16.16 Basic Protocol 5, Support Protocol 2; UNIT 16.17 Basic Protocol

4

UNIT 16.16 Basic Protocol 5, Support Protocol 2; UNIT 16.17 Basic Protocol

4

BS-C-1

CV-1

HuTK− 143B

CEF BHK-21

MVA procedures

Protocol 3 Support Protocol 1 UNIT 16.17 Basic Protocol 3 UNIT 16.17

Support Protocol 1 Protocol 1 UNIT 16.17 Basic Protocol 2 UNIT 16.17 Basic Protocol 3 UNIT 16.17 Basic

Protocol 1 Protocol 3 UNIT 16.16 Support Protocol 1 UNIT 16.16 Basic

Protocol 2 Support Protocol 1 UNIT 16.17 Basic Protocol 1 UNIT 16.16

aThe preferred use(s) for each cell line is listed first; if optional is indicated, the cell line can be used for the indicated

procedure, but the results may not be as good as those from the preferred cell line.

Protein Expression Contributed by Patricia L. Earl, Norman Cooper, Linda S. Wyatt, Bernard Moss, and Miles W. Carroll

16.16.1


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BASIC PROTOCOL 1

CULTURE OF MONOLAYER CELLS Frozen cells are thawed and grown in appropriate complete medium containing twice the maintenance amount of serum (see below). When the cells are confluent, they are treated with trypsin/EDTA, diluted, and maintained in appropriate complete medium containing 10% FBS (see Table 16.16.2). Table 16.16.2

Media Used for Growth and Maintenance of Cell Linesa

Cell line

Maintenance medium

Start-up mediuma

BHK-21 BS-C-1 CEF CV-1 HeLa S3 HuTK− 143B

Complete MEM-10 Complete MEM-10 Complete MEM-10 Complete DMEM-10 Complete spinner medium-5 Complete MEM-10/BrdU

Complete MEM-20 Complete MEM-20 Complete MEM-10 Complete DMEM-20 Complete MEM-10 Complete MEM-20/BrdU

aSee Reagents and Solutions for recipes.

Materials Frozen ampule of cells (Table 16.16.2): BS-C-1 (ATCC no. CCL26), CV-1 (ATCC no. CCL70), HuTK− 143B (ATCC no. CRL8303), or BHK-21 (ATCC no. CCL10) cells 70% ethanol Start-up medium (Table 16.16.2): complete MEM-20, complete DMEM-20, or complete MEM-20/BrdU (see recipes), 37°C Maintenance medium (Table 16.16.2): complete MEM-10, complete DMEM-10, or complete MEM-10/BrdU (see recipes), 37°C PBS (optional; APPENDIX 2) Trypsin/EDTA: 0.25% (w/v) trypsin/0.02% (w/v) EDTA, 37°C 25-cm2 and 150-cm2 tissue culture flasks Humidified, 37°C, 5% CO2 incubator Begin the culture 1. Thaw a frozen ampule of cells in a 37°C water bath. 2. Sterilize the ampule tip with 70% ethanol, break the neck, and transfer the cells with a pipet into a 25-cm2 tissue culture flask containing 5 ml start-up medium. Rotate the flask to evenly distribute the cells and place overnight in a humidified, 5% CO2 incubator at 37°C. 3. Aspirate the start-up medium and replace with appropriate maintenance medium. Return cells to the CO2 incubator at 37°C and check daily for confluency. Cells should be passaged when they become confluent. Generally, if cells are split 1:20, they reach confluence in 1 week and need not be counted.

Maintain the culture 4. When the cells are a confluent monolayer, aspirate medium.


5. Wash cells once with PBS or trypsin/EDTA to remove remaining serum from the cells by covering cells with the solution and pipetting it off.



6. Overlay cells with 37°C trypsin/EDTA using a volume that is just enough to cover the monolayer (e.g., 0.3 ml for a 25-cm2 flask). Allow to sit 30 to 40 sec (cells should become detached) and shake the flask to completely detach cells. 7. Add 1.4 ml appropriate maintenance medium. Pipet the cell suspension up and down several times to disrupt clumps (these cells are ready for passage). 8. Remove 0.5 ml cell suspension and add it to a new 150-cm2 tissue culture flask containing 30 ml maintenance medium. Rotate the flask to evenly distribute the cells and place in a CO2 incubator at 37°C until the cells are confluent (∼1 week). Maintain the cells by splitting ∼1:20 in maintenance medium at approximately weekly intervals. Cells can be maintained in smaller flasks if desired. If so, volumes should be adjusted proportionately.

CULTURE OF CELLS IN SUSPENSION HeLa S3 cells are maintained in complete spinner medium-5.

BASIC PROTOCOL 2

Materials Frozen ampule of HeLa S3 cells (ATCC no. CCL2.2) 70% ethanol Complete MEM-10 (see recipe), 37°C Trypsin/EDTA: 0.25% (w/v) trypsin/0.02% (w/v) EDTA, 37°C Complete spinner medium-5 (see recipe), 37°C 25-cm2 tissue culture flask Humidified, 37°C, 5% CO2 incubator Sorvall H-6000A rotor (or equivalent) 100- or 200-ml vented spinner bottles and caps with filters (Bellco) Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3F) Begin the culture 1. Thaw a frozen ampule of HeLa S3 cells in a 37°C water bath. 2. Sterilize the ampule tip with 70% ethanol, break the neck, and transfer the cells with a pipet to a 25-cm2 tissue culture flask containing 5 ml complete MEM-10. Rotate the flask to evenly distribute the cells and place overnight in a humidified, 5% CO2 incubator at 37°C. 3. Aspirate medium. Overlay cells with 0.5 ml of 37°C trypsin/EDTA and let sit 30 to 40 sec. Since these cells do not attach firmly to the flask, they should not be washed prior to trypsinization.

4. Add 10 ml complete spinner medium-5 and transfer cells to a 50-ml centrifuge tube. Centrifuge 5 min in a Sorvall H-6000A rotor at 2500 rpm (1800 × g), room temperature, and discard supernatant. 5. Suspend cell pellet in 5 ml complete spinner medium-5 by pipetting up and down to disrupt clumps. 6. Add 50 ml complete spinner medium-5 to a 100- or 200-ml vented spinner bottle and transfer the cell suspension to this bottle.

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7. Remove 1 ml cell suspension and count the cells using a hemacytometer (APPENDIX 3F). Add complete spinner medium-5 to adjust the cell density to 3–4 × 105 cells/ml. Place cells in a 37°C incubator without CO2 and stir continuously. The initial high density is used because some cells are not viable.

8. Grow cells for two successive days, counting cells daily and adding complete spinner medium-5 as necessary to maintain a concentration of 3–4 × 105 cells/ml. Maintain the culture 9. Remove 1 ml cell suspension and monitor the cells using a hemacytometer. 10. When the density is 4–5 × 105 cells/ml, dilute the cells to 1.5 or 2.5 × 105 cells/ml with complete spinner medium-5 for alternate day or daily feeding, respectively. 11. Place a 100- or 200-ml vented spinner bottle containing 50 ml or 100 ml cells, respectively, in 37°C incubator without CO2 and stir continuously. Passage every 1 to 2 days. HeLa S3 cells are grown and maintained in complete spinner medium-5 in vented spinner bottles at 37°C without CO2. Cells are diluted with fresh medium at 1- to 2-day intervals to keep the cell density between 1.5 × 105 and 5 × 105 cells/ml. BASIC PROTOCOL 3

PREPARATION OF A VACCINIA VIRUS STOCK To prepare a vaccinia virus stock, HeLa S3 cells from a spinner culture (see Basic Protocol 2) are plated the day before infection and allowed to attach. They are then infected with trypsinized virus. After several days, the infected cells are harvested and lysed during repeated freeze-thaw cycles. The virus stock is then aliquoted and stored at −70°C. To titer this stock, see Support Protocol 1. The protocol can be modified for monolayer cultures. Materials HeLa S3 cells from suspension culture (see Basic Protocol 2) Complete MEM-10 and -2.5 (see recipe), 37°C Vaccinia virus (ATCC no.VR1354 or equivalent) 0.25 mg/ml trypsin (2× crystallized and salt-free; Worthington; filter sterilize and store at −20°C) Sorvall H-6000A rotor (or equivalent) 150-cm2 tissue culture flask Humidified, 37°C, 5% CO2 incubator Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3F) Prepare cells 1. Count HeLa S3 cells from a suspension culture using a hemacytometer (APPENDIX 3F). 2. Centrifuge 5 × 107 cells 5 min in a Sorvall H-6000A rotor at 2500 rpm (1800 × g), room temperature, and discard supernatant. 3. Resuspend cells in 25 ml of 37°C complete MEM-10, dispense in a 150-cm2 tissue culture flask, and place overnight in a humidified, 5% CO2 incubator at 37°C.


Increase the number of HeLa cells proportionately if more than one 150-cm2 flask is to be infected. As an alternative to HeLa suspension cells, prepare 150-cm2 flasks of monolayer cells as described (see Basic Protocol 1) and continue with step 4 below.



Trypsinize virus 4. Just prior to use, mix an equal volume of vaccinia virus stock and 0.25 mg/ml trypsin, and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals throughout the incubation. Virus stocks are usually at a titer of ∼2 × 109 pfu/ml, but may be significantly lower depending on the source. Vortexing usually breaks up any clumps of cells. However, if there are still visible clumps, chill to 0°C and sonicate 30 sec on ice (UNIT 16.17). Sonication can be repeated several times but the sample should be allowed to cool on ice between sonications.

Infect cells 5. Dilute trypsinized virus in complete MEM-2.5 at 2.5–7.5 × 107 pfu/ml. Decant or aspirate medium from the 150-cm2 flask of cells and add 2 ml diluted, trypsinized virus. Place 2 hr in a CO2 incubator at 37°C, rocking flask by hand at 30-min intervals. The optimal multiplicity of infection (MOI) is 1 to 3 pfu/cell. Multiplicities of 0.1 pfu/cell may be necessary if the titer of the initial virus stock is low. The trypsinized virus must be diluted ≥10-fold to avoid detaching the cells.

6. Overlay cells with 25 ml complete MEM-2.5 and place 3 days in a CO2 incubator at 37°C. 7. Detach the infected cells from the flask by shaking and pour or pipet into a sterile plastic screw-cap tube. Centrifuge 5 min at 1800 × g, 5° to 10°C, and discard supernatant. 8. Resuspend cells in 2 ml complete MEM-2.5 (per initial 150-cm2 flask) by gently pipetting or vortexing. Harvest virus stock 9. Lyse the cell suspension by freeze-thaw cycling as follows: freeze in dry ice/ethanol, thaw in a 37°C water bath, and vortex. Carry out the freeze-thaw cycling a total of three times. 10. Keep the virus stock on ice and divide it into 0.5- to 2-ml aliquots. Store the aliquots indefinitely at −70°C. TITRATION OF VACCINIA VIRUS STOCKS BY PLAQUE ASSAY Serial dilutions of the trypsinized virus stock (see Basic Protocol 3) are used to infect the appropriate cell line. After several days growth, the medium is removed and the cells are stained with crystal violet. Plaques appear as 1- to 2-mm-diameter areas of diminished staining due to the retraction, rounding, and detachment of infected cells.

SUPPORT PROTOCOL 1

Additional Materials (also see Basic Protocols 1 and 3) BS-C-1 cells from confluent monolayer culture (see Basic Protocol 1) Virus stock (see Basic Protocol 3) 0.1% (w/v) crystal violet (Sigma) in 20% ethanol (store indefinitely at room temperature) 6-well, 35-mm2 tissue culture dishes Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3F) Protein Expression


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Prepare cell and virus stocks 1. Trypsinize confluent monolayer of BS-C-1 cells as described (see Basic Protocol 1, steps 4 to 7). 2. Count the cells using a hemacytometer (APPENDIX 3F). 3. Seed wells of a 6-well, 35-mm2 tissue culture dish with 5 × 105 cells per well BS-C-1 cells in 2 ml complete MEM-10. Place overnight in a humidified, 5% CO2 incubator at 37°C to reach confluency. 4. Trypsinize virus stock (see Basic Protocol 3, step 4). 5. Make nine 10-fold serial dilutions (UNIT 1.11) of the trypsinized virus in complete MEM-2.5, using a fresh pipet for each dilution. Perform assay 6. Remove medium from BS-C-1 cells and infect cells in duplicate wells with 0.5 ml of the 10−7, 10−8, and 10−9 trypsinized virus dilutions. Place 1 to 2 hr in a CO2 incubator at 37°C, rocking dish at 15- to 30-min intervals to spread virus uniformly and keep cells moist. 7. Overlay cells in each well with 2 ml complete MEM-2.5 and place 2 days in a CO2 incubator at 37°C. 8. Remove medium and add 0.5 ml of 0.1% crystal violet to each well. Incubate 5 min at room temperature. 9. Aspirate crystal violet and allow wells to dry. 10. Determine the titer by counting plaques within the wells and multiplying by the dilution factor. Most accurate results are obtained from wells with 20 to 80 plaques. In determining the titer, take into account the 1:1 dilution of the virus stock with trypsin. BASIC PROTOCOL 4

PREPARATION OF CHICKEN EMBRYO FIBROBLASTS The attenuated, replication-deficient modified vaccinia virus Ankara (MVA) can be used as an alternative to standard strains of vaccinia virus, for use as a vector to express foreign genes. MVA was derived from the vaccinia virus strain Ankara by multiple (>570) passages in chick embryo fibroblasts (CEF). Although MVA expresses recombinant proteins efficiently, the assembly of infectious particles is interrupted in human and most other mammalian cells lines, providing an added degree of safety to laboratory personnel. The NIH Intramural Biosafety Committee has determined that individuals do not need to be vaccinated to work with MVA, and can do so under biosafety level 1 (BL-1) conditions if no other vaccinia virus strains are being manipulated at the same location by other investigators. Recombinant MVA expressing bacteriophage T7 RNA polymerase, which can be used for high level transient expression, has also been constructed (UNIT 16.19) CEF or Syrian hamster kidney cells (BHK-21, see Basic Protocol 1) are permissive for growth of MVA. MVA is titered by immunostaining (see Support Protocol 2), because it does not form distinct plaques for accurate quantitation.


In this protocol, nine-day-old chicken embryos are trypsinized and plated in appropriate medium. After the cells form a confluent monolayer, they are transferred directly to a 31°C incubator where they can be held for 2 to 3 weeks before making secondary CEF for virus infection. With each passage, CEF require longer to become confluent; therefore use of the second passage is recommended.



Materials Nine-day-old embryonated eggs (Specific Pathogen Free Eggs, SPAFAS) 70% ethanol MEM with no additives, 37°C Trypsin/EDTA: 0.25% (w/v) trypsin/0.02% (w/v) EDTA, 37°C Complete MEM-10 (Table 16.16.2; see recipe), 37°C Sterile dissecting scissors and forceps 100-cm2 sterile petri dishes 10-ml syringes Sterile trypsinization flask with magnetic stir bar Humidified, 37° and 31°C, 5% CO2 incubators 500-ml beakers with two layers of gauze taped over tops Sorvall RC-3B centrifuge and 250-ml centrifuge bottles (or equivalent) 150-cm2 tissue culture flasks Prepare tissue 1. Position ten 9-day-old embryonated eggs with air space (blunt end) up and spray with 70% ethanol. 2. Crack the top of an egg with sterile dissecting scissors, and cut off shell to just above the membrane while keeping the latter intact. Remove membrane with sterile forceps. Repeat with remaining eggs. NOTE: Healthy eggs have well-formed blood vessels.

3. Remove the embryo from each egg and combine in a 100-cm2 sterile petri dish. 4. Remove head and feet from each embryo and place the rest of the body in a 100-cm2 sterile petri dish containing 10 ml MEM with no additives. Mince embryos by squeezing through 10-cc syringes (∼5 embryos/syringe) into a sterile trypsinization flask. Dissociate cells 5. Add 100 ml of 37°C trypsin/EDTA and incubate 5 min in a humidified, 5% CO2 incubator at 37°C, with stirring. 6. Decant fluid from the trypsinization flask into a 500-ml beaker covered with gauze. Transfer filtrate to a 250-ml centrifuge bottle. 7. Add 100 ml fresh trypsin/EDTA to the remaining tissue in the trypsinization flask, and incubate 5 min at 37°C. 8. Pour digest into a second 500-ml beaker covered with gauze, and add filtrate to the first filtrate in the 250-ml centrifuge bottle. 9. Centrifuge 10 min at 1200 × g, 4°C, in a Sorvall RC-3B centrifuge. 10. Aspirate and discard supernatant from pellet, add 10 ml complete MEM-10, resuspend by pipetting ∼10 to 15 times, and adjust volume to 100 ml. 11. Transfer to a 250-ml centrifuge bottle and centrifuge 10 min at 1200 × g, 4°C. 12. Resuspend pellet in 5 ml complete MEM-10 and adjust volume to 30 ml. Prepare confluent culture 13. Add 1 ml cell suspension to each of thirty 150-cm2 tissue culture flasks containing 30 ml complete MEM-10.

Protein Expression


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14. Incubate at 37°C for several days until confluent, and move flasks to a humidified, 31°C, 5% CO2 incubator for storage. The CEF cell cultures can be held for 2 to 3 weeks at 31°C without further attention. Primary CEF can also be used directly for virus growth, or the trypsinized stock of CEF can be frozen in liquid nitrogen for future use. BASIC PROTOCOL 5

PREPARATION OF AN MVA STOCK To prepare an MVA stock, CEF (see Basic Protocol 4) or BHK-21 (see Basic Protocol 1) cells are plated several days before infection and allowed to approach confluency. They are then infected with MVA. After several days, the infected cells are harvested, lysed by repeated freezing and thawing, sonicated, dispensed in small aliquots, and stored at −70°C. NOTE: BHK-21 cells should be acquired from ATCC, as cells from alternative sources may support lower levels of MVA replication. Materials 150-cm2 tissue culture flasks of nearly confluent CEF (see Basic Protocol 4) or BHK-21 cells (see Basic Protocol 1) Complete MEM-10 and -2.5 (see recipe), 37°C Modified vaccinia virus Ankara (MVA; A. Mayr, Institut für Med. Mikrobiologie, Munich, Germany, or B. Moss, email [email protected]) 150-cm2 tissue culture flasks Humidified, 37°C, 5% CO2 incubator Sorvall RC-3B centrifuge and sterile 250-ml centrifuge bottles (or equivalent) Additional reagents and equipment for trypsinizing cells (see Basic Protocol 1) 1. Trypsinize seven 150-cm2 tissue culture flasks of nearly confluent CEF or two of BHK-21 cells as described (see Basic Protocol 1, steps 4 to 7), using 1.5 ml trypsin/EDTA and 8.5 ml maintenance medium for the larger flask. Distribute to twenty 150-cm2 flasks. Incubate at 37°C in a humidified, 5% CO2 incubator until nearly confluent monolayers have formed (usually 2 days). CEF and BHK-21 cells are split 1:3 and 1:10, respectively. It is advisable to use monolayers of BHK-21 cells at 90% confluency, as completely confluent cultures degrade within 48 hr.

2. Remove medium and add 30 ml complete MEM-2.5 to each 150-cm2 flask. 3. Thaw MVA virus and sonicate 30 sec on ice to break up clumps. Trypsinizaton of virus is avoided with MVA because it may lower the titer.

4. Add 1 to 3 infectious units of virus/cell to the medium and place cells in a CO2 incubator for 3 days at 37°C. A monolayer of CEF or BHK-21 cells in a 150 cm2 flask contains ∼1–2 × 107 cells. If there is insufficient virus, then use fewer or smaller flasks of confluent cells.

5. Detach the cells with a cell scraper or, if possible, by shaking. Transfer cells and medium to a sterile 250-ml centrifuge bottle and centrifuge 10 min at 1200 × g, 4°C, in a Sorvall RC-3B centrifuge. Discard supernatant. 6. Resuspend cells in 1 ml complete MEM-2.5 (per 150-cm2 flask) by pipetting or vortexing. Preparation of Cell Cultures and Vaccinia Virus Stocks

7. Lyse cells by freeze-thaw cycling: freeze in dry ice/ethanol, thaw in a 37°C water bath, and vortex. Carry out the freeze-thaw cycling a total of three times.



8. Sonicate stock on ice for 1 min, pause for 1 min, and repeat sonication. Dispense in 0.5-ml aliquots and store indefinitely at −70°C. TITRATION OF MVA STOCKS BY IMMUNOSTAINING The titer of MVA is routinely determined by immunostaining, because MVA does not form distinct plaques in CEF or BHK-21 cells. To determine the MVA titer, serial dilutions of virus stock are prepared and used to infect CEF or BHK-21 cells. At 24 hr after infection, the cells are fixed with acetone/methanol and immunostained using a polyclonal vaccinia virus antiserum and a secondary peroxidase-conjugated antibody. A focus of infected cells appears as a 0.3- to 0.4-mm reddish brown–stained area. If the cells are left for two days before immunostaining, secondary foci may form by virus spread giving inaccurate titers.

SUPPORT PROTOCOL 2

Materials CEF (see Basic Protocol 4) or BHK-21 cells (see Basic Protocol 1) in a 150-cm2 tissue culture flask Complete MEM-10 and -2.5 (see recipe), 37°C MVA stock (see Basic Protocol 5) 1:1 (v/v) acetone/methanol PBS (APPENDIX 2), with and without 3% FBS Rabbit anti-vaccinia antibody (e.g., Access Biomedical & Diagnostic Research Labs or see Linscott, 1998) Horseradish peroxidase–conjugated whole anti–rabbit Ig antibody (HRP-anti-rabbit; Amersham) Dianisidine, or premade peroxidase substrate kit (Sigma) PBS/H2O2 (add 10 µl 30% H2O2 to 10 ml PBS immediately before use) 6-well, 35-mm2 tissue culture dishes Humidified, 37°C, 5% CO2 incubator Additional reagents and equipment for trypsinizing cells (see Basic Protocol 1) Infect and fix cells 1. Trypsinize a 150-cm2 tissue culture flask of CEF or BHK-21 cells as described (see Basic Protocol 1, steps 4 to 6), using 1.5 ml trypsin/EDTA for the larger flask. Resuspend cells by repeated pipetting in 5 ml complete MEM-10, and add an additional 60 ml complete MEM-10. 2. Add 2 ml cell suspension to each well of a 6-well, 35-mm2 tissue culture dish. Incubate overnight in a humidified, 5% CO2 incubator at 37°C to reach near confluency. 3. Remove medium and replace with 2 ml complete MEM-2.5. 4. Thaw MVA stock and sonicate 30 sec on ice. 5. Make eight 10-fold serial dilutions of the virus in complete MEM-2.5, using a fresh pipet for each dilution. 6. Add 0.1 ml of each of the 10−6, 10−7, and 10−8 dilutions to cells in duplicate wells. Swirl gently to mix, and incubate 24 hr in a CO2 incubator at 37°C. 7. Remove fluid and fix cells 2 min with 1 ml of 1:1 acetone/methanol. Remove fixative and add 2 ml PBS to each well. At this point, the plates can be immunostained immediately or stored at 4°C for several weeks.

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Perform immunostaining 8. Dilute rabbit anti-vaccinia antibody in PBS containing 3% FBS and add 1 ml/well. Incubate 1 hr at room temperature, with gentle rocking, if desired. The optimal dilution for each antibody must be determined empirically, but a 1:500 or 1:1000 dilution is a good place to start.

9. Wash twice with 2 ml PBS. 10. Dilute HRP-anti-rabbit secondary antibody in PBS containing 3% FBS and add 1 ml/well. Incubate for 30 to 45 min at room temperature, with gentle rocking, if desired. The optimal dilution for each antibody must be determined, but a 1:500 or 1:1000 dilution is a good place to start. If another anti-vaccinia antibody is used in step 8, the appropriate HRP-conjugated anti-species secondary antibody should be substituted here.

11. Wash twice with 2 ml PBS. 12. Make a saturated solution of dianisidine in 0.5 ml ethanol, vortex, incubate 5 min at 37°C, and clarify by microcentrifugation at maximum speed for 1 min. Add 0.2 ml dianisidine solution to 10 ml PBS/H2O2. Alternatively, use premade substrate tablets according to manufacturer’s instructions. CAUTION: Dianisidine is carcinogenic and the powder should be manipulated with gloves in a fume hood. 30% H2O2 is caustic to skin and should be handled with gloves.

13. Add 0. 5 ml dianisidine substrate solution to each well. Rotate dish gently and let stand ∼10 min. Check microscopically for foci of infected cells. When development is complete, wash dishes with water and overlay with 1 ml water to preserve stain. Weaker antibody may take longer to develop.

14. Count the number of stained foci and multiply by the dilution factor to express titer as infectious units/ml. Most accurate results are obtained from wells with 20 to 100 plaques. In determining the titer as infectious units/ml, be sure to mulitply by 10 to take into account the 0.1 ml addition of virus to the wells.


Complete DMEM-10 or -20 Dulbecco’s minimum essential medium (DMEM) containing: 10% or 20% fetal bovine serum (FBS) 0.03% glutamine 100 U/ml penicillin 100 µg/ml streptomycin sulfate Store up to several months at 4°C Add supplements from stock solutions prepared in water at the following initial concentrations: 3% glutamine (100×), 20,000 U/ml penicillin (200×), and 20 mg/ml streptomycin (200×). Filter sterilize stock solutions. Store 100× glutamine 4 months at 4°C, and 200× penicillin/streptomycin 4 months at −20°C. Preparation of Cell Cultures and Vaccinia Virus Stocks

Fetal bovine serum is added at 10% to complete medium for maintenance of cells and at 20% to encourage initial growth upon thawing. See Chapter 9 introduction for a full discussion concerning media preparation and use of serum (e.g., heat-inactivation, screening).



Complete MEM-2.5, -10, or -20 Minimum essential medium (MEM) containing: 2.5%, 10%, or 20% FBS 0.03% glutamine 100 U/ml penicillin 100 µg/ml streptomycin sulfate Store up to several months at 4°C Add supplements from stock solutions as described for complete DMEM (see recipe). Also see annotation to DMEM recipe concerning growth versus maintenance levels of serum.

Complete MEM-10/BrdU or -20/BrdU Prepare as for complete MEM-10 or -20 (see recipe) and add a 5 mg/ml solution of 5-bromodeoxyuridine (BrdU) to 25 µg/ml final. Store up to several months at 4°C. Prepare the 5 mg/ml BrdU stock solution (200×) in water and filter sterilize. Store in the dark at −20°C. After thawing 5 mg/ml BrdU, vortex to be sure it is in solution before adding to MEM.

Complete spinner medium-5 MEM spinner medium 5% horse serum Store up to several months at 4°C Supplements are in MEM spinner medium; no addition is necessary. Horse serum is used because it is cheaper than FBS and may give less cell clumping.

COMMENTARY Background Information An overview of the vaccinia life cycle and expression system is presented in UNIT 16.15. Because HeLa cells consistently give high yields of virus, they are routinely used for preparation of virus stocks. HeLa S3 cells, as obtained from the ATCC, grow well in monolayer culture but can also be put into suspension culture. After repeated passages in suspension, they do not adhere well to flasks and grow poorly in monolayer cultures. The authors prefer HeLa cells adapted to suspension culture because large numbers can be grown in a single bottle. However, suspension cultures require more maintenance than monolayer cultures and the latter may be more convenient. Suspension cells are allowed to form a monolayer before infection to increase the chances of cell-to-cell spread if not all cells are initially infected with the viral inoculum. Thus, good yields of virus may be obtained even if the inoculum is 90% of the total counts), plot the decrease in soluble 35SO4 on a log scale for typical first-order decay kinetics. Determine if single or multiple kinetic classes exist, as indicators of the number of types of sulfate esters in the sample. See Figure 17.23.1 for examples of single and multiple kinetic classes.

Preparation and Analysis of Glycoconjugates


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BASE HYDROLYSIS FOR RELEASE OF SULFATE ESTERS FROM GLYCOSAMINOGLYCANS This method will release sulfate esters that are sensitive to base hydrolysis (see Background Information). Additional Materials (also see Basic Protocol 2) Sample: 35SO4-labeled, borohydride-reduced (UNIT 17.5) oligosaccharide or glycopeptide 2 N NaOH 2 N HCl (freshly diluted from 12 N reagent-grade acid) 1. Dispense 1000 to 2000 cpm 35SO4-labeled, borohydride-reduced oligosaccharide or glycopeptide in 50 µl water into a series of at least six microcentrifuge tubes. Add 50 µl of 2 N NaOH to each tube and mix. 2. Incubate the tubes at 80°C in a heating block with wells filled with oil for 0, 1, 2, 4, 6, and 10 hr. IMPORTANT NOTE: Use prereduced oligosaccharides; otherwise the base may degrade the oligosaccharide chain. The base hydrolysis occurs at about the same rate as the loss of primary-linked sulfates (T1⁄2 = ∼1 to 2 hr).

3. At each time point remove the tube, wipe it clean, and place it on ice for a few minutes. Microcentrifuge 2 sec at 10,000 × g to collect the dispersed droplets. Add 50 µl of 2 N HCl to adjust the pH to 3 to 9, checking 1 to 2 µl of solution with pH paper. 4. After the time course has been completed for all samples and all samples have been neutralized, perform steps 5 to 8 of the acid hydrolysis protocol (see Basic Protocol 2) to precipitate the solubilized 35SO4. Completeness of any hydrolysis will be indicated by a flattening of the decay curve.


Dowex-50 X8 resin Prewash Dowex-50 resin by stirring 50 to 100 g into 2 liters of water, allowing it to stand 1 hr, then discarding water. Repeat twice, then leave overnight at room temperature. Pour off the final water and resuspend the resin in 3 to 4 vol water. The resin may be stored at 4°C for months under these conditions. COMMENTARY Background Information Solvolysis The conditions of the solvolysis reaction can be adjusted to preferentially remove either Nor O-linked sulfate esters, because N-sulfates (e.g., GlcNSO4 in heparin; not to be confused with sulfated N-linked oligosaccharides) are more sensitive than O-sulfate esters (Inoue and Nagasawa, 1976; Nagasawa et al., 1977). Analysis of Sulfate Esters by Solvolysis or Hydrolysis

Acid hydrolysis It is not possible to get an accurate compostional analysis of “sulfated sugars” by doing

an acid hydrolysis because the sulfate esters are cleaved faster than most glycoside bonds. By selecting the right conditions, acid hydrolysis can give information about the number and locations of sulfate esters because they are cleaved at different rates depending upon their location on the individual sugar residues (Freeze and Wolgast, 1984). The kinetics of sulfate liberation (see Fig. 17.23.1) can help to determine the number of classes of sulfate esters present and what their likely locations may be. These properties are not strongly influenced by other substituents on the sulfated sugar or by the position of the sugar within an oligosac-



charide chain. Since the hydrolysis can also cleave glycosidic linkages and partially destroy the sugar chain, only the release of sulfate can be measured. The acid hydrolysis procedure differentiates between sulfates linked to different hydroxyl groups on the sugar ring. Under the hydrolysis conditions in Basic Protocol 2, the T1⁄2 of sulfates linked to equatorial hydroxyl groups (e.g., glucose-3-sulfate, galactose-3-sulfate, N-acetylglucosamine-3-sulfate, and mannose-3-sulfate) is 6 to 25 min; that of those linked to axial hydroxyl groups (e.g., galactose-4-sulfate, Nacetylglucosamine-4-sulfate, and mannose-2sulfate) is 60 to 84 min, and that of those linked to primary hydroxyl groups (e.g., glucose-6-sulfate, galactose-6-sulfate, N-acetylglucosamine6-sulfate, and mannose-6-sulfate) is 90 to 120 min (Rees, 1963). All 6-OH positions are primary, and all of the hydroxyl groups in glucose are equatorial. Any hexose isomer of glucose (e.g., mannose or galactose) will change one of the equatorial positions to an axial position. Even though the time ranges for hydrolysis listed above have not been extensively documented for sulfate-labeled oligosaccharides, they are useful because they will clearly differentiate between multiple classes of sulfate es-

ters. Using the acid hydrolysis technique, it is not possible to determine which sugar is sulfated or where it is found in the chain. If other data are available on the structure of the oligsoaccharide, this information may provide likely possibilities for the location of sulfate groups, especially if certain residues are resistant to exoglycosidase digestion. Base hydrolysis Only certain types of sulfate esters are sensitive to base hydrolysis (Freeze and Wolgast, 1984; Percival, 1978). Sensitivity is determined by the location of the sulfate on the sugar residue and—in contrast to acid hydrolysis described above—by the other substituents linked to that sugar residue. Base treatment will release sulfate esters from intact oligosaccharides under two different conditions. The first is when sulfate is located in the 6 (or 3)-OH position and the 3 (or 6)-OH group of the same sugar residue is not substituted. The second is when the sulfate ester is adjacent to an unsubstituted trans OH group. In the first case, the sulfate is eliminated and an acid-labile 3,6-anhydrosugar is formed (see Fig. 17.23.2). In the second case, the sulfate is released with the formation of an epoxide

O HSO3

O

CH2

OCH2 O OH OH OH

O OH

OHOH

OCH2

O OH OH OH

CH2

O R

mild H+ OH

OCH2 O OH OH OH

O R

O OH OH

+ HOCH2 O OH OH OH O R

strong H+

CH2

OH

O O OH OH

HOCH2

+

O OH OH OH OH

Figure 17.23.2 Formation of 3,6-anhydromannose by base treatment of an oligosaccharide containing mannose-6-sulfate. The formation of the anhydro sugar makes the glycosidic linkage very sensitive to acid; mild hydrolysis causes cleavage. Stronger hydrolysis degrades the usual glycosidic linkages, but leaves the anhydrosugar. When reduced with sodium borohydride, the modified sugar can be quantified to show how much mannose-6-sulfate was originally present.

Preparation and Analysis of Glycoconjugates


Supplement 32

ring on the sugar. This is destroyed during subsequent acid hydrolysis of the oligosaccharide. The modified sugar can be identified if it is metabolically labeled—e.g., as in the discovery of mannose-6-sulfate (Freeze and Wolgast, 1984)—but this is not routinely done. On the other hand, the loss of sulfate is easily measured. When data on the acid and base hydrolysis of sulfate esters are combined with other structural information on the molecules, several potential structures can be eliminated or supported, but these procedures will not provide proof of the structure.

Critical Parameters Solvolysis is a harsh treatment that also removes sialic acids and probably fucose residues. If these sugars are normally present, their loss may be an unacceptable side effect of this prcedure. On the other hand, hexoses, hexuronic acids, and hexosamines in glycosidic linkages are mostly stable to solvolysis. The incubated control is very important because it will monitor for any nonspecific destruction of the sugar chain. For acid hydrolysis, the most critical points to observe are that the pH should be kept between 2 and 3 and that the salt concentration be kept very high during the precipitation of sufate with barium. Only then is the precipitation specific for sulfate. Under less stringent conditions, the oligosaccharides or even monosaccharide sulfates can be precipitated.

Time Considerations Solvolysis itself takes only a few hours to complete. The rate-limiting step is the time needed to remove the water after the sample has passed through the Dowex-50 column and the pyridine has been added. This will only take a few minutes using a shaker-evaporator, but will require an overnight lyophilization. Beginning with the hydrolysis step, the procedure can be completed within a day. If necessary, the procedure can be stopped after taking all of the time points and neutralizing them with base prior to precipitation. Where this is done, the reaction mixtures can be stored at 0°-4°C for 24 hr or more.

Literature Cited Inoue, Y. and Nagasawa, K. 1976. Selective N-desulfation of heparin with dimethyl sulfoxide containing water or methanol. Carbohydr. Res. 46:87-95. Nagasawa, K., Inoue, Y., and Kamata, T. 1977. Solvolytic desulfation of glycosaminoglycuronan sulfates with dimethyl sulfoxide containing water or methanol. Carbohydr. Res. 58:47-55. Freeze, H. and Wolgast, D. 1984. Structural analysis of N-linked oligosaccharides from glycoproteins secreted by Dictyostelium discoideum. J. Biol. Chem. 261:127-133. Rees, D.A. 1963. A note on the characterization of carbohydrate sulphates by acid hydrolysis. Biochem J. 88:343-346. Percival, E. 1978. Sulfated polysaccharides of the Rhodophycea. In Carbohydrate Sulfates. (R.G. Schweiger, ed.) pp. 213-224. American Chemical Society, Washington, D.C.

Anticipated Results

Expect ∼80% to 90% efficiency in desulfation, and 90% of the total protein serine/threonine phosphatase activity in many cell extracts (Shenolikar and Nairn, 1991) and demonstrate a broad in vitro substrate specificity. Therefore, PP1 and PP2A are the most commonly used reagents for in vitro dephosphorylation of phosphoserine- and phosphothreonine-containing proteins. PP1 and PP2A are inhibited by microcystin-LR as well as several other recently discovered inhibitors, including okadaic acid, calyculin A, and tautomycin, albeit with differing IC50 values. Inhibitors, such as okadaic acid (with IC50 for PP2A of 0.1 to 1.0 nM and for PP1 of 10 to 100 nM), calyculin A (with an IC50 for PP1 of 1 nM and for PP2A of 10 nM), and microcystin-LR (with an IC50 for both PP1 and PP2A of 0.1 nM) are used to exclude nonspecific effects of PP1 and PP2A. These inhibitors represent important experimental tools for establishing the physiological importance of reversible protein phosphorylation. PP2B, also called calcineurin, is a Ca2+/calmodulin-dependent phosphatase and demonstrates a narrow in vitro substrate specificity compared to PP1 or PP2A. The enzyme purified from mammalian tissues consists of a catalytic subunit that is tightly associated with a calcium-binding regulatory subunit. PP2B has no activity in the absence of 1 µM calmodulin and 1 mM CaCl2 and can be activated by either Ca2+/calmodulin or 1 mM MnCl2. Many highly purified PP2B preparations show no activity in the presence of Ca2+/calmodulin but are fully activated by Mn2+. The immunosuppressive drugs cyclosporin A and FK506 selectively inhibit PP2B in vivo and in vitro. PP2B in the presence of Ni2+ ions and PP2A in association with an endogenous regulator protein or viral antigens can dephosphorylate proteins and peptides containing phosphotyrosines. However, under the conditions described in Basic Protocol 2, PP1, PP2A, and PP2B are highly selective for dephosphorylation of phosphoserine and phosphothreonine residues. PP1 and PP2A represent the major phosphohistidine phosphatase activity measured in tissue extracts, but the regulatory importance of this modification has not yet been established.

Analysis of Protein Phosphorylation


Supplement 33

Critical Parameters Some commercial preparations of potato acid phosphatase contain contaminating protease activity. Because phosphorylation sites reside near the surface of the substrate protein, they are often accessible to proteases, so functional changes that result from limited proteolysis can be incorrectly attributed to dephosphorylation. To limit the effects of proteases, high enzyme concentrations or prolonged incubations with the general phosphatases should be avoided. Aside from including protease inhibitors in the reaction, two strategies should be considered to distinguish proteolysis from dephosphorylation. Formation of a complex between trichloroacetic acid–soluble [32P]phosphate and ammonium molybdate (Support Protocol) is particularly useful in distinguishing [32P]phosphate from 32P-labeled phosphopeptides. The use of phosphatase inhibitors in control reactions also identifies those effects that can be attributed to reactions in which there are contaminating proteolytic enzymes. Commercial preparations of PP2A and PP1 contain only the free catalytic subunits. In vivo these subunits associate with regulatory subunits, which modulate their substrate specificity. However, the catalytic subunits are the most readily purified to homogeneity and can be stored for prolonged periods without significant changes in their enzymatic properties. Both PP1 and PP2A catalytic subunits retain high activity in the absence of divalent cations; PP2A activity may be further stimulated by 1 mM Mn2+ in the enzyme reaction. Long-term storage of PP1 and PP2A results in an apparent loss of enzyme activity. However, these inactive enzymes can be fully reactivated by including 1 mM MnCl2 in the assay. The Mn2+-dependent PP1 enzyme shows slightly altered substrate specificity when compared with PP1 purified from tissues. Moreover, PP1 is potentially inhibited by several endogenous inhibitors. Thus, PP2A may be the preferred reagent for in vitro dephosphorylation of substrates in many cell extracts. Recent studies show that some tissues may contain endogenous PP2A inhibitors (Li et al., 1995). These inhibitory proteins could hinder the ability of PP2A to dephosphorylate proteins in some tissue preparations.

Troubleshooting Detection of Phosphorylation by Enzymatic Techniques

The absence of a functional change following incubation of the substrate with one or more phosphatases may indicate either that the protein was not phosphorylated or that the dephosphorylation of existing phosphates has no effect

on protein function. It is also possible that components in the reaction mixture inhibited the phosphatases. The latter possibility can be readily established if the substrate protein has been metabolically labeled with [32P]phosphate. SDS-PAGE followed by autoradiography or phosphoimage analysis can directly monitor the time-dependent loss of radiolabel from substrate protein. Coincidence between the removal of radiolabeled phosphate and change of function of the substrate protein can be used to distinguish between important regulatory modifications and “silent” phosphorylations. In analyses of unlabeled substrates, the reaction should include an internal standard or a known 32P-labeled protein or peptide. Dephosphorylation of the radiolabeled marker in the presence or absence of the sample should establish the extent to which activity of the added phosphatase has been compromised by components in the sample. If several different phosphatases fail to elicit a change in protein function under conditions that dephosphorylate the marker, the substrate protein may contain no protein-bound phosphate or phosphorylation may not be functionally important for the protein.

Anticipated Results Most phosphoproteins should be dephosphorylated by the general phosphatases, leading to complete removal of protein-bound phosphate and functional changes indicating that reversible phosphorylation controls the protein function. In a number of cases, metabolic labeling and phosphoamino acid analysis or the use of specific protein (phosphoserine/phosphothreonine or phosphotyrosine) phosphatases has successfully identified the amino acids modified. Such studies have then located the phosphorylated residues, especially in proteins whose primary structures have been determined by amino acid sequencing or cDNA cloning. A combination of biochemical studies and site-directed mutagenesis has then been used to establish the functional role of phosphorylation at individual sites. Increasing evidence points to the presence of both positive and negative regulatory phosphorylations in proteins. Thus, it is difficult to predict the functional outcome of complete in vitro dephosphorylation of many substrates.

Time Considerations Approximately 1 hr is required to complete the dephosphorylation reaction. SDS-PAGE



and analysis of the gel require 1.5 hr. The time required for functional analysis depends on the particular assay used.

Literature Cited

Shenolikar, S. and Ingebritsen, T.S. 1984. Protein (serine and threonine) phosphate phosphatases. Methods Enzymol. 107:102-129. Shenolikar, S. and Nairn, A.C. 1991. Protein phosphatases: Recent progress. Adv. Second Messenger Phosphoprotein Res. 23:1-123.

Berger, H.A., Travis, S.M., and Welsh, M.J. 1993. Regulation of the cystic fibrosis transmembrane conductance regulator Cl− channel by specific protein kinases and protein phosphatases. J. Biol. Chem. 268:2037-2047.

Swarup, G., Cohen, S., and Garbers, D.L. 1981. Selective dephosphorylation of proteins containing phosphotyrosine by alkaline phosphatases. J. Biol. Chem. 256:1447-1452.

Charbonneau, H. and Tonks, N.K. 1992. 1002 protein phosphatases? Annu. Rev. Cell Biol. 8:463493.

Van Etten, R.L. and Waymack, P.P. 1991. Substrate specificity and pH dependence of homogeneous wheat germ acid phosphatase. Arch. Biochem. Biophys. 288:634-645.

Li, M., Guo, H., and Damuni, Z. 1995. Purification and characterization of two potent heat-stable inhibitors of protein phosphatase 2A from bovine kidney. Biochemistry 34:1988-1996. Shenolikar, S. 1994. Protein serine/threonine phosphatases: New avenues for cell regulation. Annu. Rev. Cell Biol. 10:55-86.

Contributed by Shirish Shenolikar Duke University Medical Center Durham, North Carolina



Supplement 33

Production of Antibodies That Recognize Specific Tyrosine-Phosphorylated Peptides

UNIT 18.6

It is possible to produce anti-phosphopeptide antibodies (i.e., antibodies recognizing phosphorylated peptides) that recognize a protein only in its phosphorylated state, and that do not cross-react with either the cognate unphosphorylated protein or other phosphoproteins. Because the state of phosphorylation is often indicative of a protein’s functional state or activity, such antibodies provide a convenient probe of the functional state of a protein. Thus, unlike conventional antibodies, anti-phosphopeptide antibodies provide information regarding not only the abundance of a protein but also its activity. Unlike general anti–phosphoamino acid (e.g., anti-phosphotyrosine) antibodies, which have broad reactivity, anti-phosphopeptide antibodies may have unique specificity toward the cognate proteins. Such reagents not only facilitate conventional in vitro analysis of phosphoproteins, but also allow heretofore impossible applications—e.g., differential isolation of species of a particular protein that have been phosphorylated at individual phosphorylation sites, as well as analysis of the functional state of a protein in situ by immunohistochemical techniques. Basic Protocol 1 describes the production of polyclonal anti-phosphopeptide antibodies and Basic Protocol 2 describes the production of monoclonal anti-phosphopeptide antibodies. Both of these procedures are based upon immunizing animals with oligopeptides that have been synthesized containing phosphorylated tyrosine; hence, a knowledge of the sequence and phosphorylation site of the protein of interest is a prerequisite for either protocol. Such an immunization will generate an immune response with at least four components: (1) anti–carrier protein reactivity, (2) general anti-phosphotyrosine reactivity, (3) phosphorylation-independent anti-peptide reactivity, and (4) phosphorylation-dependent anti-peptide reactivity. For the production of polyclonal antibodies (see Basic Protocol 1), a multiple-step affinity chromatographic purification with several negativeselection steps is carried out to produce a final antibody preparation having the desired reactivity. For production of monoclonal antibodies (see Basic Protocol 2), ELISAs are performed to screen candidate hybridoma supernatants against the cognate phosphopeptide, as well as against related phosphorylated and nonphosphorylated peptides, until one is found with the desired reactivity and specificity. Although monoclonal antibodies have a number of advantages, production of polyclonal antibodies is likely to be more predictable, as in the authors’ experience a monoclonal hybridoma clone of stringent specificity may occur with very low frequency (see Critical Parameters). Production of monoclonal antibodies is also generally more time-consuming and expensive. The relative merits of monoclonal and polyclonal antibodies are discussed in the Chapter 11 introduction and in general antibody guides (see Key References). Support Protocols are provided for the coupling of peptides (Support Protocol 2) and phosphotyrosine (Support Protocol 3) to the affinity matrix (Affi-Gel 10); BSA-agarose affinity matrix is commercially available. General methods for the coupling of peptides to carrier proteins for use in immunization are described in UNIT 11.16. This unit describes production of antibodies against tyrosine-phosphorylated peptides, with which the authors have the most expertise, but the principles discussed here also apply to peptides phosphorylated on serine and threonine (see Key References).

Analysis of Protein Phosphorylation Contributed by Michael P. DiGiovanna, Robert R. Roussel, and David F. Stern Current Protocols in Molecular Biology (1996) 18.6.1-18.6.19 Copyright © 2000 by John Wiley & Sons, Inc.


BASIC PROTOCOL 1

PRODUCTION OF POLYCLONAL ANTI-PHOSPHOPEPTIDE ANTIBODIES Strategic Planning The first step of the planning phase is design and synthesis of the required peptides (see Support Protocol 1 and Background Information). The authors employ a peptide containing fifteen amino acids with the phosphorylation site Tyr in the center (residue 8). This peptide consists of fourteen amino acids from the actual sequence of the protein plus an N-terminal Lys to enhance coupling of the peptide to a carrier protein (BSA is used as a carrier in this protocol). The peptide, coupled to BSA, is then used to immunize rabbits. The cognate phosphopeptide is also immobilized on an affinity-chromatography column support (see Support Protocol 2); this affinity matrix will ultimately be used to purify the antibody from the rabbit serum. In addition, a series of negative-selection affinity-chromatography columns are used to adsorb antibodies from the serum that cross-react with epitopes other than the cognate phosphopeptide. These columns are composed of matrix coupled to various synthetic peptides—principally the cognate nonphosphorylated peptide (to remove phosphorylation-independent antibodies) and phosphotyrosine itself (to remove indiscriminant anti-phosphotyrosine reactivity; see Support Protocol 3 for preparation of phosphotyrosine affinity columns). Additional negative-selection steps may be used if needed, depending upon the particular target chosen, to remove antibodies that cross-react with phosphorylation sites of other proteins having known homology to the target site. This cross-reactivity occurs particularly when the target protein is a member of a family of homologous proteins—e.g., a subgroup of related receptor tyrosine protein kinases. If such cross-reactivity is anticipated, it is possible to synthesize not only the cognate phosphorylated and nonphosphorylated peptides, but also phosphopeptides based upon the related homologous sequences, to be used for negative selection. The authors also pass the serum over a column consisting of immobilized carrier protein (commercially available BSA-agarose) to remove the majority of the antibodies generated against the much larger carrier protein. Thus, a typical purification scheme might consist of the following: Negative-selection affinity-purification columns • BSA (carrier) • phosphotyrosine • cognate nonphosphopeptide • homologous phosphopeptides (if desired) Positive-selection affinity-purification column • cognate phosphopeptide.

Production of Antibodies That Recognize Phosphorylated Peptides

The second step in planning is to raise and test antisera in rabbits. To prepare the phosphopeptide-carrier conjugate for immunization of rabbits, the authors couple a 30-fold molar excess of phosphopeptide (14.4 mg dissolved in water and neutralized with 1 N NaOH) to BSA (20 mg dissolved in 0.4 M sodium phosphate buffer, pH 7.5), using glutaraldehyde as the cross-linking reagent as described in Doolittle (1986). The reaction is allowed to proceed 30 min at room temperature; the development of a yellow color in the coupling solution is an indication that the glutaraldehyde has reacted. General methods for coupling of peptides to carrier proteins for use in immunization are described in UNIT 11.16. Conjugation can be confirmed by performing an anti-phosphotyrosine immunoblot (UNIT 18.4) on a small test aliquot of the conjugate (although a smear should be expected on the blot, resulting from variation in the amount of peptide coupled per BSA molecule as well as from possible multimers of BSA). Guidelines for immunization of rabbits are provided in UNIT 11.12, and many institutions have core facilities for the immunization of



rabbits and production of antisera. The authors immunize with 1 mg of immunogen (the peptide-BSA conjugate prepared above, dissolved in PBS) in 1 ml of a 50% emulsion with complete Freunds adjuvant. This is administered subcutaneously at multiple sites, followed by four boosts at 2-week intervals using same quantity of immunogen in incomplete Freunds adjuvant. An adequate immune response can be seen as early as week 6. The specificity of the final antibody is demonstrated most simply by immunoblot analysis (UNIT 10.8) using a panel of relevant phosphorylated and nonphosphorylated proteins. Electrophoresis is carried out with purified cognate protein in both its phosphorylated and unphosphorylated states, as well as with any proteins toward which there could conceivably be cross-reactivity. These are most easily isolated by immunoprecipitation using conventional antibodies. The blot is probed with the purified anti-phosphopeptide antibody to demonstrate appropriate reactivity. This immunoblot assay may also be used after each chromatographic step to monitor the success of the purification. An assay also must be devised for estimating the titer of the immune response in the sera of the immunized rabbits. Convenient screening assays for this purpose include tests for the ability of crude sera to immunoprecipitate the target phosphoprotein from a cell lysate known to contain it, as well as immunoblotting of purified phosphorylated target protein with the crude serum. The relationship among the above procedures is illustrated by the flow chart in Figure 18.6.1. Purification Method The purification of polyclonal antibodies consists of multiple affinity-chromatography steps for the negative and positive selection of antibodies of the appropriate reactivity (see Fig. 18.6.1). All of the following chromatographic steps may be carried out at room temperature, with the solutions applied to the columns by gravity. Materials BSA-agarose affinity matrix (Sigma) packed (as in Support Protocol 2) in a 10-ml bed volume column Phosphotyrosine affinity matrix column (10-ml bed volume; see Support Protocol 3) Crude serum from rabbit immunized with phosphopeptide-BSA conjugate (refer to Strategic Planning, above) PBS/azide: PBS (APPENDIX 2) containing 0.02% (w/v) sodium azide (store indefinitely at 4°C or room temperature) Cognate nonphosphopeptide affinity matrix column (3-ml bed volume; see Support Protocols 1 and 2 and Strategic Planning) 3 M NaSCN Homologous phosphopeptide affinity matrix columns (optional; 3-ml bed volume; see Support Protocols 1 and 2 and Strategic Planning) Positive-selection phosphopeptide affinity matrix column (3-ml bed volume; see Support Protocols 1 and 2 and Strategic Planning) 3.5 M and 4.5 M MgCl2 (optional; see Critical Parameters) Spectrophotometer (optional) Dialysis tubing (MWCO 12,000 to 14,000; 10 mm width, 6.4 mm diameter; e.g., Spectra/Por 4 from Spectrum) Additional reagents and equipment for dialysis (APPENDIX 3C), protein quantitation (UNIT 10.1), and analysis of antibodies by ELISA (UNIT 11.2) or immunoblotting (UNIT 10.8)



Supplement 35

design and synthesize peptides of interest (core facilty / Support Protocol 1 / Strategic Planning)

P ~ peptide

test serum for reactivity by immunoprecipitation or immunoblotting

regenerate matrix

immunize rabbits to produce crude antiserum (core facility / UNIT 11.12 )

peptides for matrix preparation

crude serum pass serum through BSA-agarose affinity matrix (negative selection)

flowthrough regenerate matrix

pass serum through phosphotyrosine affinity matrix (negative selection)

test serum for loss of antiphosphotyrosine flowthrough reactivity by immunoblotting (optional; pass serum through repeat cognate nonphosphoUNIT 10.8 ) pass peptide affinity matrix (negative selection) regenerate matrix

prepare phosphotyrosine affinity matrix (Support Protocol 3 )

flowthrough test serum for loss of anti-nonphosphopeptide or anti–homologous phosphopeptide reactivity by immunoblotting (optional; UNIT 10.8 )

pass serum through homologous phosphopeptide matrices; (negative selection; optional )

pass serum through affinity matrix containing phosphopeptide of interest (positive selection)

prepare various peptide affinity matrices (Support Protocol 2 )

elute with NaSCN or MgCl2 ; collect fractions; dialyze vs. PBS/azide

assay by A 280 (UNIT 10.1 )


assay by ELISA (UNIT 11.2 )

assay by immunoblotting (UNIT 10.8 )

Figure 18.6.1 Flow chart for production of polyclonal anti-phosphopeptide antibodies.



Deplete reactivity of serum toward phosphotyrosine and carrier protein 1. Connect the washed BSA-agarose and phosphotyrosine affinity matrix columns in series. 2. Pass ∼15 ml crude serum through the two columns, allowing it to flow through by gravity. Wash with PBS/azide until all of the yellow color of the serum has passed through the columns (or monitor the A280 of the column effluent spectrophotometrically until baseline absorbance is reached), then wash with an additional 5 to 10 ml PBS/azide, collecting all washings in the flowthrough fraction, which will contain the antibody of interest. Aliquots of the crude serum as well as this first flowthrough may be saved for subsequent analysis and comparison of individual purification fractions. The serum may be analyzed at this time for elimination of anti-phosphotyrosine reactivity by performing immunoblotting (UNIT 10.8) of samples containing a variety of phosphotyrosyl proteins; alternatively it is possible to save an aliquot for analysis and proceed to the next step (see Troubleshooting). Expect that with each passage through a column, the volume of the serum will increase, which will prolong the amount of time needed for subsequent columns. Note that if flowthrough is monitored by the passage of the yellow color of the serum, this will become more difficult as the serum becomes more dilute. After the serum passes through, regenerate the columns by washing with 10 bed volumes of 3 M NaSCN followed by 10 bed volumes of PBS/azide, and store at 4°C in PBS/azide for future use.

Deplete reactivity of serum toward cognate nonphosphopeptide 3. Pass the flowthrough serum through the nonphosphopeptide affinity matrix column by gravity as many times as necessary to deplete cross-reactivity, regenerating the column between passes by washing with 10 bed volumes of 3 M NaSCN followed by 10 bed volumes PBS/azide. A 15-ml starting serum sample may have to be passed over this column several times to quantitatively deplete cross-reactivity (see Critical Parameters). The serum may be analyzed after each or several passes, or an aliquot can be saved for analysis at a later time.

4. If cross-reactivity with homologous phosphoprotein(s) is anticipated, pass the flowthrough serum through the homologous phosphopeptide affinity matrix column(s) using the methodology described in step 3. Refer to Strategic Planning, above, for discussion of cross-reactivity with homologous phosphoproteins.

Purify antibodies by positive-selection affinity chromatography and dialysis 5. Hydrate and wash ∼25-cm strips of dialysis tubing in advance for collection of fractions from the positive-selection affinity purification. Secure one end of each length of tubing with a dialysis clamp and check for leaks (APPENDIX 3C). Also prepare 6 liters PBS/azide and cool to 4°C in advance to use as the dialysis solution. To minimize the time that the antibodies are exposed to the elution solution, fractions (∼3 ml each) from the positive-selection affinity column will be collected directly into preprepared dialysis tubing.

6. Pass the flowthrough serum from the previous chromatography step through the positive-selection phosphopeptide affinity column three times, without washing between passes. Three passes are performed to maximize the interaction of the antibody with the affinity matrix.



Supplement 50

7. Collect the flowthrough serum from the final pass, then wash the column with 5 to 20 ml PBS/azide (depending upon the precolumn volume and column bed volume) and combine the washings with the flowthrough serum. Wash with an additional 20 ml PBS/azide, and collect the washings separately as the “wash” fraction. 8. Elute with chaotropic agent of choice: either 20 ml of 3 M NaSCN, or 10 ml of 3.5 M MgCl2 followed by 10 ml of 4.5 M MgCl2. Upon starting the elution, immediately begin collecting ∼3-ml fractions directly into the dialysis bags prepared in step 5, securing the proximal end of the tubing with dialysis clamps and dropping the bag immediately into the PBS/azide dialysis solution. Collect at least six fractions. Alternatively, ∼3-ml fractions can be collected in tubes and the liquid immediately placed into dialysis tubing upon completion of each fraction. Most of the antibody will elute in the first three fractions from a column of 2- to 3-ml bed volume. Regenerate the column as in step 3. See Critical Parameters for a discussion of the relative merits of NaSCN versus MgCl2 as eluants.

9. Dialyze all fractions collected in step 8 exhaustively against PBS/azide at 4°C (APPENDIX 3C). Analyze and store purified antibody 10. Determine protein concentration of dialyzed fractions by measuring absorbance at 280 nm or by a colorimetric protein assay (UNIT 10.1) and calculate yield. The authors have generally recovered ≥1 mg of purified antibody from a 15-ml serum sample.

11. Store the antibody in aliquots at −70°C; store working aliquot at 4°C. 12. Assay the reactivity and cross-reactivity of the final samples, as well as any aliquots saved from previous steps, using ELISA (UNIT 11.2) or immunoblotting (UNIT 10.8). BASIC PROTOCOL 2

PRODUCTION OF MONOCLONAL ANTI-PHOSPHOPEPTIDE ANTIBODIES Strategic Planning


As in the production of polyclonal anti-phosphopeptide antibodies (see Basic Protocol 1), the first step of the planning phase for anti-phosphopeptide monoclonal antibodies is the design and synthesis of the required peptides (see Support Protocol 1 and Background Information). Here too, the authors employ a peptide containing fifteen amino acids with the phosphorylation site Tyr in the center (residue 8), consisting of fourteen amino acids from the actual sequence of the protein plus an N-terminal Lys to enhance coupling to a carrier protein. A different carrier protein must be employed for immunization than will be used as substrate for the ELISA screen, to avoid detection of anti-carrier antibodies in the ELISA (see Purification Method, below). The authors use keyhole limpet hemocyanin (KLH) as the carrier for immunization and BSA as the carrier for the ELISA substrate. General methods for coupling of peptides to carrier proteins for use in immunization are described in UNIT 11.16. The cognate phosphopeptide can also be immobilized to an affinity-chromatographic column support (see Support Protocol 2) and used ultimately to affinity-purify the antibody produced by the desired hybridoma clone. A number of additional peptides must be produced for use in ELISA screening of candidate hybridoma supernatants, to screen both for the presence of the desired reactivity and the absence of undesired cross-reactivity. For ELISA screening, the peptides are coupled to a carrier protein different from the one used for immunization. Bovine serum albumin (BSA) is



used for screening, because many of the clones are likely to react against the carrier used in the immunization of the mice (KLH) and because unconjugated peptides have not been found to function well as ELISA substrates (Doolittle, 1986). In addition, because antibodies may be generated against the cross-linking reagent, the method used for coupling the phosphopeptide to the BSA to produce the substrate for ELISAs should be different from that used to link peptides to the KLH carrier for immunization (Czernik et al., 1991; Doolittle, 1986). The second step of planning is to raise antibodies in mice. General guidelines for the immunization of mice and the production of monoclonal antibodies are described in UNITS 11.4-11.11, and many institutions have core facilities that will perform that task. The authors have immunized BALB/c mice by intraperitoneal injection with 1 mg/ml of immunogen (dissolved in PBS) in a 50% emulsion with complete Freunds adjuvant on day 1 and boosted intraperitoneally on days 15 and 37 with immunogen in incomplete Freunds adjuvant. Test bleeds from day 47 were analyzed by ELISA. On day 57 the mouse with the best titer was boosted intravenously, and its spleen was harvested on day 60. Freshly harvested spleen cells were prepared for cell fusion to generate hybridoma lines, which were subsequently screened by ELISA (UNIT 11.7). The purpose of the ELISA screening is to identify clones that react with the cognate phosphopeptide and to eliminate clones that cross-react with other epitopes in addition to the cognate phosphopeptide. Epitopes screened by ELISA generally consist of the cognate nonphosphorylated peptide (to eliminate clones with phosphorylation-independent reactivities) and at least one unrelated phosphopeptide (to eliminate clones with indiscriminant anti-phosphotyrosine reactivity). In addition, depending upon the particular target chosen, it may be necessary to screen for reactivity with phosphopeptides from proteins of known homology. This occurs particularly when the target protein is a member of a family of homologous proteins—e.g., the subgroups of related receptor tyrosine protein kinases. If such cross-reactivity is anticipated, it is essential to synthesize not only the cognate phosphorylated and nonphosphorylated peptides and at least one unrelated phosphopeptide, but also phosphopeptides based upon the related homologous sequences. Thus, a typical ELISA screening scheme might consist of the following steps. 1. Screen for clones that exhibit reactivity against the cognate phosphopeptide. These clones, which will likely be 20 min to elapse from the time the resin is removed from the bottle to the time it is mixed with the phosphotyrosine solution.

4. Remove most of the excess liquid from the gel by vacuum aspiration without letting it dry completely, transfer it to a screw-cap centrifuge tube, and add the phosphotyrosine solution from step 2. 5. Incubate overnight at 4°C with end-over-end rotation. Quench and wash resin 6. Add 2 µl pure ethanolamine per ml resin and incubate 2 hr at room temperature with end-over-end rotation. Ethanolamine is added directly to the coupling reaction to quench unreacted ester groups.

7. Wash resin twice, each time by centrifuging at low speed (see Support Protocol 2, step 3), aspirating the supernatant, resuspending the resin in 5 vol of 0.4% sodium bicarbonate, then centrifuging again at low speed. 8. Wash resin twice with 0.1 M ethanolamine⋅HCl, pH 8.0, as in step 7. Remove the 0.1 M ethanolamine from the second wash and replace with fresh, incubate tube overnight at 4°C with end-over-end rotation, then centrifuge at low speed and aspirate the supernatant. 9. Wash, store, and pack resin in columns (see Support Protocol 2, steps 8 to 12). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Screening diluent 6.18 g boric acid (100 mM final) 9.54 g sodium borate (47 mM final) 4.38 g NaCl (75 mM final) 10 g bovine serum albumin (BSA; 1% w/v final) 1 g sodium azide (1% w/v final) H2O to 1 liter The pH of the solution will be 8.4 to 8.5.



Conventional antibodies have proven to be invaluable tools in numerous techniques for the biochemical analysis of proteins. In the past antibodies with specificity toward the phosphorylated forms of proteins were produced serendipitously. Using techniques such as those described in this unit, such antibodies may now be produced by design. Whereas

phosphorylated holoproteins might in general be poor candidates for immunogens in the production of such antibodies as a result of their susceptibility to dephosphorylation, it has been suggested that short phosphopeptides are relatively resistant to phosphatases (Czernik et al., 1991), thereby providing a better chance of success. A dramatic advance in the analysis of pro-



tein tyrosine phosphorylation and the regulation of signal transduction pathways by such phosphorylation occurred with the production of polyclonal and monoclonal anti-phosphotyrosine antibodies (Frackelton et al., 1983; Ross et al., 1981; reviewed in Stern, 1991). These antibodies proved capable of recognizing phosphorylated tyrosine residues in the context of virtually any flanking peptide sequence, and were shown to be exquisitely specific in the requirement for a phosphorylated tyrosine yet at the same time remarkably promiscuous in their acceptance of any peptide sequence. Techniques using polyclonal and monoclonal antiphosphotyrosine antibodies have supplanted the standard methodology of metabolically labeling proteins with 32Pi, which was cumbersome, time-consuming, hazardous, and of relatively lower sensitivity. Antibodies that recognize a specific tyrosylphosphorylated peptide as described in this unit represent a marriage of anti-phosphotyrosine technology with the stringent sequence-dependent specificity of conventional antibodies (Bangalore et al., 1992; DiGiovanna and Stern, 1995; Epstein et al., 1992; Roussel, 1990). Conceptually, they may be thought of either as anti-phosphotyrosine antibodies that have strict sequence specificity, or as conventional antibodies possessing the additional specificity of phosphorylation dependence. Anti-phosphotyrosine antibodies have been most useful in the analysis of tyrosine phosphorylation of proteins in a technique using a combination of immunoprecipitation and immunoblotting (known as the “IP-western” method). In this procedure, a protein typically is immunoprecipitated either with conventional anti-protein antibody or with anti-phosphotyrosine antibody, then immunoblotted with whichever of these two antibodies was not used for the immunoprecipitation. Because antiphosphopeptide antibodies recognize the cognate protein, but only when it is phosphorylated, they are capable of performing both functions in a single step—i.e., either an immunoprecipitation alone or an immunoblot alone is sufficient to supply the desired information. The most useful functions for such antibodies are likely to be those that in fact are not possible using traditional reagents. For example, using anti-phosphopeptide antibodies, it is possible to identify and isolate distinct phosphorylated species of a phosphoprotein containing multiple phosphorylation sites. Thus, phosphorylation at each site can be examined

independently, and a preparative separation of individual phosphospecies of a holoprotein can be achieved. Another unique application is analysis of the abundance and phosphorylation state of individual proteins in situ in preparations of cells or tissues. Because phosphorylation is a major mode of regulation of protein function, the phosphorylation state is often an indicator of the functional status of a protein. The mere identification of a protein in a cell or tissue specimen gives no indication of its functional status. The ability of anti-phosphopeptide antibodies to demonstrate the phosphorylation state, and by extrapolation the functional state, of a single protein with high specificity places these reagents in a unique class. Hence, identity and functional status may be probed simultaneously using one simple assay. In tissue sections, one may probe to determine whether a particular protein is present, and, if present, in what functional state it is found. The authors have employed this strategy in immunohistochemical staining of formalin-fixed, paraffin-embedded human breast tumors using antibody to the phosphorylated form of the receptor tyrosine kinase Neu (DiGiovanna and Stern, 1995), and were able to demonstrate that Neu is phosphorylated, and therefore functionally active, in only a subset of the tumors that overexpress this protein. Anti-phosphopeptide antibodies possess unique properties that render them capable of performing functions not possible with conventional reagents. From the technical considerations regarding their production, it would follow that antibodies with specificity for the nonphosphorylated state should also be achievable, and the production of such antibodies has also been reported (Czernik et al., 1991; Epstein, 1995; Kawakatsu et al., 1996; Nairn et al., 1982; Roussel, 1990; Tzartos et al., 1995).

Critical Parameters In the production of polyclonal anti-phosphopeptide antibodies, a major challenge is the depletion of cross-reactivity. The first consideration in achieving this is the anticipation of potential cross-reacting proteins in the planning phase of the procedure. Cross-reactivity is most likely to occur when the target protein is a member of a family of closely related homologous proteins—e.g., of the subfamilies of tyrosine protein kinases. The authors have produced a monoclonal anti-phosphopeptide antibody with specificity for the Tyr-1248 autophosphorylation site of Neu (DiGiovanna and Stern, 1995). A homologous site exists in



Supplement 35


the epidermal growth factor receptor (EGFR). The antibody produced in the authors’ laboratory does not cross-react with the homologous EGFR site in spite of having identical residues in seven of the 14 amino acids of the peptide sequence (five of which are N-terminal to the phosphotyrosine, one C-terminal, and the phosphotyrosine itself). Thus, specificity is achievable even with at least up to 50% identity. Nevertheless, such a close relationship among the phosphorylation sites of different proteins highlights the importance of considering all known related proteins. The second consideration in depleting cross-reactivity is the quantity of cross-reactive antibodies in a given aliquot to be purified relative to the capacity of the peptide columns used for the negative selection. Because of the expense of the peptides, the authors have generally prepared small columns (with 2- to 3-ml bed volumes) and passed the sera over the columns multiple times, empirically determining the number of passes necessary to quantitatively deplete cross-reacting material. In these columns, 3 µmol of ligand is coupled per milliliter of Affi-Gel 10. The manufacturer states that the resin contains ∼15 µmol of active ester per milliliter of gel, and that the gel has a capacity for 35 mg of protein or 15 to 20 µmol of a low-molecular-weight ligand per milliliter of gel. The BSA and phosphotyrosine columns are relatively inexpensive to produce; thus, a single large column of each is practical and sufficient. Because the authors elute the antibodies from the positive-selection affinity column in strongly chaotropic solutions that are potentially deleterious to the stability of the antibody (3 M NaSCN or 3.5 M followed by 4.5 M MgCl2), care is taken to collect the fractions in preprepared dialysis tubing so that they may be immediately placed into the PBS dialysate, thus minimizing the time that the antibodies are exposed to the eluting solutions. Although MgCl2 is thought to be “gentler,” the authors have found that gravity-driven flow rates from phosphopeptide columns eluted with this salt quickly become extremely slow, possibly because of precipitation of the salt in the column or an interaction of the Mg2+ ion with the phosphate groups. Thus use of NaSCN is preferred, and this salt appears to permit recovery of comparable activity. In the production of monoclonal anti-phosphotyrosyl peptide antibody, which the authors have carried out once, a major technical hurdle was the low frequency with which clones of the

desired specificity were produced (DiGiovanna and Stern, 1995). In that work, in which antibody was produced to the phosphorylated form of the receptor tyrosine kinase Neu, >1200 hybridomas (obtained from a single fusion) were screened to obtain a single clone that satisfied all requirements. The authors found 68 candidate clones that recognized the cognate phosphopeptide, of which only 20 were unreactive toward the cognate nonphosphopeptide. Of those, seven cross-reacted with an unrelated phosphopeptide (i.e, exhibited indiscriminant phosphotyrosine activity), and of the remainder three cross-reacted with the homologous EGFR. The remaining ten were subcloned by limiting dilution, after which only five continued to produce antibody. Of these five, only one reliably detected the phosphorylated holoprotein in immunoblots and immunoprecipitations. Thus, the “hit rate” for an antibody that satisfied all requirements was 150 mM. For kinases, it is a good idea to use preliminary experiments to identify the effects of pH, salt concentration, concentrations of cations such as Mg2+ or Mn2+, and temperature to optimize the assay conditions for the kinase of interest. It is often advantageous to perform kinase assays at 30°C rather than 37°C because the lower temperature makes it easier to stay within the linear range of the kinase, thus providing more control of the assay. Another factor to be considered when optimizing an assay for a newly identified kinase is the concentration of ATP in the reaction mixture. In these protocols and in general, detection of kinase activity is based on the transfer of radiolabeled phosphate from ATP to the substrate, so the concentration of ATP can be varied only a little. Also, the specific activity of the [γ-32P]ATP must be known in order to actually measure phosphotransfer. Most kinases have a Km for ATP of 1 to 100 µM. If there is too much ATP in the reaction mixture, it will be difficult to measure phosphotransfer. An ATP concentration of 50 to 100 µM tends to work well. At that concentration, the enzyme should be working at ≥50% of maximum, depending on its apparent Km for ATP, and addition of sufficient [γ-32P]ATP to measure phosphotransfer is not prohibitively expensive. Usually the substrate concentration is high so that the enzyme is working at or close to Vmax. Controls Including the correct controls for a kinase assay is critical to the success of the assay, especially when the enzyme source is a cell or tissue extract. The appropriate controls should always include a no substrate control, a no enzyme source control, a heat-denatured enzyme control, and, for enzymes that require an activator or cofactor, controls that use irrelevant activator or cofactor and controls without activator or cofactor. CAUTION: These assays use [γ-32P]ATP which should be handled and disposed of according to safety regulations. See APPENDIX 1F and the institutional Radiation Safety Office for guidelines for proper handling and disposal of 32P. BASIC PROTOCOL 1

ASSAY FOR CYCLIC NUCLEOTIDE–DEPENDENT PROTEIN KINASES Cyclic adenosine monophosphate (cAMP)–and cyclic guanosine monophosphate (cGMP)–dependent protein kinases (PrKs) are quite similar in structure and require similar assay conditions. cAMP-PrK is a heterotetramer consisting of two regulatory (R) subunits and two catalytic (C) subunits, R2C2. In each case the regulatory subunits have binding sites for two molecules of cAMP. When the cAMP concentration is elevated in cells after activation of receptor-linked adenylyl cyclases, cAMP binds to the R subunits, causing the affinity of the C subunits for R subunits to drop by about four orders of magnitude. The formation of R2cAMP4 dimers and free C-subunits is favored, and the C-subunits are now active. The case of cGMP-PrK is slightly different in that there are no free catalytic subunits released. cGMP-PrK is composed of two identical subunits. Each subunit has a regulatory domain and a catalytic domain, which are homologous to the R and C subunits of cAMP-PrK. On binding four molecules of cGMP the enzyme is activated, presumably by a conformational change.

Assays of Protein Kinases Using Exogenous Substrates

The assay procedures for these two kinases are very similar and the same protein substrate can be used for each one as detailed below. More recently, both of these kinases have been assayed using peptide substrates.



Materials 5× cyclic nucleotide–dependent kinase reaction buffer (see recipe) 10 mg/ml histone 2B in H2O [γ-32P]ATP solution (see recipe) 20× cyclic nucleotide solution: 20 µM cyclic AMP in H2O/20 µM cyclic GMP in H2O Enzyme sample containing cyclic nucleotide–dependent kinase activity (see Support Protocol 1), kept on ice until use 30° water bath Additional reagents and equipment for TCA precipitation (see Support Protocol 2), adsorption onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic analysis (see Support Protocol 4) 1. For each assay reaction, add the following to a 1.5-ml microcentrifuge tube kept on ice: 4 µl 5× cyclic nucleotide–dependent kinase reaction buffer 1 µl of 10 mg/ml histone 2B 1 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 1 µl 20× cyclic nucleotide solution 0 to 13 µl H2O. Cap tube and warm the mixture in a 30°C water bath. Perform each assay in triplicate and include no substrate, no enzyme, and no cyclic nucleotide controls. The total reaction mix volume is 20 ìl per reaction. The amount of water required depends on how much enzyme sample is used in step 2. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mixture (minus enzyme) for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.

2. Start the reaction by adding 1 to 14 µl ice-cold enzyme sample containing cyclic nucleotide–dependent kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads (UNIT 10.16), prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.

3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Use 10 µl of the reaction mix for adsorption to P81 phosphocellulose paper (see Support Protocol 3). Proceed with analysis by one of those methods.



Supplement 40

BASIC PROTOCOL 2

ASSAY FOR PROTEIN KINASE C ISOFORMS There are presently at least ten members of the protein kinase C (PKC) family of kinases. Protein kinase C was originally identified as a kinase activity that was reversibly activated by calcium (Ca2+), phospholipid (usually phosphatidylserine), and the neutral lipid diacylglycerol (DAG; Takai et al., 1979). Subsequently, these kinases were shown to be the sites of action for the tumor-promoting phorbol esters, which bind to some PKC isoforms and activate the kinase activity by substituting for DAG. The initial isoforms of PKC that were purified were called α, β, and γ, and it is these forms that are the most clearly regulated by Ca2+, phospholipid, and DAG or phorbol esters. They are termed the “conventional” subfamily of PKCs (cPKCs). Another subfamily contains δ, ε, ν, and θ, the “novel” PKCs (nPKCs); these kinases are not regulated by Ca2+ but are activated by phorbol esters and phospholipids. A third subfamily, the “atypical” PKCs (aPKCs), consist of ζ, λ, and ι; these are not regulated by Ca2+ nor do they bind phorbol esters or DAG, but there is still a requirement for phospholipids. When the isoforms are analyzed, the reaction buffer must include the appropriate activating cofactors. Materials 5× PKC reaction buffer (see recipe) 10 mg/ml histone H1 in H2O [γ-32P]ATP solution (see recipe) Enzyme sample containing PKC activity (see Support Protocol 1) 30°C water bath Additional reagents and equipment for TCA precipitation (see Support Protocol 2), adsorption onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic analysis (see Support Protocol 4) 1. For each assay reaction, add the following to a 1.5-ml microcentrifuge tube: 4 µl 5× PKC reaction buffer 1 µl 10 mg/ml histone H1 1 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 0 to 14 µl H2O. Cap tube and warm the mixture 10 min in a 30°C water bath. Perform each assay in triplicate and include no substrate and no enzyme controls. The total reaction mix volume is 20 ìl per reaction. The amount of water required depends on how much enzyme sample is used in step 2. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mixture (minus enzyme) for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.

2. Start reaction by adding 1 to 14 µl enzyme sample containing PKC activity to the warmed reaction mixture. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. Assays of Protein Kinases Using Exogenous Substrates

For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads (UNIT 10.16), prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.



3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), 10 µl, or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Use 10 µl of the reaction mix for adsorption to P81 phosphocellulose paper (see Support Protocol 3). Proceed with analysis by one of those methods. ASSAY FOR CASEIN KINASES USING β-CASEIN Casein is a very acidic milk protein available in reasonable purity and quantity. Casein kinases were named for their ability to phosphorylate casein rather than other substrates available at the time. Because of its acidic nature, the use of casein as a substrate presents a number of problems, one of which is the fact that it does not carry a net positive charge at low pH and thus does not bind to P81 phosphocellulose paper. In recent years this problem has been circumvented by using short acidic peptides engineered to bind to P81 paper as substrates for the casein kinases (see Alternate Protocol).

BASIC PROTOCOL 3

There are currently two known mammalian casein kinases, casein kinase I and casein kinase II, named for their order of elution from anion-exchange chromatography columns. Casein kinases show a marked preference for substrates containing acidic residues close to the serine or threonine to be phosphorylated. Casein kinase I is a 37-kDa monomer that phosphorylates serine and threonine residues (see Table 18.7.1) and utilizes ATP only as phosphoryl group donor. The basic phosphorylation consensus sequence is Glu/Ser(P)XXSer (where the underlined residue is the phosphate acceptor, and may be either Ser or Thr). Mammalian casein kinase II is a tetramer of α2β2 structure: the α-subunit is 24 to 28 kDa and the β-subunit is 37 to 44 kDa. Its preferred substrate has aspartate or glutamate residues C-terminal to the phosphorylation site serine or threonine and a consensus of SerGluGlu/Asp. Casein kinase II is unusual in that it can utilize both ATP and GTP as phosphoryl group donor with similar Km values, ∼10 µM for ATP and ∼30 µM for GTP. Mammalian casein kinase II is also inhibited by heparin, whereas casein kinase I is not. The IC50 for the inhibition of casein kinase II by heparin is in the 10 to 20 nM range and at these concentrations casein kinase I is not inhibited. Assays of either casein kinase I or II can be performed using casein as substrate. When isolated from milk, casein is a phosphoprotein. However, it is preferable to use dephosphorylated casein as substrate; this is commercially available in a relatively pure state as either the α or β isoform. Materials 5× casein kinase reaction buffer (see recipe) 10 mg/ml β-casein in H2O [γ-32P]ATP solution (see recipe) Enzyme sample containing casein kinase activity (see Support Protocol 1), kept on ice 30°C water bath Additional reagents and equipment for TCA precipitation (see Support Protocol 2) or electrophoretic analysis (see Support Protocol 4) Analysis of Protein Phosphorylation


Supplement 40

1. For each assay reaction, add the following to a 1.5-ml microcentrifuge tube kept on ice: 4 µl 5× casein kinase reaction buffer 1 µl 10 mg/ml β-casein 1 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 1 to 14 µl H2O. Cap tube and warm the mixture 10 min in a 30°C water bath. This reaction mix can also be used for α-casein. The total reaction mix volume is 20 ìl per reaction. The amount of water required depends on how much enzyme sample is used in step 2. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mixture (minus enzyme) for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.

2. Start the reaction by adding 1 to 14 µl enzyme sample containing casein kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads (UNIT 10.16), prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.

3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Proceed with analysis by one of those methods. ALTERNATE PROTOCOL

ASSAY FOR CASEIN KINASES USING A PEPTIDE SUBSTRATE Casein kinase I can be assayed with the peptide AspAspAspGluGluSerIleThrArgArg. Casein kinase II has most recently been assayed using specific peptide substrates, e.g., ArgArgArgGluGluGluThrGluGluGlu (where the underlined residue is the phosphate acceptor). In both cases the arginine residues allow binding to P81 phosphocellulose paper. The first peptide is relatively specific for casein kinase I, but the kinetics of its phosphorylation are not ideal; the Km is in the millimolar range. Thus, it is preferable to use this assay for casein kinase I only if the kinase is highly purified. Additional Materials (also see Basic Protocol 3) 10 mM synthetic peptide substrate solution (see recipe) for casein kinase: e.g., AspAspAspGluGluSerIleThrArgArg (for casein kinase I) or ArgArgArgGluGluGluThrGluGluGlu (for casein kinase II)


Additional reagents and equipment for TCA precipitation (see Support Protocol 2), adsorption onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic analysis (see Support Protocol 4)



1. For each assay, add the following to a microcentrifuge tube: 10 µl 5× casein kinase reaction buffer 5 µl 10 mM synthetic peptide substrate solution for casein kinase 5 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 0 to 30 µl H2O. Cap the tube and warm 10 min in a 30°C water bath. The total reaction mix volume is 50 ìl per reaction. The amount of water required depends on how much enzyme sample is based. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mix minus enzyme for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.

2. Start the reaction by adding 1 to 30 µl enzyme sample containing casein kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads (UNIT 10.16), prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.

3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Use 10 µl of the reaction mix for adsorption to P81 phosphocellulose paper (see Support Protocol 3). Proceed with analysis by one of those methods. ASSAY FOR Ca2+/CALMODULIN–DEPENDENT KINASES 2+

2+

As the name implies, Ca /calmodulin–dependent kinases are dependent on Ca and calmodulin for their activity. Ca2+/calmodulin–dependent kinase I (CaM kinase I) is a monomeric protein of ∼40 kDa. Ca2+/calmodulin–dependent kinase II (CaM kinase II) is composed of the products of four distinct genes for α, β, γ, and δ subunits. In its native form, CaM kinase II is an oligomer of Mr ∼500,000 to 600,000. As its name suggests, both Ca2+ and calmodulin are required for activity.

BASIC PROTOCOL 4

Little is known of the physiological function of CaM kinase I. Activity can be assayed using the protein synapsin 1 as substrate, but both CaM kinase 1 and CaM kinase II phosphorylate full-length synapsin 1. This problem can be overcome by using synthetic peptide substrates that include the specific phosphorylation site for the enzyme being assayed. CaM kinase I can be assayed using a peptide containing site 1 of synapsin I, TyrLeuArgArgArgLeuSerAspSerAsnPhe (where the underlined residue is the phosphate acceptor). CaM kinase II can be assayed using autocamtide, LysLysAlaLeuArgGlnGluThrValAspAlaLeu (Hanson et al., 1989), which is modeled on the autophosphorylation site of the kinase itself. Analysis of Protein Phosphorylation


Supplement 40

Materials 5× CaM kinase reaction buffer (see recipe) 10 mM synthetic peptide substrate solution (see recipe): e.g., TyrLeuArgArgArgLeuSerAspSerAsnPhe (for CaM kinase I) or LysLysAlaLeuArgGlnGluThrValAspAlaLeu (autocamtide for CaM kinase II) [γ-32P]ATP solution (see recipe) 1 mg/ml calmodulin in Milli-Q-purified water (store small aliquots at −70°C) Enzyme sample containing CaM kinase activity (see Support Protocol 1), kept on ice 30°C water bath Additional reagents and equipment for adsorption onto P81 phosphocellulose paper (see Support Protocol 3) 1. Set up and label duplicate 1.5-ml microcentrifuge tubes. For each reaction, prepare a mixture containing: 5 µl 5× CaM kinase reaction buffer 5 µl 10 mM synthetic peptide substrate solution 5 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 5 µl 1 mg/ml calmodulin or 5 µl H2O 0 to 30 µl H2O. Cap tube and warm the mixture 10 min in a 30°C water bath. The reaction mix is described for 50 ìl per reaction, but it can be carried out in volumes of 25 to 100 ìl by adjusting the amounts of the ingredients proportionally. Control reactions should include one reaction without calmodulin so that any background phosphotransfer reactions can be accounted for and only calcium2+/calmodulin–dependent phosphotransfer is measured.

2. Start each set of reactions sequentially every 30 sec by adding 1 to 25 µl enzyme sample containing CaM kinase activity to one of the tubes. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads (UNIT 10.16), prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mixture to 25 ìl immunoprecipitate bound to beads.

3. Incubate for 10 min in a 30°C water bath. 4. Once the desired incubation time has elapsed, pipet 30 µl of the contents of the tube quickly onto the surface of prepared P81 paper squares (see Support Protocol 3). Only adsorption onto P81 phosphocellulose (see Support Protocol 3) is used to measure CaM kinase–mediated transfer because peptides are not efficiently precipitated by TCA, and they require specialized techniques for analysis by PAGE.




ASSAY FOR TYROSINE KINASES Acid-denatured rabbit muscle enolase was first used as an exogenous tyrosine kinase substrate in the early 1980s, and it is still a useful substrate in a number of experimental situations. Usually enolase is used as a substrate for cytosolic tyrosine kinases such as pp60c-src. Once the reaction is complete, phosphorylation of enolase is detected by separating proteins in the reaction mixture by SDS-PAGE (UNIT 10.2).

BASIC PROTOCOL 5

The following method is optimized for pp60c-src, but may be applied to other tyrosine kinases. It is also useful as a framework for the assay of any tyrosine kinase, but optimization of the immunoprecipitation and assay buffers is required for the tyrosine kinase of interest. Materials Rabbit muscle enolase (enolase EC 4.2.1.11; ammonium sulfate precipitate or suspension purified from rabbit muscle) 1 mM DTT/50 mM HEPES, pH 7.0 Glycerol 100 mM acetic acid 5× tyrosine kinase reaction buffer (see recipe) [γ-32P]ATP solution (see recipe) Enzyme sample containing tyrosine kinase activity (see Support Protocol 1), kept on ice 2× SDS-PAGE sample buffer (UNIT 10.2), ice-cold 30°C and boiling water bath Additional reagents and equipment for immunoprecipitation (UNIT 10.16) or SDS-PAGE (see Support Protocol 4 and UNIT 10.2) 1. Microcentrifuge the equivalent of 100 µg rabbit muscle enolase 5 min at maximum speed, 4°C. Discard the supernatant. 2. Add 10 µl of 1 mM DTT/50 mM HEPES, pH 7.0, to the pellet, mix thoroughly, and incubate 30 to 60 min on ice. 3. Add 10 µl glycerol, mix, and then keep on ice if assays are to be performed the same day. This preparation can also be stored at −70°C until required.

4. Immediately prior to assay, add 20 µl of 100 mM acetic acid to the enolase solution and mix thoroughly. Incubate this mixture 5 min in a 30°C water bath, and then keep on ice until the reaction mixture is prepared. Do not store acid-denatured enolase on ice in this state for >1 hr.

5. For each reaction, add the following to a 1.5-ml microcentrifuge tube: 4 µl 5× tyrosine kinase reaction buffer 1 µl acid-denatured enolase (step 4; 2.5 µg enolase total) 1 µl [γ-32P]ATP (to give 5 µCi/µl and 5 µM ATP final) 0 to 14 µl H2O. Cap the tube and warm the mixture 10 min in a 30°C water bath. The amount of water required depends on the amount of enzyme sample to be added; the total reaction volume should be 20 ìl. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mix minus enzyme for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.



Supplement 40

6. Start the reactions by adding 1 to 14 µl enzyme sample containing tyrosine kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads (UNIT 10.16), prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mixture to 25 ìl immunoprecipitate bound to beads.

7. Incubate the tube 10 min in a 30°C water bath. 8. Stop the reaction by adding 20 µl ice-cold 2× SDS-PAGE sample buffer. Mix thoroughly and heat the tube 3 min in a boiling water bath. Analyze 20 µl of the reaction by SDS-PAGE (UNIT 10.2). Most tyrosine kinases autophosphorylate on a number of tyrosine residues and in in vitro assays are promiscuous with respect to substrate. Backgrounds are often very high so TCA precipitation and adsorption to P81 phosphocellular paper are not very useful for analysis of results. SDS-PAGE (see Support Protocol 4) gives more specific and cleaner results. BASIC PROTOCOL 6

IN-GEL PROTEIN KINASE ASSAYS In-gel protein kinase assays (Kameshita and Fujisawa, 1989) involve the copolymerization of a kinase substrate in the separating gel layer of an SDS-PAGE gel. Protein samples are run on the modified gel as usual (UNIT 10.2). After electrophoresis is complete, the separated proteins are allowed to refold by various gel treatments. The gel is then incubated with [γ-32P]ATP at 30°C for enough time to allow the ATP to penetrate the entire gel. After further washes to remove unincorporated [γ-32P]ATP, the gel is stained and fixed as normal. 32P-containing proteins are revealed by autoradiography (APPENDIX 3A). Although in general these assays are not particularly sensitive, they are sometimes very useful for identifying kinases in complex mixtures. Materials 10 mg/ml kinase substrate: e.g., myelin basic protein Enzyme sample containing kinase activity (see Support Protocol 1), kept on ice 20% (v/v) 2-propanol/50 mM Tris⋅Cl (pH 8.0 at room temperature; APPENDIX 2) 1 mM DTT/50 mM Tris⋅Cl (pH 8.0 at room temperature) 6 M guanidine⋅HCl/1 mM DTT/50 mM Tris⋅Cl (pH 8.0 at room temperature) or 8 M urea/1 mM DTT/50 mM Tris⋅Cl (pH 8.0 at room temperature) 1 mM DTT/0.05% (v/v) Tween 20/50 mM Tris⋅Cl (pH 8.0 at 4°C) Appropriate kinase reaction buffer (see recipes) 10 mM Mg/ATP solution (see recipe) 10 mCi/ml [γ-32P]ATP (3000 Ci/mmol; Amersham, DuPont NEN, or ICN Biomedicals) 5% (w/v) trichloroacetic acid (TCA) 1% (w/v) sodium pyrophosphate/5% (w/v) TCA Container for radioactive incubation: e.g., small tray with tight cover, or heat-sealable polyethylene bag (Seal-a-Meal) Seal-a-Meal apparatus (optional)


Additional reagents and equipment for SDS-PAGE (UNIT 10.2) and autoradiography (APPENDIX 3A)



CAUTION: Because the samples are radioactive, great care should be exercised in sample preparation and loading. Perform the reactions and subsequent manipulations in screwcap microcentrifuge tubes to minimize the risk of 32P contamination. Run gels until the dye front passes out of the gel, as all residual [γ-32P]ATP will migrate with the dye front. This means that the SDS-PAGE running buffer will be contaminated with 32P, and it should be disposed of as radioactive waste according to safety regulations. See APPENDIX 1F for proper handling and disposal. Separate proteins on modified gel 1. Prepare the mixture for a polyacrylamide gel of the appropriate %T (see UNIT 10.2). Add 10 mg/ml kinase substrate to give a final concentration of 1 mg/ml in the gel mixture. Polymerize as usual in a cassette prepared for casting the gel. Myelin basic protein has been used with some success for assays of the MAP kinase family of enzymes. However, any protein is suitable if it is a substrate for the kinase of interest. This assay can also be used to identify potential substrates.

2. Prepare the enzyme samples containing kinase activity for SDS-PAGE in the usual way and carry out electrophoresis (see UNIT 10.2). 3. Once separation is complete, place the gel in 200 ml of 20% 2-propanol/50 mM Tris⋅Cl, pH 8.0. Wash the gel for 20 min, change to fresh buffer, and repeat twice for a total of three 20-min washes. Washing with propanol removes SDS from the gel.

4. Wash the gel with 200 ml of 1 mM DTT/50 mM Tris⋅Cl (pH 8.0) three times, 20 min each. 5. Wash the gel with 250 ml of either 6 M guanidine⋅HCl/1 mM DTT/50 mM Tris⋅Cl (pH 8.0), or 8 M urea/1 mM DTT/50 mM Tris⋅Cl (pH 8.0) twice, 30 min each. Either guanidine⋅HCl or urea can be used to denature the proteins. Depending on the kinase, one or the other may be more successful; guanidine⋅HCl should be tried first, because it seems to work best for most kinases.

6. Wash the gel eight to ten times at 4°C over a period of 18 hr with 250 ml of 1 mM DTT/0.05% Tween 20/50 mM Tris⋅Cl (pH 8.0), to renature proteins. Assay for kinase activity 7. Incubate the gel with 250 ml of an appropriate kinase reaction buffer containing 10 mM MgCl2, for 20 min at 30°C. 8. Remove all traces of buffer and place the gel in a small tray or a Seal-a-Meal bag (the container is to be used for radioactive incubation). Add as small an amount of kinase reaction buffer as possible to just cover the gel. Add 1/4 of the reaction buffer volume of 10 mM Mg/ATP solution to give a final concentration of 50 µM ATP. Add 20 µCi/ml [γ-32P]ATP. 9. Incubate 1 hr at 30°C. The duration of this incubation can be adjusted after viewing the results of the initial experiment.

10. Incubate the gel in 250 ml 5% TCA twice, 15 min each. Wash 10 min in 500 ml of 1% sodium pyrophosphate/5% TCA. Repeat this wash until the solution contains little or no radioactivity. 11. Dry the gel onto filter paper and autoradiograph (APPENDIX 3A).



Supplement 40

SUPPORT PROTOCOL 1

PREPARING A CELL LYSATE FOR KINASE ASSAYS This protocol describes a method for detergent-induced cell lysis to prepare a crude extract containing kinase enzyme activity. Other methods of cell lysis may be appropriate (see UNIT 12.1). Hypotonic lysis and isolation of P100 and S100 fractions can also provide useful data on the recovery of a protein kinase as a soluble or membrane-bound activity. Materials Cultured cells: adherent cells at ∼70% confluence in 100-mm tissue culture dishes or suspension cells at 106 cells/ml PBS (APPENDIX 2), ice-cold Lysis buffer (see recipe) Protease inhibitor stock solutions (see recipe) Microcentrifuge, 4°C 1. Wash the cultured cells twice in ice-cold PBS. Completely aspirate the final wash solution. 2. Add 0.75 ml lysis buffer to ∼2 × 107 cells; for adherent cells check that the monolayer is completely covered. Add protease inhibitors from stock solutions to the appropriate final concentration: 1 mM PMSF, 100 µM benzamidine, 5 µg/ml leupeptin, 5 µg/ml pepstatin A, and 5 µg/ml antipain. A 0.75-ìl lysate should provide sufficient material for immunoprecipitation (use 25% to 50% of the lysate for an initial experiment and scale up or down depending on the results) or simple purification. If the purification procedure involves more than one chromatography step, prepare a larger lysate because only 10% to 20% of the activity is recovered from each chromatography step.

3. Incubate the cells 10 min at 4°C. Scrape the cells from the dish and transfer the lysate to a labeled microcentrifuge tube. 4. Microcentrifuge the lysate 10 min at maximum speed, 4°C. Carefully remove the supernatant and transfer it to a clean microcentrifuge tube. Lysates should be used immediately until proper storage conditions (e.g., −70°C, liquid nitrogen) can determined experimentally. For some kinases, it may not be possible to store the lysate at all. SUPPORT PROTOCOL 2


TCA PRECIPITATION TO DETERMINE INCORPORATION OF RADIOACTIVITY One of the classical methods for separating a reaction product from the reactants is differential precipitation. In the case of protein kinase assays using a protein substrate and [γ-32P]ATP, it is very easy to precipitate the protein and leave the [γ-32P]ATP in the soluble fraction. Most proteins are quantitatively precipitated by trichloroacetic acid (TCA), so TCA can be used to precipitate proteins phosphorylated during a kinase assay. Protein precipitate can be captured by filtration and the filter can be washed with TCA. TCA precipitation is a quick and reproducible way to determine the extent of [32P]phosphate transfer to protein substrates. However, peptides are not precipitated by TCA, so adsorption of labeled peptides to P81 phosphocellulose paper (see Support Protocol 3) is a more suitable method for analyzing the results of kinase assays that use a peptide as substrate. Materials Assay samples (see Basic Protocols 1 to 5 and Alternate Protocol) 5% and 10% (w/v) trichloroacetic acid (TCA), ice-cold 95% ethanol Diethyl ether



Whatman GF-C glass-fiber filters Vacuum manifold (e.g., Fisher) 20-ml scintillation vials Scintillation counter 1. Stop the reaction by adding 20 µl ice-cold 10% TCA. Mix thoroughly and incubate 10 min on ice to precipitate protein. 2. Pipet sample onto Whatman GF-C glass-fiber filter held in a vacuum manifold. Allow the sample to pass through the filter. Rinse the tube with 500 µl ice-cold 5% TCA. Add this wash to the appropriate filter. 3. Wash each filter with 5 ml ice-cold 5% TCA solution four times. Wash once with 10 ml of 95% ethanol, then three times with 10 ml diethyl ether. Allow each filter to dry for a few minutes. CAUTION: Diethyl ether is extremely flammable and washes should be performed in a fume hood away from any flame or heat source. Ether washes should be disposed of in an appropriate manner.

4. Place the dry filters in a 20-ml scintillation vial and count in a scintillation counter by detection of Cerenkov radiation. Alternatively, if dpm quantitation is desired, place dry filter in 10 ml scintillation fluid and count. The filter can also be counted by Cerenkov radiation after the washes with 95% ethanol. If the filters are to be counted in scintillant, however, the diethyl ether washes are essential, because TCA (which is removed by the ether) causes significant chemiluminescence when placed in scintillation fluid, making the counts produced by the scintillation counter meaningless.

ADSORPTION ONTO P81 PHOSPHOCELLULOSE PAPER One of the main methods to separate [32P]-labeled proteins or peptides from [γ-32P]ATP after a protein kinase assay reaction is by adsorbing the protein or peptide to P81 phosphocellulose paper. P81 paper is an ion-exchange matrix with net negative charge at most pHs. At low pH (such as in 75 mM orthophosphoric acid, which is used to wash the paper in this protocol), the excess [γ-32P]ATP left after a kinase assay will not bind. Under the same conditions, however, the phosphorylated peptide will bind to the P81 paper if it carries a net positive charge at low pH. In practice, this is usually achieved by tagging a peptide with two or three arginine or lysine residues at the N- or C-terminus. At the pH of 75 mM orthophosphoric acid the arginyl or lysyl side chains are positively charged and will bind to P81 paper. It is necessary, of course, that the activity of the kinase in question is not affected by the addition of basic residues to the substrate. Usually five or more residues are placed between the hydroxy-amino acid phosphorylation site and the basic residues required for binding to P81 paper.

SUPPORT PROTOCOL 3

When proteins are used as substrates for kinases, the net charge of the protein is also a consideration for adsorption to P81 paper. As for peptides, adsorption to P81 paper is more suitable for basic proteins than for acidic proteins. Under the conditions described in this protocol, histones bind very well to P81 paper because they are highly basic, but casein binds very poorly or not at all because it is a very acidic protein.



Supplement 40

fold here label here in pencil P81 paper

board of nonabsorbent material

Figure 18.7.1 Addition of sample to P81 phosphocellulose paper. The 2 × 2–cm squares of P81 paper are labeled using a pencil (ink is dissolved by the acetone wash), and 50% to 75% of the reaction product is applied to the paper.

Materials Assay samples (see Basic Protocols 1 to 5 and Alternate Protocol) 75 mM orthophosphoric acid (stored up to 6 months at room temperature) Acetone 2 × 2–cm squares of P81 phosphocellulose paper (Whatman) 500-ml plastic beaker with the bottom replaced with solvent-resistant plastic or wire mesh (see Fig. 18.7.2) 20-ml scintillation vial Scintillation counter 1. Cut 2 × 2–cm squares of P81 paper, fold the end over, and label the folded end with pencil as shown in Figure 18.7.1. Place each square on a large piece of nonabsorbent material, such as plastic or acrylic sheeting or an aluminum foil–covered fiberboard. Use a soft pencil to label the squares of P81 paper, as the final acetone wash will remove or destroy ink labels.


2. Pipet 15 or 30 µl of the contents of each assay tube quickly onto the surface of the prepared P81 paper squares. Immediately place the paper squares in a 500-ml plastic beaker with the bottom replaced with solvent-resistant plastic or wire mesh (Fig. 18.7.2). Usually 50% to 75% of the final reaction mixture is spotted onto PB1 paper to ensure that there is sufficient signal to be detected.



Figure 18.7.2 Setup for washing P81 phosphocellulose paper with reaction samples. The paper squares are washed five times for 5 min each in 75 mM orthophosphoric acid and once in acetone. They are then air dried and counted in a scintillation counter.

P81

solventresistant mesh

stir bar

magnetic stirrer

If the plastic mesh used in the beaker is not solvent-resistant, the final acetone washes will dissolve it!

3. Place this beaker into a larger beaker containing 75 mM orthophosphoric acid and incubate 5 min with stirring. 4. Pour the used orthophosphoric acid wash into a container suitable for 32P-containing liquid waste. Refill the large beaker with 75 mM orthophosphoric acid and incubate 5 min more. Repeat for a total of five 5-min washes. 5. Fill the large beaker with acetone and wash the P81 papers for 5 min. Discard the acetone and allow the papers to dry. 6. Place each square of P81 paper in a 20-ml scintillation vial and count by detection of Cerenkov radiation. Alternatively, if the absolute dpm value is desired and a 32P quenching curve is available for the scintillation counter, add scintillation fluid to the vials and count.

ELECTROPHORETIC ANALYSIS OF PHOSPHORYLATION Electrophoretic analysis of phosphorylation allows a more specific determination of protein substrate phosphorylation, because other phosphoproteins are separated by electrophoresis. It is advantageous to use the resolving power of SDS-PAGE to assay relatively crude enzyme samples; this method of analysis may also give useful information on kinase autophosphorylation.

SUPPORT PROTOCOL 4

Stop the reaction (see Basic Protocols 1 to 5 or Alternate Protocol) by adding 20 µl ice-cold 2× SDS-PAGE sample buffer (UNIT 10.2). Mix thoroughly, heat the sample 3 min in a boiling water bath, then analyze by SDS-PAGE (UNIT 10.2). Stain, fix, dry, and autoradiograph (APPENDIX 3A) the gel.



Supplement 40


[γ-32P]ATP solution Dilute 10 mM Mg/ATP solution (see recipe) 1/10 in water to give a 1 mM solution. For the assay, add 10 µl of 1 mM Mg/ATP to 80 µl water and then add 10 µl of 10 mCi/ml [γ-32P]ATP (3.3 pmol ATP). The final ATP concentration is ∼100 ìM (plus 33 nM from the [γ-32P]ATP, which is negligible), and the 100 ìCi (∼2.2 × 108 dpm) in 100 ìl gives a specific activity of 1 ìCi/ìl or ∼2.2 × 107 dpm/nmol. Suppliers for [γ-32P]ATP include Amersham, NEN, and ICN; it is usually available at a variety of specific activities. A good specific activity to use is 3000 Ci/mmol, normally provided by the manufacturer at a concentration of 10 mCi/ml. Amersham’s Redivue is a convenient [γ-32P]ATP solution to use as it is stored at 4°C instead of −20°C. As 1 Ci is ≅ 2.2 × 1012 dpm, then for a specific activity of 3000 Ci/mmol = 3 Ci/ìmol = 6.6 × 1012 dpm/ìmol = 6.6 × 106 dpm/pmol = 2.2 × 106 dpm ≅ 1 ìCi ≅ 0.33 pmol. The stock Mg/ATP solution prepared earlier can be diluted to a working ATP solution of the desired concentration (in this example, 0.1 mM).

CaM kinase reaction buffer, 5× 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 25 mM MgCl2 1 mM CaCl2 Store up to 6 months at −20°C Casein kinase reaction buffer, 5× 100 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2) 25 mM MgCl2 2.5 mM DTT 750 mM KCl Store up to 6 months at −20°C Cyclic nucleotide–dependent protein kinase reaction buffer, 5× 250 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2) 25 mM MgCl2 Store up to 6 months at −20°C Lysis buffer 50 mM HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), pH 7.4 100 mM NaCl 50 mM sodium fluoride 5 mM β-glycerophosphate 2 mM EDTA 2 mM EGTA 1 mM sodium vanadate (Na3VO4) 1% (v/v) Nonidet P-40 (NP-40) or Triton X-100 Store up to 6 months at −20°C Assays of Protein Kinases Using Exogenous Substrates

To stabilize kinase activity, specific kinase inhibitors, protease inhibitors, and reducing agents may be added to the lysis buffer (see Strategic Planning, Enzyme Sources).



Mg/ATP solution, 10 mM Stock solutions: 20 mM MgCl2 or MgSO4 20 mM Na2ATP Working solution: Mix equal volumes of the two stock solutions and measure the pH. Adjust the pH slowly to pH 7.4 with 100 mM HCl or 100 mM NaOH. Store up to 1 year in small aliquots at −20°C. If a precipitate forms, the solution should be stirred continually and the pH maintained at 7.4. Adenosine trisphosphate is normally purchased as a sodium salt. Most kinases require ATP to be complexed with magnesium for its efficient utilization as a substrate. In some cases manganese can substitute for Mg2+.

PKC reaction buffer, 5× Solvent: 100 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2) 25 mM MgCl2 1 mM CaCl2 Store up to 6 months at −20°C Lipids: Just before the buffer is to be used, transfer enough 5 µg/ml phosphatidylserine stock solution in chloroform to provide 100 µg phosphatidylserine/ml buffer (final concentration) to a clean tube. Add sufficient 0.5 µg/ml Diolein stock solution (Sigma) in chloroform to provide 10 µg Diolein/ml buffer (final concentration) to the same tube and mix. Dry the lipids under a stream of nitrogen. Working solution: Add the required volume solvent (see above) and place on ice for 10 min. Sonicate the mixture at medium power for 1 min while the sample is on ice. Let the sample sit on ice 2 min. Repeat sonication 4 more times for a total of 5 min. Discard any unused buffer. Diolein is the old name for various preparations that contain mixed isomers of diacylglycerol, the cofactor for PKC. Diolein is a suitable source for the cofactor in these assays, and it is less expensive than pure preparations of 1,2-diacylglycerol.

Protease inhibitor stock solutions 100 mM phenylmethylsulfonyl fluoride (PMSF) in 100% ethanol 100 mM benzamidine 1 mg/ml leupeptin 1 mg/ml pepstatin A 1 mg/ml antipain Store up to 6 months at −20°C Synthetic peptide substrate solution Prepare 10 mM solutions of each peptide (highest quality available) in Milli-Q water and adjust pH to 7.4 before storage at −70°C or use. If the peptide is insoluble, check the sequence—a peptide with a high proportion of acidic residues may be more soluble at alkaline pH, and one with a high proportion of basic residues may be more soluble at acidic pH. Hydrophobic peptides may be solubilized by adding up to 20% (v/v) acetonitrile. Whatever is added must not compromise the assay conditions: e.g., if a peptide must be dissolved at pH 10, adding the solution to the reaction mix must not change the pH of the reaction. Most synthetic peptides are quite stable when stored at −70°C; the stability of individual peptide substrates should be determined for the storage conditions used, however. If peptides are custom synthesized they should ideally be purified by HPLC before use.



Supplement 40

Tyrosine kinase reaction buffer, 5× 100 mM HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), pH 7.4 5 mM MnCl2 5 mM DTT 500 µM sodium vanadate (Na3VO4) Store up to 6 months at −20°C COMMENTARY Background Information


In recent years there has been an explosion of interest in protein kinases. It is now obvious that this class of enzyme plays many essential roles in a large array of biological processes, including the cellular responses to hormones, control of most stages of the cell cycle, and development of the nervous system in systems as diverse as fruit fly and man. Because of this interest, a very large number of protein kinase and protein phosphatase genes have been identified; current estimates are that there are well over 2000 genes encoding these enzymes in mammalian genomes. Obviously the study of protein kinases and phosphatases can provide fundamental information regarding the regulation of many diverse systems; the protocols provided in this unit include enough information to enable researchers new to this type of experiment to embark on the study of protein kinases. Protein kinases can be assayed using two types of substrate, intact proteins or synthetic peptides, and there are advantages and disadvantages to the use of each. A number of intact proteins may be used as general protein kinase substrates, and those in common use are available in relatively pure form from a number of manufacturers at reasonable prices. The main disadvantage to the use of proteins as substrates is the lack of specificity, sometimes making interpretation of experimental data difficult. Intact proteins such as myelin basic protein (MBP) contain many different phosphorylation sites. MBP can be phosphorylated by MAP kinase, some protein kinase C isoforms, and Ca2+/calmodulin kinase II, each at one or more sites. Hence, it is often impossible to differentiate which kinase is responsible for phosphorylating MBP in a complex mixture like a whole cell lysate. Some proteins cause headaches because they cannot be used with simple experimental protocols, e.g., casein. The use of synthetic peptides improves the specificity of the reaction by providing a single, well-defined phosphorylation site. They also allow experiments to define the concensus se-

quence for phosphorylation by a particular protein kinase and make it possible to design a substrate that has excellent kinetic properties for the kinase of interest. Peptides are often easier to use than their protein counterparts as in the case of casein kinases, where peptides can be designed that allow the use of phosphocellulose paper rather than other more involved methods for assessment of experimental results. Use of peptides is not without disadvantages though, the most significant being the cost of producing reasonable quantities of large numbers of peptides. When this is coupled with the observation that sometimes peptides are not recognized by the kinase because some structural element is missing, using peptide substrates can be a frustrating and expensive business. However, this is the exception rather than the rule, and the use of specific peptides to assay protein kinases is ultimately the best choice. One unit of kinase activity is usually defined as the amount of enzyme required to transfer 1 µmol of phosphate to a substrate from ATP per minute. From the following example it is possible to calculate a number of the reaction parameters: enzyme + [γ-32P]ATP + substrate → enzyme + ADP + [32P]product For this example, 1 mg protein (based on protein assay; see UNIT 10.1A) was used for the assay. The [γ-32P]ATP in the assay mixture had a specific activity of 106 dpm/nmol. After the 10-min reaction the 32P-labeled product had a specific activity of 105 dpm; therefore, there were (105/106) nmol of 32P-labeled product, and the concentration was 10−1 nmol or 100 pmol. Hence, the enzyme transferred 10 pmol of phosphate per minute, and represents 0.01 U of enzyme activity when measured with this substrate. Put another way, this represents an enzyme specific activity of 10 pmol/min/mg of protein. This latter expression of enzyme activity is a useful measure when comparing different preparations of a kinase or, during purification of a kinase, as a measure of the extent of purification through different steps.



Critical Parameters Although it may seem obvious, the most critical element in performing kinase assays is using the correct controls. This is especially true when performing assays on tissue samples or cell extracts. Controls should always include addition of no substrate, addition of no enzyme source, addition of heat-denatured enzyme source, and, for reactions that require activator or cofactor, addition of any activator or cofactor and addition of no activator or cofactor. Another important aspect is the concentration of ATP in the assay. The ATP concentration can vary depending on the type of experiment to be performed. For derivation of kinetic data, the concentrations of both ATP and peptide or protein substrate are critical. If kinase detection during purification is required, assay linearity is not as important as long as the right controls are present. For mapping in vitro phosphorylation sites on a kinase substrate, very often the ATP concentration is kept extremely low but the specific activity very high. When designing potential peptide substrates for individual protein kinases, there are a number of considerations to remember. For example, phosphorylated peptide will bind to the P81 phosphocellulose paper. To be able to analyze the assay with this paper, it is necessary that the peptide carry a net positive charge at low pH. In practice this is usually achieved by tagging a peptide with two or three arginine or lysine residues at the N- or C-terminus. At the pH of 75 mM orthophosphoric acid, the arginyl or lysyl side chains are positively charged and will bind to the P81 paper. It is important to determine that the activity of the kinase of interest is relatively unaffected by the addition of basic residues to its peptide substrates in this way. To help ensure this is the case, it is a good idea to place five or more residues between the hydroxy-amino acid to be phosphorylated and the basic residues required for binding to P81 paper. Net charge considerations also pertain to some extent to proteins used as substrates for protein kinases. Under the conditions described in Support Protocol 3, histones will bind very well to P81 paper, because they are highly basic, but casein, which is very acidic, binds very poorly or not at all. In these cases, the P81 paper assay is very good when histones are used to assay kinase activity, but worthless when casein is used. When crude enzyme sources, such as cell lysates, are analyzed, many other proteins are present and phosphorylation of multiple proteins in the crude mixture is possible, as is

autophosphorylation of the protein kinase. Because many proteins will be precipitated by trichloroacetic acid (TCA) or will bind to P81 paper, there can be elevated backgrounds and severe overestimates of peptide phosphorylation when Support Protocols 2 and 3 are used. When performing kinetic studies, this overestimation of phosphorylation can be very misleading, as substrates other than the peptide under study will undoubtedly be phosphorylated and with different kinetics.

Troubleshooting Any troubleshooting of kinase assay procedures should be very straightforward. If the assay does not work, verify that both the [γ-32P]ATP and the peptide or protein substrate have been added at the correct concentrations. It is also very important to keep the enzyme source on ice before use and to store it appropriately between experiments. Very often when this type of experiment does not work it is because of a labile enzyme, poor storage of the enzyme, or a combination. It is important to know how long a particular enzyme is stable under the conditions used and the rate of inactivation or denaturation, if relevant. It is also essential to know if the enzyme activity is comparable between different preparations, especially for purified proteins.

Anticipated Results Depending on the requirements, information can be obtained about the specific activity of a particular enzyme preparation, kinetic parameters for the reaction catalyzed, or whether a particular kinase has been identified or substantially purified.

Time Considerations Kinase assays are quite simple to set up and perform. If the samples for assay are available, it is a matter of only ∼3 to 4 hr to go from assay set up to data interpretation when either TCA precipitation (Support Protocol 2) or adsorption to phosphocellulose paper (Support Protocol 3) is used to separate phosphorylated protein from contaminating [γ-32P]ATP. With SDS-PAGE analysis (Support Protocol 4), the assays can be set up, performed, and the samples run on a SDS-PAGE gel in ∼4 to 5 hr. Gel drying and autoradiography can take from 2 hr to a number of days depending on the level of protein phosphorylation. In general, if individual protein bands have ∼20,000 cpm of 32P associated with them, they will be visible with 1 hr of autoradiography.



Supplement 40

Literature Cited Hanson, P.I., Kapiloff, M.S., Lou, L.L., Rosenfeld, M.G., and Shulman, H. 1989. Expression of a multifunctional Ca2+/calmodulin–dependent protein kinase and mutational analysis of its autoregulation. Neuron 3:59-70. Kameshita, I. and Fujisawa, H. 1989. A sensitive method for detection of calmodulin-dependent protein kinase II activity in sodium dodecyl sulfate polyacrylamide gel. Anal. Biochem. 183:139-143. Pearson, R.B., and Kemp, B.E. 1991. Protein kinase phosphorylation site sequences and consensus specificity motifs. Methods Enzymol. 200:62-81. Songyang, Z., Carraway, K.L. III, Eck, M.J., Harrison, S.C., Feldman, R.A., Mohammadi, M.,

Schlessinger, J., Hubbard, S.R., Smith, D.P., Eng, C., Lorenzo, M.J., Ponder, B.A.J., Mayer, B.J., and Cantley, L.C. 1995. Catalytic specificity of protein tyrosine kinases is critical for selective signalling. Nature 373:536-539. Takai, Y., Kishimoto, A., Kikkawa, U., Mori, T., and Nishizuka, Y. 1979. Unsaturated diacylglycerol as a possible messenger for the activation of calcium-activated, phospholipid-dependent protein kinase system. Biochem. Biophys. Res. Commun. 91:1218-1224.

Contributed by A. Nigel Carter The Salk Institute for Biological Studies La Jolla, California




Permeabilization Strategies to Study Protein Phosphorylation

UNIT 18.8

This unit deals with the use of nucleotide triphosphates to label proteins in vitro in permeabilized cells and isolated cellular fractions. These experiments generally utilize [γ-32P]ATP as an exogenously added phosphate donor, although [γ-32P]GTP can be used in specific cases. The method is very straightforward, although numerous considerations must be made before applying it to each new system. describes labeling of phosphoproteins in intact cells with 32P-labeled inorganic phosphate (32Pi). After ascertaining that the protein of interest is a phosphoprotein using the protocols detailed in that unit, the phosphorylation event itself may be studied either in cell-free systems or in a more intact cellular system. In vitro, most protein kinases are capable of phosphorylating many substrates in addition to those phosphorylated in vivo, thus making it desirable to identify and study a protein kinase–catalyzed phosphorylation reaction in a more intact system than a cell lysate or homogenate. Permeabilization of intact cells provides a powerful experimental approach for accomplishing this. Brief permeabilization of cells is possible using a variety of agents. This procedure gives access to the intracellular milieu but still allows signal transduction events to be initiated by hormone stimulation of the cells via cell-surface receptors. UNIT 18.2

Brief chemical permeabilization of cells allows the introduction of cell-impermeable reagents into the intracellular compartment in order to study particular signal-transduction processes. In this type of experiment, relatively large molecules such as the Fab′ fragments of specific antibodies may be introduced into the permeabilized cell, although the kinetics of entry may be too slow for the study of some processes. This technique is often more useful when introducing relatively small molecules into the permeabilized cell, for example non-cell-permeable activators (e.g., hydrophilic or lipophobic small molecules that cannot partition into the membrane) or inhibitors of protein kinases or phosphatases (see, e.g., Calbiochem catalog). The kinetics of entry into permeabilized cells for small molecules such as [γ-32P]ATP are fast, and the system equilibrates the cellular ATP and [γ-32P]ATP with intracellular processes within 2 to 3 min. Figure 18.8.1 shows the 40,000

20,000

32

P (cpm)

30,000

10,000 A 0 0

5

10

15 20 Time (min)

25

30

Figure 18.8.1 Equilibration of the monoester phosphates of the lipid phosphatidylinositol-4,5bisphosphate with the ATP pool after permeabilization of human platelets. The letter A marks the point of addition of [γ-32P]ATP. Contributed by A. Nigel Carter Current Protocols in Molecular Biology (1997) 18.8.1-18.8.19 Copyright © 1997 by John Wiley & Sons, Inc.



equilibration of the monoester phosphates of the lipid phosphatidylinositol-4,5-bisphosphate with the ATP pool after permeabilization of human platelets. As these monoester phosphates are known to equilibrate with the ATP pool extremely quickly, this experiment provides a good indication of the speed of equilibration of cellular ATP pools with intracellular processes after addition of [γ-32P]ATP to permeabilized cells. Procedures are outlined for performing a protein phosphorylation experiment using permeabilized cells (see Basic Protocol 1) and isolated intracellular organelles (see Basic Protocol 2). Both of these procedures result in lysates from which the protein of interest may be easily immunoprecipitated; however alternative techniques are described for preparing the final lysate from either Basic Protocol 1 or Basic Protocol 2 for electrophoretic analysis (see Alternate Protocols 1, 2, 3, and 4). A related procedure that does not involve permeabilization is outlined for direct analysis of cytosolic or membranebound kinases (see Alternate Protocol 5). Two different methods for determining the specific radioactivity of 32P-containing compounds are also included (see Support Protocols 1 and 2). CAUTION: It cannot be emphasized enough that each investigator should be fully familiarized with their local regulations regarding the safe handling and disposal of 32P and also have the appropriate equipment for dealing with the containment of 32P. See APPENDIX 1F for more information on the safe use of radioisotopes. NOTE: Milli-Q purified water or its equivalent should be used to prepare all solutions used throughout this unit. BASIC PROTOCOL 1

Permeabilizatin Strategies to Study Protein Phosphorylation

ANALYSIS OF PROTEIN PHOSPHORYLATION IN PERMEABILIZED CELLS When setting out to study a protein phosphorylation event via a permeabilization protocol, the general procedure is as follows. First, cells are incubated in a buffer which has been designed to mimic the composition of the intracellular compartment and which contains a permeabilizing agent. The preferred permeabilization agent of choice is streptolysin O, as this agent allows many signal-transduction systems to remain intact for a number of minutes after its addition (Stephens et al., 1994). This is certainly not true of all permeabilization agents or protocols; if another agent (e.g., saponin) is to be tried, the methods detailed below should be used as a general guideline only. Depending on the experiment to be performed, the permeabilization buffer may also contain [γ32-P]ATP and the particular reagents under study. The [32γ-P]ATP provides the “hot” phosphate, and will equilibrate with the intracellular processes utilizing ATP within 2 to 3 min (A.N. Carter, unpub. observ.). After a brief time the experimental reagents, such as inhibitors, can be introduced into the permeabilization medium and hence into the inside of the cell. At this point the cells may be stimulated with the appropriate agent in the same way as for intact cell preparations. The phosphorylation of the protein under study is initially investigated by immunoprecipitation as described in UNIT 10.16 and UNIT 18.2, followed by phosphopeptide mapping as described in UNIT 18.3. Materials Cell culture to be labeled Cell culture medium appropriate for cells being studied, buffered with 20 mM HEPES, pH 7.2, prewarmed to 37°C Cells under study (normally cultured cell lines or isolated primary cells in culture) Extracellular buffer (see recipe), 37°C Permeabilization buffer (see recipe), 37°C 60 U/ml streptolysin O working solution (see recipe), freshly prepared 10 mM cold ATP stock solution (see recipe) 10 mCi/ml [γ-32P]ATP (3000 Ci/mmol; e.g., DuPont NEN)



Pharmacological agents, peptides, or Fab′ fragments to be studied Appropriate receptor agonists 2× lysis buffer for immunoprecipitation (see recipe), ice-cold Protease inhibitor stock solutions (in absolute ethanol; store up to 6 months at −20°C): 100 mM phenylmethylsulfonyl fluoride (PMSF) 100 mM benzamidine 1 mg/ml pepstatin A 1 mg/ml leupeptin 1 mg/ml antipain 100 mM DTT Dry ice/ethanol bath Noncirculating 37°C water bath containing a flat shelf (preferably made of wire mesh) with space for the number of cell plates in the experiment 50-ml centrifuge tubes Tabletop centrifuge Cell scrapers Screw-cap microcentrifuge tubes Permeabilize cells For monolayer cultures 1a. Approximately 1 hr before starting the experiment, change the medium in the cell culture plates to be labeled to equivalent medium containing 20 mM HEPES. Return cells to incubator and allow the cells to equilibrate with the new medium before use. The number of cells used for any experiment of this type is totally dependent on how easy it is to detect the end product being studied. In some cases, a 60-mm dish of cells that has just reached confluency, with ∼1 × 106 cells, is adequate; in others a 100-mm dish of cells with ∼1 × 107 cells is required. It is advisable to start with ∼1 × 106 cells and see if the signal is detectable. Use of a 60-mm dish allows a smaller volume of permeabilization medium (normally of the order of 0.5 to 2 ml) to be used. Standard cell culture medium is buffered by bicarbonate ions in the medium and the CO2 in the atmosphere of the incubator. Outside the incubator this buffering system is ineffective and the pH will quickly rise if the culture is left in the normal atmosphere. The pH of HEPES buffers are minimally affected by temperature (unlike Tris buffers), hence they provide good pH control for cell culture medium. One drawback is that HEPES may be toxic to some cells above a concentration of ∼20 mM, so it is important to check possible cellular toxicity before the use of HEPES-buffered culture media.

2a. Place the required number of plates containing cells in the 37°C water bath and allow 10 min for the medium to reequilibrate. The cells are stable in this state for an hour or so, depending on the rate of evaporation of the culture medium.

3a. Aspirate the culture medium from each plate and quickly rinse the cells twice, each time with 10 ml of 37°C extracellular buffer. Aspirate the extracellular buffer, then add 0.5 to 2 ml permeabilization buffer to each plate. Be careful when rinsing, as some cell lines are less adherent than others. Usually, most epithelial or mesenchymal lines adhere well and will not slough off the dish unless extreme force is used. Transformed cells usually adhere poorly and can easily be rinsed off the dish if care is not taken.

4a. Add streptolysin O (from the 60 U/ml working solution) to each plate to a final concentration of 0.6 U/ml. Add cold ATP (from the 10 mM stock solution) to a final concentration of 100 µM.



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For suspension cultures 1b. Approximately 1 hr before starting the experiment, transfer cells to sterile 50-ml centrifuge tubes and centrifuge 5 min at 1000 × g, room temperature. Remove the supernatant, then rinse cells twice, each time by adding ∼40 ml of 37°C extracellular buffer, centrifuging 5 min at 1000 × g, room temperature, and removing the supernatant. The cell suspension may be split into aliquots, if desired, after the addition of the final rinse of extracellular medium. To do this, add the final (40-ml) rinse of extracellular medium, resuspend cells gently, and dispense aliquots into fresh tubes. Centrifuge the aliquots and remove the supernatants to complete the final rinse before proceeding to step 2b.

2b. Resuspend the cells in 37°C extracellular buffer at a density of 1 × 107 cells/ml. Allow the cells to equilibrate 15 to 30 min at 37°C with gentle swirling. This step may have to be modified if a cell is used that could be activated by this procedure, (e.g., platelets). If a sensitive cell is used, equilibration time may be kept down to 5 to 10 min and the agitation may be omitted.

3b. Centrifuge cells again for 5 min at 1000 × g, room temperature, and aspirate the buffer. 4b. Add streptolysin O (from the 60 U/ml working solution) to fresh 37°C permeabilization buffer to a final concentration of 0.6 U/ml. Add cold ATP (from the 10 mM stock solution) to a final concentration of 100 µM. Add this permeabilization solution containing streptolysin O and ATP to the cells and mix gently but thoroughly. Addition of cold ATP at the same time as streptolysin O allows cellular ATP levels to be maintained and will therefore preserve cell viability. One important consideration is whether the cells under study possess cell-surface receptors for ATP. The ATP acts as a full agonist for some types of purinergic receptors, and this can obviously cause confusion when analyzing signal-transduction mechanisms. Prior to performing permeabilization studies, one should first determine, using intact cells, whether extracellular ATP activates signal-transduction mechanisms that interfere with the system under investigation. Stephens et al. (1994) have developed a way of inactivating extracellular receptors for ATP in neutrophils prior to permeabilization. The utility of this method for other cell types is not known to date, although it may be used as a guideline if required. Readers are referred to the original paper for full discussions of this method.

Permeabilize and label cells 5. Permeabilize cells by incubating 1 to 5 min at 37°C. The time of incubation depends very much on the particular system under study and the cell type, and must be determined empirically. Permeabilization for >5 min usually reduces hormone responsiveness of cells dramatically. This is probably due to a number of processes, including the loss of cytosolic components into the extracellular pearmeabilization buffer and physical uncoupling of receptors from signal-transduction processes.

6. Add 10 mCi/ml [γ-32P]ATP to a final concentration of 50 to 500 µCi/ml. [γ-32P]ATP is usually purchased at a specific activity of ∼3000 Ci/mmol, and a concentration of 10 mCi/ml. Hence 5 ìl of the stock is 50 ìCi [γ-32P]ATP. The label may be used at this concentration. If the stock [γ-32P]ATP is to be diluted into another solution it is better to do this directly before use. The criterion for the amount of radiolabeled ATP to add (between 50 and 500 ìCi/ml) is the detection limit. Initial experiments must be performed to see how much label must be added. Permeabilizatin Strategies to Study Protein Phosphorylation

The alternative to this separate labeling step is to add the [γ-32P]ATP at the same time as the streptolysin O and cold ATP in step 4a or b. The type of experiment to be performed will determine at which point the [γ-32P]ATP is added to the cells.



7. Add the pharmacological agents, peptides, or Fab′ fragments under study and allow time for their action as required. Many pharmacological agents amenable to testing by this protocol are listed in the Calbiochem catalog. For example, a peptide containing residues 280 to 305 of the CaM kinase II α subunit can be a potent inhibitor of CaM kinase II and is non-cell-permeable. There are other peptides modeled on pseudosubstrate inhibitors of a variety of protein kinases that can be used as specific inhibitors, all of which are non-cell-permeable. Any Fab′ fragment may be tested by this protocol as long as it has an effect on enzyme activity, either positive or negative. Similarly, if an antibody stops an enzyme from reaching its site of action, it may also be tested via this procedure. The time for a particular agent to act as required should be determined experimentally as part of the initial experimental protocols. As with the [γ-32P]ATP addition, these agents may also be added at the same time as streptolysin O and cold ATP.

8. Add receptor agonists as required. The same considerations are true for receptor agonist additions as for inhibitor and [γ-32P]ATP additions to the cells. One must perform preliminary experiments to empirically determine the timings for these additions for each cell type under study. The remaining steps of this protocol are to prepare the cells for analysis by immunoprecipitation. If electrophoretic analysis is to be performed, see Alternate Protocols 1 and 2.

Prepare cell lysates for immunoprecipitation 9a. For monolayer cultures: Stop the reactions by rapidly aspirating the permeabilization buffer and adding 0.75 ml ice-cold 2× lysis buffer for immunoprecipitation to each plate. Add protease inhibitors and DTT (as stock solutions) to the following final concentrations: PMSF to 1 mM final Benzamidine to 1 mM Pepstatin A to 5 µg/ml final Leupeptin to 5 µg/ml final Antipain to 5 µg/ml final DTT to 1 mM final. Place plates on ice for 10 min, then scrape the cell debris from the bottoms of the plates into the buffer using a separate cell scraper for each plate. 9b. For suspension cultures: Stop the reactions by adding an equal volume of ice-cold 2× lysis buffer for immunoprecipitation. Add protease inhibitors and DTT to the final concentrations in step 9a, then place the tubes of cells on ice for 10 min. 10. Transfer lysates to screw-cap microcentrifuge tubes (one plate or cell aliquot/tube). Microcentrifuge lysates at maximum speed, 4°C, then transfer the supernatants to fresh screw-cap microcentrifuge tubes and freeze immediately in a dry ice/ethanol bath. Store at −70°C until needed. The lysates are now ready for immunoprecipitation (UNIT 10.16). CAUTION: The permeabilization buffer in the latter steps of this protocol contains [γ-32P]ATP, and great care should be taken to remove the medium safely and dispose of it correctly as liquid radioactive waste. The lysates and cell scrapers are also very radioactive at this point. Screw-cap tubes should be used at all subsequent stages of processing the cell lysates as these provide the best containment of the 32P-containing samples. The same microcentrifuge should be used for all centrifugation steps; after use it will undoubtedly be contaminated with 32P. It can then be set aside for future use with 32P-containing samples with the appropriate safety precautions.



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INTACT CELL SAMPLE PREPARATION FOR ELECTROPHORETIC ANALYSIS OF PROTEIN PHOSPHORYLATION The preceding protocol (see Basic Protocol 1) is set up to allow immunoprecipitation of specific proteins after the permeabilization experiment. Two alternatives to this approach, described below, involve using the separating power of polyacrylamide gel electrophoresis (PAGE) to fractionate proteins, instead of immunoprecipitation. Fractionation can be either by one-dimensional SDS-PAGE (UNIT 10.2), or by two-dimensional PAGE in which isoelectric focusing is performed as the first dimension, followed by SDS-PAGE (UNIT 10.3). In each case, sample loading must be optimized so that best resolution of the protein of interest is achieved. These protocols are written for both monolayer cells and suspension cells. To perform the procedure with suspension cells, the cells must be centrifuged down as quickly as possible after reacting with the appropriate pharmacological agents/receptor agonists, then treated as described for monolayer cultures. Obviously, the timing of the reactions is less precise this way, but experiments can be done successfully and reproducibly if reaction and centrifugation times are kept exactly the same. CAUTION: Samples for SDS-PAGE or isoelectric focusing will still contain a great deal 32 P at the latter stages of the protocol. Great care should be exercised in sample preparation. In particular do not use a probe sonicator to shear DNA, as 32P-containing aerosols will be produced. In the case of SDS-PAGE, the gels should be run until the dye front passes out of the gel. This means that the SDS-PAGE running buffer will be also be contaminated with 32P. In the case of isoelectric focusing the 32P will focus out of the gel during electrophoresis; hence the running buffer will be contaminated with 32P. These solutions should be disposed of as radioactive waste as local regulations dictate. ALTERNATE PROTOCOL 1

Intact Cell Sample Preparation for SDS-PAGE Additional Materials (also see Basic Protocol 1) 2× SDS-PAGE sample buffer (see recipe), 4°C Bath sonicator Boiling water bath 1. Prepare cells and perform permeabilization and labeling (see Basic Protocol 1, steps 1 through 8). 2a. For monolayer cultures: Stop the reactions by rapidly aspirating the permeabilization buffer and adding 100 to 250 µl of 4°C 2× SDS-PAGE sample buffer to each plate. Quickly scrape the cells into the buffer using a separate cell scraper for each plate and quantitatively transfer the sample to a screw-cap microcentrifuge tube. 2b. For suspension cultures: Microcentrifuge the reaction tubes 1 min at maximum speed, room temperature. Quickly aspirate the supernatants, taking care not to disturb the cell pellets. Add 100 to 250 µl of 2× SDS-PAGE sample buffer to each tube and vortex thoroughly. When performing this step for suspension cultures it is very important to use exactly the same times for centrifugations if reproducible data are to be obtained.

3. If samples are very viscous as a result of DNA being released from cell nuclei, shear the DNA by placing the capped sample tubes in a bath sonicator filled with ice water and sonicating 5 min. Permeabilizatin Strategies to Study Protein Phosphorylation

4. Boil samples 3 min in a boiling water bath. To perform the analysis, carefully apply up to 50 ìl of the samples to the sample wells of a precast SDS-PAGE gel. Proceed as described in UNIT 10.2.



Intact Cell Sample Preparation for Isoelectric Focusing


Additional Materials (also see Basic Protocol 1) Two-dimensional-PAGE lysis buffer (see recipe), 4°C Two-dimensional-PAGE sample buffer (see recipe) Bath sonicator Dry ice/ethanol bath Lyophilizer 1. Prepare cells and perform permeabilization and labeling (see Basic Protocol 1, steps 1 through 8). 2a. For monolayer cultures: Stop the reactions by rapidly aspirating the permeabilization buffer and adding 100 to 250 µl of 4°C two-dimensional-PAGE lysis buffer. Scrape the cells into the buffer using a separate cell scraper for each plate and transfer the sample to a screw-cap microcentrifuge tube. 2b. For suspension cultures: Microcentrifuge the reaction tubes 1 min at maximum speed, room temperature. Quickly aspirate the supernatants, taking care not to disturb the cell pellets. Add 100 to 250 µl of two-dimensional PAGE lysis buffer to each tube and vortex thoroughly. When performing this step for suspension cultures it is very important to use exactly the same times for centrifugations if reproducible data are to be obtained.

3. If samples are very viscous as a result of DNA being released from cell nuclei, shear the DNA by placing the samples in a bath sonicator filled with ice-water and sonicating 5 min. Freeze samples in a dry ice/ethanol bath, then lyophilize. 4. Redissolve the lyophilized samples in 50 to 100 ml of two-dimensional-PAGE sample buffer. Warm the samples to 37°C for 3 to 5 min. To perform the analysis, apply 10 to 20 ìl of each sample to a prepared isoelectric focusing gel of the required pH gradient (see UNIT 10.3). Samples for isoelectric focusing must have a low salt concentration because a high salt concentration often causes band broadening during isolectric focusing.

ANALYSIS OF PROTEIN PHOSPHORYLATION USING ISOLATED SUBCELLULAR FRACTIONS

BASIC PROTOCOL 2

When the protein under study exists in a particular organelle and is phosphorylated in intact cells, one consideration when studying the phosphorylation event in vitro in isolated organelles is whether the protein is accessible to exogenously added kinases or phosphatases, or whether it is accessible to endogenous kinases or phosphatases. For instance, if it is a protein of the endoplasmic reticulum, is its location transmembrane or intraluminal? If transmembrane, is it normally phosphorylated on the cytosolic side or luminal side of the membrane? Permeabilization protocols can also be used with intact intracellular organelles as well as intact cells. This is obviously an important consideration when dealing with proteins found inside the organelle in question. Another consideration here is whether the organelle can be isolated in an intact condition, or whether its contents will be partially or completely lost during purification. In the case of mitochondria this does not pose a problem, but for more fragile organelles such as the Golgi apparatus this may be more problematic. The study of protein phosphorylation in isolated organelles (see Castle, 1995 for isolation procedures) can be a straightforward extension of the permeabilization protocols de-



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scribed above (see Basic Protocol 1 and Alternate Protocols 1 and 2). The isolated organelle is treated in the same way as one would treat cultured cells in suspension. However, as there is little or no requirement for the maintenance of signal-transduction mechanisms in isolated organelles, the permeabilization procedure can be more prolonged than for intact cells. This allows the introduction of larger molecules such as purified protein kinases into the organelles under study. Materials Suspension of isolated intracellular organelles Intracellular buffer (see recipe), 4°C Permeabilization buffer (see recipe), 37°C 60 U/ml streptolysin O working solution (see recipe) 10 mM cold ATP stock solution (see recipe) 10 mCi/ml [γ-32P]ATP (3000 Ci/mmol; e.g., DuPont NEN) Reagents under study: e.g., kinase/phosphatase inhibitors, protein kinases, or phosphatases 2× lysis buffer for immunoprecipitation (see recipe), 4°C Screw-cap microcentrifuge tubes Benchtop ultracentrifuge (e.g., Beckman TL-100 or Airfuge) 37°C circulating water bath 1. Wash the organelle preparation three times with 4°C intracellular buffer, each time by ultracentrifuging 10 min at ≥100,000 × g. Transfer the equivalent of 1 mg of protein to individual screw-cap microcentrifuge tubes and keep on ice. The best way to wash the organelle preparations is to use a Beckman TL-100 benchtop ultracentrifuge or the older Beckman Airfuge. Both of these instruments will produce g-forces >100,000 × g. Because of the difficulty in centrifuging some organelles (e.g., plasma membrane preparations) it is essential to use ultracentrifugation for all organelles except nuclei, which can be recovered by microcentrifuging 30 sec at maximum speed. See Castle (1995) for details of organelle isolation.

2. Ultracentrifuge the organelle preparation 10 min at ≥100,000 × g, 4°C, and aspirate the supernatant. Add 200 ml 37°C permeabilization buffer, mix gently, and transfer to the 37°C circulating water bath. Work quickly at this point to minimize any proteolytic and other unwanted side reactions.

3. Add streptolysin O (from the 60 U/ml working solution) to each tube to a final concentration of 0.6 U/ml. Add cold ATP (from the 10 mM stock solution) to a final concentration of 100 µM. Exactly the same considerations apply when performing permeabilization of organelles as when performing permeabilization of intact cells (see Basic Protocol 1). However, because there is no intact signal-transduction machinery in isolated organelle preparations, the range of possible experiments is fewer, so the system is usually more robust than for intact-cell experiments.

4. Add 10 mCi/ml [γ-32P]ATP to a final concentration of 50 to 500 µCi/ml. The criterion for the amount of radiolabeled ATP to add (between 50 and 500 ìCi/ml) is the detection limit. Initial experiments must be performed to see how much label must be added.


5. Add the reagents under study—e.g., kinase/phosphatase inhibitors, protein kinases, or phosphatases.



6. Add 200 ml of 4°C 2× lysis buffer to each tube to lyse the organelles. Mix well and place on ice for 5 min. 7. Microcentrifuge the samples 10 min at maximum speed, 4°C. Store lysates at −70°C. The lysates are now ready for immunoprecipitation (UNIT 10.16).

ORGANELLE SAMPLE PREPARATION FOR ELECTROPHORETIC ANALYSIS OF PROTEIN PHOSPHORYLATION Exactly the same considerations apply to the electrophoretic analysis of protein phosphorylation in isolated organelles as apply to permeabilized cells (see Alternate Protocols 1 and 2). Organelle preparations must first be ultracentrifuged and the supernatant removed before adding the appropriate buffer for electrophoresis. For this, the use of a Beckman TL-100 benchtop ultracentrifuge, which can provide g-forces >300,000 × g, is recommended. Complete removal of the supernatant is especially important if two-dimensionalPAGE (Alternate Protocol 4) is used, as the salt concentration must be kept low. CAUTION: Samples for SDS-PAGE or isoelectric focusing will contain a great deal 32P at the latter stages of the protocol. Great care should be exercised in sample preparation. In the case of SDS-PAGE, the gels should be run until the dye front passes out of the gel. This means that the SDS-PAGE running buffer will be also be contaminated with 32P. In the case of isoelectric focusing, the 32P will focus out of the gel during electrophoresis; hence the running buffer will be contaminated with 32P. These solutions should be disposed of as radioactive waste as local regulations dictate. Organelle Sample Preparation for SDS-PAGE


Additional Materials (also see Basic Protocol 2) 2× SDS-PAGE sample buffer (see recipe) Boiling water bath 1. Prepare organelles and perform permeabilization and labeling (see Basic Protocol 2, steps 1 through 5). 2. Ultracentrifuge samples 10 min at 100,000 × g, 4°C, and aspirate the supernatant. Add 50 µl of 2× SDS-PAGE sample buffer and mix well. 3. Boil the samples 3 min in a boiling water bath. The samples are now ready for analysis by SDS-PAGE (UNIT 10.2).

Organelle Sample Preparation for Isoelectric Focusing


Additional Materials (also see Basic Protocol 2) Two-dimensional-PAGE lysis buffer (see recipe) Lyophilizer 1. Prepare organelles and perform permeabilization and labeling (see Basic Protocol 2, steps 1 through 5). 2. Ultracentrifuge samples 10 min at >100,000 × g, 4°C, and aspirate the supernatant. Add 200 µl of two-dimensional-PAGE lysis buffer and mix well. 3. Lyophilize the samples overnight. The samples are now ready for analysis by isoelectric focusing (UNIT 10.3). Samples for isoelectric focusing must have a low salt concentration because a high salt concentration often causes band broadening during the isoelectric focusing dimension.



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DIRECT ANALYSIS OF CYTOSOLIC OR MEMBRANE-BOUND KINASES This protocol utilizes isolated subcellular fractions in straightforward kinase assays by adding exogenous, purified protein kinases to the reaction mixture containing the organelle in question. This works only if the protein under study is phosphorylated by a cytosolic or membrane-bound kinase and its substrate is on the cytosolic face of the organelle. One can then determine the phosphorylation of proteins by analyzing the reaction mixture by SDS-PAGE. It is a good idea to perform the reactions and subsequent manipulations in screw-cap microcentrifuge tubes to minimize the risk of [32P] contamination. This protocol may be used to extend findings based on the use of the permeabilization strategies and assays with isolated organelles detailed elsewhere in this unit. Hence, if a protein of interest is phosphorylated, it can be useful to determine the subcellular localizations of the kinase(s) responsible, as this may make it possible to infer clues as to the identity of the kinase(s). Materials Appropriate kinase assay buffer (UNIT 18.7), 4°C [γ-32P]ATP solution (see recipe in UNIT 18.7) Suspension of isolated intracellular organelles, 4°C Purified protein kinase of interest (keep on ice) 2× SDS-PAGE sample buffer (see recipe), ice-cold Screw-cap microcentrifuge tubes 30°C and boiling water baths 1. Prepare the following reaction mixture in a screw-cap microcentrifuge tube at 4°C: 10 µl 5× kinase assay buffer 5 µl [γ-32P]ATP solution (5 µCi/5 µM ATP final) 1 to 35 µl organelle suspension (i.e., volume equivalent to 1 mg protein) H2O to 50 to 100 µl. After reaction mix has been assembled, warm to 30°C for 5 min. 2. Start reaction by addition of the purified protein kinase of interest. Incubate 15 min at 30°C. In the case of immunoprecipitated enzymes adsorbed onto beads, the assay mix minus the enzyme source can be made up complete first. This can then be dispensed, prewarmed, onto the immunoprecipitates. Incubation time can be adjusted as necessary depending on the extent of protein phosphorylation observed and the linearity of the reaction.

3. Stop the reactions by the addition of 100 µl of ice-cold 2× SDS-PAGE sample buffer. After thorough mixing, heat the samples 3 min in a boiling water bath. The samples are then ready for resolution by SDS-PAGE (UNIT 10.2). Samples for SDS-PAGE contain all the 32P originally in the assays. Great care should be exercised in sample preparation and loading. The gels should be run until the dye front passes out of the gel. This means that the SDS-PAGE running buffer will be contaminated with 32P, and this should be disposed of as radioactive waste as local regulations dictate.




DETERMINATION OF SPECIFIC RADIOACTIVITY The specific radioactivity (specific activity) of any radioactive compound is defined as the number of atoms of the radioactive isotope in a molecule as a proportion of the total number of atoms of the same type. The structure of ATP is shown in Figure 18.8.2, and the form used in the study of protein kinases is [γ-32P]ATP, indicating that only the γ phosphate group has the radioactive 32P atom. There are, however, two other atoms of 31P. Hence, to attain the maximum possible specific activity, one should replace 100% of all phosphorus atoms with the 32P isotope. This is obviously not necessary in the case of kinase phosphotransfer reactions, as only the γ phosphate is transferred to a substrate; therefore in practice it is irrelevant whether the α and β phosphate groups have the 32P label. The units used by vendors of [γ-32P]ATP are curies per mole (Ci/mol), even though the curie is no longer the international unit of radioactivity.

SUPPORT PROTOCOL 1

NH2 N

N O H

O

P HO

γ

O O

P OH

β

O O

P

O

CH2

N

N O

OH

α HO

OH

Figure 18.8.2 Structure of ATP highlighting the α, β, and γ phosphates/phosphorus atoms.

It is important to know the specific radioactivity of the ATP in a kinase assay, as important data can be derived from this information. Consequently this can be determined as required, although one can make assumptions based on the volume of [γ-32P]ATP added to cold ATP prior to the assay. Obviously, the specific activity of [γ-32P]ATP cannot be assumed when adding [γ-32P]ATP to permeabilized cells or after labeling intact cells with 32 Pi. In these cases, one can determine the specific activity experimentally as described below. The specific radioactivity of [γ-32P]ATP can be determined by a number of methods, but the following are reliable and relatively simple. Methodologies for the extraction of ATP from either 32Pi-labeled cells or from cell lysates are detailed below (Stephens and Downes, 1990). The basis for the extraction is the same in each case, but slightly different alternative steps are required. Protein or cell debris is first precipitated with perchloric acid and removed by centrifugation. The supernatants are then neutralized with a mixture of tri-n-octylamine and 1,1,2-trichlorotrifluoroethane (Freon). The aqueous phase from this step can then be directly analyzed for ATP concentration and 32P content. ATP may be quantified using anion-exchange HPLC to separate nucleotides and a UV detector to detect nucleotides via UV absorption at 254 nm. The peak areas associated with injection of different amounts of ATP onto the column are then calculated and a standard curve constructed. The radioactivity associated with the ATP peak is easily determined by scintillation counting. Analysis of Protein Phosphorylation


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Materials 32 P-labeled cells or cell lysates Phosphate-buffered saline (PBS; APPENDIX 2) Standard ATP samples of known concentration in 4% perchloric acid 4% or 8% (v/v) perchloric acid, ice-cold Tri-n-octylamine (Sigma) 1,1,2-trichlorotrifluoroethane (Freon; e.g., Aldrich, Sigma) Linear gradient consisting of: Solution A: Milli-Q water filtered through 0.2-mm filter Solution B: 1.25 M NH4H2PO4, pH 3.8 (adjust pH with H3PO4; filter through 0.2-mm filter) Nucleotide standard mix: e.g., 1 mM each of AMP, ADP, ATP, GMP, GDP, and GTP Cell scrapers Polypropylene screw-cap microcentrifuge tubes or 5-ml polypropylene tubes with very tight-fitting caps Tabletop centrifuge HPLC system consisting of: Whatman Partisphere SAX HPLC column and guard column Apparatus for producing linear gradient UV detector set at 254 nm Fraction collector Whatman low-dead-volume 0.2-mm syringe filters Additional reagents and equipment for HPLC (UNITS 10.12-10.14) Extract ATP from cells or cell lysates For labeled cells 1a. Aspirate the medium from the labeled cells and rinse twice with PBS. Aspirate the final wash thoroughly and place the cells on ice. Quickly add 0.5 ml of ice-cold 4% perchloric acid and mix gently to lyse the cells. Incubate 5 min on ice. 2a. Scrape the cells into the acid solution using a cell scraper and quantitatively transfer the lysate to a screw-cap microcentrifuge tube. Microcentrifuge 5 min at maximum speed, 4°C, to bring down the cell debris. 3a. Transfer the supernatant to a fresh tube and keep on ice. Freshly prepare a mixture of 1.2 parts tri-n-octylamine to 1 part Freon (on a v/v basis) and mix well. Add 0.6 ml of this mixture to the perchloric acid supernatants, cap the tubes tightly, and vortex for at least 1 min. Extract four to six standard ATP samples of known concentration in parallel with the test supernatants. It is very important that the samples be mixed extremely thoroughly at this point to ensure complete neutralization.

For labeled cell lysates 1b. In a 5-ml polypropylene tube add an equal volume of ice-cold 8% perchloric to the amount of labeled cell lysate required and place on ice. Mix thoroughly and incubate 5 min on ice. 2b. Centrifuge 5 min (or until protein is completely pelleted) at 3800 × g, 4°C. Permeabilizatin Strategies to Study Protein Phosphorylation

3b. Transfer the supernatant to a fresh tube and keep on ice. Freshly prepare a mixture of 1.2 parts tri-n-octylamine to 1 part Freon (on a v/v basis) and mix well. Add 1.2 vol of this mixture to the volume of sample remaining after protein precipitation and



vortex extremely well. Extract four to six standard ATP samples of known concentration in parallel with the test supernatants. It is very important that the samples be mixed extremely thoroughly at this point to ensure complete neutralization.

Recover neutralized aqueous sample 4. After complete mixing, centrifuge 5 min at 3800 × g, room temperature. Three layers should be evident following centrifugation. The bottom layer is excess tri-n-octylamine/Freon, the central layer is octylamine perchlorate (which is often yellow in color), and the upper layer is the neutralized aqueous sample.

5. Transfer the upper layer to a fresh tube and measure the pH using pH paper. The pH should be pH 6 to 7. If the pH of the sample is 90% of the radioactivity from the cellulose.

12. Remove the elution tip from each of the microcentrifuge tubes, being careful to leave all the eluate in the tube (some may cling to the sides of the tip as drops, which should be removed and added back to the contents of the tube). Save the elution tip. If eluting more than one spot, keep track of which tip was used for which peptide. 13. Clarify the eluate(s) by microcentrifuging 5 min at full speed (a small cellulose pellet will be visible after centrifugation; its size will depend on how snugly the sintered disk fits into the elution tip). Transfer the supernatant to a fresh microcentrifuge tube. It is very important to remove all traces of cellulose at this point, as contamination of the phosphopeptide with cellulose can ruin further analyses.

14. Count both the eluate and the “empty” elution tips by Cerenkov counting. ∼90% of the radioactivity should be in the eluates, with little remaining in the cellulose left in the tips. Given the pain and frustration involved in their manufacture, a good elution tip should be saved and reused. To clean these tips, apply a vacuum to the small end of the tip and suck the cellulose out (into a vacuum flask) while aspirating ∼10 ml elution buffer or deionized water through the tip to rinse it. Dry and then count the tips on a scintillation counter before reusing them.



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15. Lyophilize the eluates in a SpeedVac, then count them by Cerenkov counting. The counts here should be slightly higher than those of the liquid eluate. The number of cpm in this final sample of eluted peptide will often determine how it can be analyzed further. BASIC PROTOCOL 2

DETERMINATION OF THE POSITION OF THE PHOSPHORYLATED AMINO ACID IN THE PEPTIDE BY MANUAL EDMAN DEGRADATION If insufficient material is available for direct sequencing, a manual Edman degradation of the peptide can be performed to determine at which position the phosphorylated amino acid is present in the peptide. During each cycle of Edman degradation, the most amino-terminal amino acid residue is released from the peptide, and a sample from the reaction mixture is taken after each cycle. Phosphoserine or phosphothreonine is released as a derivative of serine or threonine and free phosphate; in contrast, phosphotyrosine is released as the anilinothiazolinone derivative of phosphotyrosine. Free phosphate and the PTH derivative of phosphotyrosine can be separated from the peptide by electrophoresis on a TLC plate. This approach indicates at which cycle the radioactivity and thus the phosphorylated amino acid is released from the peptide. Materials Eluted phosphopeptide (see Support Protocol 1) 5% (v/v) phenylisothiocyanate (PITC) in pyridine 10:1 (v/v) heptane/ethyl acetate—mix 10 parts heptane with 1 part ethyl acetate 2:1 (v/v) heptane/ethyl acetate—mix 2 parts heptane with 1 part ethyl acetate 100% (w/v) trifluoroacetic acid (TFA) Electrophoresis buffer, pH 1.9 (see recipe) 200 to 500 cpm 32P (prepared by diluting 32P orthophosphate with deionized water) or 2 mg/ml PTH-phosphotyrosine (see recipe) Microcentrifuge tubes (Myriad Industries) 45°C water bath Scintillation counter appropriate for Cerenkov counting Glass-backed TLC plates (20 × 20 cm, 100 µm cellulose; EM Science) 65°C drying oven or fan Additional reagents and equipment for electrophoresis of peptides on a TLC plate (see Basic Protocol 1 and Figure 18.9.3) and autoradiography (APPENDIX 3A) Determine experimental parameters 1. Decide the number of cycles to be run based on the list of candidate peptides. The number of cycles is designated as X. The starting volume for each cycle will be 20 ìl.

2. Dissolve the eluted peptide in 20 µl deionized water in what will be called the reaction tube (a microcentrifuge tube). 3. Remove a sample equal to 20/(X + 1) µl to a new tube; this is the starting material sample. Store this at 4°C. This sample will be lyophilized with the other cycle fractions at a later point.

Phosphopeptide Mapping and Identification of Phosphorylation Sites

Perform the Edman reactions 4. Add enough deionized water to the reaction tube to restore the volume to 20 µl. Count the sample at this point: a. to insure that the expected number of cpm have in fact been removed from the initial sample (as the starting material sample);



b. to check the cpm at the start of each given cycle. 5. Add 20 µl of 5% phenylisothiocyanate in pyridine to each reaction tube, vortex well, spin briefly in a microcentrifuge to collect the sample at the bottom, and incubate at 45°C for 30 min. 6. Add 200 µl of 10:1 heptane/ethyl acetate to each reaction tube and vortex for 15 sec. Microcentrifuge 1 min at full speed to separate the two phases. The pyridine will partition into the (upper) organic phase.

7. Carefully remove the upper organic phase using a plastic transfer pipet. Reextract the (bottom) aqueous phase a second time with 10:1 heptane/ethyl acetate as in step 6. 8. Extract the aqueous phase two more times as in step 6, this time using 2:1 heptane/ethyl acetate. 9. Freeze the aqueous phase on dry ice and lyophilize in a SpeedVac evaporator. 10. Dissolve the dried sample in 50 µl of 100% trifluoroacetic acid (TFA) and incubate this at 45°C for 10 min. 11. Lyophilize the sample in a SpeedVac evaporator. 12. Count the sample by Cerenkov counting. There should be the same number of cpm as at the beginning of the cycle (i.e., at step 4 in this case).

13. Add 20 µl deionized water to the reaction tube, vortex, and microcentrifuge briefly. Remove a portion for analysis of the first-cycle products that is equal to 20/X. Store this at 4°C with the starting material sample. 14. Add deionized water to restore the sample volume to 20 µl to start the second cycle. Repeat steps 5 to 12. 15. After the second cycle, add 20 µl deionized water, resuspend the remaining sample, and remove 20/(X−1) µl to a new tube for analysis of the second-cycle products. Repeat steps 4 to 12. 16. Continue repeating steps 4 to 12 until the desired number of cycles have been run. For each new cycle, the amount of the sample to be removed is 20/X−Y where Y equals the cycle number minus 1.

Analyze the Edman products 17. Lyophilize all samples to dryness in a SpeedVac evaporator. Count all final samples by Cerenkov counting, 18. If lyophilized, dissolve the samples in 5 µl of pH 1.9 electrophoresis buffer or deionized water. Microcentrifuge 2 min at maximum speed to bring down any insoluble material. Alternatively, if the sample volumes removed after each cycle are small enough, skip steps 17 and 18 and load the samples directly onto the TLC plate.

19. Spot all samples from the analysis of a given phosphopeptide at least 1 cm apart on a line of origins running vertically through the center of the TLC plate (Fig. 18.9.5). As a marker, depending on the phosphoamino acid content of the peptide under investigation, spot 50 to 200 cpm of [32P]phosphate or 1 to 2 µg PTH-phosphotyrosine (0.5 to 1.0 µl of 2 mg/ml PTH-phosphotyrosine) at an origin on that same vertical



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A

B marker origin cycle 6 cycle 5 cycle 4

4.5 cm

2 cm

15 cm

cycle 3 cycle 2

15.5 cm

cycle 1 starting material 3 cm 10 cm

5.0 cm

10 cm 12.5 cm

12.5 cm

Figure 18.9.5 Sample and dye origins and blotter dimensions for analysis of manual Edman degradation products at pH 1.9. (A) Location of the sample and standard origins. To mark a TLC plate, the plate is placed on top of a life-size template on top of a light box and the origins are marked on the cellulose side using a very blunt extra-soft pencil. (B) Dimensions of the blotter and the location of the slot that fits over multiple sample and marker origins. The blotter is soaked in electrophoresis buffer, blotted with a sheet of Whatman paper to remove most of the buffer, and placed on top of the TLC plate so that the origins are in the middle of the slot.

line. Load 1⁄3 to 1⁄2 µl of sample at a time, air drying the sample between applications (see Basic Protocol 1, step 25). 20. Wet the plate as described in Figure 18.9.5. 21. Prepare the HTLE 7000 apparatus and electrophorese the samples for 25 min at 1.0 kV in pH 1.9 electrophoresis buffer (see Basic Protocol 1 and Figure 18.9.3). 22. After drying the plate (either in a 65°C oven or with a fan) mark it appropriately with radioactive or fluorescent markers and expose it to presensitized film with an intensifying screen at −70°C (autoradiography; APPENDIX 3A). BASIC PROTOCOL 3

DIAGNOSTIC SECONDARY DIGESTS TO TEST FOR THE PRESENCE OF SPECIFIC AMINO ACIDS IN THE PHOSPHOPEPTIDE Further information about a phosphopeptide of interest can be obtained by digestion with sequence-specific proteases or cleavage by site-specific chemicals. After incubation with a protease or chemical, the peptide is analyzed by separation in two dimensions on a TLC plate. A change in mobility upon treatment with a particular reagent indicates that the peptide was susceptible to cleavage, and consequently that the amino acid or amino acid sequence that confers susceptibility to cleavage by this reagent must be present in the peptide.


Materials Eluted phosphopeptide (see Support Protocol 1) Enzyme to be used for digestion and appropriate buffer (see Table 18.9.1) 2-mercaptoethanol



Table 18.9.1

Specificities and Digestion Conditions for Enzymes and Other Cleavage Reagents

Enzyme or reagent Specificitya

Digestion conditions

Comments

TPCK-trypsin

K—X; R—X

pH 8.0-8.3

α-Chymotrypsin Thermolysin

F—X; W—X; Y—X X—L; X—I; X—V

pH 8.3 pH 8.0, 1 mM CaCl2, 55°C

Does not cut K/R-P; cuts inefficiently at K/R-XP.Ser/P.Thr and K/R-D/E; cuts wells at K/RP.Ser/P.Thr; cuts X-K/R-K/R-K/R incompletely Does not cleave F/W/Y-P or P.Tyr-X Will recognize most apolar residues to some extent; CaCl2 may affect the electrophoretic mobility —

Proline-specific P—X endopeptidase Cyanogen bromide M—X (CNBr)

pH 7.6

V8 protease

E—X

50 mg/ml CNBr in 70% formic acid, 90 min, 23°C pH 7.6

Endoproteinase Asp-N Formic acid

X—CSO3H; X—D

pH 7.6

D—P

70% formic acid, 37°C, 24-48 hr

CNBr is toxic; will only cleave unoxidized methionine V8 will not cleave at every E in whole proteins; does give a consistent pattern Will cleave X-E at high enzyme concentrations —

aThe dash indicates the cleavage site. See APPENDIX 1C for definitions of the one-letter abbreviations for amino acids.

Electrophoresis buffer of appropriate pH (see recipe) Water bath at appropriate temperature for enzyme digestion Glass-backed TLC plates (20 × 20 cm, 100 µm cellulose; EM Science) Additional reagents and equipment for chromatography and electrophoresis of phosphopeptides (see Basic Protocol, steps 24 to 31) 1. Dissolve the eluted phosphopeptide in 50 µl of the appropriate buffer in a microcentrifuge tube and microcentrifuge briefly to collect all solution at the bottom of the tube. Check the pH of the peptide solution by spotting 1 µl on a piece of pH paper to be sure that this final pH will allow enzyme activity. If the buffer’s pH has been altered dramatically by addition of the peptide, adjust it before adding enzyme. 2. Remove a portion of the sample (usually 50%) to be run both as an undigested control and as a mix with a portion of the digested sample. 3. Add 1 to 2 µg enzyme to the portion of the sample to be digested, vortex, and concentrate the sample in the bottom of the tube by microcentrifuging briefly. 4. Incubate all tube(s) in a water bath at the appropriate temperature for at least 1 hr. 5. Add another aliquot of enzyme and continue the incubation step for an additional hour. 6. Add 1 µl of 2-mercaptoethanol to each sample and boil 5 min to inactivate the enzyme Do this to all samples to ensure uniformity of sample preparation. It is necessary to completely inactivate the enzyme prior to loading the sample on the plate when analyzing a mix of digested and undigested peptide, since the undigested sample may be rapidly digested when the two samples are mixed.



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7. Lyophilize the samples in a SpeedVac evaporator. 8. Resuspend the samples by vortexing vigorously in 6 µl of electrophoresis buffer of the appropriate pH. Microcentrifuge at full speed to bring down insoluble material. 9. Load half of the undigested sample on a single TLC plate. Load half of the digested sample on each of two TLC plates; on one of these load the other half of the corresponding undigested sample as a mix. 10. Perform electrophoresis and chromatography on the plate as described above in Basic Protocol 1, steps 24 to 31. Based on the position where the particular phosphopeptide being analyzed ran in the original map, choose a pH and running time that will allow good separation of the peptide from its potential cleavage products but will ensure retention of the smaller cleavage products on the plate. SUPPORT PROTOCOL 2

PREPARATION OF PHOSPHOPEPTIDES FOR MICROSEQUENCE DETERMINATION OR MASS SPECTROMETRY The following is a general protocol and list of considerations for generating enough material for analysis by mass spectrometry or microsequencing starting with either intact cells or an in vitro system. 1. Optimize 32P labeling of the protein of interest. If the site of interest is seen only in stimulated cells, a time course of phosphorylation following stimulation may be helpful, as would determination of the optimal concentration of agonist. If an in vitro system is being employed, determine the optimum conditions (time, and ratio of kinase and substrate concentrations) for the kinase reaction. Include 1 mM cold ATP in the reactions to maximize the stoichiometry. UNIT 18.7, which deals with in vitro phosphorylation reactions, provides a detailed discussion of these parameters and how to manipulate them.

2. Calculate the number of cells or amount of substrate needed to isolate 10 pmol phosphorylated material. Even under optimal conditions, it is often not possible to achieve more than 25% stoichiometry of phosphorylation in vitro. In intact cells, the stoichiometry may be even less. It cannot hurt to overestimate the amount of starting material required, as the losses taken during the isolation procedures will always exceed expectation.

3. When calculating how to scale up the reactions, consider the following points.


a. The radioactivity of these samples is only used for visualization purposes—i.e., to determine which gel band to isolate, which phosphopeptide to isolate from the TLC plate, and which HPLC fraction(s) to use for final analysis. Thus, the majority of the material can be unlabeled, as only ∼1000 cpm per map spot are necessary for analysis at the time when the preparative HPLC is run. When isolating overexpressed protein from cells, labeling only 2 or 3 dishes of the 20 needed to generate enough material may be sufficient. When using an in vitro system, perform only one reaction using γ [32P]-ATP (include only the very minimum amount of cold ATP necessary for kinase activity) to generate the labeled material. To generate enough material for further analysis, perform an additional kinase reaction with unlabeled ATP only. For visualization, mix the labeled and unlabeled samples before resolving them by SDS-PAGE.



b. The efficiency of protein elution decreases as the amount of gel increases, so try to keep the number of lanes on the preparative gel(s) to a minimum. About four lanes/slices of acrylamide gel can be successfully extracted per tube. c. If ∼20 µg substrate protein is present in each elution sample, it will not be necessary to add carrier protein at the TCA precipitation step (see Basic Protocol 1 step 11). This will result in a cleaner sample, as the tryptic fragments of the carrier protein will be eliminated from the mix of fragments run on the TLC plate. d. The 20 µg trypsin used to digest map samples in Basic Protocol 1 is in vast excess. While it is important that the digestion go as far to completion as possible, it is probably not necessary to scale up the amount of trypsin used. Instead, consider pooling several like samples at the performic acid digestion step (at the end of the 60 min incubation in order to give the protein the maximum time to dissolve). Adding 1 to 5 µg (total) trypsin to even 50 µg of protein for digestion is not unreasonable. Minimizing the amount of trypsin used will in turn minimize the amount of “extra” protein loaded on each TLC plate and ensure that the sample does not streak due to overloading. e. Determine the number of TLC plates to be run based on the amount of total protein to be analyzed—total protein includes the amount of trypsin and the amount (if any) of carrier protein used as well as the amount of the protein of interest. Even though the capacity of the TLC plates is ∼100 µg, to ensure good separation, no more than 60 µg of protein should be run on each plate. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Chromatography buffers Phosphochromatography buffer: 750 ml n-butanol 500 ml pyridine 150 ml glacial acetic acid 600 ml deionized water Store at room temperature Isobutyric acid buffer: 1250 ml isobutyric acid 38 ml n-butanol 96 ml pyridine 58 ml acetic acid 558 ml deionized water Regular chromatography buffer: 785 ml n-butanol 607 ml pyridine 122 ml glacial acetic acid 486 ml deionized water Store all of the above buffers up to 6 months at room temperature.



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Electrophoresis buffers For each of these buffers, mix well and check the pH. Record the pH and the date on the bottle; if the pH is more than a tenth of a unit off, remake the solution. Do not adjust the pH. Store all buffers at room temperature. pH 1.9 buffer: 50 ml formic acid (88% w/v) 156 ml glacial acetic acid 1794 ml deionized water pH 3.5 buffer 100 ml glacial acetic acid 10 ml pyridine 1890 ml deionized water pH 4.72 buffer 100 ml n-butanol 50 ml pyridine 50 ml glacial acetic acid 1800 ml deionized water pH 6.5 buffer 8 ml glacial acetic acid 200 ml pyridine 1792 ml deionized water pH 8.9 buffer 20 g ammonium carbonate 2000 ml deionized water Green marker dye Prepare a solution containing 5 mg/ml ε-dinitrophenyllysine (yellow) and 1 mg/ml xylene cyanol FF (blue) in pH 4.72 electrophoresis buffer (see recipe) diluted 1:1 with deionized water. Store up to 1 year at room temperature. PTH-phosphotyrosine, 2 mg/ml Combine 20 µl of 100 mg/ml phosphotyrosine with 20 µl of 5% (v/v) phenylisothiocyanate in pyridine. Incubate 30 min at 45°C. Extract twice with 200 µl of 10:1 (v/v) heptane/ethyl acetate, then twice with 200 µl of 2:1 (v/v) heptane/ethyl acetate (see Basic Protocol 2, steps 6 to 8, for extraction technique). Freeze the aqueous phase and lyophilize in a SpeedVac evaporator. Dissolve the sample in 0.1 N HCl, incubate 20 min at 80°C, and lyophilize again in a SpeedVac evaporator. Dissolve in 1 ml pH 1.9 buffer (see recipe for electrophoresis buffers). PVP-360 in 100 mM acetic acid 0.5 g PVP-360 (Sigma) 575 µl glacial acetic acid 99.4 ml deionized water Store up to 1 year at room temperature




COMMENTARY Background Information Phosphopeptide mapping is a very sensitive technique that can help the investigator answer a variety of questions about a protein of interest. For some, phosphopeptide mapping is a tool to find out whether a particular protein is phosphorylated on one or more sites. This question can be answered by simply running a phosphopeptide map of the protein labeled in living cells. Other investigators want to know whether the increase in phosphorylation seen when cells are treated with a particular agent is restricted to one or more specific sites or whether it is evenly distributed over all phosphorylation sites present in the protein. Finally, detailed analysis of phosphopeptides isolated from a TLC plate can be used to identify the residues that are phosphorylated in a protein of interest. Several different strategies may be followed to identify the phosphorylation site represented by a particular spot on a phosphopeptide map. This is most definitively accomplished by eluting phosphopeptides from cellulose plates for either direct sequencing or for analysis by mass spectrometry (see Support Protocol; Fischer et al., 1991; Wang et al., 1993; Mitchelhill et al., 1997). While these two techniques require expensive instruments and expertise not found in most laboratories, such analysis can often be arranged by collaboration. However, sometimes it is not possible to take advantage of these techniques, since they require 1 to 10 pmole of material for analysis. For a 50-kDa protein one would need 0.1 to 1.0 µg starting material, assuming 50% recovery and 100% stoichiometry of phosphorylation at the site of interest. Use of an in vitro phosphorylation system that mimics the situation in intact cells will simplify matters greatly. Further considerations and strategies for preparation of samples for these two techniques are discussed at the end of this chapter in Support Protocol 2. Another approach to phosphorylation-site identification is to make an educated guess as to the identity of the site. Clues to a site’s identity include phosphoamino acid analysis of the individual phosphopeptide (UNIT 18.3); the result of manual Edman degradation of a phosphopeptide providing the cycle at which the phosphate is released and thus the position of the phosphorylated residue in the peptide (see Basic Protocol 2); and secondary enzymatic digests of the phosphopeptide that can be used in a diagnostic sense to determine the presence of other specific amino acids in the peptide (see

Basic Protocol 3). All three of these techniques are easily accomplished in a laboratory that is already set up for phosphopeptide mapping. The first step in all three is the isolation of the phosphopeptide from the cellulose plate (see Support Protocol 1). The validity of one’s guess can be tested by phosphopeptide mapping of a mutant protein lacking a phosphate acceptor at the site in question. Alternatively the guess can be substantiated by synthesizing the tryptic phosphopeptide and testing it for comigration with the phosphopeptide isolated from the peptide map.

Critical Parameters and Troubleshooting Generating phosphopeptide maps Keep in mind that the sort of analyses presented throughout this unit will give information regarding only the acid-stable forms of phosphoamino acids (i.e., phosphoserine, phosphothreonine, and phosphotyrosine) and will essentially ignore other forms such as phosphohistidine and phosphoaspartate, should they be present. Carrier Protein. The authors prefer to use RNase as carrier protein during TCA precipitation, particularly when analyzing proteins labeled in intact cells, because it degrades 32P-labeled RNA species that may have copurified with the protein of interest. The nucleotides generated by the degradation of RNA do not precipitate in TCA. Cleaving the protein. In order to generate a phosphopeptide map, the 32P-labeled protein needs to be cleaved into smaller fragments that can be separated by electrophoresis and chromatography on TLC plates. To do this requires an enzyme or chemical agent that cleaves reproducably and with a certain frequency. If not enough cleavage sites are present, the fragments generated will be too large and will not be separated easily by electrophoresis and chromatography on TLC plates. In addition, large fragments may contain multiple phosphorylation sites. This leads to maps that are less informative and more difficult to analyze. The authors routinely use trypsin and chymotrypsin. Other reagents are available (Table 18.9.1), but most of them cut much less frequently and some of them do not work very efficiently on full-length proteins. Removing ammonium bicarbonate. Following digestion, repeated cycles of lyophilization



Supplement 48


are carried out to evaporate all ammonium bicarbonate. The presence of salts in the sample will interfere with the electrophoretic separation of the peptides. After lyophilization, the protein digest appears as an invisible film at the bottom of the tube. The presence of any crystalline material indicates the presence of salts, most likely ammonium bicarbonate that can be removed by additional rounds of lyophilization. Controlling oxidation. Both cysteine and methionine can give rise to several oxidized derivatives. The oxidation state of these amino acids affects the mobility of peptides during chromatography, resulting in separation of oxidation state isomers. This complicates the interpretation of the phosphopeptide map. To prevent this, the protein or peptides are oxidized by incubation in performic acid at 4°C. Incubation at higher temperatures may give rise to unwanted side reactions and should be avoided. Elution and TCA precipitation or transfer to a membrane? In Basic Protocol 1, the 32P-labeled protein is isolated from a small slice of a dried polyacrylamide gel by rehydrating and grinding up the gel followed by elution in a buffer containing SDS and 2-mercaptoethanol. The protein is subsequently TCA precipitated, oxidized, and digested with trypsin. This is a time-consuming and laborious procedure. Yields are variable and usually not better than 50%. The alternative is to transfer the protein to a PVDF membrane; any unoccupied proteinbinding sites on the strips of membrane containing the protein of interest are blocked by incubation with PVP-360 in acetic acid before incubation with trypsin. Most peptides dislodge from the membrane during the digestion. This protocol is much faster and less laborious, and does not require the use of additional carrier proteins that may lead to overloading of the TLC plate and to streaky maps. Obviously this method is a poor choice for proteins that transfer poorly from the polyacrylamide gel to a membrane. In addition, it is possible that certain peptides that are generated during protease digestion retain a high affinity for the membrane and therefore fail to elute. If those peptides contain a phosphorylation site, this site will not be represented on the peptide map. This can lead to misinterpretations of the results. It is therefore advisable to first compare maps generated with Basic Protocol 1 and the Alternate Protocol. If these maps are identical, and if the protein transfers well from the gel to the membrane, the Alternate Protocol should be the protocol of choice.

Amount of sample. The authors like to load at least 1000 cpm on a plate for a peptide map. If the final sample has many more than 1000 cpm and a “pretty-looking” map is desired, it may be better to load only a fraction of the sample. Remember that overloading can lead to streaky maps. If a preparative map from which a particular peptide will be isolated is being run, it may be best to run the entire sample on two (or more) separate plates. Theoretically, it should be possible to separate 100 µg of material on a single TLC plate; this is often not the case in practice. Check the rate at which the first drop spotted sinks into the cellulose; as more sample is spotted, this rate will decrease. If, while spotting, one gets to a point where the sample drop just sits on the origin and does not spread into the cellulose, stop loading. Peptide diffusion. Peptides diffuse during the electrophoresis and chromatography, and this leads to a reduction in resolution and sensitivity. To counteract this, the authors try to keep the area on the TLC plate onto which the sample is spotted as small as possible by spotting only a small amount at a time (i.e., less than 1 µl) and drying the sample origin between spottings. In addition, the sample is concentrated by wetting the TLC plates with electrophoresis buffer using a blotter with holes cut out around the origin (Figs. 18.9.1 and 18.9.3). Pressing the edges of the hole onto the plate results in buffer moving from the blotter towards the center of the hole. This concentrates the sample on the origin. For this process to work well, the origin has to be precisely in the center of the hole. In addition, the buffer has to move with similar speed from the entire circumference towards the origin. The sample will inevitably streak if the buffer takes a long time to wet the spot, or moves unevenly through the spot. Electrophoresis system. In the authors’ laboratories the HTLE 7000 electrophoresis system (Fig. 18.9.2) is used. This system features water cooling and an inflatable airbag that presses the TLC plate against the cooling plate. Water cooling prevents overheating during electrophoresis. The inflatable airbag presses excess buffer from the TLC plate; this limits diffusion of the peptides and improves resolution. Buffers. Three different buffers are typically used for electrophoresis: pH 1.9, pH 4.72, and pH 8.9. Less often used are pH 3.5 and pH 6.5 electrophoresis buffer. To find out which buffer gives the best separation of peptides generated from a particular protein, all three buffers should be tested. If possible, the authors prefer



to work with pH 1.9 buffer. Most peptides dissolve well at this pH. In addition, use of this buffer results less often in streaky maps. The authors usually spot the digest on the origins as marked in Figures 18.9.1 and 18.9.3. For optimal separation of the phosphopeptides generated from a particular protein, the position of the origin and the electrophoresis time may need to be changed. We prefer to change the time of electrophoresis rather than changing the voltage. Chromatographic process. Chromatography usually takes 12 to 15 hr, but the exact time may vary depending on the age of the chromatography buffer, the batch of plates, the buffer system, the quality of reagents used in the buffer, and the temperature in the room. Three different chromatography buffers are commonly used: isobutyric acid buffer, regular chromatography buffer, or phosphochromatography buffer (see Reagents and Solutions). Pyridine, which is present in all three chromatography buffers, oxidizes and turns yellow over time. Do not use oxidized pyridine to make up chromatography buffers. To find out which buffer gives the best separation of the peptides generated from a particular protein, all three chromatography buffers should be compared. Most investigators prefer not to use isobutyric acid buffer because it is particularly foul smelling. During the chromatography run, the air space in the tank saturates with buffer and this makes it possible for the volatile chromatography buffer to run all the way to the top of the TLC plate. When the chromatography tank is opened, most of the buffer-saturated air will escape from the tank. This makes it counterproductive to extend the run after the tank has been opened. Therefore do not open the tank when chromatography is in progress, and take all plates out when the chromatography tank is opened. Separation of the yellow and blue dye functions as a control for successful electrophoresis and allows one to follow the progress during chromatography. The dyes can also be used as standards relative to which the mobility of phosphopeptides of interest can be described, and can be used as markers for the comparison of one plate to another. The yellow compound is neutral at pH 4.72 and pH 8.9 and defines the position to which neutral peptides migrate; at p H 1.9 ε-dinitrophenyllysine is positively charged.

Phosphopeptide identification After running several phosphopeptide maps, it may become apparent that particular phosphopeptides present on the map change in intensity upon treatment of the cells with specific reagents. Such observations often lead to the next question—what is the identity of peptide “A” that becomes phosphorylated following treatment of the cells with factor “B”? If approximately 1 to 10 pmole of phosphorylated peptide can be generated, the peptide is isolated from the TLC plate, purified by HPLC, and identified by mass spectrometry or microsequencing. In many cases, it is not possible to obtain a phosphorylated peptide in such quantities. The investigator is then forced to learn as much about the phosphopeptide as possible before making an educated guess. We find it useful to make a list of all possible candidate peptides including some of their properties. The next step is to eliminate as many candidates as possible using mobility predictions and the results of relatively simple experiments that can be performed on the minute amounts of labeled peptides isolated from TLC plates. Making a list of candidate peptides and eliminating the first candidates Make a list of all possible phosphopeptides that could be generated from the protein of interest given the enzyme used in the primary digest; be sure to include partial cleavage products on this list. This list of peptides should include the nature and position of amino acids that can be phosphorylated and the peptides’ susceptibility to further cleavage by proteases or chemicals (for an example, see van der Geer and Hunter, 1990) After making such a list, first calculate and then plot the predicted mobilities of all candidate phosphopeptides. See Table 18.9.2 for values that can be used to do this. To calculate electrophoretic mobility. The mobility of a peptide in the electrophoresis dimension is dependent on its charge (e) to mass (M) ratio, as Mr = keM−2/3. When calculating relative mobilities (Mr) the simplified equation Mr = eM−1 can be used with good success. The net charge on a peptide is calculated by summing the charges of the N and C termini and those of the side chains of its amino acids at a given pH, and dividing by either the actual mass of the peptide or simply by the number of amino acids in it. Approximate charge values at the specific pHs commonly



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Table 18.9.2 Approximate Charge Values at Specific pHs Commonly Used for Electrophoresisa

Amino-terminal NH2 Carboxy-terminal COOH Arginine Aspartate Cysteine (oxidized) Histidine Glutamate Lysine Phosphoserine Phosphothreonine Phosphotyrosine

pH 1.9

pH 3.5

pH 4.7

pH 6.5

pH 8.9

+1 N +1 N −1 +1 N +1 −1 −1 −1

+1 −0.5 +1 N −1 +1 N +1 −1 −1 −1

+1 −1 +1 −0.7 −1 +1 −0.5 +1 −1 −1 −1

+1 −1 +1 −1 −1 +0.5 −1 +1 −1.3 −1.3 −1.3

+0.5 −1 +1 −1 −1 N −1 +1 −2 −2 −2

aN indicates neutral.


used for electrophoresis are given in Table 18.9.2. To calculate chromatographic mobility. A peptide’s mobility in the chromatographic dimension is dependent on its hydrophobicity, and thus on its amino acid sequence. The order of the amino acids will also change the peptide’s mobility; thus, two peptides of identical sequence which are phosphorylated at one or the other of two possible sites may migrate different distances in the second map dimension, although they migrate identically in the first dimension since their charge:mass ratio is the same. While it is not possible to exactly predict chromatographic mobilities, relative mobilities can be plotted with some success by calculating an average mobility for the peptide based on migratory values of its constituent amino acids. This is not ideal, since the calculated Rf values of individual amino acids are significantly influenced by the presence of their charged amino and carboxy termini, which of course are noncontributory in the context of a peptide. This accounts in part for the compression of the calculated maps compared with the observed peptide migrations. Values for chromatographic mobilities of amino acids have been published in (Boyle et al., 1991); these were determined for each individual amino acid relative to the ε-DNP-lysine (yellow) marker using cellulose plates available twenty years ago. The quality of the cellulose used in such plates has changed markedly over the years; contact Ned Lamb for values that have been empirically determined more recently (http://www.genestream.org).

Bear in mind that these calculations can also be accomplished using a computer program. Ned Lamb (CNRS, Paris) has constructed a Web site for analysis of phosphopeptide maps, which may be found at http://www.genestream. org. Phospepsort 4, the program that he has developed based on an earlier version which originated at the Salk Institute, gives the biophysical characteristics as well as the electrophoretic and chromatographic mobilities of each proteolytic fragment. Alternatively, predicted peptide mobilities can be visualized using the graphical interface to Phospepsort 4: Mobility. In addition, Ned Lamb is working on a program that fits the calculated mobility values to the actual values for peptides of known composition. The Resolve program then reads the position of a spot on the actual map and calculates which peptide(s) derived from the protein being mapped could have the mobility of that spot. It is imperative to note that to date there is no program that accurately predicts the mobility of all peptides of a protein. This may be explained by the fact that mobilities are calculated using values established for single amino acids rather than for peptides. Plotting predicted phosphopeptide mobilities on a graph using linear axes results in a greatly compressed map as compared to that observed in one’s autoradiograms, especially in the chromatographic dimension. Therefore, do not despair if the predicted map of all phosphopeptides in the protein of interest looks nothing like the actual map that was generated experimentally. The value of such predictions comes from the fact that the relative positions of phosphopep-



tides are predicted with great accuracy by such programs/calculations. Thus, if the predicted mobility of a hypothetical phosphopeptide places it on the anode side of a cluster of phosphopeptide candidates, while the phosphopeptide of interest on the actual map is present on the cathode side of the cluster, the hypothetical phosphopeptide may be eliminated from further consideration. This example illustrates how careful use of predicted peptide mobility maps may lead to elimination of candidate peptides. Isolating peptides from TLC plates Phosphopeptides isolated by elution from cellulose, as described above, can be used without further purification for certain types of analysis. What many people overlook, however, is that this sample is by no means necessarily pure. It includes, in addition to the radioactive phosphopeptide in question, any unlabeled tryptic fragments that may have comigrated with it on the cellulose plate—these peptides are generated from trypsin itself and from the carrier protein used in the TCA precipitation during sample preparation. For this reason, the sample is usually further purified by HPLC to clean it up before analysis by mass spectrometry or microsequencing; the column fractions are counted in a scintillation counter to determine which ones to use for further analysis. However, for manual Edman degradation (see Basic Protocol 2), secondary cleavage (see Basic Protocol 3), and phosphoamino acid analysis (UNIT 18.3) no further purification is necessary, as the interpretation of the results relies solely on the visualization of the resultant 32P-containing reaction products. The presence of unlabeled contaminants does not interfere with the interpretation of the results. Phosphoamino acid analysis Perhaps the most obvious step to take with an unidentified phosphopeptide in hand is to determine the phosphoamino acid content of the peptide of interest. This will eliminate many candidate peptides from consideration; this step is obviously not necessary if a long exposure of the phosphoamino acid analysis of the labeled protein in question indicates that only one species of phosphorylated amino acid is present. For phosphoamino acid analysis only 50 cpm of purified phosphopeptide is needed (though in peptide mapping and related protocols one can never have enough cpm). The details of such analysis have been discussed in detail in UNIT 18.3. Briefly, the phosphopeptide

eluted from the TLC plate is hydrolyzed by incubation for 60 min at 110°C in 30 µl of 6 N HCl. The appearance of yellow to brown color in the sample during hydrolysis indicates that some cellulose remained despite efforts to clarify the phosphopeptide eluate. This sometimes causes the sample to streak. After the sample is lyophilized, it is resolved with stainable standards in two dimensions by electrophoresis as described (UNIT 18.3). The phosphoamino acid composition is determined by matching the resultant spot(s) on the autoradiogram with the ninhydrin-stained standards on the cellulose plate. Manual Edman degradation At pH 8 to 9, phenylisothiocyanate reacts with the free amino group(s) of a peptide to form a corresponding phenylthiocarbamyl peptide. Treatment of these PTC-peptides with acid (TFA) results in the cleavage of the derivatized amino-terminal amino acid and its release as an anilinothiazolinone molecule. This latter species is not stable, and will cyclize to yield the phenylthiohydantoin (PTH) derivative of the amino acid in aqueous acid. If a phosphoserine or phosphothreonine residue is present, a βelimination during the cyclization releases free phosphate. Phosphotyrosine, however, is simply released as the anilinothiazolinone derivative. This may be converted to the phenylhydantoin form for analysis by incubating it in 0.1 N HCl at 80°C for 20 min. PTH-phosphotyrosine to use as a marker is easily synthesized by reacting phosphotyrosine with phenylisothiocyanate and then heating it in acid (see Basic Protocol 2, step 5); it can be visualized as a dark spot using a hand-held UV light. While the protocol is relatively simple, each cycle takes ∼2 hr to complete and requires at least 100 cpm for an unambiguous result. It is important to run a portion of the starting material out on the TLC plate to show how much, if any, free phosphate is there, since some hydrolysis of the peptide may have occurred during its isolation. Because at each cycle the reaction may not go to completion, one should plan to do at least one more cycle than is predicted to be necessary to release the phosphate (i.e., if all the candidate peptides are phosphorylated at or before the third residue from the N terminus, at least 4 cycles should be run). Thus, how many cpm are in the eluted map spot may determine how many cycles are run. In any case, we generally do not attempt to perform more than 5 or 6 cycles. If no phosphate is released during the course of these



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cycles, this conclusion in itself will usually greatly reduce the number of candidate sites. In addition to the position of the phosphorylated residue, clues to a peptide’s sequence may also be gleaned from Edman degradation. The way in which the residual peptide shifts its electrophoretic position on the plate after each cycle will indicate whether an acidic or basic amino acid has just been removed. If the tryptic peptide’s carboxy-terminal residue is a lysine, the lysine’s ε-amino group will be derivatized in the first cycle, and so a positive charge will be lost in that instance as well. If an automated peptide sequencer is available, 20 or more Edman cycles may be analyzed by coupling the phosphopeptide via carboxyl groups to a Sequelon membrane (Millipore) and letting the machine do the work. At the end, the fractions are counted to see where the radioactivity is released. This method obviously requires far fewer cpm in a phosphopeptide sample than does the manual Edman protocol, since one only analyzes the released material for radioactivity rather than a portion of the whole sample at the end of each cycle. A method for adapting an automated sequencer for such a purpose is discussed in Mitchelhill et al. (1997). An adaptation of the protocol presented above is found in Fischer et al. (1997). This protocol uses a volatile isothiocyanate (trifluoroethyl isothiocyanate) and volatile buffers; as a consequence the extraction steps can be eliminated and this results in shorter cycle times (∼45 min).


Secondary digests In the past, secondary digests of tryptic phosphopeptides represented a large part of the further analysis of these peptides. The utility of these digests, however, is totally dependent on the sequence of the protein in question. A list of enzymes commonly used in such digests is found in Table 18.9.1, along with their cleavage site(s) and optimal pH and temperature. As is true for trypsin, these enzymes sometimes cleave inefficiently when encountering a cleavage site in a certain sequence context—some of these problematic sites are listed as well. In our experience, the enzymes/reagents that cleave only one amino acid (i.e., proline-specific endopeptidase, V8, or cyanogen bromide) tend to be more useful and give less ambiguous results. Do not expect that the results of one enzyme digest will eliminate more than a few candidate sites. As it is most likely that the enzyme in question will not cleave the phosphopeptide, it

is imperative that a positive control be included in the digest—i.e., a peptide whose sequence is known and which the enzyme will cleave. If such a peptide is not available, a portion of the primary digest might be used as a positive control—with luck this will contain at least one peptide whose migration will be changed by this secondary digestion. Running out the mix of undigested and digested peptides is very important, since failure to comigrate with the original peptide will unequivocally demonstrate a change in the phosphopeptide’s mobility. A sample treated exactly as the digested sample, but without the cleavage reagent, should be analyzed in parallel. This is to ensure that changes in mobility seen in the digested peptide are truly due to the presence of the cleavage reagent. When deciding where to spot the samples and what conditions to use to run the plates, keep in mind that if the peptide did not migrate very far in the original map, it may be possible to load two sample on a single plate. Also keep in mind, however, that free phosphate may be released during the enzymatic digestion (due to elevated temperatures and resulting hydrolysis). Free 32P-phosphate originating from the sample loaded to the right may complicate the interpretation of the sample loaded on the left (anode) side of the plate. It should be noted that additional information about the other amino acids in a phosphopeptide may be gleaned just by running a tryptic digest in the electrophoretic dimension at another pH—identification of the different peptides by their mobility in the chromatography dimension will allow one to compare the mobility of peptides when run at different pHs and decide if a particular spot has changed its migration in the first dimension. Such a change in migration would be affected only if the phosphopeptide contained an amino acid whose charge was changed at the second pH used. Comigration of a synthesized phosphopeptide with a phosphopeptide isolated from a peptide map After eliminating all but a few of the candidate peptides, comigration of a synthetic phosphopeptide with the 32P-labeled peptide generated by digesting a protein from labeled cells may provide convincing evidence as to the latter’s identity. Obviously, care must be taken that the synthetic peptide be quite pure (generating only one spot on the cellulose plate) and that, while enough is loaded to be easily visualized, the plate not be overloaded (which



would cause streaking and therefore render the comigration ambiguous). If the peptide is synthesized as a phosphopeptide, ∼5 to 25 µg of pure peptide is needed for each comigration, as it will need to be ninhydrin stained for visualization. This is by far the easier approach—starting with an unphosphorylated peptide entails not only finding a kinase that will phosphorylate it, but also purifying the phosphopeptide first before running the comigration. In either case, synthesis of peptides, and in particular those with special residues such as phosphorylated amino acids, is not inexpensive, and so this type of experiment is usually attempted after one has accumulated several other clues regarding a site’s identity. The alternative is to mutate the remaining candidate phosphorylation sites. An epitopetagged version of the mutant protein can then be expressed in cells and mapped to check whether the spot in question disappears from the map. This approach of course assumes that the sequence of the protein in question is known and that one has a clone in hand for mutagenesis. While satisfying to many, this mutagenesis approach is not definitive, as it can be argued that the mutated form of the protein may not fold correctly and therefore may not be phosphorylated correctly, resulting in a misleading loss of map spots. Another potential problem is that, if the phosphorylation of the site in question is an ordered event, dependent on another site’s state of phosphorylation, mutation of this other site could lead to the erroneous conclusion that the disappearance of a particular spot is the direct result of the mutation introduced, rather than an event of secondary consequence. Mutagenesis then should be taken as a supporting argument for the presumed identity of a phosphorylation site, rather than as definitive proof. Taken in conjunction with other evidence, it can nevertheless be quite convincing. Scaling up phosphopeptide mapping to isolate peptides for mass spectrometry and microsequencing The most definitive methods for determining the site(s) of phosphorylation in a protein are the determination of the amino acid sequence or mass of the phosphopeptides. While the sensitivity of the instruments used in both these techniques has improved over the past 5 years, both methods require on the order of 1 to 10 pmole of material for analysis. When analyzing the phosphorylation of a receptor protein-tyrosine kinase of which ∼10,000

molecules are present on the cell surface, one would need to grow 60 10-cm dishes of cells to isolate the 6 × 1011 molecules (1 pmol) required for a successful analysis. In this calculation, we have assumed 100% recovery and 100% stoichiometry of phosphorylation at the sites of interest. However, since it is very possible that not every molecule is phosphorylated at the sites in question, and that one is likely to take at least a 50% loss of material over the entire protocol, it would be best to start then with at least 240 dishes of cells. The use of overexpressed protein in cells would obviously facilitate the accumulation of sufficient amounts of protein for analysis. Isolation of enough material for analysis becomes less difficult if an in vitro system can be used to generate the phosphopeptides in question. Protein kinases are notoriously promiscuous in vitro, so it may be possible to generate the sites of interest using a kinase that is actually not the one responsible for the phosphorylation in vivo. The protein of interest produced in bacteria makes a good substrate, as it is most likely not phosphorylated to start with. It is first necessary to show that the phosphopeptides generated by incubation of recombinant protein with a purified kinase in vitro comigrate exactly with those purified from labeled cells. If there is any doubt about this, it is reassuring to run the maps for a longer time than usual in the electrophoresis dimension, or at a different pH to further demonstrate comigration. Given the specificity of the two-dimensional separation on these maps, it is unlikely that two phosphopeptides that truly comigrate will not be identical.

Anticipated Results Upon developing the first autoradiogram of an initial peptide map, depending on the number of spots seen, the researcher will most likely be left wondering what it all means. There are several things to keep in mind when studying the pattern of spots on a peptide map. First, migration in the electrophoresis dimension is a function of the charge to mass ratio of a peptide. Migration in the chromatography dimension is related to the hydrophobicity of a peptide. The more hydrophobic a peptide, the further it migrates in the chromatographic dimension. In general, most phosphopeptides of the protein studied will be represented by one spot in the map. It is possible then to determine the relative stoichiometry of phosphorylation at different sites by comparing the intensity of each spot on the autoradiogram. This, however, relies on the



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Figure 18.9.6 An example of a tryptic phosphopeptide map based on that of human Nck-alpha. For the first (horizontal) dimension, electrophoresis was run at pH 1.9 for 25 min at 1.0 kV; the anode is at the left. Ascending chromatography was run for 15 hr in phosphochromo buffer. The sites represented by spots 1 to 7, with the exception of spot 2, have been identified. Spot 1 is an 11-amino-acid, phosphotyrosine-containing peptide. While spot 2 also contains phosphotyrosine, and runs in a position likely to be the doubly phosphorylated version of this peptide, it turns out to be unrelated to spot 1. Spot 3 represents a 5-amino-acid, phosphoserine-containing peptide; this same peptide with an amino-terminal arginine runs as spot 4; thus it can be seen that in this case the tryptic cleavage is largely incomplete. Spot 5 represents a 20-amino-acid phosphoserine-containing peptide which, with an amino-terminal lysine, runs as spot 6. Spot 7 represents a peptide that is unrelated to 5 and 6. Spot 8 represents free phosphate, released during sample preparation.


assumption that all tryptic peptides are recovered during the the entire protocol with similar efficiencies. There are several cases, however, in which this one phosphopeptide:one spot rule does not hold. One such case is when the enzyme used to digest the protein of interest has not worked to completion, yielding both partial and complete digestion products seen in the map. Multiple digestion products are also generated at sites where a run of basic residues is present. Trypsin works very efficiently as an endopeptidase, hydrolysing peptide bonds following basic residues. In contrast, trypsin works poorly as an exopeptidase. Trypsin will

cleave randomly within the run of basic residues and is unable to take off additional basic residues that may have been left. This results in a series of digestion products differing by the number of basic residues at their amino- or carboxy-terminus. Addition of such a basic residue to a phosphopeptide changes its migration in both the electrophoresis and the chromatography dimension (it is more positively charged and runs further towards the cathode, it is also more hydrophilic and as a consequence runs less far relative to the buffer front in phosphochromo buffer). Similarly, a given peptide will migrate differently if phosphorylated at two positions



rather than one. Addition of another negative charge will again make the peptide more hydrophilic and thus it will migrate less far in the chromatography dimension. This time, however, it will migrate further towards the anode in the electrophoresis dimension, giving a diagonal pattern descending in the opposite direction to that seen with the addition of a lysine or an arginine. It is important to remember that the electrophoretic mobility of a peptide is dependent on its mass. For larger peptides, the slope of the diagonal seen with the addition of either positive or negative charges will be steeper, since the addition of another charge when divided by the mass will make less of a difference to the distance the peptide travels. While trypsin has been traditionally used in peptide mapping, it may not be the enzyme of choice for proteins phosphorylated by PKA, PKC, or other protein kinases whose recognition sequence involves multiple arginines or lysines, as trypsin often fails to cleave after all such residues when they are present in runs. In addition, trypsin cleaves inefficiently at arginines or lysines two residues amino-terminal of a phosphoserine or phosphothreonine (i.e., R/K-X-P.Ser). Sometimes an individual peptide may appear to have an electrophoretic partner that migrates directly above or below it in the chromatographic dimension. This sort of pattern may be observed as the result of two different scenarios: (1) it may be the result of incomplete oxidation of the peptide if it contains a methionine residue (in which case the lower spot is the oxidized form), or (2) it may be the result of methylation of the peptide running in the lower position. Such a methylation may occur during the performic acid oxidation and is dependent on the 1.5-ml microcentrifuge tubes being used. Historically we have found that certain tubes are more apt than others to produce such unwanted side reactions; for this reason it is advisable to stock certain lots of tubes that do not produce such artifacts in the final maps. Similarly, brands and batches of tubes appear to differ in the extent to which peptides “stick” to them during the final steps of the protocol. An exemplary tryptic phosphopeptide map based on that of a real protein (Nck) is shown in Figure 18.9.6. This map illustrates the points mentioned above. Perhaps most importantly, it also illustrates the fact that just because two spots appear to be on a diagonal it is not a foregone conclusion that they are related. Although peptides 1 and 2 appear to represent the

singly and doubly phosphorylated forms of a single tryptic peptide, in this case it turned out that they represent two completely different peptides. Peptides 3 and 4 and 5 and 6, respectively, represent two sets of peptides that are related and differ only by the addition of a basic residue. Spot 8 represents free phosphate, liberated by hydrolysis of phosphoester bonds that has occurred during sample preparation. It is useful to both compare the amount of free phosphate generated in different samples and to use the phosphate spot as another standard marker when comparing peptide mobilities on different plates.

Time Considerations To generate a two-dimensional phosphopeptide map, at least 9 days will elapse from the time the 32P label is added to the cells until the autoradiogram of the map is in hand. The typical researcher, intrigued by one or more particular spots that appear or disappear from such maps depending on how the cells or samples were treated, may rush to attempt to identify the phosphorylation site represented by such spot(s). Please be advised that this will take at least 4 months of hard work and effort, assuming that everything goes well. There are several different strategies to follow, which are outlined throughout the unit (especially see Background Information). Choice of a particular course will depend on the reagents and the equipment available for analysis.

Literature Cited Boyle, W.J., van der Geer P., and Hunter, T. 1991. Phosphopeptide mapping and phosphoamino acid analysis by two-dimensional separation on thin-layer cellulose plates. Methods Enzymol. 201:110-148. Fischer, W.H., Karr, D., Jackson, B., Park, M., and Vale, W. 1991. Microsequence analysis of proteins purified by gel electrophoresis. Methods Neurosci. 6:69-84. Fischer, W.H., Hoeger, C.A., Meisenhelder, J. Hunter, T., and Craig, A.G. 1997. Determination of phosphorylation sites in peptides and proteins employing a volatile Edman reagent. J. Protein Chem. 16:329-333. Mitchelhill, K.I., Michell, B.J., House, C.M., Stapleton, D., Dyck, J., Gamble, J., Ullrich, C., Witters, L.A., and Kemp, B.E. 1997. Posttranslational modifications of the 5′-AMP-activated protein kinase β1 subunit. J. Biol. Chem. 272:24475-24479. van der Geer, P. and Hunter, T. 1990. Identification of tyrosine 706 in the kinase insert as the major colony-stimulated factor 1 (CSF-1)–stimulated autophosphorylation site in the CSF-1 receptor



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in a murine macrophage cell line. Mol. Cell. Biol. 10:2991-3002. Wang, Y.K., Liao, P.-C., Allison, J., Gage, D.A., Andrews, P.C., Lubman, D.M., Hanash, S.M., and Strahler, J.R. 1993. Phorbol 12-myristate 13-acetate-induced phosphorylation of op18 in Jurkat T cells. J. Biol. Chem. 268:14269-14277.

Internet Resources http://www.genestream.org This Web site contains a program for calculating the mobility of a peptides of known composition and a program that reads the position of a spot on the actual map and calculates which peptide(s) derived from the protein being mapped could have the mobility of that spot.

Key References Boyle et al., 1991. See above. van der Geer, P., Luo, K. Sefton, B.M., and Hunter, T. 1993. Phosphopeptide mapping and phosphoamino acid analysis on cellulose thin-layer plates. In Protein Phosphorylation; a Practical Approach (D.G. Hardie, ed.) pp. 31-59, IRL Press, Oxford. Both of these papers discuss many of the protocols described in this unit.

Contributed by Jill Meisenhelder and Tony Hunter The Salk Institute for Biological Studies La Jolla, California Peter van der Geer University of California, San Diego La Jolla, California




Use of Protein Phosphatase Inhibitors

UNIT 18.10

Reversible protein phosphorylation is recognized as a major mechanism regulating the physiology of plant and animal cells. Virtually every biochemical process within eukaryotic cells is controlled by the covalent modification of key regulatory proteins. This in turn dictates the cellular response to a variety of physiological and environmental stimuli; errors in signals transduced by phosphoproteins contribute to many human diseases. Thus, defining protein phosphorylation events, and specifically, the phosphoproteins involved, is crucial for obtaining a better understanding of the physiological events that distinguish normal and diseased states. In studying protein phosphorylation, two common experimental problems arise that mandate the use of protein phosphatase inhibitors. The first problem arises when the goal of the experiment is to decipher physiological events regulated by reversible protein phosphorylation but the hormonal stimuli or signaling pathways involved are not known. This situation is further complicated by the fact that many protein phosphorylations are rapid and transient, thereby eluding investigation. One solution is to employ cell-permeable compounds that inhibit cellular protein phosphatases in order to amplify the basal activity of protein kinases. Inhibition of protein phosphatases also prevents the turnover of protein-bound phosphate, thereby enhancing the detection of phosphoproteins by standard techniques such as SDS-PAGE (UNIT 10.2A) and autoradiography (APPENDIX 3A) or phosphorimaging. In other situations, the aim may be to analyze the impact of hormones and other physiological stimuli on the function of a specific phosphoprotein. Such detailed biochemical analysis is only made possible after the protein of interest has been separated from other cellular components that potentially interfere with these studies. In this circumstance, the cells or tissues exposed to hormones must be homogenized in the presence of protein phosphatase inhibitors to preserve the cellular phosphorylation events promoted by the hormones. The goal is to utilize the most effective inhibitors to abolish the activity of protein phosphatases that may dephosphorylate the proteins of interest during the lengthy protein isolation procedures. This unit describes the general or basic protocols for inhibiting the cellular PP1/PP2A activity with okadaic acid (see Basic Protocol 1) as well as in vitro with microcystin-LR (see Basic Protocol 2). Basic Protocols 3 and 4 describe approaches recommended for inhibiting cellular and in vitro activity, respectively, of PP2B/calcineurin. Finally, Basic Protocol 5 outlines a widely utilized strategy for inhibiting protein tyrosine phosphatases. The reader should be aware that these Basic Protocols are deliberately designed to cover the widest range of experimental situations and therefore represent prototypic procedures that can be readily used with many phosphatase inhibitors that are not specifically mentioned in this unit. A variety of very effective inhibitors, particularly those targeting protein serine/threonine phosphatases, are currently commercially available. Some of these compounds are cell-permeable and yet others are better utilized in vitro to suppress phosphatase activity in lysates and/or cell fractions. This unit describes methods for defining physiologically relevant protein phosphorylation events. Thus, in some cases, cells in culture are treated with cell-permeable phosphatase inhibitors to detect and establish the in vivo phosphorylation events. In other cases, the cells are stimulated with known physiological stimuli that modify cellular protein phosphorylation. In either situation, subsequent analyses require that the cells be homogenized in buffers containing phosphatase inhibitors that preserve the protein modifications through the steps of protein purification and analysis. This in turn facilitates the identification of the particular amino acids modified and sets Contributed by Douglas C. Weiser and Shirish Shenolikar Current Protocols in Molecular Biology (2003) 18.10.1-18.10.13 Copyright © 2003 by John Wiley & Sons, Inc.



the stage for more defined molecular approaches, such as site-directed mutagenesis, that will allow the investigator to establish the functional role of individual protein phosphorylation events. PROTEIN SERINE/THREONINE PHOSPHATASES Serines and threonines constitute the major sites of protein phosphorylation in all eukaryotic cells. Varying estimates suggest that even in cells transformed using oncogenic viral protein tyrosine kinases, >97% of the protein-bound phosphate appears on serines and threonines. Thus, it is not surprising that numerous microorganisms produce toxins and natural products that elicit their deleterious effects by inhibiting cellular protein serine/threonine phosphatases (McCluskey et al., 2002). Many of these compounds show some degree of selectivity for subgroups of protein serine/threonine phosphatases and may provide insight into the cellular phosphatases that catalyze protein dephosphorylation events. The specificity of some of these compounds has been determined in vitro as differences in their concentration-response curves for the inhibition of purified protein phosphatases. When used in vivo, the chemical properties of the compounds, many of which are hydrophobic, do not allow accurate estimates of cellular concentration or predict their subcellular distribution. Thus, the in vitro specificity of the phosphatase inhibitors may be lost and other more direct molecular approaches may be required to define the protein phosphatases that regulate specific cellular events. The principal use of phosphatase inhibitors, as described in this unit, is to complement the in vitro use of purified phosphatases (UNIT 18.5) to define the regulatory importance of reversible protein phosphorylation, rather than to identify the physiologically relevant protein phosphatases. Early biochemical studies classified eukaryotic protein serine/threonine phosphatases into two major groups, termed type-1 (PP1) and type-2 protein phosphatases (Shenolikar and Nairn, 1991). The type-2 phosphatases are further separated into three distinct groups, PP2A, PP2B, and PP2C, showing very different profiles of inhibition by the available compounds (Table 18.10.1). Molecular cloning of mammalian phosphatase catalytic subunits further expanded this family of enzymes, which to date includes up to five additional phosphatase catalytic subunits numbered PP3 to PP7. Of these five, all but PP7 share a highly conserved catalytic site, and therefore are similarly inhibited by various compounds. Yet others, such as PP2C or PP7, show complete insensitivity to the known phosphatase inhibitors and there are no currently available tools to inhibit these phosphatases. Table 18.10.1


Commercially Available PP1/PP2A Inhibitorsa

Compound

Phosphatases

Recommended use

Okadaic acid Calyculin A Tautomycin Cantharidin A Microcystin-LR Nodularin Fostreicin

PP2A > PP1 PP1 ≥ PP2A PP1 > PP2A PP2A > PP1 PP1 = PP2A PP1 = PP2A PP2A>>PP1

In vivo In vivo In vivo In vivo In vitro In vitro In vitro

Supplier(s) Alexis Biochemicals, Sigma Alexis Biochemicals, Sigma Biomol Research Laboratories — Alexis Biochemicals, Sigma — A.B. Scientific, Sigma, Alexis Biochemicals

aAll compounds listed in this table also inhibit PP3, PP4, PP5, and PP6. Their shared sensitivity to these compounds results in their inclusion in the category of PP2A-like enzymes. In contrast, these compounds show no significant inhibitory activity towards PP2B, PP2C, and PP7. As the compounds possess little structural homology, several different inhibitors may be used to validate the physiological effects of inhibiting their shared targets, serine/threonine phosphatases.



Table 18.10.2

Compound Cyclosporin Ab FK506 Cypermethrinc Deltamethrin

Commercially Available PP2B (Calcineurin)a Inhibitors

Phosphatases

Recommended use

PP2B PP2B PP2B PP2B

In vivo In vivo In vivo In vitro

Suppliers Alexis Biochemicals, Sigma Alexis Biochemicals Alexis Biochemicals Alexis Biochemicals

aPP2B (calcineurin) is a Ca2+/calmodulin-activated protein serine/threonine phosphatase and, like many Ca2+/calmodulin-activated enzymes, is inhibited by the calcium chelator, EGTA. Inhibition of PP2B activity using

EGTA is not a very viable approach in cells and the compounds listed in this table more effectively suppress cellular PP2B activity. bA number of inactive and partially active compounds related to cyclosporin are available. Rapamycin, which shares common target proteins with FK506, namely FK506-binding proteins or FKBPs, does not inhibit PP2B activity and is used as control for FK506. cCompounds related to cypermethrin, such as resmethrin, show weak or no activity as PP2B inhibitors and can function controls for the pyrethroid inhibitors.

The compounds listed in Table 18.10.1 inhibit several classes of protein serine/threonine phosphatases. Enzymes such as PP3, PP4, PP5, and PP6 share structural homology to PP2A and therefore show similar sensitivity to these compounds. In contrast, PP2B is inhibited by a distinct group of compounds (Table 18.10.2) that show no activity against either PP1 or the PP2A-like phosphatases. PP1/PP2A Classes of Protein Serine/Threonine Phosphatases PP1 and PP2A together account for >90% of protein serine/threonine phosphatase activity in most eukaryotic cells. With the exception of mammalian skeletal muscle, where PP1 predominates, most cells and tissues possess equivalent amounts of PP1 and PP2A. This has made okadaic acid and calyculin A, both potent cell-permeable PP1/PP2A inhibitors, the most widely utilized compounds for studying cellular protein phosphorylation. INHIBITION OF CELLULAR PP1/PP2A ACTIVITY WITH OKADAIC ACID Okadaic acid (OA), a polyether produced by marine dinoflagellates, is concentrated by shellfish and is a common cause of diarrhetic shellfish poisoning in humans. OA is commercially available as a free acid or as a potassium, sodium, or ammonium salt. Prevailing evidence favors the sodium salt of OA as the most readily permeable across cell membranes.

BASIC PROTOCOL 1

Biochemical studies demonstrate that OA inhibits the purified PP2A catalytic subunit with an IC50 value of 0.2 to 1.0 nM (Cohen et al., 1989). This concentration of OA, however, has little effect on PP2A activity in cell lysates, which typically exhibit 30- to 150-fold higher IC50 values than the purified enzyme. Extensive dilution of cell extracts enhances the efficacy of OA as a PP2A inhibitor. This may suggest that OA binds, albeit with low affinity, to other cellular components in cell extracts. This may be more of an issue in intact cells where significantly higher OA concentrations (10 to 100 nM) are required for effective PP2A inhibition. PP1, unlike PP2A, has a higher IC50 value for OA (10 to 100 nM; Cohen et al., 1989), whether assayed with purified enzyme or cell lysates, and the sensitivity of PP1 activity to OA is not modified by dilution of cell lysates. In some cellular studies, OA concentrations between 10 and 50 nM resulted in significant enhancement of protein phosphorylation, presumably reflecting the inhibition of PP2A and other PP2A-like enzymes. In contrast, OA concentrations >10 µM are needed to effectively inhibit cellular PP1 activity.



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This difference in affinity can provide an opportunity to discriminate between the physiological functions of the two major classes of cellular protein serine/threonine phosphatases, namely PP1 and PP2A. Numerous studies suggest that the predominant site of OA accumulation is the lipid bilayer of the plasma membrane (Suganuma et al., 1989; Nam et al., 1990; Namboodiripad and Jennings, 1996). OA transfers slowly from this site to other compartments and this may account for the slow onset of phosphatase inhibition by OA in most cells (up to 90 min) as well as the requirement of 10- to 100-fold higher concentrations of OA to inhibit PP2A activity in intact cells compared to cell lysates. Thus, it is surprising to find that OA can also be washed out of cells, albeit thorough washing can take hours. This reversible behavior may allow investigators to analyze the reversal of phosphorylation events triggered by OA. Duration of OA treatment to elicit maximal changes in cellular protein phosphorylation is largely empirically determined and must be defined for each individual cell type. In general, cells display a maximal response to micromolar concentrations of OA within 10 to 30 min. Lower OA concentrations require significantly longer exposure times (up to 2 hr). Due to their remarkable ability to activate multiple cellular signaling pathways, OA and other cell-permeable phosphatase inhibitors display some cytotoxicity. This is partially alleviated by lowering OA concentrations and reducing exposure times. Thus, OA treatments for several hours or days are not recommended. If longer times are necessary, calyculin A (CA), which inhibits PP1 and PP2A with near equal potency, may be a better substitute. While the literature on calyculin A is less extensive, this compound can be used at much lower concentrations (0.1 to 1.0 µM) than OA to yield similar physiological effects. OA induces errors in DNA repair that are not mediated by phosphatase inhibition, (Nakagama et al., 1997) which suggests the presence of additional targets for OA. This behavior mandates that appropriate controls be used in cellular studies. In this regard, the treatment of cells with vehicle (DMSO) is commonly used as a control for OA, CA, and other phosphatase inhibitors that are dissolved in this solvent. There are, however, a number of commercially available inactive OA analogs that may serve as better controls. The best of these is 1-norokadaone, whose structure closely matches that of OA and maintains some of the non-phosphatase-mediated effects elicited by OA. Structure-function studies that established a critical role for the acidic side-chain of OA in phosphatase inhibition also generated a number of inactive OA analogs in which the critical carboxylate group is esterified (Nashiwaki et al., 1991). Such compounds, like methyl okadaate, show low or no activity in vitro as PP1/PP2A inhibitors. However, these compounds may not be effective negative controls, as they are slowly de-esterified in cells, yielding the active OA. Finally, OA, CA, and several other phosphatase inhibitors show significant activity as tumor promoters in mouse skin and gut (Fujuki et al., 1988; Suganuma et al., 1990). Thus, it is important that the investigator takes extreme care in handling and disposing of solutions containing these compounds.


Materials Cultured mammalian cells grown to 1 hr, as very few physiological events respond to this drug in periods under 30 min.



IN VITRO INHIBITION OF PP2B/CALCINEURIN ACTIVITY WITH CYPERMETHRIN

BASIC PROTOCOL 4

Cypermethrin and other type II pyrethroids are synthetic insecticides that show activity as PP2B inhibitors and are available from many commercial sources. In addition, a number of pyrethroid analogs, resmethrin, allethrin, and permethrin, show reduced or no activity as PP2B inhibitors and can be used as controls. This is particularly important for cellular studies using cypermethrin, as many of the type-II pyrethroids have a wide range of physiological effects including oxidative stress, cytotoxicity, and tumorigenesis. Only a few of these are likely to be mediated by PP2B inhibition. Given the availability of cyclosporin A, a better characterized and more potent PP2B inhibitor, the value of using pyrethroids in mammalian cells is questionable. Materials 1.0 mM cypermethrin (Alexis Biochemicals) in DMSO and store in dark glass containers pretreated with 1% polyethylene glycol (PEG) Sample cell extracts or subcellular fractions NOTE: Cypermethrin and other type II pyrethroids are sold in both solid and liquid form. The liquid form represents cypermethrin dissolved in organic solvents, such as DMSO, acetone, or ethanol. Hydrophobicity of these compounds means that long-term storage in plastic or polyethylene containers should be avoided because these compounds adhere to such surfaces and may be slowly extracted from solution. Many pyrethroids are also light-sensitive. If stored in dark glass containers pretreated with 1% (w/v) PEG to prevent surface adherence, stock solutions of cypermethrin are stable for many months. For cellular studies using cypermethrin, the compound should be added directly to the culture medium as its PP2B inhibitory activity is reduced when made up in fresh medium. While the underlying basis for a time-dependent loss of cypermethrin activity is not clear, this provides a strong argument for using cypermethrin and the pyrethroid inhibitors solely for in vitro studies of PP2B function. 1. Add 1.0 mM cypermethrin (10 to 20 µM final concentration) to cell extracts or subcellular fractions and analyze phosphoproteins within a period not exceeding 60 min. Cypermethrin addition in vitro should result in rapid PP2B inhibition that is evident within minutes. Prolonged incubations with this compound are not recommended.

INHIBITION OF PROTEIN TYROSINE PHOSPHATASES WITH SODIUM ORTHOVANADATE

BASIC PROTOCOL 5

Protein tyrosine phosphatases (PTPases) represent a larger family of enzymes derived from a common ancestral gene and share no structural homology with the protein serine/threonine phosphatases (Sefton and Shenolikar, 1995). Thus, PTPases are not sensitive to any of the compounds described above. PTPases share a common catalytic mechanism that involves the formation of an enzyme phospho-intermediate with the modification of a conserved cysteine within the catalytic cleft. While cysteine-modifying reagents, such as phenyl arsene oxide, and metals such as zinc chloride, potently inhibit PTPases, these reagents are non-specific and likely target many proteins other than PTPases and therefore, will not be further discussed here. Currently, the most useful reagent for inhibiting PTPases in vivo and in vitro is vanadate, a phosphate analog. One might predict that vanadate would also be non-specific and inhibit many phosphotransferases and phosphohydrolases, including protein serine/threonine phosphatases. However, the remarkable chemistry of the vanadium metal



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ion (Gordon, 1991) can be exploited to produce a preferential and effective PTPase inhibitor. Materials Sodium orthovanadate (Sigma) 0.1 M NaOH Sample cells Boiling water bath or heating block Additional reagents and equipment for metabolic labeling (Sefton, 1997) or immunoblotting with an anti-phosphotyrosine antibody (DiGiovanna et al., 1996) 1. Dissolve sodium orthovanadate (200 mM final concentration) in sterile, distilled water (often a yellowish solution) and adjust pH of the solution to 10.0 with 0.1 NaOH. Place solution in a boiling water bath until the solution clarifies or becomes translucent. Readjust pH to 10.0 and store the stock solution in a flint glass container at room temperature or dispense into aliquots in plastic containers and store at −20°C. Sodium orthovanadate, whether solid or in solution at neutral pH, slowly oxidizes and acquires a yellow color. The presence of thiols, often used to maintain protein structure and function following the homogenization of tissues or cells, promotes a rapid (100 µM fostreicin is needed to inhibit the structurally related serine/threonine phosphatase, PP1. Use of fostreicin is, however, limited by its propensity to be inactivated by oxidation and thus must be stored in the presence of a large excess of antioxidants. Commercially available fostreicin, therefore, contains numerous additives, making it very difficult to use as a specific reagent in cellular studies. Nevertheless, it is likely that selective PP1 and PP2A inhibitors can and will be developed (see Key References). Compared to most



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current serine/threonine phosphatase inhibitors, which display cytotoxicity and tumor promoter activity, clinical trials established fostreicin as a potential anti-neoplastic drug. This also hints at the possibility that more selective PP1 or PP2A inhibitors may be better experimental tools than currently available compounds whose cytotoxicity may be a result of inhibiting many different protein serine/threonine phosphatases. On the other hand, it is the remarkable chemistry of the vanadium metal ion that makes the current compounds such effective tools for analyzing highly labile and difficult-to-detect cellular phosphorylation events. Finally, a number of broad-acting or nonspecific reagents have been used to inhibit protein phosphatases. These include sodium fluoride (50 mM) and sodium pyrophosphate or glycerophosphate (10 mM). These compounds suppress protein dephosphorylations in cell lysates and fractions, although not as effectively as other inhibitors. They can serve as alternatives for inhibiting phosphatases in vitro but show no utility in phosphoprotein analysis in intact cells.

Critical Parameters


Treatment of cells with the cell-permeable phosphatase inhibitors described in this unit is often so effective that it saturates and occludes the physiological signaling pathways, making studies that utilize combinations of hormones and phosphatase inhibitors difficult to interpret. Many mammalian proteins are regulated both positively and negatively by phosphorylation, generally occurring at distinct sites. Thus, the treatment of cells with phosphatase inhibitors may facilitate both types of modifications and yield a compound functional readout, which may bear little or no relationship to the physiological mechanisms that orchestrate the orderly control of protein function. Even when utilizing phosphatase inhibitors in vitro, artifactual or post-homogenization phosphorylations can result from inappropriate exposure of proteins to kinases normally located in distinct subcellular compartments and therefore are not normally physiological regulators of these proteins. Thus, this unit stresses the detection of cellular protein phosphorylation and events bearing on the functional role of such modifications. The pharmacological approach to studying cellular protein phosphorylation as described in this unit can be employed by investigators from many backgrounds. However, further op-

timization of the outlined experimental strategy may be necessary for individual cell types, where permeability of the compounds and the array of phosphatases inhibited may dictate unique physiological consequences. Thus, the investigator is encouraged to undertake pilot studies that determine the optimal concentrations of the compound and the duration of cell exposure required to obtain a measurable and meaningful cellular response. Also, as emphasized above, while the cell-permeable phosphatase inhibitors display some target specificity, they cannot unequivocally define the phosphatases that regulate specific phosphoproteins or physiological events. Finally, the investigator should be aware that many of the compounds described above display some capacity to bind proteins other than phosphatases albeit with much lower affinity. Thus, control experiments should utilize compounds, such as the inactive okadaic acid or cyclosporin analogs, that fail to inhibit the target phosphatases. These controls should provide a more compelling argument supporting the biological effects uniquely observed with the active compounds as reflecting the inhibition of the target protein phosphatases. To define the specific phosphatases, the investigator must turn to more molecular approaches that may involve the overexpression of phosphatases (currently a very challenging task), or the endogenous protein inhibitors that show exquisite selectivity for individual phosphatases (Oliver and Shenolikar, 1998). An alternate to overexpression may be the microinjection of phosphatases and the inhibitor proteins, an approach that has been successful in modulating cellular phosphatase activity (Alberts et al., 1993; Mulkey et al., 1994; Morishita et al., 2001). The difficulty of introducing protein inhibitors into cells, whether by overexpression or microinjection, is that most, if not all, of these proteins are tightly regulated by covalent modification. The functions of various inhibitors may be enhanced or dampened by reversible protein phosphorylation, making it difficult to predict their effects on phosphatases. Yet another approach is the use of antisense oligonucleotides or interference RNAs to “knock-down” expression of specific protein phosphatases. While this approach harbors tremendous specificity, it is difficult to control the effectiveness of these inhibitors which lower phosphatase expression, and elicit appropriate physiological responses that cannot be readily predicted. The use of dominantnegative reagents, including inactive phos-



phatase catalytic subunits or fragments of phosphatases, has shown some promise in suppressing phosphatase functions, but these reagents most likely rely on their competition with endogenous phosphatases for regulators rather than substrates, thereby making it difficult to predict the physiological outcome. However, catalytically inactive phosphatases have proven very useful in deciphering the functions of PTPases. These inactive phosphatases function as substrate traps and form stable complexes with cellular phosphoproteins, allowing identification of the potential physiological substrates of the PTPase. Even in this situation, there are serious concerns that overexpression of inactive PTPases may distort the natural specificity of these enzymes and yield spurious or unreliable data. Protein serine/threonine phosphatases themselves also occasionally form stable complexes with cellular substrates and such complexes can be isolated using affinity chromatography based on phosphatase inhibitors, such as microcystin-LR, immobilized on Sepharose or other dextran surfaces. Such affinity matrices are not commercially available but a number of publications describe the synthesis and use of such resins. The reader may refer to Campos et al. (1996) or Damer et al. (1998). However, this approach has thus far been more successful in identifying phosphatase regulators than substrates.

Troubleshooting The use of phosphatase inhibitors as described in this unit is associated with several problematic issues regarding compound stability and permeability. First and foremost, many of these compounds are highly hydrophobic. Some are light-sensitive and others difficult to store for long periods. These factors contribute to the problem of defining their precise concentration in culture media and buffers and predicting their effects on target phosphatases. In cellular studies, there are additional problems in that external inhibitor concentration is not an accurate predictor of the intracellular concentration of compounds, which are not evenly distributed throughout the cell. Thus, the literature reports widely varying IC50 values for inhibition of purified phosphatases and the sensitivity of known PP1 and/or PP2A-regulated events in cells. Thus, the protocols described in this unit deliberately err on the side of using excess phosphatase inhibitors. While this ensures effective phosphatase inhibition, additional problems may arise in terms of the cytotoxicity elicited by these reagents, which varies

widely among different cell types. Quiescent, confluent and highly differentiated cells appear more susceptible to programmed cell death in response to phosphatase inhibitors such as OA and CA, although the prolonged overactivation of multiple signaling pathways by these inhibitors may also contribute to the cytotoxicity observed in actively growing cells. Thus, investigators are encouraged to determine the lowest concentrations of inhibitors and shortest duration of cell exposure necessary for a consistent and measurable physiological response.

Anticipated Results Phosphatase inhibitors that inhibit the major mammalian protein serine/threonine or tyrosine phosphatases unmask an array of kinase cascades and thereby enhance the phosphorylation of a wide range of proteins. This in turn permits the investigator to detect and analyze novel phosphorylation events. By selecting specific phosphatase inhibitors and restricting their concentrations, the investigator can also inhibit a subset of cellular phosphatases yielding more defined results by not only limiting the panel of phosphoproteins visualized but also the sites modified on the given phosphoprotein. For an example of a specific phosphatase inhibition experiment, refer to Haystead et al. (1989). Functional effects associated with the rapid phosphorylation-dephosphorylation of cellular proteins are more readily visualized in the presence of phosphatase inhibitors. These experiments can establish the importance of protein phosphorylation in physiological events where the precise protein target or phosphorylation event has not been identified. However, readouts such as gene expression that result from increased phosphorylation of nuclear proteins often require long exposure of cells to phosphatase inhibitors and thereby risk the potentially confounding effects of cytotoxicity. Here, a compromise must be reached and inhibitor concentrations lowered to levels that maintain cell viability and still promote the transcription activation of target genes.

Time Considerations A common scenario for the experiments described above is that cells are metabolically labeled using 32P-orthophosphate for periods between 90 and 120 min (Sefton, 1997). Phosphatase inhibitors are then added to the media and the cells incubated for a further 30 min. Following homogenization, the phosphoproteins are analyzed using SDS-PAGE (UNIT 10.2A)



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and autoradiography or phosphorimaging, which adds an additional 2 to 3 hr. Thus, the total time required may be minimally estimated at 6 hr. In the absence of metabolic labeling, SDSPAGE may be followed by electrophoretic transfer and analysis of changes in protein mobility reflecting increased phosphorylation or the modification of specific proteins. Phosphorylation sites may be identified by western immunoblotting using phosphospecific antibodies. These approaches provide different pieces of information but carry similar total time considerations.

Literature Cited Alberts, A.S., Thorburn, A.M., Shenolikar, S., Mumby, M.C., and Feramisco, J.R. 1993. Regulation of cell cycle progression and nuclear affinity of the retinoblastoma protein by protein phosphatases. Proc. Natl. Acad. Sci. U.S.A. 90:388-392. Campos, M., Fadden, P., Alms, G., Qian, Z., and Haystead, T.A. 1996. Identification of protein phosphatase-1-binding proteins by microcystinbiotin affinity chromatography. J. Biol. Chem. 271:28478-28484. Cohen, P.T.W. 1997. Novel protein serine/threonine phosphatases: Variety is the spice of life. Trends Biochem. Sci. 22:245-251. Cohen, P.T. 2002. Protein phosphatase 1- targeted in many directions. J. Cell Sci. 115:241-256. Cohen, P., Klumpp, S., and Schelling, D.L. 1989. An improved procedure for identifying and quantitating protein phosphatases in mammalian tissues. FEBS Lett. 250:596-600. Damer, C.K., Partridge, J., Pearson, W.R., and Haystead, T.A. 1998. Rapid identification of protein phosphatase 1-binding proteins by mixed peptide sequencing and data base searching. Characterization of a novel holoenzymic form of protein phosphatase 1. J. Biol. Chem. 273:2439624405. DiGiovanna, M.P., Roussel, R.R., and Stern, D.F. 1996. Production of antibodies that recognize specific tyrosine-phoshorylated peptides. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, D.W. Specher, and P.T. Wingfield, eds.), pp. 13.6.1-13.6.13. John Wiley & Sons, New York. Fischer, E.H., Charbonneau, H., and Tonks, N.K. 1991. Protein tyrosine phosphatases: A diverse family of intracellular and transmembrane enzymes. Science 253:401-406. Fujuki, H., Suganuma, M., Suguri, H., Yoshizawa, S., Takagi, K., Uda, N., Wakanatsu, K., Yamada, K., Murata, M., and Yasumoto, T. 1988. Diarrhetic shellfish toxin, dinophysistoxin-1, is a potent tumor promoter on mouse skin. Jap. J. Cancer Res. 79:1089-1093. Use of Protein Phosphatase Inhibitors

Goldstein, B.J. 2001. Protein-tyrosine phosphatase 1B (PTP1B): A novel therapeutic target for type

2 diabetes mellitus, obesity and related states of insulin resistance. Curr. Drug Target–Immun. Endocrin. Metab. Disorders 1:265-275. Gordon, J.A. 1991. Use of vanadate as protein-phosphotyrosine phosphatase inhibitor. Methods Enzymol. 201:477-482. Haystead, T.A., Sim, A.T., Carling, D., Honnor, R.C., Tsukitani, Y., Cohen, P., and Hardie, D.G. 1989. Effects of the tumour promoter okadaic acid on intracellular protein phosphorylation and metabolism. Nature 337:78-81. Kiguchi, K., Glesne, D., Chubb, C.H., Fujiki, H., and Huberman, E. 1994. Differential induction of apoptosis in human breast tumor cells by okadaic acid and related inhibitors of protein phosphatases 1 and 2A. Cell Growth & Differentiation 5: 995-1004. Maehama, T., Taylor, G.S., and Dixon, J.E. 2001. PTEN and myotubularin: Novel phosphoinositide phosphatases. Annu. Rev. Biochem. 70:24779. McCluskey, A., Sim, A.T.R., and Sakoff, J.A. 2002. Serine-threonine protein phosphatase inhibitors: Development of potential therapeutic strategies. J. Medicinal Chem. 45:1151-1175. Morimoto, Y., Ohba, T., Kobayashi, S., and Haneli, T. 1997. The protein phosphatase inhibitors okadaic acid and calyculin A induce apoptosis in human osteoblastic cells. Exp. Cell Res. 230:181-186. Morishita, W., Connor, J.H., Xia, H., Quinlan, E.M., Shenolikar, S., and Malenka, R.C. 2001. Regulation of synaptic strength by protein phosphatase 1. Neuron 32:1133-1148. Mulkey, R.M., Endo, S., Shenolikar, S., and Malenka, R.C. 1994. Involvement of a calcineurin/inhibitor-1 phosphatase cascade in hippocampal long-term depression. Nature 369:486-488. Nakagama, H., Kaneko, S., Shima, H., Fukuda, H., Kominami, R., Sugimura, T., and Nagao, M. 1997. Induction of minisatellite mutation in NIH3T3 cells by treatment with the tumor promoter okadaic acid. Proc. Natl. Acad. Sci. U.S.A. 94:10813-10816. Nam, K.Y., Hiro, M., Kimura, S., Fujiki, H., and Imanishi, Y. 1990. Permeability of a non-TPAtype tumor promoter, okadaic acid, through lipid bilayer memebrane. Carcinogenesis 11:11711174. Namboodiripad, A.N. and Jennings, M.L. 1996. Permeability characteristics of erythrocyte membrane to okadaic acid and calyculin A. Am. J. Physiol. 270:C449- C456. Nashiwaki, S., Fujiki, H., Suganuma, M., FuruyaSuguri, H., Matsushima, R., Iida, Y., Ojika, M., Yamada, K., Uemura, D., and Yasumoto, T. 1991. Structure-activity relationship within a series of okadaic acid derivatives. Carcinogenesis 11:1837-1841. Oliver, C.J. and Shenolikar, S. 1998. Physiologic importance of protein phosphatase inhibitors. Frontiers in Biosci. 3:D961-D972.



Schonthal, A.H. 1998. Analyzing gene expression with the use of serine/threonine phosphatase inhibitors. Methods Mol. Biol. 93:35-40. Sefton, B.M. 1997. Labeling cultured cells with 32Pi and preparing cell lysates for immunoprecipitation. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, D.W. Specher, and P.T. Wingfield, eds.), pp. 13.2.1-13.2.8. John Wiley & Sons, New York. Sefton, B.M and Shenolikar, S. 1995. Overview of protein phosphorylatyion. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, D.W. Specher, and P.T. Wingfield, eds.), pp. 13.1.1-13.1.5. John Wiley & Sons, New York. Shenolikar, S. and Nairn, A.C. 1991. Protein phosphatases: Recent progress. Adv. Second Messenger Phosphoprotein Res. 23:1-121. Shenolikar, S. and Brautigan, D.L. 2000. Targeting protein phosphatases: Medicines for the new millenium. Science (STKE) Nov. 2000. http://stke.sciencemag.org/. Suganuma, M., Suttajit, M., Suguri, H., Ojika, M., Yamada, K., and Fujiki, H. 1989. Specific bind-

ing of okadaic acid, a new tumor promoter in mouse skin. FEBS Lett. 250:615-618. Suganuma, M., Fujiki, H., Furuya-Suguri, H., Yoshizawa, S., Yasumoto, S., Kato, Y., Fusetani, N., and Sugimura, T. 1990. Calyculin A, an inhibitor of protein phosphatases, is a potent tumor promoter on CD-1 mouse skin. Cancer Res. 15:3521-3525.

Key References Current Medicinal Chemistry. November, 2002, vol. 9. A special “hot topics” issue, which is devoted to the synthesis of serine/threonine phosphatase inhibitors.

Contributed by Douglas C. Weiser and Shirish Shenolikar Duke University Medical Center Durham, North Carolina



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Design and Use of Analog-Sensitive Protein Kinases

UNIT 18.11

Many protein kinases can be engineered to accept analogs of ATP that are not efficiently used by wild-type kinases. These engineered kinases, which are referred to as “analogsensitive” or “–as” alleles, are also often sensitive to protein kinase inhibitor variants that do not block the activity of nonmutant kinases. Selective in vitro use of radiolabeled ATP analogs by –as kinases can be exploited to identify the direct phosphorylation targets of individual kinases in complex extracts. In organisms in which it is practical to replace wild-type kinase genes with engineered alleles, the in vivo activity of an –as kinase can be reversibly blocked with an allele-specific inhibitor. Thus, analog-sensitive kinases can be effective tools for discovery of the cellular functions and phosphorylation targets of individual enzymes. A theoretical background for the design and use of these alleles is discussed in the Commentary section at the end of this unit, as are strategies for construction of candidate –as alleles of any kinase. In vivo use of analog-sensitive kinase alleles requires access to kinase inhibitor analogs. This unit describes protocols for synthesis of 1NM-PP1 and 1NA-PP1 (see Basic Protocols 1 and 2), which are among the most effective allele-specific inhibitors of –as kinases. These protocols assume knowledge of organic synthesis, and it is advisable to work with a chemist that can provide assistance if one is not familiar with organic chemistry methods. Under no circumstances should one attempt these protocols if not comfortable with safety procedures fundamental to synthetic organic chemistry. Much of the equipment required for these procedures is uncommon in molecular biology groups but readily accessible in chemistry laboratories. The authors have provided information relevant to analysis of NMR spectra for researchers familiar with this technique. A detailed description of product analysis is, however, beyond the scope of this section. The authors have also supplied a protocol for production of γ-32P-labeled N6(benzyl)ATP, an ATP analog that is commonly selectively used by –as kinases, from the ADP analog (see Basic Protocol 3 and Support Protocols 1 and 2). This procedure employs enzymatic phosphorylation of N6(benzyl)ADP by immobilized nucleoside diphosphate kinase (NDPK), and can be used to produce other γ-labeled ATP analogs from ADP analogs (Mourad and Parks, 1966; Kraybill et al., 2002). Commercial availability of N6(benzyl)ADP, as well as other ADP analogs, is still somewhat limited. A method for synthesis of N6(benzyl)ADP has been published (Shah et al., 1997; Shah and Shokat, 2002). Labeling of ADP analogs by the procedure provided here requires moderately large quantities (∼2.5 mCi) of [γ-32P]ATP, and it is therefore advisable to ensure that this raises no issues with one’s license to use radioactive materials. A final protocol (see Basic Protocol 4) describes the assays for testing inhibition of analog-sensitive kinases in yeast. NOTE: Although the protocols described in this unit are directly derived from published material, it is important to note that the use of analog-sensitive kinases, analog-sensitive kinase inhibitors, and ATP analogs is protected by U.S. patents 6,610,483; 6,521,417; 6,390,821 and 6,383,790, as well as patents in other countries, which are owned by Princeton University and exclusively licensed to Cellular Genomics Incorporated (CGI) of Branford, Connecticut. Information regarding accessing rights to make and use the technology can be obtained by contacting CGI. Analysis of Protein Phosphorylation Contributed by Justin Blethrow, Chao Zhang, Kevan M. Shokat, and Eric L. Weiss Current Protocols in Molecular Biology (2004) 18.11.1-18.11.19 Copyright © 2004 by John Wiley & Sons, Inc.


BASIC PROTOCOL 1

SYNTHESIS OF 1NM-PP1 This protocol and the one that follows for 1NA-PP1 (see Basic Protocol 2) are based on previously published methods (Bishop et al., 1998; Bishop et al., 1999). The synthetic schemes are diagrammed in Figure 18.11.1. A representative NMR spectrum is presented in Figure 18.11.2. Materials 1-naphthylacetic acid (Acros Organics) Hexane N,N-dimethylformamide (DMF; Aldrich) Oxalyl chloride (Aldrich) Tetrahydrofuran (THF; Aldrich) Sodium hydride (Aldrich) Malononitrile (Aldrich) 1 N H2SO4 Ethyl acetate MgSO4, anhydrous 1,4-dioxane (Aldrich) Sodium bicarbonate (Aldrich) Dimethyl sulfate (Aldrich) Diethyl ether (Et2O) Silica Gel 60 (pore size 0.040 to 0.063 mm; Merck) Ethanol (Aldrich) Triethylamine (Aldrich) tert-butylhydrazine hydrochloride (Aldrich) Chloroform (CHCl3) 1% (v/v) methanol in CHCl3 Formamide (Aldrich) Activated charcoal Celite Büchi Rotavapor Model R-200 or equivalent rotary evaporator Ice bath Separatory funnel Oil baths, 80°, 100°C, 180°C 10-in. (25.4-cm) length × 2-in. (5.0-cm) i.d. chromatography column TLC plates (Silica Gel F254; EM Science) and tank Reflux condenser Filter paper Buchner funnel Synthesize 1-naphthylacetylmalononitrile 1. Dissolve 3.72 g 1-naphthylacetic acid (20.0 mmol) in 100 ml hexane. 2. Add 146 mg N,N-dimethylformamide (DMF; 2.0 mmol). 3. Add, dropwise, 12.7 g oxalyl chloride (100 mmol). 4. Wait for gas evolution to stop (∼30 min), then remove solvent under reduced pressure in a Büchi Rotavapor or equivalent rotary evaporator. 5. Redissolve the residue (containing the acid chloride) in 30 ml tetrahydrofuran (THF).


6. Suspend 1.06 g sodium hydride (44.0 mmol) in 10 ml THF.



CN O

O

OH

O

CI ii

i

CN O

CN

CN

iii

iv

NH2

NC H2N

vi

N N

N N

N N

1NM-PP1

O

O CI

NH2

O CN iii

ii

CN

CN

v

NC N N

H2N

CN

vi

N N

N N

1NA-PP1

Figure 18.11.1 Synthetic schemes of 1NM-PP1 and 1NA-PP1. Conditions: (i) 5 eq oxalyl chloride, 0.1 eq DMF, room temperature, 1 hr; (ii) 2 eq NaH, 1 eq malononitrile, THF, 0°C to room temperature, 1 hr; (iii) 8 eq NaHCO3, 5 eq dimethyl sulfate, dioxane/water (6:1), reflux, 1 hr; (iv) 2 eq triethylamine, 1 eq tert-butylhydrazine hydrochloride, EtOH, reflux, 1 hr; (v) 1 eq tert-butylhydrazine, DMF, room temperature, 1 hr; (vi) formamide, 180°C, 10 hr.

9

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5

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Figure 18.11.2 NMR data for 1-tert-butyl-3-naphthalen-1ylmethyl-1H-pyrazolo[3,4-d]pyrimidin-4-ylamine (1NMPP1). Compound is white powder; 1H NMR (CDCl3, 400 Mhz) δ 1.82 (s, 9H), 4.73 (s, 2H), 4.87 (br s, 2H), 7.17 (d, J = 7 Hz, 1H), 7.37 (t, J = 8 Hz, 1H), 7.52 (m, 2H), 7.78 (d, J = 8 Hz, 1H), 7.87 (m, 1H), 8.19 (m, 1H), 8.23 (s, 1H); 13C NMR (CDCl3, 100 Mhz) δ 29.2, 32.7, 60.0, 101.1, 123.5, 125.6, 125.8, 126.2, 126.6, 128.2, 128.9, 131.9, 133.9, 134.0, 140.5, 154.5, 154.7, 157.6, HRMS (EI) molecular ion calculated for C20H21N5 331.17993, found 331.17951.


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7. Dissolve 1.45 g malononitrile (22.0 mmol) in 10 ml THF and add dropwise to the NaH/THF suspension (step 6) on an ice bath. 8. Add the THF solution of acid chloride from step 5 dropwise to the malononitrile solution with vigorous stirring. Stir for 1 hr. 9. Add ∼15 ml of 1 N H2SO4 and 50 ml water. 10. Extract three times, each time with 100 ml ethyl acetate, in a separatory funnel. The ethyl acetate phase is the top phase.

11. Combine ethyl acetate extracts and dry by addition of ~5 g anhydrous MgSO4. 12. Evaporate the solvent under reduced pressure in a Büchi Rotavapor or equivalent rotary evaporator, to yield 1-naphthylacetylmalononitrile, a yellow solid. Yield should be ∼70%.

Synthesize 1-naphthylmethyl(methoxy)methylidenemalononitrile 13. Dissolve 2.23 g 1-naphthylacetylmalononitrile (from step 12; 10.0 mmol) in a mixture of 18 ml dioxane and 3 ml water. 14. Add 6.8 g sodium bicarbonate. 15. Add 4.9 ml of dimethyl sulfate with vigorous stirring. Heat the reaction mixture at 100°C on an oil bath for 2 hr. 16. Cool the reaction mixture to room temperature. 17. Dilute with 70 ml water and extract three times, each time with 100 ml Et2O, in a separatory funnel. 18. Combine the Et2O extracts and dry over anhydrous MgSO4. 19. Evaporate the Et2O under reduced pressure in a Büchi Rotavapor or equivalent rotary evaporator to give an oil. 20. Prepare a 10-in. (25.4-cm) × 2-in. (5.0-cm) column containing 200 g Silica Gel 60, saturated with 1:1 Et2O/hexane (the mobile phase). 21. Dissolve the oil from step 19 in a small amount of mobile phase and apply to column. Elute with mobile phase. Sample the eluate as it emerges from the column and analyze by thin-layer chromatography (TLC) using 1:1 Et2O/hexanes as the mobile phase. Pool fractions containing a species with Rf of ∼0.24. 22. Evaporate solvent under reduced pressure in a Büchi Rotavapor or equivalent rotary evaporator to produce a white crystalline solid 1-naphthylmethyl(methoxy)methylidenemalononitrile. Yield should be ∼75%.

Synthesize 5-amino-1-tert-butyl-3-(1′-naphthylmethyl)-4-cyano-1H-pyrazole 23. Dissolve 1.74 g of 1-naphthylmethyl(methoxy)methylidenemalononitrile (from step 22; 7.0 mmol) in 50 ml ethanol. 24. Add 1.51 g triethylamine (15.0 mmol) and 0.92 g tert-butylhydrazine hydrochloride (7.4 mmol). Reflux at 80°C on an oil bath for 1 hr. 25. Cool the mixture and evaporate the solvent under reduced pressure. 26. Suspend the residue in water and extract three times, each time with with 50 ml CHCl3, in a separatory funnel. Design and Use of Analog-Sensitive Protein Kinases

The CHCl3 phase is the top phase.



27. Combine the CHCl3 extracts, dry over anhydrous MgSO4, and evaporate the solvent under reduced pressure in a Büchi Rotavapor or equivalent rotary evaporator, to give a yellow solid. 28. Prepare a 10-in. (25.4-cm) × 2-in. (5.0-cm) column containing 100 g Silica Gel 60, saturated with CHCl3 (the mobile phase). 29. Dissolve the yellow solid in a minimal volume of CHCl3 and apply to the column. Elute with 1% methanol in CHCl3. Sample the eluate as it emerges from the column and analyze by thin-layer chromatography (TLC) using 1% methanol in CHCl3 as the mobile phase. Pool fractions containing a species with Rf of ∼0.1. 30. Evaporate solvent to obtain a white solid, 5-amino-1-tert-butyl-3-(1′-naphthylmethyl)-4-cyano-1H-pyrazole. Yield should be ∼75%.

Synthesize 4-amino-1-tert-butyl-3-(1′-naphthylmethyl)-1H-pyrazolo[3,4-d]pyrimidine (1NM-PP1) 31. Suspend 0.9 g 5-amino-1-tert-butyl-3-(1′-naphthylmethyl)-4-cyano-1H-pyrazole (from step 30; 3.4 mmol) in 20 ml of formamide. 32. Heat this suspension at 180°C on an oil bath, stirring with a reflux condenser for 10 hr. 33. Cool reaction mixture and dilute with 80 ml water. 34. Collect precipitate by filtration on paper in a Buchner funnel. 35. Dissolve precipitate in 40 ml room temperature ethanol. 36. Add 1 g powdered activated charcoal. Boil at 80°C for 10 min. 37. Pour a celite pad on a Buchner funnel using a slurry of celite in ethanol. Filter the hot mixture through this celite pad. Collect the filtrate. 38. Evaporate the solvent to obtain 1NM-PP1 as a white powder. Expect yields of ∼75%.

SYNTHESIS OF 1NA-PP1 1NA-PP1 can be synthesized in the same manner as 1NM-PP1, with two differences. First, the acid chloride (1-naphthoyl chloride) is directly used in the first step of the synthesis, since it is commercially available. Second, a different condition is used in the reaction that forms the pyrazole ring, giving much better yields than the original procedure. A representative NMR spectrum is presented in Figure 18.11.3. Materials 1-naphthoyl chloride (Aldrich) Tetrahydrofuran (THF; Aldrich) Sodium hydride (Aldrich) Malononitrile (Aldrich) 1 N H2SO4 Ethyl acetate MgSO4, anhydrous 1,4-dioxane (Aldrich) Sodium bicarbonate (Aldrich) Dimethyl sulfate (Aldrich)

BASIC PROTOCOL 2



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9

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3

2

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ppm

Figure 18.11.3 NMR data for 1-tert-butyl-3-naphthalen-1-yl-1H-pyrazolo[3,4-d]pyrimidin-4-ylamine (1NA-PP1). Compound is white powder; 1H NMR (CDCl3, 400 Mhz) δ 1.87 (s, 9H), 5.04 (br s, 2H), 7.05 (m, 2H), 7.58 (t, J = 8 Hz, 1H, 7.64 (d, J = 7 Hz, 1H), 7.92 (m, 2H), 7.95 (d, J = 8 Hz, 1H, 8.36 (s, 1H); 13C NMR (CDCl3, 100 Mhz) δ 29.3, 60.6, 101.5, 125.5, 125.6, 126.5, 127.1, 128.4, 128.4, 129.6, 130.6, 131.8, 134.0, 140.4, 153.9, 154.7, 157.7, HRMS (EI) molecular ion calculated for C19H19N5 317.16427, found 317.16247.

Diethyl ether (Et2O) Silica Gel 60 (pore size 0.040 to 0.063 mm; Merck) Sodium ethoxide (Acros Organics) tert-butylhydrazine hydrochloride (Aldrich) N,N-dimethylformamide (DMF; Aldrich) Chloroform (CHCl3) 1% (v/v) methanol in CHCl3 Formamide (Aldrich) Ethanol (Aldrich) Activated charcoal Celite


Separatory funnel Oil baths, 80°, 100°, and 180°C 10-in. (25.4-cm) length × 2-in. (5.0-cm) i.d. and 8-in. (20.3-cm) length × 1.5-in. (3.8-cm) i.d. chromatography columns TLC plates (Silica Gel 60 F254; EM Science) and tank Reflux condenser Filter paper Buchner funnel



Synthesize 1-naphthoylmalononitrile 1. Dissolve 3.80 g (20 mmol) 1-naphthoyl chloride in 30 ml tetrahydrofuran (THF). 2. Dissolve 1.06 g sodium hydride (44.0 mmol) in 10 ml THF. 3. Dissolve 1.45 g malononitrile (22.0 mmol) in 10 ml THF and add dropwise to the NaH/THF suspension on an ice bath. 4. Add the THF solution of 1-naphthoyl chloride from step 1 dropwise to the malononitrile solution with vigorous stirring. Stir for 1 hr. 5. Add ∼15 ml of 1 N H2SO4 and 50 ml water. 6. Extract three times, each time with 100 ml ethyl acetate in separatory funnel. The ethyl acetate phase is the top phase.

7. Combine ethyl acetate extracts and dry by addition of anhydrous MgSO4. 8. Evaporate the solvent to yield 1-naphthoylmalononitrile, a yellow solid. Yield should be approximately 70%.

Synthesize 1-naphthyl(methoxy)methylidenemalononitrile 9. Dissolve 2.20 g 1-naphthoylmalononitrile (from step 8; 10.0 mmol) in a mixture of 18 ml dioxane and 3 ml water. 10. Add 6.8 g sodium bicarbonate. 11. Add 4.9 ml of dimethyl sulfate with vigorous stirring. Heat the reaction mixture at 100°C on an oil bath for 2 hr. 12. Cool the reaction mixture to room temperature. 13. Dilute with 70 ml water and extract three times, each time with 100 ml Et2O. 14. Combine the Et2O extracts and dry over anhydrous MgSO4. 15. Evaporate the Et2O under reduced pressure to give an oil. 16. Prepare a 10-in. (25.4-cm) × 2-in. (5.0-cm) column containing 200 g Silica Gel 60, saturated with 1:1 Et2O/hexane (the mobile phase). 17. Dissolve the oil from step 15 in a small amount of mobile phase and apply to column. Elute with mobile phase. Sample the eluate as it emerges from the column and analyze by thin-layer chromatography (TLC) using 1:1 Et2O/hexanes as the mobile phase. Pool fractions containing a species with Rf of ∼0.22. 18. Evaporate solvent under reduced pressure in a Büchi Rotavapor or equivalent rotary evaporator, to produce a white crystalline solid, 1-naphthyl(methoxy)methylidenemalononitrile. Yield should be ∼75%.

Synthesize 5-amino-1-tert-butyl-3-(1′-naphthyl)-4-cyano-1H-pyrazole 19. Add 0.38 g sodium ethoxide (5.5 mmol) and 0.68 g tert-butylhydrazine hydrochloride (5.5 mmol) to 10 ml DMF. Stir mixture at room temperature for 0.5 hr. 20. Filter the mixture. Save the filtrate, which contains tert-butylhydrazine. 21. Dissolve 1.17 g 1-naphthyl(methoxy)methylidenemalononitrile (from step 18; 5.0 mmol) in 20 ml DMF. Add the tert-butylhydrazine-containing filtrate from step 20. Stir the reaction mixture at room temperature for 1 hr. 22. Evaporate the solvent under reduced pressure.



Supplement 66

23. Suspend the residue in water and extract three times, each time with 50 ml CHCl3, in a separatory funnel. 24. Combine the CHCl3 extracts, dry over anhydrous MgSO4, and evaporate the solvent to give a yellow solid. 25. Prepare an 80-in. (20.3-cm) × 1.5-in. (3.8-cm) column containing 100 g Silica Gel 60, saturated with CHCl3 (the mobile phase). 26. Dissolve the yellow solid in a minimal volume of CHCl3 and apply to the column. Elute with 1% methanol in CHCl3. Sample the eluate as it emerges from the column and analyze by thin-layer chromatography (TLC) using 1% methanol in CHCl3 as the mobile phase. Pool fractions containing a species with Rf of ∼0.1. 27. Evaporate solvent to produce a white solid, 5-amino-1-tert-butyl-3-(1′-naphthyl)-4cyano-1H-pyrazole. Yield should be ∼75%.

Synthesize 4-amino-1-tert-butyl-3-(1′-naphthyl)-1H-pyrazolo[3,4-d]pyrimidine (1NA-PP1) 28. Suspend 0.99 g 5-amino-1-tert-butyl-3-(1′-naphthyl)-4-cyano-1H-pyrazole (3.4 mmol) in 20 ml of formamide. 29. Heat this suspension at 180°C on an oil bath, stirring with a reflux condenser for 10 hr. 30. Cool the reaction mixture and dilute with 80 ml water. 31. Collect the precipitate by filtration on paper in a Buchner funnel. 32. Dissolve precipitate in 40 ml room temperature ethanol. Add 1 g powdered activated charcoal. Boil at 80°C for 10 min. 33. Pour a celite pad on a Buchner funnel using a slurry of celite in ethanol. Filter the hot mixture through this celite pad. Collect the filtrate. 34. Evaporate the solvent to obtain 1NA-PP1 as a white powder. Expect yields of ∼75%. BASIC PROTOCOL 3

PREPARATION OF γ-32P-LABELED N6(BENZYL)ATP NDPK is a ubiquitous enzyme that catalyzes the exchange of γ phosphate between nucleoside triphosphate species and nucleoside diphosphate species by means of a bi-bi-ping-pong mechanism involving a phospho-enzyme intermediate (Weaver, 1962; Mourad and Parks, 1966). In this protocol, 6× His–tagged NDPK is immobilized in the solid phase by metal affinity chromatography, and the column-bound NDPK is equilibrated with [γ -32P]ATP in low magnesium to generate the labeled phospho-enzyme. The column is then washed to remove residual ATP, and a ten-fold molar excess of N6(benzyl)ADP is added in the presence of larger amounts of magnesium. Eluate from this stage contains a mixture of [γ -32P]N6(benzyl)ATP and N6(benzyl) ADP, and can be used directly in kinase reactions. This protocol can also be used to generate γ-35S-labeled nucleoside triphosphates.




Materials 1:1 slurry of cobalt affinity resin (IDA-Co2+-Sepharose) in HBS/0.02% sodium azide (see Support Protocol 1) HEPES buffered saline (HBS): 150 mM NaCl/100 mM HEPES, pH 7.4 Purified NDPK-6×His (see Support Protocol 2) Phosphate-buffered saline (PBS): 150 M NaCl/100 mM sodium phosphate pH 7.4 (see APPENDIX 2 for sodium phosphate buffer) [γ -32P]ATP (∼3000 Ci/mmol or ∼7000 Ci/mmol) 1 mM N 6(benzyl)ADP (Shah et al., 1997; Shah and Shokat, 2002) PBS (see above) containing 5 mM MgCl2 2-mm glass beads (VWR) 1-ml disposable pipet tip Stand and clamp to accommodate 1-ml disposable pipet tip NOTE: All steps are performed at room temperature. Prepare cobalt affinity column 1. Place a 2-mm glass bead inside a 1-ml disposable pipet tip and flick the tip to make sure the bead is firmly lodged near the end. Use scissors to remove the end of the tip where it extends beyond the glass bead. Mount this miniature column in a clamp on a stand. 2. Add 200 µl of 1:1 slurry of cobalt affinity resin in HBS/0.02% sodium azide (prepared as in Support Protocol 1) to the column prepared in step 1. 3. Add 1 ml HBS and allow the buffer to drain until level with the top of the beads. If desired, a 1-ml pipet can be used to drive fluid flow through the beads. If this is done, be careful to avoid introducing a negative relative pressure in the column when removing the pipet, as this can disturb the beads.

Add NDPK-6×His and radiolabeled ATP to column 4. Add ∼165 µg purified NDPK-6×His. If the NDPK is in a buffer other than the recommended HKG buffer (see Support Protocol 2), it must be freed of any chelators or reducing agents, including EGTA, before addition to the column.

5. When this has flowed into the column, follow with 1 ml PBS. If desired, retain the eluate to quantify protein retention. PBS is used in place of HBS in all steps subsequent to addition of NDPK. IDA-Co2+ appears to have a weak affinity for the various nucleoside phosphate species; PBS competitively blocks these interactions. A slight lavender color may be noticed on the initial addition of PBS. This is insoluble cobaltous phosphate. It has not been found to interfere with any subsequent steps, nor has the use of PBS caused any significant loss of enzyme from the column. NDPK-6×His has an A280 extinction coefficient of 1.31 for a 1 mg/ml solution, as calculated using the Gill equation.

6. Dilute 833 pmol [γ -32P]ATP (2.5 mCi at 3000 Ci/mmol, 5.8 mCi at 7000 Ci/mmol) with PBS to contain no greater than 1 mM reducing agent. Commercial [γ -32P]ATP may contain an appreciable quantity of DTT or other reducing agent. This should be diluted so that enzyme is not lost from the column due to reduction of cobalt. The reactions can be scaled for use of smaller quantities of label. Analysis of Protein Phosphorylation


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7. Add the radioisotope to the column and allow it to flow through, driving the flow if necessary. A moderate darkening of the column due to a low level of metal reduction may be observed, and a small amount of reduced metal may be present in the eluate. This should not normally be cause for concern. Small samples of the load and eluate may be taken to quantify label retention on the column by liquid scintillation counting.

8. Wash the column at least twice, each time with 1 ml PBS. The eluate should be sampled to quantify label release in these steps, to ensure that residual [γ -32P]ATP has been cleared from the column.

Add N6(benzyl)ADP to column and elute N6(benzyl)ATP 9. Add 8 µl of 1 mM N6(benzyl)ADP (8 nmol) to 32 µl PBS containing 5 mM MgCl2. This ADP analog and others can be synthesized using published procedures (Shah et al., 1997; Shah and Shokat, 2002).

10. Carefully add this to the column, and then gently force it into the beads until the liquid is again level with the top of the beads. 11. Add 250 µl PBS containing 5 mM MgCl2 and gently force it through the beads until level, collecting the eluate. A small sample of the eluate should be taken for liquid scintillation counting to determine yield. SUPPORT PROTOCOL 1

PREPARATION OF COBALT AFFINITY RESIN (IDA-Co2+-SEPHAROSE) Materials Iminodiacetic acid (IDA)–Sepharose slurry (Sigma) 200 mM cobalt chloride HEPES-buffered saline (HBS): 150 mM NaCl/100 mM HEPES, pH 7.4 HBS (see above) containing 0.02% sodium azide 15-ml conical tubes 1. Place ∼3 ml of IDA-Sepharose bead slurry in a 15-ml conical tube. 2. Wash several times with water, each time by filling the 15-ml conical tube and then decanting. 3. Wash once with 200 mM cobalt chloride. The binding is essentially instantaneous.

4. Wash four times with HBS. 5. Store beads at 4°C in a bead volume equivalent of HBS containing 0.02% sodium azide. SUPPORT PROTOCOL 2


EXPRESSION AND PURIFICATION OF NDPK A 6× His–tagged NDPK from S. cerevisiae is expressed in BL21(DE3) E. coli and purified by metal-affinity chromatography. Materials pJDB1 expression plasmid: available from the Shokat Laboratory ([email protected]) or Weiss Laboratory ([email protected]) E. coli strain BL21(DE3) (Novagen) IPTG Cobalt affinity resin (IDA-Co2+-Sepharose; see Support Protocol 1) HIK200 buffer (see recipe), 4°C



HEK10 buffer (see recipe), 4°C HKG buffer (see recipe), 4°C Liquid nitrogen 30°C shaking incubator 5-ml chromatography column MWCO 15,000 dialysis membrane Additional reagents and equipment for transformation of E. coli (UNIT 1.8), growing bacterial cultures (UNIT 1.2), preparation of bacterial lysates (UNIT 1.7), dialysis (APPENDIX 3C), and determination of protein concentration (UNIT 10.1A) Grow transformed bacteria 1. Transform (UNIT 1.8) the pJDB1 expression plasmid into E. coli strain BL21(DE3). This plasmid is a derivative of pET19b containing the S. cerevisiae gene YNK1 with a C-terminal 6×-His tag. This plasmid confers resistance to kanamycin. Cells carrying this plasmid should be cultured in medium containing 30 ìg/ml kanamycin for the transformation procedure (see UNIT 1.2).

2. Grow transformed strain to mid-log phase (OD600 = 0.5 to 1.0) at 37°C (UNIT 1.2). 3. Add IPTG to the culture for a final concentration of 0.4 mM. 4. Incubate with shaking at 37°C for 3 hr. Prepare bacterial lysate The following steps are performed at 4°C, using chilled buffers. 5. Isolate E. coli cells by centrifugation (UNIT 1.7). 6. Produce clarified lysate by method of choice (UNIT 1.7), ensuring that the salt concentration is not less than 150 mM in the final extract. 7. Prepare a 5-ml column of IDA-Co2+-Sepharose. A 5-ml column is appropriate for a 1-liter-scale expression.

Purify lysate by cobalt affinity chromatography and dialysis 8. Apply the lysate (from step 7) to the column. 9. Wash with 4 column volumes HIK200 buffer. 10. Elute with 2.5 column volumes HEK10 buffer. 11. Dialyze (APPENDIX 3C) against three changes of HKG buffer, each time for 2 hr, using a MWCO 15,000 dialysis membrane. 12. Determine protein concentration by measuring A280 (UNIT 10.1A), divide into aliquots, and freeze using liquid nitrogen. Store at −80°C. NDPK-6×His has an A280 extinction coefficient of 1.31 for a 1 mg/ml solution, as calculated using the Gill equation (also see UNIT 10.1A). The purified protein has a tendency to precipitate in buffers containing quit 221-You have transferred 230379 bytes in 1 files. 221-Total traffic for this session was 231859 bytes in 1 transfers. 221-Thank you for using the FTP service on iubio.bio.indiana.edu. 221 Goodbye.

Figure 19.1.4 Using UNIX FTP to download a file. An anonymous FTP session is established with the molecular biology FTP server at the University of Indiana to download the ClustalW alignment program. The user inputs are shown in boldface.

Although FTP occurs within the UNIX environment, Macintosh and PC users can employ programs that utilize graphical user interfaces (GUI, pronounced “gooey”) to navigate through the UNIX directories on the FTP server. Users need not have any knowledge of UNIX commands to download files; they can instead rely on pop-up menus and the ability to point-and-click their way through the UNIX file structure. The most popular FTP program on the Macintosh platform for FTP sessions is Fetch. Internet Basics for Biologists



Figure 19.1.5 Using Fetch to download a file. An anonymous FTP session is established with the molecular biology FTP server at the University of Indiana (top) to download the ClustalW alignment program (bottom). Notice the difference between this GUI-based program and the UNIX equivalent illustrated in Figure A.3J.4.

A sample Fetch window is shown in Figure 19.1.5 to illustrate the difference between using a GUI-based FTP program and the equivalent UNIX FTP in Figure 19.1.4. In the figure, notice that the Automatic radio button (near the bottom of the second window under the Get File button) is selected, meaning that Fetch will determine the appropriate type of file transfer to perform. This may be manually overridden by selecting either Text or Binary, depending on the nature of the file being transferred. As a rule, text files should be transferred as Text, programs or executables as Binary, and graphic format files such as PICT and TIFF files as Raw Data. THE WORLD WIDE WEB While FTP is of tremendous use in the transfer of files from one computer to another, it does suffer from some limitations. When working with FTP, once a user enters a particular directory, they can only see the names of the directories or files. In order to actually view

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what is within the files, it is necessary to physically download them onto one’s own computer. This inherent drawback led to the development of a number of distributed document delivery systems (DDDS), interactive client-server applications that allowed information to be viewed without having to perform a download. The first generation of DDDS development led to programs like Gopher, which allowed plain text to be viewed directly through a client-server application. From this evolved the most widely known and widely used DDDS, namely the World Wide Web. The Web is an outgrowth of research performed at the European Nuclear Research Council (CERN) in 1989 that was aimed at sharing research data between several locations. That work led to a medium through which text, images, sounds, and videos could be delivered to users on demand, anywhere in the world. Navigation on the World Wide Web Navigation on the Web does not require advance knowledge of the location of the information being sought. Instead, users can navigate by clicking on specific text, buttons, or pictures. These clickable items are collectively known as hyperlinks. Once one of these hyperlinks is clicked, the user is taken to another Web location, which could be at the same site or halfway around the world. Each document displayed on the Web is called a Web page, and all of the related Web pages on a particular server are collectively called a Web site. Navigation strictly through the use of hyperlinks has been nicknamed “Web surfing.” Users can take a more direct approach to finding information by entering a specific address. One of the strengths of the Web is that the programs used to view Web pages (appropriately termed browsers) can be used to visit Gopher and FTP sites as well, somewhat obviating the need for separate Gopher or FTP applications. As such, a unified naming convention was introduced to indicate to the browser program both the location of the remote site and, more importantly, the type of information at that remote location so that the browser could properly display the data. This standard-form address is known as a uniform resource locator (URL), and takes the general form protocol://computer. domain, where protocol specifies the type of site and computer.domain specifies the location (Table 19.1.2). The http used for the protocol in World Wide Web URLs stands for hypertext transfer protocol, the method used in transferring Web files from the host computer to the client. Browsers Browsers, which are used to look at Web pages, are client-server applications that connect to a remote site, download the requested information at that site, display the information on the user’s monitor, then disconnect from the remote host. The information retrieved from the remote host is in a platform-independent format called hypertext markup language (HTML). HTML code is strictly text-based, and any associated graphics or sound for that document exist as separate files in a common format. For example, images Table 19.1.2 Uniform Resource Locator (URL) Format for Each Type of Transfer Protocol

Internet Basics for Biologists

Site

URL format (example)

General form

protocol://computer.domain

FTP site Gopher site

ftp://ftp.ncbi.nlm.nih.gov gopher://gopher.iubio.indiana.edu

Web site

http://www.nhgri.nih.gov



may be stored and transferred in GIF format, developed by CompuServe for the quick and efficient transfer of graphics; while GIF format is most commonly used for graphics, other formats such as JPEG and BMP may also be used. Because of this, a browser can display any Web page on any type of computer, whether it be a Macintosh, IBM-compatible, Linux, or UNIX machine. The text is usually displayed first, then the remaining elements are placed on the page as they are downloaded. With minor exceptions, a given Web page will look the same when the same browser is used on any of the above platforms. The two major players in the area of browser software are Netscape, with their Communicator product, and Microsoft, with Internet Explorer. As with many other areas where multiple software products are available, the choice between Netscape and Internet Explorer comes down to one of personal preference. While the computer literati will debate the fine points of difference between these two packages, for the average user, both packages perform equally well and offer the same types of features, adequately addressing the Web-browser needs of most users. It is worth mentioning that, while the Web is by definition a visually based medium, it is also possible to travel through Web space and view documents without the associated graphics. For users limited to line-by-line terminals, a browser called Lynx is available. Developed at the University of Kansas, Lynx allows users to use their keyboard arrow keys to highlight and select hyperlinks, using their return key the same way that Netscape and Internet Explorer users would click their mouse. Internet versus Intranet The World Wide Web is normally thought of as a way to communicate with people at a distance, but the same infrastructure can be used to connect people within an organization. Such intranets provide an easily accessible repository of relevant information, capitalizing on the simplicity of the Web interface. It also provides another channel for broadcast or confidential communication within the organization. Having an intranet is of particular value when members of an organization are physically separated, whether it be in different buildings or different cities. Intranets are protected in such a way that people who are not on the organization’s network are prohibited from accessing the internal Web pages; additional protections through the use of passwords are also common. Finding Information on the World Wide Web Most people find information on the Web the old fashioned way—by word of mouth either by using lists such as Table 19.1.3 or by simply following hyperlinks put in place by Web authors. Continuously clicking from page to page can be a highly ineffective way of finding information, especially when the information sought is of a very focused nature. One way of finding interesting and relevant Web sites is to consult virtual libraries, which are curated lists of Web resources arranged by subject. Virtual libraries of special interest to biologists include the WWW Virtual Library, maintained by Keith Robison at Harvard and the EBI BioCatalog, based at the European Bioinformatics Institutes. The URLs for these sites can be found in Table 19.1.3. It is also possible to directly search the Web by using search engines such as Alta Vista and Excite, among others. A search engine is simply a specialized program that can perform full-text or keyword searches on databases that catalog Web content. The result of a search is a hyperlinked list of Web sites fitting the search criteria from which the user can visit any or all of the found sites. However, search engines use slightly different methods in compiling their databases. One variation is the attempt to capture most or all of the text of every Web page that the search engine is able to find and catalog (“Web crawling”). Another technique is to catalog only the title of each Web page rather than its

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Table 19.1.3

World Wide Web Sites of Interest

Site

URL

Domain names gTLD-MOU Internet Software Consortium

http://www.gtld-mou.org http://www.isc.org

Electronic mail and newsgroups BIOSCI Newsgroups http://www.bio.net/docs/biosci.FAQ.html Eudora http://www.eudora.com Microsoft Exchange http://www.microsoft.com/exchange/ NewsWatcher ftp://ftp.acns.nwu.edu/pub/newswatcher/ File Transfer Protocol Fetch 3.0/Mac FTP Voyager

http://www.dartmouth.edu/pages/softdev/fetch.html http://ftpvoyager.deerfield.com

Internet access America Online AT&T Bell Atlantic Bell Canada CompuServe MCI Ricochet Telus

http://www.aol.com http://www.att.com/worldnet http://www.bellatlantic.net http://www.bell.ca http://www.compuserve.com http://www.mci.com http://www.ricochet.net http://telus.com

Virtual libraries EBI BioCatalog Amos’ WWW Links Page NAR Database Collection WWW Virtual Library

http://www.ebi.ac.uk/biocat/biocat.html http://www.expasy.ch/alinks.html http://www3.oup.co.uk/nar/Volume_28/Issue_01/introduction/ http://mcb.harvard.edu/BioLinks.html

World Wide Web browsers Internet Explorer http://www.microsoft.com/insider/ie5/default.htm Lynx ftp://ftp2.cc.ukans.edu/pub/lynx Netscape Navigator http://home.netscape.com World Wide Web search engines AltaVista http://www.altavista.com Excite http://www.excite.com HotBot http://hotbot.lycos.com Infoseek http://infoseek.go.com Lycos http://www.lycos.com Northern Light http://www.northernlight.com World Wide Web meta-search engines MetaCrawler http://www.metracrawler.com Savvy Search http://www.savvysearch.com




Table 19.1.4 Number of Hits Returned for Four Defined Search Queries on Some of the More Popular Search and Meta-Search Engines

Search term genetic mapping human genome positional cloning prostate cancer

Search Engine

Meta-search engine

HotBot

Excite

Infoseek

Lycos

MetaCrawler SavvySearch

478 13,231 279 14,044

1,040 34,760 735 53,940

4,326 15,980 1,143 24,376

9,395 19,536 666 33,538

62 42 40 60

58 54 52 57

entire text. A third is to consider words that must appear next to each other or only relatively close to one another. Because of these differences in search-engine algorithms, the results returned by issuing the same query to a number of different search engines can produce wildly different results (Table 19.1.4). It may also be noted from Table 19.1.4 that most of the numbers are exceedingly large, reflecting the overall size of the World Wide Web. Unless a particular search engine ranks its results by relevance (for example, by scoring words in a title higher than words in the body of the Web page), the results obtained may not be particularly useful. It is also necessary to keep in mind that, depending on the indexing scheme that the search engine is using, the found pages may actually no longer exist, leading the user to the dreaded “404 Not Found” error. Compounding this problem is the issue of coverage—the number of Web pages that any given search engine is actually able to survey and analyze. A comprehensive study by Lawrence and Giles (1998) indicates that the coverage provided by any of the search engines studied is both small and highly variable. For example, the HotBot engine produced 57.5% coverage of what was estimated to be the size of the “indexable Web,” while Lycos had only 4.41% coverage, a full order of magnitude less than HotBot. The most important conclusion from this study was the extent of coverage increased as the number of search engines are increased and the results from those individual searches are combined. Combining the results obtained from the six search engines examined in this study produced coverage approaching 100%. To address this point, a new class of search engines called meta-search engines have been developed. These programs will take the user’s query and poll anywhere from five to ten of the “traditional” search engines. The meta-search engine will then collect the results, filter out duplicates, and return a single, annotated list to the user. One big advantage is that the meta-search engines take relevance statistics into account, returning much smaller lists of results. Even though the hit list is substantially smaller, it is much more likely to contain sites that directly address the original query. Since the programs must poll a number of different search engines, searches conducted this way obviously take longer to perform, but the higher degree of confidence in the compiled results for a given query outweighs the extra few minutes (sometimes only seconds) of search time. Reliable and easy-to-use meta-search engines include SavvySearch and MetaCrawler (see Table 19.1.3). LITERATURE CITED Froehlich, F. and Kent, A. 1991. ARPANET, the Defense Data Network, and Internet. In Encyclopedia of Communications. Marcel Dekker, New York. Krol, E. and Klopfenstein, B.C. 1996. The Whole Internet User’s Guide and Catalog. O’Reilly and Associates, Sebastopol, Calif. Kennedy, A.J. 1999. The Internet: Rough Guide 2000. Rough Guides, London.

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Lawrence, S. and Giles, C.L. 1998. Searching the World Wide Web. Science 280:98-100. Rankin, B. 1996. Dr. Bob’s Painless Guide to the Internet and Amazing Things You Can Do with E-mail. No Starch Press, San Francisco. Quarterman, J. 1990. The Matrix: Computer Networks and Conferencing Systems Worldwide. Digital Press, Bedford, Mass.

Contributed by Andreas D. Baxevanis National Human Genome Research Institute, NIH Bethesda, Maryland B.F. Francis Ouellette Centre for Molecular Medicine and Therapeutics University of British Columbia Vancouver, British Columbia




Sequence Databases: Integrated Information Retrieval and Data Submission

UNIT 19.2

UNIT 19.1 discusses general tools for electronic information retrieval, giving only brief glimpses of the databases of interest to molecular biologists. This unit provides a more thorough overview of biomedical information resources, focusing on sequence data, structure information, and the associated literature—and also discusses how nucleotide sequence data gets into the databases in the first place—an often neglected but nevertheless important topic. UNIT 19.3 covers sequence homology searching in detail.

First it is important to appreciate not only the variety of available data but also how rapidly it is expanding (Fig. 19.2.1A). The biomedical literature, as represented by MEDLINE, is growing at a rate of 400,000 articles per year. MEDLINE, produced by the U.S. National Library of Medicine (NLM), currently contains bibliographic information and journal abstracts from more than 11,000,000 publications indexed from ∼4,300 biomedical journals. Genetic and physical mapping data are expanding rapidly, as are protein crystal and NMR structures; DNA and protein sequence data have been growing exponentially for years (Fig. 19.2.1B to D). Even faster growth rates are anticipated for cDNA and concerted genome sequencing as technology advances. In mid-1997 complete sequences of Saccharomyces cerevisiae and seven bacterial genomes were available and the sequencing phase of the Human Genome Project was initiated. By January 2000, there were 24 bacterial and archaeal genomes completed and another 70 in progress. Drosophila melanogaster and Caenorhabditis elegans were both essentially done. Other eukaryotes for which substantial genomic sequencing had been done included human, mouse, Arabidopsis thaliana, rice, corn and the malaria parasite (Plasmodium falciparum). GenBank is a comprehensive repository of sequence data and associated annotation, built and distributed by the National Center for Biotechnology Information (NCBI) at the NLM in Bethesda, Maryland. GenBank is part of an international collaboration with the DNA Database of Japan (DDBJ) in Mishima and the European Molecular Biology Laboratory (EMBL) Data Library at the European Bioinformatics Institute near Cambridge, United Kingdom. Because these partners exchange data daily, what we refer to here as simply “GenBank” actually contains sequences from GenBank, EMBL, and DDBJ. In the 1990s, new types of sequence data became a major part of the database entries. For example, “expressed sequence tags” or ESTs are partial sequences that are derived from automated, “single-pass” sequencing on clones that are randomly selected from cDNA libraries, usually with the intent of surveying expressed genes (Adams et al., 1991; Kahn et al., 1992; Okubo et al., 1992; Waterston et al., 1992). These data present certain analytic challenges (Boguski et al., 1993) that are dealt with in UNIT 19.3, and they also require special procedures for batch submission to sequence databases (see discussion of submitting expressed sequence tags (EST), sequence tag site (STS), or genome survey sequence (GSS) data). Short genomic sequences, including those derived from “exon trapping” or “exon amplification” and genomic survey sequences (Church et al., 1993; Smith et al., 1994) present analytical challenges similar to those for ESTs. There are special procedures for submission of these “GSS” entries, similar to the procedures for EST submission. Large-scale sequencing of complete genomes has required additional methods for submission and has spurred development of new ways to access and display the data. Informatics for Molecular Biologists Contributed by Jane M. Weisemann, Mark S. Boguski, and B.F. Francis Ouellette Current Protocols in Molecular Biology (2000) 19.2.1-19.2.20 Copyright © 2004 by John Wiley & Sons, Inc.


A

12 Articles (×106)

10

C

6 4 2 0

Protein sequences (× 105) Nucleotide sequences (× 106)

B

8

'65 '68

'72

'76

'80

'84

'88

'92

'96 2000

'65 '68

'72

'76

'80

'84

'88

'92

'96 2000

'65 '68

'72

'76

'80

'84

'88

'92

'96 2000

'65 '68

'72

'76

'80 '84 Years

'88

'92

'96 2000

6 5 4 3 2 1 0 6 5 4 3 2 1

D

Structure entries (× 103)

0 14 12 10 8 6 4 2

Figure 19.2.1 The cumulative growth of biomedical research data. (A) Growth of MEDLINE. (B) Growth in the number of nucleotide sequences in GenBank (Benson et al., 2000). (C) Growth in the number of protein sequences in the “nonredundant” set of proteins in GenBank. (D) Growth of protein and nucleic acid three-dimensional structures represented in the Brookhaven Protein Data Bank, April 2000 (http://www.rcsb.org/pdb/). These figures occasionally induce panic or despair in individuals who fear that it is impossible to make sense of so much data; however, modern information retrieval systems such as Entrez (Fig. 19.2.2) have succeeded in integrating this data, reducing redundancy, and providing powerful and convenient user interfaces.

Sequence Databases



Retrieval of records by electronic mail and anonymous FTP are covered in UNIT 19.1, as well as access to databases via the World Wide Web. Database homology searching is described in detail in UNIT 19.3. The present unit focuses on the wide variety of sequence, structure, and bibliographic databases—as well as how they have been integrated (Fig. 19.2.2) into a single, easy-to-use retrieval environment called Entrez, which provides users with “one stop shopping” over the Internet. INTRODUCTION TO ENTREZ Entrez (Fig. 19.2.2) ties together a diverse set of information resources: it accesses nucleotide and protein sequences from a number of databases as well as genome data from the NCBI Genomes division, three-dimensional structures from MMDB (Molecular Modeling Database of NCBI), human disease data from OMIM (Online Mendelian Inheritance in Man), genetic locus data from LocusLink, and bibliographic citations from PubMed (discussed below). Entrez is available through the World Wide Web (http://www.ncbi.nlm.nih.gov/Entrez/) and in a client/server version (available by FTP, see http://ncbi.nlm.nih.gov/Entrez/Network/nentrez.overview.html or ftp://ncbi.nlm.nih. gov/entrez/README_1). In the case of the client/server application, Network Entrez (NetEntrez), the client runs on the user’s local machine and interacts with the server at NCBI. Versions of the client program are available for Macintosh, Microsoft Windows, Unix/X-windows and some other systems. Entrez for the World Wide Web (WebEntrez) works through standard WWW browsers. NetEntrez and WebEntrez have similar func-

OMIM

GenBank EMBL DDBJ dbEST dbSTS dbGSS patent

maps and genomes

PubMed

full-text electronic journals

GenBank nucleotide sequences

3-dimensional structures

protein sequences

taxonomy

Swiss-Prot PIR PRF PDB GenBank EMBL DDBJ patent

Figure 19.2.2 Data sources and links that constitute the Entrez system.

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tionality, but there are differences in the way that they are used. WebEntrez has the advantage of additional links—e.g., to WebBLAST outputs and to full-text articles for some journals. The NetEntrez interface is more intuitive and is often easier to use for complex queries. Entrez can be used in many ways and some common scenarios are presented below. Suppose we have just done a similarity search and have observed a number of promising matches on the “hit list.” The information on this list is often insufficient to allow us to reach even preliminary conclusions about the biological implications of our discovery. Thus, we want to find the publication describing this sequence, which meant, until recently, that we had to use a separate retrieval system to obtain the database record corresponding to the sequence. In this database record we would find the literature citation, and then we would have to take a trip down to the library to actually pull the printed copy of the journal. The “activation energy” for this process is high enough that one sometimes decides it is not worth it for “borderline” matches. Even if our motivation is high enough, perhaps there is no literature citation in the database record, or it involves a journal to which our library does not subscribe. In addition to finding the corresponding publication to confirm our search results, we may also want to retrieve some of the matching sequences and perform additional searches and alignments with these. All together, this is quite an involved process. Entrez substantially lowers the activation energy by combining all of these activities in an easy-to-use system with a mouse-driven, graphical user interface. Entrez establishes all of the requisite links between DNA sequences, the proteins they encode, and the literature that describes their biology. The literature component of Entrez comes from the PubMed database, which includes abstracts. While an abstract is not a substitute for the full text of a paper, one can frequently go an amazingly long way in the analysis and interpretation of sequence homology results by the information contained in an abstract. For an increasing list of journals, the Web version of Entrez also has links to full-text articles. For many human disease genes, there are also links from PubMed records to the description of the disease in the Online Mendelian Inheritance in Man (OMIM) database. The PubMed database of bibliographic information contains citations from MEDLINE and PreMEDLINE (preliminary citations that eventually become MEDLINE records). Entrez not only contains explicit links between different data sources, but also implicit or computed links within the data itself. Daily, the entire set of DNA and protein sequence databases are compared among one another and all significant homologies are computed and stored in the system. Thus, it is possible to instantly retrieve the homologs of any sequence in Entrez with the click of a mouse, without having to manually perform a new database search. Thus Entrez contains within it thousands of answers to questions that have not yet been asked. In one interesting case, this automated system actually “discovered” a significant homology prior to its recognition by a human biologist (Boguski and McEntyre, 1994).

Sequence Databases

In addition to precomputed sequence homologies, Entrez performs a similar operation on PubMed records. It is possible to compute a statistical relationship between two articles based on their frequency of use of significant terms. To indicate what constitutes a significant term, consider two negative examples: i.e., the terms “novel” and “important” are not significant terms because they appear in most scientific papers and thus are not of use in distinguishing one paper from another in terms of specific content. The set of “related articles” (sometimes referred to as “neighbors”) is recomputed daily. This constitutes a very powerful information resource. You can enter Entrez via an author name or keyword and retrieve not only a specific paper but an entire bibliography of related material. You can then branch out (or “neighbor out”) to other areas of the information



space and effortlessly go back and forth between the literature and the sequences. Specific examples have appeared in reviews (Baxevanis et al., 1997; Cockerill, 1994; Harper, 1994; Schuler et al., 1996). Further examples can also be found on the NCBI Coffee Break web site (http://www.ncbi.nlm.nih.gov/Coffeebreak/). These pages describe recent biological discoveries and include tutorials demonstrating the linkage between various components of Entrez and other bioinformatics tools. Finding information on sequence homologs using the Web version of Entrez is made even easier by the connections between Entrez and Web-based BLAST output. For example, it is possible to do a BLAST search using WebBLAST, and the hit list will have hypertext links to protein or nucleotide entries in Entrez. Other features of Entrez include taxonomy-based sequence retrieval; from the Web page for the NCBI Taxonomy Browser you can scan through organism names and phylogenetic trees and select the full set of nucleotide or protein sequences for any species. There are now links in Entrez from sequences and the literature to protein three-dimensional structures in NCBI’s Molecular Modeling Database (MMDB). A three-dimensional structure viewer called Cn3D has also been added to the system. Cn3D can be run as a client/server program or as a “Helper” application integrated into a standard Web browser. Another special viewer is available for the Entrez Genomes division. The genome-level views presented by Entrez are built from sequences of complete chromosomes, from composites of sequence fragments, and from integrated genetic and physical maps. Entrez allows the user to visualize the sequence information at varying levels of detail, either graphically or as text. The chromosome views are tightly linked to the other Entrez component databases, allowing the user to jump effortlessly between maps, sequences, and bibliographic components. DATA SUBMISSION: GENERAL CONSIDERATIONS During the late 1970s, when DNA sequencing technology was still new, the Journal of Biological Chemistry required that any submitted manuscript containing sequence data be accompanied by the actual autoradiograms, so that reviewers could check the accuracy of the sequence data. We have come a long way since then. Currently, journals prefer not to even publish sequence data because it takes up a great deal of space and is much more usable in electronic form. Most journals, however, require proof in the form of a database accession number that the authors of a publication have sent their data to GenBank, EMBL, or DDBJ. Accession numbers are unique identifiers (consisting of one letter followed by five digits or two letters followed by six digits) that serve as confirmation that the data has been submitted and allow the scientific community to retrieve it. Making sequences available by depositing them in the public databases is recognized as an important responsibility of researchers to the community. GenBank, the EMBL Data Library, and the DDBJ are partners in the international database collaboration; authors may submit their sequences to whichever database is most convenient without regard to where papers describing those sequences may be published. Sequences should be submitted to a single site; data submitted to one site are exchanged with the others on a daily basis. Sequences by themselves have little meaning; hence, thorough, accurate biological annotation is very important. One important source of annotation is the published article referring to the sequence; indeed, NCBI has made every effort to link citations to the sequences via MEDLINE and Entrez. However, electronic annotation obtained from authors at the time of submission is becoming even more critical as the amount of data



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grows (Fig. 19.2.1B), because the only realistic way of searching through this much data is by computer. The database staff are very helpful in assisting submitters in the proper annotation of their data. Sequences submitted to any of the sequence databases go through a series of validation checks conducted by the database staff. Accession numbers are returned to the authors, usually within 24 hr, or less, if there are no problems with the submission(s). The accession number should be included in any manuscript that deals with the sequence, preferably in a footnote on the first page of the article (or in the manner prescribed by the individual journal). The full text of the provisional record is passed back to the authors for their review prior to being released. Authors may request that the data be kept confidential until publication. See Internet Resources for URLs and e-mail addresses of the sequence databases. SUBMITTING A SEQUENCE TO THE NUCLEOTIDE DATABASE Submission Methods As mentioned above, a sequence can be submitted to any of the three sequence databases (see discussion of Data Submission: General Considerations; also see Internet Resources). A submission to GenBank is described in this section, but readers should keep in mind that the example could apply to DDBJ or EMBL submissions. Researchers should submit their sequences to whichever database is most convenient. The important point is to only submit to one database. If the researcher has access to the World Wide Web, the easiest way to submit data to GenBank is to use the BankIt form on the NCBI home page (or for DDBJ and EMBL submissions, use Sakura or WebIn, respectively; see Internet Resources). BankIt (http://www.ncbi.nlm.nih.gov/BankIt/) allows one to simply copy and paste most of the submission information and data directly out of one’s word processor or sequence-analysis package into an electronic form. BankIt was designed to handle the most common types of GenBank submissions, such as mRNAs or short genomic records. NCBI also has developed a platform-independent submission program called Sequin, which runs stand-alone or over the network. Sequin is suitable for a wide range of sequence lengths and complexities, including traditional (gene-sized) nucleotide sequences, segmented entries (e.g., genomic sequences of a spliced gene where not all of the intronic sequences have been determined), long (genome-sized) sequences with many annotated features, and sets of related sequences (i.e., population, phylogenetic, or mutation studies of a particular gene, region, or viral genome). It also has a number of built-in validation functions for enhanced quality assurance. The validator checks such things as missing organism information, incorrect coding-region lengths (compared to the attached protein sequence), internal stop codons in coding regions, mismatched amino acids, or nonconsensus splice sites. Sequin can present various views of the data, including GenBank-, FASTA-, and EMBL-formatted files, and a graphical view of the sequence including features and alignments. A sequence editor is built into Sequin; this editor automatically adjusts feature intervals as the sequence is edited. Additional capabilities include sequence-analysis functions—e.g., a coding region translator, an ORF finder, and a function that will replace a sequence with the sequence of the complementary strand and simultaneously adjust the positions of the features.

Sequence Databases

Sequin is available from NCBI by anonymous FTP (UNIT 19.1); see http://www.ncbi. nlm.nih.gov/Sequin/index.html for further details on Sequin, including full detailed documentation.



BankIt is best suited for submitting one or a small number of sequences. Sequin provides more of a data “workbench” environment that can accommodate large as well as small contigs with detailed annotation that can be added over time and which can finally be used to submit data to GenBank. In addition to these methods, NCBI provides streamlined submission procedures for large data sets—e.g., expressed sequence tags (ESTs), sequence-tagged sites (STSs), genome survey sequences (GSSs), and high-throughput genome sequences (HTGSs). NCBI provides a means for interacting with the laboratory information-management systems of genome-sequencing centers to ensure the efficient, timely, and accurate submission of their data. These options are discussed in more detail in subsequent sections (see discussion of Submitting EST, STS, or GSS Data, and Submitting High-Throughput Genome Sequences). Instructions and Tips for Preparing Sequence Submissions The most important item is, of course, the new DNA sequence itself. It should be free of sequencing errors, cloning artifacts, etc.; standard things to check for in this regard are described below. In the special case of “single-pass” sequences such as ESTs, the data often contain ambiguities and sequencing errors; in these cases it is important to note it as “single-pass” data. What does one need to know about a sequence to submit it to GenBank? The sequence record contains references to the article in which the sequence is published, information on the submitters, and information on the source organism. The record also should describe the sequence as fully as possible. Significant regions of the sequence, such as coding region, promoter region, and transcription boundaries, are labeled with defined feature names. These features are detailed in the GenBank Feature Table document, available by anonymous FTP (UNIT 19.1) from ftp://ncbi.nlm.nih.gov in the directory /genbank/docs, or on the Web at http://www.ncbi.nlm.nih.gov/collab/FT/index.html. This list of features (along with their syntax and qualifiers) is somewhat overwhelming, and it is much easier to familiarize oneself with this system by simply looking at some well-done examples of existing GenBank records (see Appendix to this unit). The DEFINITION line (see Appendix to this unit for illustrations of this and other elements of the GenBank record) is a short description of the biological sequence in the record and the organism from which it was derived. The LOCUS field contains the length of the sequence, the type of sequence, and the date that it was released to the public (or re-released following any updates, corrections, or modifications). The ORGANISM field in the SOURCE section contains a complete phylogenetic classification. The REFERENCE section contains one or more citations, the name and address of the submitter, and the date that the sequence was originally submitted to the database. The COMMENT field may be used to include annotations that cannot be accommodated in the FEATURES table. REFERENCEs and COMMENTs that apply to the complete record are referred to as “descriptors.” Whenever possible, annotations should be expressed as “legal” FEATURES, using the proper syntax, because this is the only manner in which they are computer-readable. A more complex example, illustrating detailed annotation of a human breakpoint cluster region, is the GenBank record with the accession number U07000, which can be obtained from Entrez (see Introduction to Entrez, above) or the e-mail server. A few other examples are: accession numbers AF010398-AF010400, a segmented set with three segments (see Appendix to this unit, Example 4); AF018073, a bacterial sequence with multiple genes; and AF015226, a mitochondrial sequence with multiple genes (see Appendix to this unit, Example 3). Informatics for Molecular Biologists


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Further details on submission to the sequence databases can be found on the database Web sites and in Kans and Ouellette (1998). The following are seven important questions that should be asked when preparing a sequence for submission. 1. What organism is the source of this DNA? Taxonomic classification of the organism from which the sequence was isolated is very important when using GenBank to ensure that the correct genetic code is used when translating the DNA to generate the protein product from a coding sequence (CDS) feature (see item 4). If the wrong taxa are used, or if a mitochondrial origin is not indicated, then an erroneous translation will be produced and may go undetected. Correct taxonomic classifications are also important for retrieval purposes via Entrez. Over 60,000 species are represented in GenBank. The ten most common at this time are: Homo sapiens (human), Mus musculus (mouse), Drosophila melanogaster (fruit fly), Caenorhabditis elegans (nematode worm), Arabidopsis thaliana (thale cress), Oryza sativa (rice), Rattus norvegicus (lab rat), Danio rerio (zebrafish), Saccharomyces cerevisiae (baker’s yeast), and Lycopersicon esculentum (tomato). A sequence submission in most cases need only include the genus and species, which the database staff use to obtain the complete classification from NCBI’s Taxonomy Database. If precise information is not available, the researcher should send in the best conjecture as to the genus, species, and/or common name, and the database staff will attempt to determine the proper classification. Because there is great biodiversity among microbial organisms used for research, it is often important to include the strain name or number with the DNA sequence for microbial submissions. These will appear as a qualifier, for example, /strain=“S288c”, and are quite useful when doing sequence comparisons or trying to explain certain sequence conflicts. Strain information is also important in the context of genetic mapping data from the mouse (e.g., /strain=“BALB/c”) and other organisms. 2. Is this a genomic or mRNA (cDNA) sequence? The sequenced molecule is most often DNA, but the annotation should indicate the biological identity of the starting material. For example, cDNA (i.e., an mRNA that was made into a DNA molecule using reverse transcriptase so that it could be cloned) should be annotated in the database as an mRNA. HIV is an interesting case. HIV sequences, as well as those from other retroviruses, may be classified in different ways: as cDNA corresponding to genomic RNA (therefore as “RNA”); as nonintegrated, nonproviral DNA (therefore as single-stranded DNA or “ssDNA”); or as integrated, proviral DNA, (therefore as double-stranded DNA or “dsDNA”). 3. If this is a genomic sequence, are there introns for which the sequence has not been determined?

Sequence Databases

In many cases only parts of a gene have been sequenced—e.g., the exons with only a few nucleotides of flanking intron. In this case it is necessary to create what is known as a “segmented entry.” What this means is that the researcher will create individual submissions for each discrete sequence and indicate the relationships between them—i.e., their linear order. Additionally, if the approximate distances (in base pairs or kilobase pairs) between the segments of a set are known, this information should be included. This will permit the complete gene to be reassembled automatically by software programs. It should be noted that this is an example of a type of entry for which Sequin is the best tool.



4. Is there a coding sequence (CDS) or other RNA product (rRNA or tRNA) on this DNA? If so, what are their positions in the DNA sequence? If this is an mRNA, or if it is a DNA molecule encoding a protein product, it is important to indicate that a coding region is present and to supply its start and stop codons. RNA products also have boundaries or coordinates that should be indicated. An important check that is done with all submissions is to verify that the coordinates submitted correspond with the indicated feature(s). If there are any discrepancies, these are resolved by the database annotation staff in consultation with the submitter. As described above under question 1, a common problem occurs when the taxonomic information is incorrect or incomplete, leading to the use of an improper genetic code. Fifteen different genetic codes are presently in use, as outlined in the GenBank Feature Table Document (also see information on BLAST parameters in the BLAST unit, see UNIT 19.3.) Any codon usage that is not represented in this list must use the translation exception qualifier (/transl_except) to indicate the discrepancy. Sequin has a built-in tool for analyzing open reading frames. The user can set the minimum size of the open reading frames and the genetic code and the “ORF Finder” will graphically display all open reading frames in a sequence. There is also a separate Web version of the ORF Finder (http://www.ncbi.nlm.nih.gov/gorf/gorf.html). 5. If there is a CDS, what is gene name and the name of the encoded protein? If the gene product is an enzyme, what is the EC number? Putting a protein or gene product name on a CDS is the best way of labeling the protein encoded by the DNA. It is possible to add more than one qualifier, but a /product, /gene, and /EC_number (if the product is an enzyme) are quite sufficient. This is the label that will be carried over to the protein databases—e.g., GenPept or Swiss-Prot. Enzyme Commission (EC) numbers can be obtained or verified using the World Wide Web (UNIT 19.1) at http://www.expasy.ch/enzyme/enzyme-searchec.html. 6. Is there information about your sequence for which there does not seem to be an appropriate place on the forms? There are sometimes problems finding the appropriate place to put a given feature in the BankIt or Sequin forms. In such cases, just attach a note to the submission and the database staff will find the correct place or contact you for more information. 7. Is this sequence ready for submission? Has it been tested for the presence of vector contamination, mitochondrial DNA, ribosomal and tRNAs, and repetitive (e.g., Alu) elements? Potential problems can be detected by using the BLAST programs to compare a new sequence against special data sets that represent common “contaminants.” The network-aware version of Sequin will allow you to run BLASTN against vector and mitochondrial databases. The VecScreen system has recently been developed for quickly finding vector contamination in sequences. VecScreen is automatically run on BankIt sequences and it can be run from within Sequin. It can also be used independently from its own web page (http://www.ncbi.nlm.nih.gov/VecScreen/VecScreen.html). Other validation routines are built into both the Sequin and BankIt submission tools. These routines include a comparison of the conceptual translation of a coding region with the submitted amino acid sequence, a check that actual data length matches the given length, a check for correct use of qualifiers on features, and a check for illegal characters. Although the database staff applies a number of



Supplement 51

validation tests to each new sequence, it is more expeditious for authors themselves to catch any problems prior to submission. SUBMITTING AN UPDATE OR CORRECTION TO AN EXISTING GENBANK ENTRY Authors are encouraged to submit updates or corrections to their sequence records. In fact anyone who notices a problem, error or omission in a database record should bring this to the attention of the database staff. It is not uncommon for the authors to forget to notify the databases that a previously confidential sequence may now be released. If an accession number is seen in a published article and the sequence cannot be retrieved, this is almost certainly the case. The databases will release the data if the complete journal citation is supplied, including the full title of the paper, or if a copy of the title page plus the page showing the accession number is faxed to the databases (see Internet Resources for contact information). Update information consisting of a simple revision, such as a citation change or release of information, can be submitted by sending an e-mail message in paragraph form explaining the change. The message must include the accession number of the sequence to be updated along with all update, correction, or publication information. Updates can also be submitted using BankIt or Sequin. See the BankIt or Sequin Web pages for details (see Submitting a Sequence to the Nucleotide Database, Submission Methods). SUBMITTING EST, STS, OR GSS DATA Expressed-sequence tags (ESTs), sequence-tagged sites (STSs), and genome survey sequences (GSSs) are usually submitted to GenBank and the specialized databases dbEST, dbSTS, or dbGSS as batches of dozens to thousands of entries, with a great deal of redundancy in the citation, submitter, and library information. For these data, there are special procedures designed to improve the efficiency of the submission process. A special “tagged flat file” input format is used. Documents describing the format can be obtained by sending a request to [email protected]. They are also available from the dbEST, dbSTS, or dbGSS Web pages at: http://www.ncbi.nlm.nih.gov/dbEST/, http://www. ncbi.nlm.nih.gov/dbSTS/, and http://www.ncbi.nlm.nih.gov/dbGSS/, respectively. Once the data are ready to submit, they should be e-mailed to [email protected]. For large data sets, an account can be set up into which the data can be deposited by FTP. For more information write to [email protected]. SUBMITTING HIGH-THROUGHPUT GENOME SEQUENCES (HTGS) Some groups may find that their needs are not conveniently met by the standard means for data submission—e.g., a genome center that has its own internal information system or a group assembling a very large contig. The staff of NCBI, EBI, or DDBJ are happy to discuss these issues and make special arrangements with such groups so that their data can be conveniently incorporated into the databases. This includes setting up automatic exchange of data, creation of special FTP accounts, and the generation of tools to ensure data exchange in the most useful format. At NCBI, for example, FTP accounts have been set up for all submitting genome sequencing centers and a variety of tools have been created for accelerating the submission of high-throughput genome sequences (HTGS). Sequence Databases

For large sequencing operations that wish to develop an integrated direct submission system, or for those researchers who have a data submission problem that does not seem



to be addressed by the standard tools, e-mail can be sent to [email protected]. The situation should be explained in as much detail as possible and a phone number and postal address should be provided so that the GenBank staff can contact the investigators to discuss their situation. CONCLUSION The Handbook of Nucleic Acid Sequences (Barrell and Clark, 1974) contained all known DNA and RNA sequences available at that time. The book was only about half an inch thick and contained 51 tRNA sequences, 11 5S rRNA sequences, some assorted viral and mRNAs, and 9 DNA sequences—the longest of which was an 89-nucleotide sequence from bacteriophage f1. All together, this “database” amounted to ∼10,000 nucleotides. If the current data in GenBank (5.8 billion nucleotides in 5.6 million entries) were published in a comparable printed form the stack of books would be about the height of Mount McKinley (20,320 feet). The early history of electronic sequence databases is covered in part by Smith (1990). Developments since 1990 can be surveyed by consulting the annual database issues of Nucleic Acids Research (e.g., Benson et al., 2000) and the authors recommend scanning this special issue occasionally to learn about new databases and services. There has been a great proliferation of electronic data resources and a revolution in their delivery to the scientific community via the Internet, as described in UNIT 19.1. This unit has discussed retrieval and submission of sequence data and associated literature as well as access to precomputed homology results via the Entrez system. But the real power of sequences is the possibility of making new and sometimes profound discoveries by computing on the data. The information content of sequences is a fundamental link between genotype and phenotype, and it is possible to infer much about genes and proteins—their functions, structures, evolution, and regulation—by the simple discovery of a sequence homology. For detailed discussion of how to effectively search for homologies in sequence databases, consult the BLAST unit (UNIT 19.3). LITERATURE CITED Adams, M.D., Kelley, J.M., Gocayne, J.D., Dubnick, M., Polymeropoulos, M.H., Xiao, H., Merril, C.R., Wu, A., Olde, B., Moreno, R.F., Kerlavage, A.R., McCombie, W.R., and Venter, J.C. 1991. Complementary DNA sequencing: Expressed sequence tags and human genome project. Science 252:1651-1656. Barrell, B.G. and Clark, B.F.C. 1974. Handbook of Nucleic Acid Sequences. Joynson-Bruvvers, Oxford. Baxevanis, A.D., Boguski, M.S., and Ouellette, B.F.F. 1997. Computational analysis of DNA and protein sequences. In Genome Analysis: A Laboratory Manual (B. Birren, E.D. Green, S. Kapholz, R.M. Myers, and J. Roskams, eds.) pp. 533-586. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Benson, D.A., Karsch-Mizrachi, I., Lipman, D.J., Ostell, J., Rapp, B.A., Wheeler, D.L. 2000. GenBank. Nucl. Acids. Res. 28:15-18. Boguski, M. and McEntyre, J. 1994. I think therefore I publish. Trends Biochem. Sci. 19:71. Boguski, M.S., Lowe, T.M., and Tolstoshev, C.M. 1993. dbEST: Database for “Expressed Sequence Tags”. Nature Genet. 4:332-333. Church, D.M., Stotler, C.J., Rutter, J.L., Murrell, J.R., Trofatter, J.A., and Buckler, A.J. 1993. Isolation of genes from complex sources of mammalian genomic DNA using exon amplification. Nature Genet. 6:98-105. Cockerill, M. 1994. A versatile tool for retrieving molecular sequences. Trends Biochem. Sci. 19:94-96. Harper, R. 1994. Access to DNA and protein databases on the Internet. Current Opin. Biotechnol. 5:4-18. Kahn, A.S., Wilcox, A.S., Polymeropoulos, M.H., Hopkins, J.A., Stevens, T.J., Robinson, M., Orpana, A.K., and Sikela, J.M. 1992. Single pass sequencing and physical and genetic mapping of human brain cDNAs. Nature Genet. 2:180-185.



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Kans, J.A. and Ouellette, B.F.F. 1998. Submitting DNA sequences to the databases. In Bioinformatics: A Practical Guide to the Analysis of Genes and Proteins (A.D. Baxevanis and B.F.F. Ouellette, eds.) pp. 319-353. John Wiley & Sons, New York. Okubo, K., Hori, N., Matoba, R., Niiyama, T., Fukushima, A., Kojima, Y., and Matsubara, K. 1992. Large scale cDNA sequencing for analysis of quantitative and qualitative aspects of gene expression. Nature Genet. 2:173-179. Schuler, G.D., Epstein, J.A., Ohkawa, H., and Kans, J.A. 1996. Entrez: Molecular biology database and retrieval system. Methods Enzymol. 266:141-162. Smith, M.W., Holmsen, A.L., Wei, Y.H., Peterson, M., and Evans, G.A. 1994. Genomic sequence sampling: A strategy to high resolution sequence-based physical mapping of complex genomes. Nature Genet. 7:40-47. Smith, T.F. 1990. The history of the genetic sequence databases. Genomics 6:702-707. Waterston, R., Martin, C., Craxton, M., Huynh, C., Coulson, A., Hillier, L., Durbin, R., Green, P., Shownkeen, R., Halloran, N., Metzstein, M., Hawkins, T., Wilson, R., Berks, M., Du, Z., Thierry-Mieg, J., and Sulston, J. 1992. A survey of expressed genes in Caenorhabditis elegans. Nature Genet. 1:114-123.

INTERNET RESOURCES DNA Data Bank of Japan (DDBJ; Center for Information Biology, National Institute of Genetics), 1111 Yata, Mishima, Shiznoka 411, Japan; Fax 81-559-81-6849. e-mail submissions: [email protected] updates: ddbjupdt@ ddbj.nig.ac.jp information: ddbj@ ddbj.nig.ac.jp home page: http://www.ddbj.nig.ac.jp/ WWW submissions: http://sakura.ddbj.nig.ac.jp/ European Molecular Biology Laboratory (EMBL), EMBL Outstation, European Bioinformatics Institutes (EBI), Welcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SD, United Kingdom. e-mail submissions: [email protected] updates: [email protected] information: [email protected] home page: http://www.ebi.ac.uk WWW submissions: http://www.ebi.ac.uk/Submissions/index.html WebIn: http://www.ebi.ac.uk/embl/Submission/webin.html National Center for Biotechnology Information (NCBI), National Library of Medicine (NLM), Bldg. 38A, Room 8N-803, 8600 Rockville Pike, Bethesda, Maryland 20894; Telephone: 301-496-2475; Fax: 301480-9241. e-mail submissions: [email protected] EST/GSS/STS: [email protected] updates: [email protected] information: [email protected] home page: http://www.ncbi.nlm.nih.gov/ WWW submissions: http://www.ncbi.nlm.nih.gov/Genbank/index.html BankIt: http://www.ncbi.nlm.nih.gov/BankIt/

Contributed by Jane M. Weisemann and Mark S. Boguski National Center for Biotechnology Information Bethesda, Maryland B.F. Francis Ouellette Centre for Molecular Medicine and Therapeutics University of British Columbia Vancouver, Canada

Sequence Databases



APPENDIX: SAMPLE GENBANK RECORDS Note that these examples present data in a format called the “GenBank report.” This file format is available with Sequin or Entrez, but other formats for viewing the data are also available, including the “EMBL” format, FASTA format (see UNIT 19.3), and a graphical view. Example 1: A Human mRNA Sequence Record with the Feature Labels and Other Landmarks Underlined This is a typical record for an mRNA sequence with a coding region (CDS) and gene features. The CDS feature includes /gene, standard_name, and /function qualifiers, which are a few of many qualifiers that could be used. There is also the /translation qualifier with the amino acid sequence for the coding region. The position of the coding region on the nucleotide sequence is given by the interval 731..3616. The record also has a source feature. This feature is required for all GenBank entries. A source feature must at a minimum include an /organism qualifier. There are two citations associated with the record. REFERENCE 1 is the publication in which the sequence is described and the accession number is published. REFERENCE 2 shows the name and address of the submitter of the record and the date of submission. The ORGANISM field contains the name of the organism (the same as that given in the /organism qualifier) and the phylogenetic classification. The DEFINITION gives a brief summary of the identity of the sequence. The typical basic DEFINITION consists of the organism name, gene product, gene symbol in parentheses, molecule type (e.g., mRNA), and a statement about the completeness of the coding region. The LOCUS field gives the length of the sequence in the record, the molecule type (mRNA, DNA, RNA, etc.), and the date the current version of the record was released to the public. Note that GenBank report qualifiers are underlined in Example 1 for ease of identification, but not in actual GenBank reports. LOCUS DEFINITION ACCESSION VERSION KEYWORDS SOURCE ORGANISM

HSU11690 4266 bp mRNA PRI 10-DEC-1994 Human faciogenital dysplasia (FGD1) mRNA, complete cds. U11690 U11690.1 GI:595424 . human. Homo sapiens Eukaryota; Metazoa; Chordata; Craniata; Vertebrata; Mammalia; Eutheria; Primates; Catarrhini; Hominidae; Homo. REFERENCE 1 (bases 1 to 4266) AUTHORS Pasteris,G.N., Cadle,A., Logie,L.J., Porteous,M.E.M., Schwartz,C.E., Stevenson,R.E., Glover,T.W., Wilroy,R.S. and Gorski,J.L. TITLE Isolation and characterization of the faciogenital dysplasia (Aarskog-Scott syndrome) gene: a putative rho/rac guanine nucleotide exchange factor JOURNAL Cell 79, 669-678 (1994) MEDLINE 95042764 REFERENCE 2 (bases 1 to 4266) AUTHORS Pasteris,N.G. TITLE Direct Submission JOURNAL Submitted (29-JUN-1994) Noe G. Pasteris, Pediatric Genetics, University of Michigan, 1150 W. Medical Center Dr., Ann Arbor, MI 48109, USA FEATURES Location/Qualifiers source 1..4266 /organism="Homo sapiens" /db_xref="taxon:9606" /chromosome="X" /map="Xp11.21" /clone="pFCF1.1,pFCF3.85"



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gene CDS

/tissue_type="craniofacial" /clone_lib="lambda ZAP II 16 week human fetal craniofacial cDNA library" /dev_stage="fetal" 731..3616 /gene="FGD1" 731..3616 /gene="FGD1" /standard_name="faciogenital dysplasia" /function="putative guanine nucleotide exchange factor" /codon_start=1 /protein_id="AAA57004.1" /db_xref="GI:595425" /translation="MHGHRAPGGRRAFGARTPGHEPAGAAPPACADSDPGASEPGLLA RRGSGSALGGPLDPQFVGPSDTSLGAAPGHRVLPCGPSPQHHRALRFSYHLEGSQPRP —————sequence omitted for brevity—————————— GAPQDVKAQRSLPLIGFEVGPPEAGERPDRRHVFKITQSHLSWYFSPETEELQRRWMA VLGRAGRGDTFCPGPTLSEDREMEEAPVAALGATAEPPESPQTRDKT" 858 a 1442 c 1203 g 763 t

BASE COUNT ORIGIN 1 cagggctcct tcccgcccgc cgccgccgag acccaggctg aagctgggga ggacggtgga 61 gcccggccca ggccgttctc ctgcccccgc ggctggagca tcgtctggga ctactggctg ———————sequence omitted for brevity———————————4201 ttttttgtgt ttttttgtgt gttttttttt ttttaagaaa aatacagttt atttcaggct 4261 ttacag //

Example 2: A Bacterial Genomic Sequence Record This bacterial genomic sequence contains several genes. The record has three CDS features for two complete protein coding regions (mutS and mutL) and one partial coding region (cotE). There are additional features in the record: ribosome binding sites (RBS), promoter regions (-10_signal)and stem_loop regions. The DEFINITION includes the name of all three genes and indication of the completeness of the coding regions. REFERENCE 2 is the main citation for this entry. REFERENCE 1 cites a paper that pertains only to nucleotides 1 to 81 of the sequence. LOCUS DEFINITION

ACCESSIO VERSION KEYWORDS . SOURCE ORGANISM

REFERENCE AUTHORS TITLE JOURNAL MEDLINE REFERENCE AUTHORS TITLE

Sequence Databases

JOURNAL MEDLINE REFERENCE AUTHORS TITLE

BSU27343 Bacillus mismatch complete U27343 U27343.1

4740 bp DNA BCT 21-NOV-1996 subtilis spore coat protein (cotE) gene, partial cds, and repair recognition proteins (mutS) and (mutL) genes, cds. GI:1002518

Bacillus subtilis. Bacillus subtilis Bacteria; Firmicutes; Bacillus/Clostridium group; Bacillus/Staphylococcus group; Bacillus. 1 (bases 1 to 81) Zheng,L.B., Donovan,W.P., Fitz-James,P.C. and Losick,R. Gene encoding a morphogenic protein required in the assembly of the outer coat of the Bacillus subtilis endospore Genes Dev. 2 (8), 1047-1054 (1988) 89006215 2 (bases 1 to 4740) Ginetti,F., Perego,M., Albertini,A.M. and Galizzi,A. Bacillus subtilis mutS mutL operon: identification, nucleotide sequence and mutagenesis Microbiology 142 (Pt 8), 2021-2029 (1996) 96349107 3 (bases 1 to 4740) Albertini,A.M. Direct Submission



JOURNAL Submitted (22-MAY-1995) Alessandra M. Albertini, Dipartimento di Genetica e Microbiologia, University of Pavia, 207 via Abbiategrasso, Pavia, 27100, Italy FEATURES Location/Qualifiers source 1..4740 /organism="Bacillus subtilis" /strain="168" /db_xref="taxon:1423" /map="170 degrees" /clone="pFG2752, pFG2764, pFG2767" gene 1..57 /gene="cotE" CDS N (IN REF. 5). Informatics


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where SIGNAL represents the extent of a signal sequence (prepeptide), MOD_RES indicates a post-translationally modified residue (sulfatation, in this case), CARBOHYD shows the glycosylation site, DISULFID means that a disulfide bond exists between the two indicated residues (293 and 430), and CONFLICT shows that different papers report differing sequences. PROTEIN STRUCTURE DATABASES The three-dimensional structures of proteins not only define their biological functions, but also hold a key in rational drug design. Traditionally, protein structures were solved at a low-throughput mode. However, recent advances in new technologies, such as synchrotron radiation sources and high-resolution nuclear magnetic resonance (NMR), accelerate the rate of protein structure determination substantially. There is an overwhelming consensus in the structural biology community that protein structures can be solved en masse (an effort called structural genomics), in a similar fashion as for determining DNA sequences, and that impact of this approach can be compared with that of the Human Genome Project. Searching structure databases is becoming more and more popular in molecular biology. The only international repository for the processing and distribution of protein structures is the PDB (Bernstein et al., 1977). Most of the structures in the PDB were determined experimentally by X-ray crystallography (∼82%) and NMR (∼16%). A small number of structures (∼2%) were derived from theoretical models, which may provide approximate structures but may not be accurate. The PDB also contains some structures of chemical ligands and nucleotides. Each PDB entry is represented by a four-character identifier (PDB ID), where the first character is always a number from 0 to 9 (e.g., 1cau, 256b). The PDB can be accessed through the home server (http://www.rcsb.org/pdb/ in the USA) or through one of its mirror sites from around the world, e.g., http://molmod.angis.org.au/pdb/ (Australia) http://www.pdb.ufmg.br (Brazil) http://www.ipc.pku.edu.cn/npdb/ (China) http://pdb.weizmann.ac.il/ (Israel) http://pdb.ccdc.cam.ac.uk/ (United Kingdom). The PDB offers three search methods: search by PDB ID, by SearchLite, and by SearchFields. SearchLite is a simple key-word search that uses, for instance, protein name or author’s name. A search using SearchFields, as an advanced search engine, allows a user to specify features of the protein, such as Enzyme Commission (EC) number, name of binding ligand, range of protein size, range of resolution in the X-ray structure, and secondary structure content. The PDB stores structural information in two formats: the PDB file format (Bernstein et al., 1977) and the macromolecular crystallographic information file (mmCIF) format (Bourne et al., 1997). The PDB file format is still the dominant format used in the protein community. It contains three parts: annotations, coordinates, and connectivities. The connectivity part, which shows chemical connectivities between atoms, is optional. It is listed at the end of the PDB file, beginning the line with the key word CONECT. The coordinate part uses each line for a three-dimensional coordinate of an atom, starting from ATOM (for standard amino acids) or HETATM (for nonstandard groups). The following shows an example of the PDB file format: Protein Databases on the Internet



HEADER OXIDOREDUCTASE (OXYGEN(A)) COMPND GLYCOLATE OXIDASE (E.C.1.1.3.1) ... ATOM 232 N ALA 29 54.035 4.332 ATOM 233 CA ALA 29 52.992 65.356 ATOM 234 C ALA 29 53.519 66.762 ATOM 235 O ALA 29 54.648 67.179 ATOM 236 C BALA 29 52.433 65.340 ... HETATM 3165 O HOH 658 62.480 62.480 CONECT 2837 2838 2854

14-JUN-89

1GOX

1GOX 3 1GOX 4

19.352 19.569 19.309 19.655 20.993

1.00 1.00 1.00 1.00 1.00

23.93 24.74 25.43 25.66 24.54

1GOX 1GOX 1GOX 1GOX 1GOX

374 375 376 377 378

0.000

0.50

65.79

1GOX3170 1GOX3171

Each line shows the atom serial number, atom type, residue type, chain identifier (in case of multichain structure), residue serial number, orthogonal coordinates (three values), occupancy, temperature factor, and segment identifier. The annotation part of the PDB file format contains dozens of possible record types, including: HEADER (name of protein and release date), COMPND (molecular contents of the entry), SOURCE (biological source), AUTHOR (list of contributors), SSBOND (disulfide bonds), SLTBRG (salt bridges), SITE (groups comprising important sites), HET (nonstandard groups or residues [heterogens]), MODRES (modifications to standard residues), SEQRES (primary sequence of backbone residues), HELIX (helical substructures), SHEET (sheet substructures), and REMARK (other information and comments). The PDB allows a user to view a molecule structure interactively through a Virtual Reality Modeling Language (VRML) viewer, RasMol (Sayle and Milner-White, 1995), Chime, or QuickPDB (a Java applet for viewing sequence and structure) when the browser is configured to support these free rendering tools. The PDB provides related information about the protein, such as secondary structure assignment and geometry. Each PDB entry also links to a wide range of annotations from secondary databases, including (1) summary and display databases such as Graphical Representation and Analysis of Structure Server (GRASS; Nayal et al., 1999), Image Library (Shnel, 1996), Molecular Modelling Database (MMDB; Marchler-Bauer et al., 1999) in Entrez, PDBsum (Laskowski et al., 1997), and Sequence to and within Graphics (STING); (2) domain partition information from 3Dee (Siddiqui and Barton, 1995); (3) the MEDLINE bibliography; (4) structure quality assessment in PDBREPORT from WHAT IF (Vriend, 1990); (5) protein movements recorded in Database of Macromolecular Movement (MolMovDB; Gerstein and Krebs, 1998); and (6) structure families (CATH, CE, FSSP, SCOP, and VAST, as discussed later in this unit). Several structure databases that are not linked by the PDB can also provide useful information. WPDB (Shindyalov and Bourne, 1995) can be used to visualize and analyze a PDB entry from Microsoft Windows. BioMagResBank (University of Wisconsin, 1999) is a repository for NMR spectroscopy data on proteins, peptides, and nucleic acids. Particularly, it provides partial NMR data (e.g., chemical shifts) before the full structure can be solved. PROTEIN FAMILY DATABASES Proteins can be classified according to their evolutionary, structural, or functional relationships. A protein in the context of its family is much more informative than the single protein itself. For example, residues conserved across the family often indicate special functional roles. Two proteins classified in the same functional family may suggest that they share similar structures, even when their sequences do not have significant similarity. Informatics


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There is no unique way to classify proteins into families. Boundaries between different families may be subjective. The choice of classification system depends in part on the problem; in general, the authors suggest looking into classification systems from different databases and comparing them. Three types of classification methods are widely adopted, based upon either sequence similarity, structure, or function. Sequence-based methods are applicable to any proteins whose sequences are known, while structure-based methods are limited to the proteins of known structures, and function-based methods depend on the functions of proteins being annotated. Sequence- and structure-based classifications can be automated and are scalable to high-throughput data, whereas function-based classification is typically carried out manually. Structure- and function-based methods are more reliable, while sequence-based methods may result in a false positive result when sequence similarity is weak (i.e., two proteins are classified into one family by chance rather than by any biological significance). In addition, since protein structure and function are better conserved than sequence, two proteins having similar structures or similar functions may not be identified through sequence-based methods. Databases for Sequence-Based Protein Families Sequence-based protein families are classified according to a profile derived from a multiple-sequence alignment. The profile can be shown across a long domain (typically 100 residues or more) or can be revealed in short sequence motifs. Classification methods based on profiles across long domains tend to be more reliable but less sensitive than those based on short sequence motifs. Several sequence-based methods focus more on profiles across long domains, including Pfam (Bateman et al., 1999), ProDom (Corpet et al., 1999), SBASE (Murvai et al., 1999), and Clusters of Orthologous Group (COG; Tatusov et al., 1997). These methods differ in the techniques used to construct families. Pfam builds multiple-sequence alignments of many common protein domains using hidden Markov models. The ProDom protein domain database consists of homologous domains based on recursive PSI-BLAST searches (UNIT 19.3). SBASE is organized through BLAST neighbors and is grouped by standard protein names that designate various functional and structural domains of protein sequences. COG aims toward finding ancient conserved domains by delineating families of orthologs across a wide phylogenetic range. The following shows an example of Pfam for the GRIP domain (accession number PF01465). Pfam lists some useful information for the entry as follows: The GRIP (golgin-97, RanBP2alpha, Imh1p and p230/golgin-245) domain is found in many large coiled-coil proteins. It has been shown to be sufficient for targeting to the Golgi. The GRIP domain contains a completely conserved tyrosine residue.

The references of the above annotation are also given. In addition, Pfam gives the alignment between the family members: 015045/1511-1558 YNP9_CAEEL/633-681 Q06704/864-909 Q92805/691-737 O42657/703-748 O70365/1161-1205 Q21071/692-741D Q18013/574-623

Protein Databases on the Internet

SAANLEYLKNVLLQFIFLKPG—SERERLLPVINTMLQLSPEEKGKLAAV O15045 NEKNMEYLKNVFVQFLKPESVP-AERDQLVIVLQRVLHLSPKEVEILKAA P34562 KNEKIAYIKNVLLGFLEHKE——QRNQLLPVISMLLQLDSTDEKRLVMS Q06704 REINFEYLKHVVLKFMSCRES—-EAFHLIKAVSVLLNFSQEEENMLKET Q92805 MLIDKEYTRNILFQFLEQRD——RRPEIVNLLSILLDLSEEQKQKLLSV O42657 EPTEFEYLRKVMFEYMMGR——-ETKTMAKVITTVLKFPDDQAQKILER O70365 PAEAEYLRNVLYRYMTNRESLGKESVTLARVIGTVARFDESQMKNVISS Q21071 STSEIDYLRNIFTQFLHSMGSPNAASKAILKAMGSVLKVPMAEMKIIDKK Q18013

The alignment shows accession numbers and the range of each sequence. One can identify some features of the family through this pattern (i.e., from particularly conserved residues at specific alignment positions).



Some methods are based on “fingerprints” of small conserved motifs in sequences, as with PROSITE (Hofmann et al., 1999), PRINTS (Attwood et al., 1999), and BLOCKS (Heniko et al., 1999). In protein sequence families, some regions have been better conserved than others during evolution. These regions are generally important for the function of a protein or for the maintenance of its three-dimensional structure, and hence are suitable for fingerprinting. The fingerprints can be used to assign a newly sequenced protein to a specific family. Fingerprints are derived from gapped alignments in PROSITE and PRINTS, but are derived from ungapped alignments (corresponding to the highly conserved regions in proteins) in BLOCKS. A fingerprint in PRINTS may contain several motifs from PROSITE, and thus may be more flexible and powerful than a single PROSITE motif. Therefore, PRINTS can provide a useful adjunct to PROSITE. It should be noted that some functionally unrelated proteins may be classified together due to chance matches in short motifs. Other sequence-based protein family databases consist of multiple sources. The ProClass database (Wu et al., 1999) is a nonredundant protein database organized according to family relationships as defined collectively by PROSITE patterns and PIR superfamilies. The MEGACLASS server (States et al., 1993) provides classifications by different methods, including Pfam, BLOCKS, PRINTS, ProDom, and SBASE. The MOTIF search engine at http://www.motif.genome.ad.jp/ includes PROSITE, BLOCKS, ProDom, and PRINTS. Databases for Structure-Based Protein Families The hierarchical relationship among proteins can be clearly revealed in structures through structure-structure comparison. Structure families often provide more information on the relationship between proteins than what sequence families can offer, particularly when two proteins share a similar structure but no significant sequence identity. Figure 19.4.2 shows an example of a structure-structure alignment between two proteins. Sometimes, sequence similarity between two proteins exists but is not strong enough to produce an unambiguous alignment. In this case, the alignment between two structures can generate better alignment in terms of biological significance, and thus may pinpoint the active sites more accurately. Different structure-structure comparison methods yield different structure families. CATH (Class, Architecture, Topology and Homologous superfamily; Orengo et al., 1997) is a hierarchical classification of protein domain structures. CE (Combinatorial Extension of the optimal path; Shindyalov and Bourne, 1998) provides structural neighbors of the PDB entries with structure-structure alignments and three-dimensional superpositions. FSSP (Fold classification based on Structure-Structure alignment of Proteins; Holm and Sander, 1996) features a protein family tree and a domain dictionary, in addition to whole-chain-based classification, sequence neighbors, and multiple structure alignments. SCOP (Structural Classification of Proteins; Murzin et al., 1995) uses augmented manual classification, class, fold, superfamily, and family classification. VAST (Vector Alignment Search Tool; Gibrat et al., 1996) contains representative structure alignments and threedimensional superpositions. Among these five databases, SCOP provides more functionrelated information. However, due to the manual work involved, SCOP is not updated as frequently as the others (as of October, 1999, it was last updated August 1, 1998), whereas FSSP and CATH follow the PDB updates closely. SCOP is used here as an example to show the features of structure-based families. SCOP can be accessed through its home server in the UK (http://scop.mrc-lmb.cam.ac.uk/scop/). It is also widely mirrored around the world: Informatics


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Figure 19.4.2 Structure superposition between glycolate oxidase (1gox, in black) and inosine monophosphate dehydrogenase (1ak5, in gray). The sequence identity between the two proteins is 17%, and the root mean square deviation between the two structures is 2.5 Å for the Cα atoms. The structure superposition is obtained through VAST and the figure is made using MOLSCRIPT (Kraulis, 1991).

http://pdb.wehi.edu.au/scop/ (Australia) http://www.ipc.pku.edu.cn/scop/ (China) http://pdb.weizmann.ac.il/scop/ (Israel) http://phys.protres.ru/scop/ (Russia) http://scop.stanford.edu/scop/ (USA) However, only the home server has the BLAST sequence-similarity search capacity. SCOP describes the hierarchical relationship among proteins through the major levels of (homologous) family, superfamily, and fold. Proteins are clustered together into a (homologous) family if they have significant sequence similarity. Different families that have low sequence similarity but whose structural and functional features suggest a common evolutionary origin are placed together in a superfamily. Different superfamilies are categorized into a fold if they have the same major secondary structures in the same arrangement and with the same topological connections (the peripheral elements of secondary structure and turn regions may differ in size and conformation). Two superfamilies in the same fold may not have a common evolutionary origin. Their structural similarities may arise from the physics and chemistry of proteins favoring certain packing arrangements and chain topologies (Murzin et al., 1995). Figure 19.4.3 shows the SCOP interface using an example of protein 1gox in the PDB. Protein Databases on the Internet



Figure 19.4.3 An example of the SCOP interface when searching the structure of 1gox in the PDB. By clicking the boxes followed by the entry name, one can obtain pictures of the protein in JPEG format, interactive graphics within RasMol or Chime, and a link to Entrez. The neighbors of 1gox can be explored at different levels by clicking the links to family, superfamily, and fold.

Databases for Function-Based Protein Families There are various protein functional families classified from different perspectives. The ENZYME data bank (Bairoch, 1993) contains the following data for each enzyme: EC number, recommended name, alternative names, catalytic activity, cofactors, pointers to the SwissProt entry, and pointers to any disease associated with a deficiency of the enzyme. PROCAT is a database of three-dimensional enzyme active site templates (Wallace et al., 1996). PDD (Protein Disease Database; Lemkin et al., 1995; Merril et al., 1995) correlates diseases with proteins observable in serum, urine, and other common human body fluids based on biomedical literature. There is also a growing number of databases dedicated to special types of proteins, such as antibodies, G-protein-coupled receptors, HIV proteases, glycoproteins, and RNases, as shown in Table 19.4.1. OTHER DATABASES Protein Binding Databases Protein binding includes protein-substrate docking and protein-protein association. ReLiBase (Hendlich, 1998) is a database system for analyzing receptor-ligand complexes in the PDB. DIP (Database of Interacting Proteins) records protein pairs that are known Informatics


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to bind with each other. The information in DIP may provide information related to signaling pathways, multiple interactions, and complex systems. Protein Energetics Databases There are few databases for protein energetics, due to the low-throughput nature of the data source. One useful energetics database can be found in ProTherm (Thermodynamic Database for Proteins and Mutants; Gromiha et al., 1999). It contains thermodynamic data on mutations, including Gibbs free energy, enthalpy, heat capacity, and transition temperature. These data are important for understanding the structure and stability of proteins. Bibliographic Databases Searching for protein information through traditional bibliographic databases, such as MEDLINE or Grateful Med, can be rewarding. In addition, some bibliographic reference databases dedicated to proteins may provide certain information more directly. For example, SeqAnalRef stores papers dealing with sequence analysis. COMBINED DATABASES By integrating different types of protein databases together, a database of databases (or a data warehouse) can be built. Such combined databases not only serve as “one-stop shopping,” but also provide cross-references between entries in different databases. Two combined databases, Entrez and SRS, have been very successful. Entrez Entrez (Schuler et al., 1996) is a combined database consisting of literature, protein sequence and structure, nucleotide sequence, and taxonomy. Different types of information are interconnected through the grouping of sequences/structures and references by computed similarity scores. Entrez can be used through a variety of media, including CD-ROM, a custom Graphical Interface client, a World Wide Web browser, a command line browser (CLEVER), and the National Center for Biotechnology Information’s (NCBI’s) toolkit written in C. SRS SRS (Sequence Retrieval System; Etzold et al., 1996) is the most comprehensive database for molecular biology. The home server at http://srs.ebi.ac.uk contains 392 biological databases (as of October, 1999), including almost all the major protein/genetic databases. As an indexing system, it provides fast access to different databases through searches by sequence or by key words from various data fields. SRS also builds indices using cross-references between databases. An entry from one database can be linked to other databases that contain the entry. SRS is widely available on-line worldwide. The Web page http://srs.ebi.ac.uk/srs5list.html shows a list of dozens of mirror sites, e.g., http://ash.lsd.ornl.gov:80/srs5/ (USA), http://www.virus.kyoto-u.ac.jp:80/srs5/ (Japan), and http://www.bic.nus.edu.sg:80/srs5/ (Singapore). However, it should be noted that the contents of SRS may lag behind the other databases in updating (i.e., some new entries in the original databases may not be included in SRS). SUMMARY Protein Databases on the Internet

This unit reviews several major protein databases on the Internet, and shows what kind of information users can expect from protein databases. Although all technical procedures



cannot be described here, most of the protein databases are easy to use and provide detailed on-line manuals so that even users with little computer skills can learn them quickly. Protein databases may not always be easily accessible or usable through the Internet. Sometimes a database server may be down. Some structures or image files are very large (several megabytes), and the download time may be long. It can be helpful to use a mirror site of the database at a close location in order to accelerate the access speed. For a frequent user, it may be worthwhile to install the database on a local machine. On the other hand, it must be kept in mind that a mirror site or a local copy may contain an older version of the database than the one on the home server. It is important to assess the quality of the data. There are three types of data in protein databases. (1) Experimental data are generally very reliable. However, some entries may contain errors (e.g., some protein sequences) or may be based on low-resolution data (e.g., some protein structures determined by NMR). (2) Annotation data uses computational techniques on experimental data, for example, secondary structure assignment and domain partition in structure. These data depend on the quality of the experimental data and the computational methods used. Different methods may yield different results. (3) Prediction data includes, for example, sequence domain parsing and three-dimensional structure prediction. No matter how good the method, the results are still predictions and should be subjected to experimental verification. In addition, different methods typically give different predictions. In summary, caution is needed when using the data from databases to draw a conclusion. It is worthwhile to check the same type of data from different databases and compare them. It is sometimes necessary to use additional computational tools (e.g., tools to assess the quality of a structure) for further analysis. LITERATURE CITED Altschul, S.F., Madden, T.L., Schäffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. 1997. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucl. Acids Res. 25:3389-3402. Attwood, T.K., Flower, D.R., Lewis, A.P., Mabey, J.E., Morgan, S.R., Scordis, P., Selley, J., and Wright, W. 1999. PRINTS prepares for the new millennium. Nucl. Acids Res. 27:220-225. Bairoch, A. 1993. The ENZYME data bank. Nucl. Acids Res. 21:3155-3156. Bairoch, A. and Apweiler, R. 1999. The SwissProt protein sequence data bank and its supplement TrEMBL in 1999. Nucl. Acids Res. 27:49-54. Barker, W.C., Garavelli, J.S., McGarvey, P.B., Marzec, C.R., Orcutt, B.C., Srinivasarao, G.Y., Yeh, L.L., Ledley, R.S., Mewes, H., Pfeiffer, F., Tsugita, A., and Wu, C. 1999. The PIR-international protein sequence database. Nucl. Acids Res. 27:39-42. Bateman, A., Birney, E., Durbin, R., Eddy, S.R., Finn, F.D., and Sonnhammer, E.L.L. 1999. Pfam 3.1: 1313 multiple alignments match the majority of proteins. Nucl. Acids Res. 27:260-262. Benson, D.A., Boguski, M.S., Lipman, D.J., Ostell, J., Ouellette, B.F., Rapp, B.A., and Wheeler, D.L. 1999. Genbank. Nucl. Acids Res. 27:12-17. Bernstein, F.C., Koetzle, T.F., Williams, G.J.B., Meyer, E.F., Brice, M.D., Rodgers, J.R., Kennard, O., Shimanouchi, T., and Tasumi, M. 1977. The protein data bank: A computer based archival file for macromolecular structures. J. Mol. Biol. 112:535-542. Bourne, P., Berman, H., Watenpaugh, K., Westbrook, J., and Fitzgerald, P. 1997. The macromolecular crystallographic information file (mmCIF). Methods Enzymol. 277:571-590. Corpet, F., Gouzy, J., and Kahn, D. 1999. Recent improvements of the ProDom database of protein domain families. Nucl. Acids Res. 27:263-267. Etzold, T., Ulyanov, A., and Argos, P. 1996. SRS: Information retrieval system for molecular biology data banks. Methods Enzymol. 266:114-128. Gerstein, M. and Krebs, W. 1998. A database of macromolecular motions. Nucl. Acids Res. 26:4280-4290. Informatics


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Gibrat, J.F., Madej, T., and Bryant, S.H. 1996. Surprising similarities in structure comparison. Curr. Opinion Struct. Biol. 6:377-385. Gromiha, M.M., An, J., Kono, H., Oobatake, M., Uedaira, H., and Sarai, A. 1999. Protherm: Thermodynamic database for proteins and mutants. Nucl. Acids Res. 27:286-288. Hendlich, M. 1998. Databases for protein-ligand complexes. Acta Crystallogr., Sect. D 1:1178-1182. Heniko, J.G., Heniko, S., and Pietrokovski, S. 1999. New features of the blocks database servers. Nucl. Acids Res. 27:226-228. Hofmann, K., Bucher, P., Falquet, L., and Bairoch, A. 1999. The PROSITE database, its status in 1999. Nucl. Acids Res. 27:215-219. Holm, L. and Sander, C. 1996. Mapping the protein universe. Science 273:595-602. Kraulis, P. 1991. MOLSCRIPT—a program to produce both detailed and schematic plots of protein structures. J. Appl. Crystallogr. 24:946-950. Laskowski, R.A., Hutchinson, E.G., Michie, A.D., Wallace, A.C., Jones, M.L., and Thornton, J.M. 1997. PDBsum: A web-based database of summaries and analyses of all PDB structures. Trends Biochem. Sci. 22:488-490. Lemkin, P.F., Orr, G.A., Goldstein, M.P., Creed, G.J., Myrick, J.E., and Merril, C.R. 1995. The protein disease database of human body fluids: I. Computer methods and data issues. Appl. Theor. Electrophor. 5:55-72. Marchler-Bauer, A., Addess, K.J., Chappey, C., Geer, L., Madej, T., Matsuo, Y., Wang, Y., and Bryant, S.H. 1999. MMDB: Entrez’s 3D structure database. Nucl. Acids Res. 27:240-243. Merril, C.R., Goldstein, M.P., Myrick, J.E., Creed, G.J., and Lemkin, P.F. 1995. The protein disease database of human body fluids: I. Rationale for the development of this database. Appl. Theor. Electrophor. 5:49-54. Mural, R.J., Parang, M., Shah, M., Snoddy, J., and Uberbacher, E.C. 1999. The Genome Channel: A browser to a uniform first-pass annotation of genomic DNA. Trends Genet. 15:38-39. Murvai, J., Vlahovicek, K., Barta, E., Szepesvari, C., Acatrinei, C., and Pongor, S. 1999. The SBASE protein domain library, release 6.0: A collection of annotated protein sequence segments. Nucl. Acids Res. 27:257-259. Murzin, A.G., Brenner, S.E., Hubbard, T., and Chothia, C. 1995. SCOP: A structural classification of proteins database for the investigation of sequences and structures. J. Mol. Biol. 247:536-540. Nayal, M., Hitz, B.C., and Honig, B. 1999. GRASS: A server for the graphical representation and analysis of structures. Protein Sci. 8:676-679. Ogata, H., Goto, S., Sato, K., Fujibuchi, W., Bono, H., and Kanehisa, M. 1999. KEGG: Kyoto encyclopedia of genes and genomes. Nucl. Acids Res. 27:29-34. Orengo, C.A., Michie, A.D., Jones, D.T., Swindells, M.B., and Thornton, J.M. 1997. CATH—a hierarchic classification of protein domain structures. Structure 5:1093-1108. Rebhan, M., Chalifa-Caspi, V., Prilusky, J., and Lancet, D. 1998. GeneCards: A novel functional genomics compendium with automated data mining and query reformulation support. Bioinformatics 14:656-664. Rodriguez-Tom, P., Stoehr, P.J., Cameron, G.N., and Flores, T.P. 1996. The European Bioinformatics Institute (EBI) databases. Nucl. Acids Res. 24:6-13. Sayle, R.A. and Milner-White, E.J. 1995. RASMOL: Biomolecular graphics for all. Trends Biochem. Sci. 20:374-376. Schuler, G.D., Epstein, J.A., Ohkawa, H., and Kans, J.A. 1996. Entrez: Molecular biology database and retrieval system. Methods Enzymol. 266:141-162. Shindyalov, I.N. and Bourne, P.E. 1995. WPDB: A PC-based tool for analyzing protein structure. J. Appl. Crystallogr. 28:847-852. Shindyalov, I.N. and Bourne, P.E. 1998. Protein structure alignment by incremental combinatorial extension (CE) of the optimal path. Protein Eng. 11:739-747. Shnel, J. 1996. Image library of biological macromolecules. Comput. Appl. Biosci. 12:227-229. Siddiqui, A.S. and Barton, G.J. 1995. Continuous and discontinuous domains: An algorithm for the automatic generation of reliable protein domain definitions. Protein Sci. 4:872-884. States, D.J., Harris, N.L., and Hunter, L. 1993. Computationally efficient representation of classes in protein sequence megaclassification. Proc. Intel. Syst. Mol. Biol. 1:387-394. Tatusov, R.L., Koonin, E.V., and Lipman, D.J. 1997. A genomic perspective on protein families. Science 278:631-637. University of Wisconsin. 1999. BioMagResBank. University of Wisconsin, Madison, Wis. Vriend, G. 1990. WHAT IF: A molecular modelling and drug design program. J. Mol. Graphics 8:52-56. Protein Databases on the Internet



Wallace, A.C., Laskowski, R.A., and Thornton, J.M. 1996. Derivation of 3D coordinate templates for searching structural databases: Application to Ser-His-Asp catalytic triads in the serine proteinases and lipases. Protein Sci. 5:1001-1013. Wu, C., Shivakumar, S., and Huang, H. 1999. Proclass protein family database. Nucl. Acids Res. 27:272-274.

INTERNET RESOURCES The Web addresses of the databases mentioned in this unit are listed in Table 19.4.1. Readers can find more protein databases and their related tools in the following Web pages, which collect a large number of useful links. http://compbio.ornl.gov/structure/resource/ Oak Ridge National Laboratory’s resources of protein modeling tools. http://gnomic.stanford.edu/∼luciano/bioservers.html Pedro’s biomolecular research tools. http://www-biol.univ-mrs.fr/english/logligne.html University of Provence’s on-line analysis tools. http://www.expasy.ch/alinks.html Amos’ WWW links page.

Contributed by Dong Xu and Ying Xu Oak Ridge National Laboratory Oak Ridge, Tennessee

The authors thank Drs. Edward C. Uberbacher, Michael A. Unseren, Jay Snoddy, and Gwo-liang Chen for helpful discussions. This research was sponsored by the Office of Biological and Environmental Research, U.S. Department of Energy, under contract no. DE-AC05-96OR22464 with Lockheed Martin Energy Research Corporation.

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CHAPTER 20 Analysis of Protein Interactions INTRODUCTION

M

ost of the work of living organisms is performed by proteins. Proteins do their work by acting on other macromolecules: nucleic acids, carbohydrates, lipids, and, especially, other proteins. Inside cells, protein-protein interactions are instrumental in enzymatic actions upon protein substrates and in the fleeting and lasting protein assemblies that govern signal transduction, cell division, DNA replication, and transcription initiation. Outside cells, protein interactions allow cells to talk with each other: ligands expressed on the cell surface often bind protein receptors expressed on adjacent cells, and secreted protein ligands bind receptors on distant cells. Techniques to detect and study protein-protein interactions have become steadily more easy to use in recent years. This chapter lays them out. BASIC CONCEPTS Most of the work of living organisms is performed by proteins. Proteins do their work by acting on other macromolecules: nucleic acids, carbohydrates, lipids, and, especially, other proteins. Inside cells, protein-protein interactions are instrumental in enzymatic actions upon protein substrates and in the fleeting and lasting protein assemblies that govern signal transduction, cell division, DNA replication, and transcription initiation. Outside cells, protein interactions allow cells to talk with each other: ligands expressed on the cell surface often bind protein receptors expressed on adjacent cells, and secreted protein ligands bind receptors on distant cells. Techniques to detect and study proteinprotein interactions have become steadily more easy to use in recent years. This unit, and those that follow, lay them out.

Qualitative statements about protein-protein interactions inform much of the current biological literature. This unit presents the basics for understanding such qualitative statements—i.e., the quantitative concepts by which these can be evaluated. Colloquially, protein interactions are often referred to as either stable or transient. The statement that a certain interaction is stable is typically used when the interacting proteins form a complex. Membership in a protein complex is most commonly defined operationally— based on detection of the protein after its coprecipitation with a reagent specific for another member of the complex—but is sometimes defined based on a strong and biologically plausible interaction in a two-hybrid experiment. Proteins that form such complexes with one another are often referred to as partners. By contrast, the term transient is used to denote, e.g., interactions between an enzyme and a protein substrate. As the preceding example suggests, there is no particular correlation between the stability of a protein interaction and its biological significance. It is also worth noting that there are exceptions to commonly held assumptions about stable and transient interactions: some interactions observed in coprecipitation experiments have no biological validity, whereas some enzyme-substrate interactions last for minutes. For these reasons, it is often useful to think of protein interactions in more formal terms. The strength, or affinity, of an interaction between two proteins is described by equilibrium parameters. The speed at which an interaction occurs, and at which interacting proteins come apart, is given by kinetic parameters. Introduction Contributed by Roger Brent Current Protocols in Molecular Biology (1999) 20.0.1-20.0.6 Copyright © 1999 by John Wiley & Sons, Inc.


Table 20.0.1 Interactions

Dissociation Constants for Some Biologically Significant

Interaction

Kd

Streptavidin-biotin binding Antibody-antigen interaction with good antibody DNA binding protein with specific site Antibody-antigen interaction with weak antibody Enzyme-substrate interactions Cooperative interaction between DNA-bound phage repressors Cooperative interaction between phage repressor and E. coli RNA polymerase

10−14 M 10−8 to 10−10 M 10−8 to 10−10 M 10−6 M 10−4 to 10−10 M 10−4 M 10−2 M

EQUILIBRIUM PARAMETERS The strength of an interaction can be given as the equilibrium dissociation constant, Kd, or by its reciprocal, Ka (discussed below). When two proteins A and B associate, the Kd is given by

Kd =

[A][B] [AB]

where [AB] is the concentration of the complexed species and [A] and [B] are the concentrations of the noncomplexed species. Concentrations are given in molar terms, as is the Kd. There are two useful special cases to keep in mind. First, when A and B are present in equal concentration, the concentration at which half of each protein is present in the form of AB complex is the Kd. Second, when the concentration of A vastly exceeds the concentration of B (say, by 100-fold)—typically indicated [A] >> [B]—the concentration of A at which half of the B is found in the AB complex approximates Kd. Typical Kd values for biologically significant interactions are given in Table 20.0.1. Note that many significant interactions are weak. Alternatively, affinity may be given as an equilibrium association constant, Ka. The Ka is simply

Ka =

[AB] [A][B]

In other words, it is the reciprocal of the equilibrium dissociation constant: Ka = 1/Kd. Association constants are used less often than Kd, but do appear in the literature of some subfields—for example, in descriptions of antibody-antigen interactions. The strength of an interaction is directly proportional to the change in Gibbs free energy (∆G) when A and B interact, which is given by ∆G = ∆H − T∆S Analysis of Protein Interactions

where T is the temperature in degrees Kelvin, ∆S is the change in entropy (S), and ∆H is the change in enthalpy (H). ∆G is given in units of kilocalories/mole (kcal mol−1). An



increase in the free energy of an interaction by 1 kcal decreases the Kd by a factor of ∼7. Note that the temperature affects the entropy term of the equation. As the temperature decreases, reactions driven by enthalpy are less affected by losses in entropy, and are usually favored. By contrast, reactions whose free energy is derived largely from favorable changes in entropy are disfavored at lower temperatures. This means that the dependence of a protein association on temperature can often provide a clue to which term predominately contributes to the free energy change involved in that association. The relationship between Ka and ∆G is defined as follows: ∆G 0 = − RT ln

[ AB] [ A][B]

∆G 0 = − RT ln Ka = − RT ln

1 = −RT ln K K

d

d

where ∆G0 is the free energy change under standard conditions (25oC); R is the universal gas constant (1.9872 calmole−1K–1); and T is the temperature in degrees Kelvin (25°C is 289.1 K). Therefore,

∆G0 = 0.588 ln Kd and, since ln x = 2.303 log10 x, ∆G0 = 1.36 log10 Kd. For example: (a ) K d = 1 × 10 −14 , then ∆G 0 = −19.04 kcal mol −1 ( b) K d = 1 × 10 −2 , then ∆G 0 = −2.72 kcal mol −1

The streptavidin-biotin reaction (a) is intrinsically more favorable in the direction of binding, [AB] formation, than the low-affinity interaction (b) involving phage repressor (see Table 20.0.1). It should be pointed out that the DG values calculated above assume that the molar ratio of reactants [A] [B] and product [AB] is 1 M (standard state). Many (but by no means all) biologically important protein interactions seem to be largely driven by ∆H, or changes in enthalpy. That is fortunate, in that changes in entropy on binding are very hard to quantitate, or even think about precisely. For example, proteins in solution are surrounded by water molecules that form hydrogen bonds with surface residues. When two proteins interact, the ordered arrangement of the water molecules that surrounded the interacting surfaces of the proteins is often disrupted, and this loss of order provides an entropically favorable term to the free energy of the interaction. Such changes in the free energy due to entropy are very hard to predict. By contrast, enthalpic changes are easier to understand. If formation of one hydrogen bond liberates about −1 kcal M−1, and formation of a particular ionic contact liberates about −2 kcal M−1, then the energies of these changes are additive such that the formation of both bonds usually liberates −3 kcal M−1—which means that the Kd is decreased >100-fold by the enthalpic changes. Kinetic Parameters The above descriptions of protein interactions make no reference to the speed at which association or dissociation occurs. These speeds are given by kinetic parameters. The dissociation rate constant, kdissoc, gives the speed at which the AB complex dissociates into A and B (AB → A + B). kdissoc is a first-order rate constant—i.e., one that Introduction


Supplement 65

is dependent on the concentration of one species, in this case the AB complex—and is given by the rate of decrease in the concentration of AB: kdissoc [A][B] =

− d[ AB] dt

Its units are those of reciprocal time (t in the equation), usually given in sec−1. For example, a dissociation rate constant of 10−4 sec−1 means that one in 104 of the AB complexes present comes apart each second. Similarly, the association rate constant, kassoc, gives the speed at which A and B associate to form an AB complex (A + B → AB). This is a second-order reaction—i.e., its speed depends on the concentrations of both A and B—and its rate constant is is given by kassoc [A][B] =

+ d[ AB] dt

Its units are those of reciprocal concentration × reciprocal time, typically given in M−1 sec−1. For example, suppose that protein A is present at a concentration of 10−6 M in a cell, that protein B is injected to a nuclear concentration of 10−5 M, and that the rate constant for this antibody-antigen association is 10−4 M−1 sec−1. After injection, the concentration of AB will be [AB] = [A] × [B] × Kassoc = (10 −6 M)(10 −5 M)(10 −4 M) = 10 −7 M sec −1 That is, in the first second after mixing, 10−7 M of AB complex will form. Every 10-fold increase in the concentration of either reactant increases the rate of product formation 10-fold. Note that, at equilibrium, by definition, d[AB] =0 dt and, therefore, for the AB interaction, Kd =

kdissoc kassoc

The fact that the strength of equilibrium interactions reflects the speed of associations and dissociations has important consequences. To understand this, imagine two pairs of proteins that interact with the same Kd. Proteins A and B come together slowly, but, once the AB complex forms, it takes a long time to come apart. By contrast, proteins C and D come together rapidly, and the CD complex dissociates rapidly. There are two common cases in which these kinetic differences in the AB and CD associations would be significant.

Analysis of Protein Interactions

One is in measurement. Many techniques, such as the “pulldown” and immunological coprecipitation techniques described in this chapter, rely on the the fact that proteins remain associated during some sequence of steps, while they are being separated from other proteins in a mixture and while the isolated complex is being rinsed. No matter how tightly the proteins associate, if they come apart before their complex can be separated and rinsed, the complex will not be detected. Moreover, if the AB and CD interactions



have the same Kd, but CD comes apart more rapidly, then a coprecipitation experiment can falsely suggest that the CD association is weaker. The second case concerns the biological effects. Many biological phenomena, such as the transcription phenotypes resulting from protein-protein interactions in two-hybrid experiments (UNIT 19.3), seem to be well-described by consideration of equilibrium measurements. However, it is worth keeping in mind that any biological process that results from the association of two proteins requires a minimum time to occur. For some enzyme-substrate interactions, the minimum time may be on the order of microseconds, but for others, such as the initiation of DNA replication, it may be measured in seconds. If the complex dissociates faster than the minimum time, then, no matter how tight the interaction, the process will not occur. If AB and CD have the same Kd, but CD dissociates faster, the association may not produce a biological effect. WHAT THIS CHAPTER DESCRIBES To a contemporary biologist, the phrase “analysis of protein interactions” encompasses both the identification of interacting proteins and the measurement of the strength and rates of their interactions. This chapter covers both. It does not a number of classical biochemical techniques, including those that separate bound species from free species on chromatographic media, in velocity gradients, or by equilibrium dialysis. It has been designed to present most of the techniques currently used for identification and the most widely used techniques for measurement. This chapter features several units that provide quite different approaches for identifying interacting proteins. The first approach, described in UNIT 20.1, presents recent developments in two-hybrid methods. These approaches allow detection of proteins that interact with a DNA-bound “bait” protein, and isolation of the genes that encode them, based on their biological effect on transcription of reporter genes in yeast. This chapter also adds a second method (UNIT 20.6) for detecting interacting proteins. This method (called the “far-Western”) depends on proteins in a liquid phase (e.g., a cell extract interacting with “bait” proteins separated electrophoretically on a gel). Proteins that interact and whose interaction persists through the washing steps are detected by antibodies against them. This method favors detection of protein interactions with long-off rates. A third procedure, detailed in UNIT 20.2, provides a biochemical method for detecting associations between a GST-fused bait protein and a mixture of proteins that may contain an interacting partner. In this coprecipitation or “pulldown” approach, interacting proteins are identified and can be purified using standard affinity methods for GST-containing proteins. In the related technique of co-immunoprecipitation (UNIT 20.5), cell-free extracts are incubated with an antibody to a desired protein (or to an epitope tag) in order to coprecipitate associated proteins. This approach is used to isolate multiprotein complexes that presumably were present in the intact cell. Another molecular biological approach that directly yields a clone encoding the interacting protein is described in UNIT 20.3. Interaction cloning (also known as expression cloning) is a technique to identify and clone genes which encode proteins that interact with a protein of interest, or “bait” protein. Phage-based interaction cloning requires a gene encoding the bait protein and an appropriate expression library constructed in a bacteriophage expression vector, such as λgt11. Introduction


Supplement 46

An in vitro technique called surface plasmon resonance (SPR) can simultaneously detect interactions between unmodified proteins and measure kinetic parameters of the interaction. The availability of user-friendly instruments has facilitated the use of SPR as a means of studying macromolecular interactions. This technology is presented in UNIT 20.4. Roger Brent




Interaction Trap/Two-Hybrid System to Identify Interacting Proteins

UNIT 20.1

To understand the function of a particular protein, it is often useful to identify other proteins with which it associates. This can be done by a selection or screen in which novel proteins that specifically interact with a target protein of interest are isolated from a library. One particularly useful approach to detect novel interacting proteins—the twohybrid system or interaction trap (see Figs. 20.1.1 and 20.1.2)—uses yeast as a “test tube” and transcriptional activation of a reporter system to identify associating proteins (see Background Information). This approach can also be used specifically to test complex formation between two proteins for which there is a prior reason to expect an interaction. In the basic version of this method (see Fig. 20.1.2), the plasmid pEG202 or a related vector (see Fig. 20.1.3 and Table 20.1.1) is used to express the probe or “bait” protein as a fusion to the heterologous DNA-binding protein LexA. Many proteins, including transcription factors, kinases, and phosphatases, have been successfully used as bait proteins. The major requirements for the bait protein are that it should not be actively excluded from the yeast nucleus, and it should not possess an intrinsic ability to strongly activate transcription. The plasmid expressing the LexA-fused bait protein (see Table 20.1.1) is used to transform yeast possessing a dual reporter system responsive to transcriptional activation through the LexA operator. In one such example, the yeast strain EGY48 (see Table 20.1.2) contains the reporter plasmid pSH18-34. In this case, binding sites for LexA are located upstream of two reporter genes. In the EGY48 strain, the upstream activating sequences of the chromosomal LEU2 gene—required in the biosynthetic pathway for leucine (Leu)—are replaced with LexA operators (DNA binding sites). pSH18-34 contains a LexA operator–lacZ fusion gene. These two reporters allow selection for transcriptional activation by permitting selection for viability when cells are plated on medium lacking Leu, and discrimination based on color when the yeast is grown on medium containing Xgal (UNIT 13.6). In Basic Protocol 1, EGY48/pSH18-34 transformed with a bait is characterized for its ability to express protein (Support Protocol 1), growth on medium lacking Leu, and for the level of transcriptional activation of lacZ (see Fig. 20.1.2A). A number of alternative strains, plasmids, and strategies are presented which can be employed if a bait proves to have an unacceptably high level of background transcriptional activation. In an interactor hunt (Basic Protocol 2), the strain EGY48/pSH18-34 containing the bait expression plasmid is transformed (along with carrier DNA made as described in Support Protocol 2) with a conditionally expressed library made in the vector pJG4-5 (see Fig. 20.1.6 and Table 20.1.3). This library uses the inducible yeast GAL1 promoter to express proteins as fusions to an acidic domain (“acid blob”) that functions as a portable transcriptional activation motif (act) and to other useful moieties. Expression of libraryencoded proteins is induced by plating transformants on medium containing galactose (Gal), so yeast cells containing library proteins that do not interact specifically with the bait protein will fail to grow in the absence of Leu (see Fig. 20.1.2B). Yeast cells containing library proteins that interact with the bait protein will form colonies within 2 to 5 days, and the colonies will turn blue when the cells are streaked on medium containing Xgal (see Fig. 20.1.2C). The DNA from interaction trap positive colonies can be analyzed by polymerase chain reaction (PCR) to streamline screening and detect redundant clones in cases where many positives are obtained in screening (see Alternate Protocol 1). The plasmids are isolated and characterized by a series of tests to confirm specificity of the interaction with the initial bait protein (Support Protocols 3 to 5). Those found to be specific are ready for further analysis (e.g., sequencing).


Contributed by Erica A. Golemis, Ilya Serebriiskii, Russell L. Finley, Jr., Mikhail G. Kolonin, Jeno Gyuris, and Roger Brent

20.1.1


Supplement 46

prepare bait strain(s) for interaction mating (Alternate Protocol 2, steps 1-3)

construct bait protein plasmid and transform yeast (Basic Protocol 1, step 1)

characterize bait protein expression and activity

transform cDNA library into lexA operatorLEU2 strain to make pretransformed library strain (Alternate Protocol 2, steps 4-11)

obtain cDNA library in pJG4-5

transform cDNA library into lexA-operator-LEU2/lexA-operator-lacZ/pBait yeast (Basic Protocol 2, steps 1-7 ) introduce cDNA library into bait strain(s) by interaction mating (Alternate Protocol 2 , select for library plasmid steps 12-20 ) (Basic Protocol 2, step 8)

assess transcriptional activity (Basic Protocol 1, steps 4-7 ) assess repressor activity (Basic Protocol 1, steps 8 -11) test for Leu requirement (Basic Protocol 1, steps 12 -13) assess protein synthesis (Support Protocol 1)

freeze and replate transformants (Basic Protocol 2, steps 9 -15)

select for interacting proteins (Basic Protocol 2, steps 16 -19)

transform E. coli (UNIT 5.10 and Basic Protocol 2, steps 20 -22)

test for specificity (Basic Protocol 2, steps 23 - 27, and Support Protocol 5 )

analyze and sequence positive isolates (Basic Protocol 2, step 28, and Support Protocol 5 )

assess whether clones are independent by restriction mapping or by filter hybridization (Support Protocol 3)

or obtain profile of independent interactors by microplate plasmid rescue (Support Protocol 4) or

warehouse clones and repeat screen with less sensitive strain analyze positive clones by PCR and restriction endonuclease digestion (Alternate Protoco l 1)

Figure 20.1.1 Flow chart for performing an interaction trap.



A

B

C act act

bait

bait LEU2

bait LEU2

LEU2

act

act bait

bait lacZ

bait lacZ

lacZ

Figure 20.1.2 The interaction trap. (A) An EGY48 yeast cell containing two LexA operator–responsive reporters, one a chromosomally integrated copy of the LEU2 gene (required for growth on −Leu medium), the second a plasmid bearing a GAL1 promoter–lacZ fusion gene (causing yeast to turn blue on medium containing Xgal). The cell also contains a constitutively expressed chimeric protein, consisting of the DNA-binding domain of LexA fused to the probe or bait protein, shown as being unable to activate either of the two reporters. (B) and (C), EGY48/pSH18-34/pbait-containing yeast have been additionally transformed with an activation domain (act)–fused cDNA library in pJG4-5, and the library has been induced. In (B), the encoded protein does not interact specifically with the bait protein and the two reporters are not activated. In (C), a positive interaction is shown in which the library-encoded protein interacts with bait protein, resulting in activation of the two reporters (arrow), thus causing growth on medium lacking Leu and blue color on medium containing Xgal. Symbols: black rectangle, LexA operator sequence; open circle, LexA protein; open pentagon, bait protein; open rectangle, library protein; shaded box, activator protein (acid blob in Fig. 20.1.6).

When more than one bait will be used to screen a single library, significant time and resources can be saved by performing the interactor hunt by interaction mating (see Alternate Protocol 2). In this protocol, EGY48 is transformed with library DNA and the transformants are collected and frozen in aliquots. For each interactor hunt, an aliquot of the pretransformed EGY48/library strain is thawed and mixed with an aliquot of a bait strain transformed with the bait expression plasmid and pSH18-34. Overnight incubation of the mixture on a YPD plate results in fusion of the two strains to form diploids. The diploids are then exposed to galactose to induce expression of the library-encoded proteins, and interactors are selected in the same manner as in Basic Protocol 2. The advantage to this approach is that it requires only one high-efficiency library transformation for multiple hunts with different baits. It is also useful for bait proteins that are somewhat toxic to yeast; yeast expressing toxic baits can be difficult to transform with the library DNA. CHARACTERIZING A BAIT PROTEIN The first step in an interactor hunt is to construct a plasmid that expresses LexA fused to the protein of interest. This construct is transformed into reporter yeast strains containing LEU2 and lacZ reporter genes, and a series of control experiments is performed to establish whether the construct is suitable as is or must be modified, and whether alternative yeast reporter conditions should be used. These controls establish that the bait protein is made as a stable protein in yeast, that it is capable of entering the nucleus and binding LexA operator sites, and that it does not appreciably activate transcription of the LexA operator–based reporter genes. This last is the most important constraint on use of this system. The LexA-fused bait protein must not activate transcription of either re-

BASIC PROTOCOL 1



Supplement 46

Table 20.1.1

Interaction Trap Componentsa,b

Plasmid name/source

Selection In yeast

Comment/description

In E. coli

LexA fusion plasmids HIS3 pEG202c,d,e HIS3 pJK202

Apr Apr

pNLexAe

HIS3

Apr

pGildad

HIS3

Apr

pEE202I

HIS3

Apr

pRFHM1e,f (control)

HIS3

Apr

pSH17-4e,f (control) pMW101f

HIS3

Apr

HIS3

Cmr

pMW103f

HIS3

Kmr

pHybLex/Zeof,g Zeor

Zeor

Activation domain fusion plasmids pJG4-5c,d,e,f TRP1 Apr

pJG4-5I

TRP1

Apr

pYESTrpg

TRP1

Apr

pMW102f

TRP1

Kmr

pMW104f

TRP1

Cmr

LacZ reporter plasmids URA3 pSH18-34d,e,f

Apr

pJK103e

URA3

Apr

pRB1840e

URA3

Apr

pMW112f pMW109f

URA3 URA3

Kmr Kmr

Contains an ADH promoter that expresses LexA followed by polylinker Like pEG202, but incorporates nuclear localization sequences between LexA and polylinker; used to enhance translocation of bait to nucleus Contains an ADH promoter that expresses polylinker followed by LexA; for use with baits where amino-terminal residues must remain unblocked Contains a GAL1 promoter that expresses same LexA and polylinker cassette as pEG202; for use with baits whose continuous presence is toxic to yeast An integrating form of pEG202 that can be targeted into HIS3 following digestion with KpnI; for use where physiological screen requires lower levels of bait to be expressed Contains an ADH promoter that expresses LexA fused to the homeodomain of bicoid to produce nonactivating fusion; used as positive control for repression assay, negative control for activation and interaction assays ADH promoter expresses LexA fused to GAL4 activation domain; used as a positive control for transcriptional activation Same as pEG202, but with altered antibiotic resistance markers; basic plasmid used for cloning bait Same as pEG202, but with altered antibiotic resistance markers; basic plasmid used for cloning bait Bait cloning vector compatible with interaction trap and all other two-hybrid systems; minimal ADH promotor expresses LexA followed by extended polylinker Contains a GAL1 promoter that expresses nuclear localization domain, transcriptional activation domain, HA epitope tag, cloning sites; used to express cDNA libraries An integrating form of pJG4-5 that can be targeted into TRP1 by digestion with Bsu36I (New England Biolabs); to be used with pEE202I to study interactions that occur physiologically at low protein concentrations Contains a GAL1 promoter that expresses nuclear localization domain, transcriptional activation domain, V5 epitope tag, multiple cloning sites; contains f1 ori and T7 promoter/flanking site; used to express cDNA libraries (Invitrogen) Same as pJG4-5, but with altered antibiotic resistance markers; no libraries yet available Same as pJG4-5, but with altered antibiotic resistance markers; no libraries yet available Contains 8 LexA operators that direct transcription of the lacZ gene; one of the most sensitive indicator plasmids for transcriptional activation Contains two LexA operators that direct transcription of the lacZ gene; an intermediate reporter plasmid for transcriptional activation Contains 1 LexA operator that directs transcription of the lacZ gene; one of the most stringent reporters for transcriptional activation Same as pSH18-34, but with altered antibiotic resistance marker Same as pJK103, but with altered antibiotic resistance marker continued



Table 20.1.1

Plasmid name/source pMW111f pMW107f pMW108f pMW110f pJK101e,f (control)

Interaction Trap Componentsa,b, continued

Selection In yeast

In E. coli

URA3 URA3 URA3 URA3 URA3

Kmr Cmr Cmr Cmr Apr

Comment/description Same as pRB1840, but with altered antibiotic resistance marker Same as pSH18-34, but with altered antibiotic resistance marker Same as pJK103, but with altered antibiotic resistance marker Same as pRB1840, but with altered antibiotic resistance marker Contains a GAL1 upstream activating sequence followed by two lexA operators followed by lacZ gene; used in repression assay to assess bait binding to operator sequences

aAll plasmids contain a 2µm origin for maintenance in yeast, as well as a bacterial origin of replication, except where noted (pEE202I, pJG4.5I). bInteraction Trap reagents represent the work of many contributors: the original basic reagents were developed in the Brent laboratory (Gyuris et al.,

1993). Plasmids with altered antibiotic resistance markers (all pMW plasmids) were constructed at Glaxo in Research Triangle Park, N.C. (Watson et al., 1996). Plasmids and strains for specialized applications have been developed by the following individuals: E. Golemis, Fox Chase Cancer Center, Philadelphia, Pa. (pEG202); J. Kamens, BASF, Worcester, Mass. (pJK202); cumulative efforts of I. York, Dana-Farber Cancer Center, Boston, Mass. and M. Sainz and S. Nottwehr, U. Oregon (pNLexA); D.A. Shaywitz, MIT Center for Cancer Research, Cambridge, Mass. (pGilda); R. Buckholz, Glaxo, Research Triangle Park, N.C. (pEE2021, pJG4-51); J. Gyuris, Mitotix, Cambridge, Mass. (pJG4-5); S. Hanes, Wadsworth Institute, Albany, N.Y. (pSH17-4); R.L. Finley, Wayne State University School of Medicine, Detroit, Mich. (pRFHM1); S. Hanes, Wadsworth Institute, Albany, N.Y. (pSH18-34); J. Kamens, BASF, Worcester, Mass. (pJK101, pJK103); R. Brent, The Molecular Sciences Institute, Berkeley, Calif. (pRB1840). Specialized plasmids not yet commercially available can be obtained by contacting the Brent laboratory at (510) 647-0690 or [email protected], or the Golemis laboratory, (215) 728-2860 or [email protected]. cSequence data are available for pEG202 (pLexA) accession number pending. dPlasmids commercially available from Clontech and OriGene; for Clontech pEG202 is listed as pLexA, pJG4-5 as pB42AD, and pSH18-34 as p8op-LacZ. ePlasmids and strains available from OriGene. fIn pMW plasmids the ampicillin resistance gene (Apr) is replaced with the chloramphenicol resistance gene (Cmr) and the kanamycin resistance gene (Kmr) from pBC SK(+) and pBK-CMV (Stratagene), respectively. The choice between Kmr and Cmr or Apr plasmids is a matter of personal taste; use of basic Apr plasmids is described in the basic protocols. Use of the more recently developed reagents would facilitate the purification of library plasmid in later steps by eliminating the need for passage through KC8 bacteria, with substantial saving of time and effort. Apr has been maintained as marker of choice for the library plasmid because of the existence of multiple libraries already possessing this marker. These plasmids are the basic set of plasmids recommended for use. gPlasmids commercially available from Invitrogen as components of a Hybrid Hunter kit; this kit also includes all necessary positive and negative controls (not listed in this table). See Background Information for further details on commercially available reagents.

porter—the EGY48 strain (or related strain EGY191) that expresses the LexA fusion protein should not grow on medium lacking Leu, and the colonies should be white on medium containing Xgal. The characterized bait protein plasmid is used for Basic Protocol 2 to screen a library for interacting proteins. Materials DNA encoding the protein of interest Plasmids (see Table 20.1.1): pEG202 (see Fig. 20.1.3), pSH18-34 (see Fig. 20.1.4), pSH17-4, pRFHM1, and pJK101 for basic characterization; other plasmids for specific circumstances as described (Clontech, Invitrogen, OriGene, or R. Brent) Yeast strain EGY48 (ura3 trp1 his3 3LexA-operator-LEU2), or EGY191 (ura3 trp1 his3 1LexA-operator-LEU2; Table 20.1.2) Complete minimal (CM) medium dropout plates (UNIT 13.1), supplemented with 2% (w/v) of the indicated sugars (glucose or galactose), in 100-mm plates: Glu/CM, −Ura, −His Gal/CM, −Ura, −His Gal/CM, −Ura, −His, −Leu Z buffer (UNIT.13.6) with 1 mg/ml 5-bromo-4-chloro-3-indolyl-β-D-galactosidase (Xgal)



Supplement 46

PstI 8672

Aat II 7995 StuI 7835 NruI 7765 PstI 7671 PstI 7590

Tth111I 10059 NarI 180 NaeI 246 SphI 1110 HindIII 1514 MluI 1616 PmeI 2056 pBR backbone ADHpro EcoRI 2144 BamHI r SalI* Ap NcoI lexA NotI XhoI Sal I *2182 ADH ter PstI 2188 pEG202 SphI 2396 10166 bp

HIS3 HindIII 6464 BstXI 6380 HindIII 6277 PstI 6089 BssHII 5960

2µm ori

SacI 5113

XbaI 3487

PstI 4780 Avr II 5004

Polylinker sequence SalI* NotI SalI* EcoRI BamHI NcoI XhoI GAA TTC CCG GGG ATC CGT CGA CCA TGG CGG CCG CTC GAG TCG AC

Figure 20.1.3 LexA-fusion plasmids: pEG202. The strong constitutive ADH promoter is used to express bait proteins as fusions to the DNA-binding protein LexA. Restriction sites shown in this map are based on recently compiled pEG202 sequence data and include selected sites suitable for diagnostic restriction endonuclease digests. A number of restriction sites are available for insertion of coding sequences to produce protein fusions with LexA; the polylinker sequence and reading frame relative to LexA are shown below the map with unique sites marked in bold type. The sequence 5′-CGT CAG CAG AGC TTC ACC ATT G-3′ can be used to design a primer to confirm correct reading frame for LexA fusions. Plasmids contain the HIS3 selectable marker and the 2µm origin of replication to allow propagation in yeast, and the Apr antibiotic resistance gene and the pBR origin of replication to allow propagation in E. coli. In the recently developed LexA-expression plasmids pMW101 and pMW103, the ampicillin resistance gene (Apr) has been replaced with the chloramphenicol resistance gene (Cmr) and the kanamycin resistance gene (Kmr), respectively (see Table 19.2.1 for details).

Gal/CM dropout liquid medium (UNIT 13.1) supplemented with 2% Gal Antibody to LexA or fusion domain: monoclonal antibody to LexA (Clontech, Invitrogen) or polyclonal antibody to LexA (available by request from R. Brent or E. Golemis) H2O, sterile 30°C incubator Nylon membrane Whatman 3MM filter paper Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

Additional reagents and equipment for subcloning DNA fragments (UNIT 3.16), lithium acetate transformation of yeast (UNIT 13.7), liquid assay for β-galactosidase (UNIT 13.6), preparation of protein extracts for immunoblot analysis (see Support Protocol 1), and immunoblotting and immunodetection (UNIT 10.8)



Smal 0.00 Bam HI 0.01 EcoRI 0.01 XhoI 0.28

PstI 9.42 Hind III 9.22 EcoRI 9.19

EcoRI 0.60 Hind III 0.61 Bam HI 0.62

PstI 8.95 URA3

lexA op GAL1pro

lacZ

2µm LacZ reporter 10.3 kb

SacI 2.63

Apr

Hind III 7.05 Eco RI 6.95

pBR ori

Eco RI 3.70

Pst I 6.20

Figure 20.1.4 LacZ reporter plasmid. pRB1840, pJK103, and pSH18-34 are all derivatives of LR1∆1 (West et al., 1984) containing eight, two, or one operator for LexA (LexAop) binding inserted into the unique XhoI site located in the minimal GAL1 promoter (GAL1pro; 0.28 on map). The plasmid contains the URA3 selectable marker, the 2µm origin to allow propagation in yeast, the ampicillin resistance (Apr) gene, and the pBR322 origin (ori) to allow propagation in E. coli. Numbers indicate relative map positions. In the recently developed derivatives, the ampicillin resistance gene (Apr) has been replaced with the chloramphenicol or kanamycin resistance genes (see Table 19.2.1 for details).

NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly. Transform yeast with the bait protein plasmid 1. Using standard subcloning techniques (UNIT 3.16), insert the DNA encoding the protein of interest into the polylinker of pEG202 (see Fig. 20.1.3) or other LexA fusion plasmid to make an in-frame protein fusion. The LexA fusion protein is expressed from the strong alcohol dehydrogenase (ADH) promoter. pEG202 also contains a HIS3 selectable marker and a 2ìm origin for propagation in yeast. pEG202 with the DNA encoding the protein of interest inserted is designated pBait. Uses of alternative LexA fusion plasmids are described in Background Information.

2. Perform three separate lithium acetate transformations (UNIT 13.7) of EGY48 using the following combinations of plasmids: pBait + pSH18-34 (test) pSH17-4 + pSH18-34 (positive control for activation) pRFHM1 + pSH18-34 (negative control for activation). Use of the two LexA fusions as positive and negative controls allows a rough assessment of the transcriptional activation profile of LexA bait proteins. pEG202 itself is not a good negative control because the peptide encoded by the uninterrupted polylinker sequences is itself capable of very weakly activating transcription.



Supplement 46

Table 20.1.2

Interaction Trap Yeast Selection Strainsa

Strain

Relevant genotype

EGY48b,c,d

MATα trp1, his3, ura3, lexAops-LEU2

6

EGY191

MATα trp1, his3, ura3, lexAops-LEU2

2

L40c

MATα trpl, leu2, ade2, GAL4, lexAops-HIS34, lexAops-lacZ8

Number of operators

Comments/description lexA operators direct transcription from the LEU2 gene; EGY48 is a basic strain used to select for interacting clones from a cDNA library EGY191 provides a more stringent selection than EGY48, producing lower background with baits with instrinsic ability to activate transcription Expression driven from GAL1 promoter is constitutive in L40 (inducible in EGY strains); selection is for HIS prototrophy. Integrated lacZ reporter is considerably less sensitive than pSH18-34 maintained in EGY strains

aInteraction Trap reagents represent the work of many contributors: the original basic reagents were developed in the Brent laboratory

(Gyuris et al., 1993). Strains for specialized applications have been developed by the following individuals: E. Golemis, Fox Chase Cancer Center, Philadelphia, Pa. (EGY48, EGY191); A.B. Vojtek and S.M. Hollenberg, Fred Hutchinson Cancer Research Center, Seattle, Wash. (L40). Specialized strains not yet commercially available can be obtained by contacting the Brent laboratory at The Molecular Sciences Institute, Berkeley, (510) 647-0690 or [email protected], or the Golemis laboratory, (215) 728-2860 or [email protected]. bStrains commercially available from Clontech. cStrains commercially available from Invitrogen as components of a Hybrid Hunter kit; the kit also includes all necessary positive and

negative controls (not listed in this table). See Background Information for further details on commercially available reagents. dStrains commercially available from OriGene.

pSH18-34 contains a 2ìm origin and a URA3 selectable marker for maintenance in yeast, as well as a bacterial origin of replication and ampicillin-resistance gene. It is the most sensitive lacZ reporter available and will detect any potential ability to activate lacZ transcription. pSH17-4 is a HIS3 2ìm plasmid encoding LexA fused to the activation domain of the yeast activator protein GAL4. This fusion protein strongly activates transcription. pRFHM1 is a HIS3 2ìm plasmid encoding LexA fused to the N-terminus of the Drosophila protein bicoid. This fusion protein has no ability to activate transcription.

3. Plate each transformation mixture on Glu/CM −Ura, −His dropout plates. Incubate 2 days at 30°C to select for yeast that contain both plasmids. Colonies obtained can be used simultaneously in tests for the activation of lacZ (steps 4 to 7) and LEU2 (steps 12 to 13) reporters.

Assay lacZ gene activation by β-galactosidase assay 4. Streak a Glu/CM −Ura, −His master dropout plate with at least five or six independent colonies obtained from each of the three transformations in step 3 (test, positive control, and negative control) and incubate overnight at 30°C. The filter assay described in Steps 5a to 7a (based on Breeden and Nasmyth, 1985) provides a rapid assay for β-galactosidase transcription. Alternatively, a liquid assay (UNIT 13.6) or a plate assay (described in Steps 5b to 7b) may be used.

Perform filter assay for β-galactosidase activity: 5a. Lift colonies by gently placing a nylon membrane on the yeast plate and allowing it to become wet through. Remove the membrane and air dry 5 min. Chill the membrane, colony side up, 10 min at −70°C. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

Whatman 3MM filters can be cut to the size of the yeast plate as a more economical alternative to nylon membranes for performing lifts. In addition, two or three 5-min temperature cycles (−70°C to room temperature) can be used instead of a single cycle to promote better lysis; this may be worth doing if there is difficulty visualizing blue color.



6a. Cut a piece of Whatman 3MM filter paper slightly larger than the colony membrane and soak it in Z buffer containing 1 mg/ml Xgal. Place colony membrane, colony side up, on Whatman 3MM paper, or float it in the lid of a petri dish containing ∼2 ml Z buffer with 1 mg/ml Xgal. Acceptable results may be obtained using as little as 300 ìg/ml Xgal.

7a. Incubate at 30°C and monitor for color changes. It is generally useful to check the membrane after 20 min, and again after 2 to 3 hr. Strong activators will produce a blue color in 5 to 10 min, and a bait protein (LexA fusion protein) that does so is unsuitable for use in an interactor hunt using this lacZ reporter plasmid. Weak activators will produce a blue color in 1 to 6 hr (compare versus negative control pRFHMI which will itself produce a faint blue color with time) and may or may not be suitable. Weak activators should be tested using the repressor assay described in steps 8 to 11.

Perform Xgal plate assay for lacZ activation: 5b. Prepare Z buffer Xgal plates as described in UNIT 13.1. For activation assays, plates should be prepared with glucose as a sugar source. For repression assays (steps 8 to 11), galactose should be used as a sugar source. In our experience, when patching from a master plate to Xgal plates, sufficient yeast are transferred that plasmid loss is not a major problem even in the absence of selection; this is balanced by the desire to assay sets of constructs on the same plate to eliminate batch variation in Xgal potency. Hence, plates should be made either with complete minimal amino acid mix, or by dropping out only uracil (−Ura), to make the plates universally useful.

A +++ endogenous

GAL4 GALUAS

ops

lacZ

plasmid JK101

B

P1

endogenous

L e x A ops

GAL4 GALUAS

+ lacZ

plasmid JK101

Figure 20.1.5 Repression assay for DNA binding. (A) The plasmid JK101 contains the upstream activating sequence (UAS) from the GAL1 gene followed by LexA operators upstream of the lacZ coding sequence. Thus, yeast containing pJK101 will have significant β-galactosidase activity when grown on medium in which galactose is the sole carbon source because of binding of endogenous yeast GAL4 to the GALUAS (B). LexA-fused proteins (P1-LexA) that are made, enter the nucleus, and bind the LexA operator sequences (ops) will block activation from the GALUAS, repressing β-galactosidase activity (+) 3- to 5-fold. On glucose/Xgal medium, yeast containing pJK101 should be white because GALUAS transcription is repressed.



Supplement 46

6b. Streak yeast from master plate to Xgal plate and incubate at 30°C. 7b. Examine plates for color development at intervals over the next 2 to 3 days. Strongly activating fusions should be visibly blue on the plate within 12 to 24 hr; moderate activators will be visibly blue after ∼2 days. When a bait protein appreciably activates transcription under these conditions, there are several recourses. The first and simplest is to switch to a less sensitive lacZ reporter plasmid; use of pJK103 and pRB1840 may be sufficient to reduce background to manageable levels. If this fails to work, it is frequently possible to generate a truncated LexA fusion that does not activate transcription.

Confirm fusion-protein synthesis by repression assay For LexA fusions that do not activate transcription, confirm by performing a repression assay (Brent and Ptashne, 1984) that the LexA fusion protein is being synthesized in yeast (some proteins are not) and that it is capable of binding LexA operator sequences (Fig. 20.1.5). The following steps can be performed concurrently with the activation assay. 8. Transform EGY48 yeast with the following combinations of plasmids (three transformations): pBait + pJK101 (test) pRFHM1 + pJK101 (positive control for repression) pJK101 alone (negative control for repression). 9. Plate each transformation mix on Glu/CM −Ura, −His dropout plates or Glu/CM −Ura dropout plates as appropriate to select yeast cells that contain the indicated plasmids. Incubate 2 to 3 days at 30°C until colonies appear. 10. Streak colonies to a Glu/CM −Ura, −His or Glu/CM −Ura dropout master plate and incubate overnight at 30°C. 11. Assay β-galactosidase activity of the three transformed strains (test, positive control, and negative control) by liquid assay (using Gal/CM dropout liquid medium), filter assay (steps 5a to 7a, first restreaking to Gal/CM plates to grow overnight), or plate assay (steps 5b to 7b, using Gal/CM −Ura XGal plates). This assay should not be run for more than 1 to 2 hr for membranes, or 36 hr for Xgal plates, as the high basal lacZ activity will make differential activation of pJK101 impossible to see with longer incubations. Use of Xgal plates, and inspection 12 to 24 hr after streaking, is generally most effective. The plasmid pJK101 contains the galactose upstream activating sequence (UAS) followed by LexA operators upstream of the lacZ coding sequence. Thus, yeast containing pJK101 will have significant β-galactosidase activity when grown on medium in which galactose is the sole carbon source because of binding of endogenous yeast GAL4 to the GALUAS. LexA-fused proteins that are made, enter the nucleus, and bind the LexA operator sequences block activation from the GALUAS, repressing β-galactosidase activity 3- to 20-fold. Note that on Glu/Xgal medium, yeast containing pJK101 should be white, because GALUAS transcription is repressed.

12. If a bait protein neither activates nor represses transcription, perform immunoblot analysis by probing an immunoblot of a crude lysate with antibodies against LexA or the fusion domain to test for protein synthesis (see Support Protocol 1).

Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

Even if a bait protein represses transcription, it is generally a good idea to assay for the production of full-length LexA fusions, as occasionally some fusion proteins will be proteolytically cleaved by endogenous yeast proteases. If the protein is made but does not repress, it may be necessary to clone the sequence into a LexA fusion vector that contains a nuclear localization motif, e.g., pJK202 (see Table 20.1.1), or to modify or truncate the fusion domain to remove motifs that target it to other cellular compartments (e.g., myristoylation signals).



Test for Leu requirement These steps can be performed concurrently with the lacZ activation and repression assays. 13. Disperse a colony of EGY48 containing pBait and pSH18-34 reporter plasmids into 500 µl sterile water. Dilute 100 µl of suspension into 1 ml sterile water. Make a series of 1/10 dilutions in sterile water to cover a 1000-fold concentration range. 14. Plate 100 µl from each tube (undiluted, 1/10, 1/100, and 1/1000) on Gal/CM −Ura, −His dropout plates and on Gal/CM −Ura, −His, −Leu dropout plates. Incubate overnight at 30°C. There will be a total of eight plates. Gal/CM −Ura, −His dropout plates should show a concentration range from 10 to 10,000 colonies and Gal/CM −Ura, −His, −Leu dropout plates should have no colonies. Actual selection in the interactor hunt is based on the ability of the bait protein and acid-fusion pair, but not the bait protein alone, to activate transcription of the LexA operator-LEU2 gene and allow growth on medium lacking Leu. Thus, the test for the Leu requirement is the most important test of whether the bait protein is likely to have an unworkably high background. The LEU2 reporter in EGY48 is more sensitive than the pSH18-34 reporter for some baits, so it is possible that a bait protein that gives little or no signal in a β-galactosidase assay would nevertheless permit some level of growth on −Leu medium. If this occurs, there are several options for proceeding, the most immediate of which is to substitute EGY191 (see Table 20.1.2), a less sensitive screening strain, and repeat the assay. As outlined in this protocol, the authors recommend the strategy of performing the initial screening using the most sensitive reporters and then, if activation is detected, screening with increasingly less sensitive reporters (see Critical Parameters for further discussion).

PERFORMING AN INTERACTOR HUNT An interactor hunt involves two successive large platings of yeast containing LexA-fused probes and reporters and libraries in pJG4-5 (Fig. 20.1.6, Table 20.1.3) with a cDNA expression cassette under control of the GAL promoter. In the first plating, yeast are plated on complete minimal (CM) medium −Ura, −His, −Trp dropout plates with glucose (Glu) as a sugar source to select for the library plasmid. In the second plating, which selects for yeast that contain interacting proteins, a slurry of primary transformants is plated on CM −Ura, −His, −Trp, −Leu dropout plates with galactose/raffinose (Gal/Raff) as the sugar source. This two-step selection is encouraged for two reasons. First, a number of interesting cDNA-encoded proteins may be deleterious to the growth of yeast that bear them; these would be competed out in an initial mass plating. Second, it seems likely that immediately after simultaneous transformation and Gal induction, yeast bearing particular interacting proteins may not be able to initially express sufficient levels of these proteins to support growth on medium lacking Leu. Library plasmids from colonies identified in the second plating are purified by bacterial transformation and used to transform yeast cells for the final specificity screen.

BASIC PROTOCOL 2

A list of libraries currently available for use with this system is provided in Table 20.1.3. The protocol outlined below describes the steps used to perform a single-step screen that should saturate a library derived from a mammalian cell. For screens with libraries derived from lower eukaryotes with less complex genomes, fewer plates will be required. Occasionally, baits that seemed well-behaved during preliminary tests produce unworkably high backgrounds of “positives” during an actual screen (see Background Information and Critical Parameters). To forestall the waste of time and materials performing a screen with such a bait would entail, an alternative approach is to perform a scaled-back



Supplement 46

screen when working with a new bait (e.g., 5 rather than 30 plates of primary transformants). The results can be assessed before doing a full screen; it is then possible to switch to lower-sensitivity reporter strains and plasmids, if appropriate. Although individual baits will vary, the authors’ current default preference is to use the lacZ reporter pJK103 in conjunction with either EGY48 or EGY191. Polymerase chain reaction (PCR) can also be used in a rapid screening approach that may be preferable if a large number of positions are obtained in a library screen (see Alternate Protocol 1).

SacI 6440 PvuII 6253 Afl III 6075

KpnI 6446

HindIII 528 EcoRI 849 XhoI 861 GAL pro Alw NI 5661 HindIII 867 fusion SphI 1191 cassette BamHI 1330 pUC backbone XbaI 1336 ADH ter SalI 1342 NotI 1350 PstI 1364 Ap HindIII 1474 pJG4-5 6449 bp ScaI 4704

2µm ori

AatII 4264

XbaI 2072

TRP1 XbaI 4002 HindIII 3573

PstI 3365

Fusion cassette NLS

HA Tag

B42 domain

EcoRI

XhoI

ATG GGT GCT CCT CCA AAA AAG AAG ... CCC GAA TTC GGC CGA CTC GAG AAG CTT ... M G A P P K K K ... P E F G R L E K L ...


Figure 20.1.6 Library plasmids: pJG4-5. Library plasmids express cDNAs or other coding sequences inserted into unique EcoRI and XhoI sites as a translational fusion to a cassette consisting of the SV40 nuclear localization sequence (NLS; PPKKKRKVA), the acid blob B42 domain (Ruden et al, 1991), and the hemagglutinin (HA) epitope tag (YPYDVPDYA). Expression of cassette sequences is under the control of the GAL1 galactose-inducible promoter. This map is based on the sequence data available for pJG4-5, and includes selected sites suitable for diagnostic restriction digests (shown in bold). The sequence 5′-CTG AGT GGA GAT GCC TCC-3′ can be used as a primer to identify inserts or to confirm correct reading frame. The pJG4-5 plasmid contains the TRP1 selectable marker and the 2µm origin to allow propagation in yeast, and the antibiotic resistance gene and the pUC origin to allow propagation in E. coli. In the recently developed pJG4-5 derivative plasmids pMW104 and pMW102, the ampicillin resistance gene (Apr) has been replaced with the chloramphenicol resistance gene (Cmr) and the kanamycin resistance gene (Kmr), respectively (see Table 19.2.2 for details). Currently existing libraries are all made in the pJG4-5 plasmid (Gyuris et al., 1993) shown on this figure. Unique sites are marked in bold type.



Materials Yeast containing appropriate combinations of plasmids (see Table 20.1.1 and Table 20.1.2): EGY48 containing LexA-operator-lacZ reporter and pBait (see Basic Protocol 1) EGY48 containing LexA-operator-lacZ reporter and pRFHM-1 EGY48 containing LexA-operator-lacZ reporter and any nonspecific bait Complete minimal (CM) dropout liquid medium (UNIT 13.1) supplemented with sugars (glucose, galactose, and/or raffinose) as indicated [2% (w/v) Glu, or 2% (w/v) Gal + 1% (w/v) Raff]: Glu/CM −Ura, −His Glu/CM −Trp Gal/Raff/CM −Ura, −His, −Trp H2O, sterile TE buffer (pH 7.5; APPENDIX 2)/0.1 M lithium acetate Library DNA in pJG4-5 (Table 20.1.3 and Fig. 20.1.6) High-quality sheared salmon sperm DNA (see Support Protocol 2) 40% (w/v) polyethylene glycol 4000 (PEG 4000; filter sterilized)/0.1 M lithium acetate/TE buffer (pH 7.5) Dimethyl sulfoxide (DMSO) Complete minimal (CM) medium dropout plates (UNIT 13.1) supplemented with sugars and Xgal (20 µg/ml) as indicated [2% (w/v) Glu, and 2% (w/v) Gal + 1% (w/v) Raff]: Glu/CM −Ura, −His, −Trp, 24 × 24–cm (Nunc) and 100-mm Gal/Raff/CM −Ura, −His, −Trp, 100-mm Gal/Raff/CM −Ura, −His, −Trp, −Leu, 100-mm Glu/Xgal/CM −Ura, −His, −Trp, 100-mm Gal/Raff/Xgal/CM −Ura, −His, −Trp, 100-mm Glu/CM −Ura, −His, −Trp, −Leu, 100-mm Glu/CM −Ura, −His, 100-mm Gal/CM −Ura, −His, −Trp, −Leu, 100-mm TE buffer (pH 7.5), sterile (optional) Glycerol solution (see recipe) E. coli KC8 (pyrF leuB600 trpC hisB463; constructed by K. Struhl and available from R. Brent) LB/ampicillin plates (UNIT 1.1) E. coli DH5α or other strain suitable for preparation of DNA for sequencing Bacterial defined minimal A medium plates: 1× A medium plates containing 0.5 µg/ml vitamin B1 (UNIT 1.1) and supplemented with 40 µg/ml each Ura, His, and Leu 30°C incubator, with and without shaking Low-speed centrifuge and rotor 50-ml conical tubes, sterile 1.5-ml microcentrifuge tubes, sterile 42°C heating block Glass microscope slides, sterile Additional reagents and equipment for rapid miniprep isolation of yeast DNA (UNIT 13.11), transformation of bacteria by electroporation (UNIT 1.8), miniprep isolation of bacterial DNA (UNIT 1.6), restriction endonuclease digestion (UNIT 3.1; optional), and agarose gel electrophoresis (UNIT 2.5A; optional) NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly.



Supplement 46

Table 20.1.3 Libraries Compatible with the Interaction Trap Systema

Source of RNA/DNA Cell lines HeLa cells (human cervical carcinoma)

Independent clones

Insert size (average)b Contact information

JG

9.6 × 106

0.3-3.5 kb (1.5 kb)

Y JG

3.7 × 106 5.7 × 106

0.3-1.2 kb 0.3-3.5 kb (1.5 kb)

JG

4.0 × 106

0.7-2.8 kb (1.5 kb)

R. Brent, Clontech, Invitrogen, OriGene Invitrogen R. Brent, Clontech, OriGene R. Brent

Y Y JG JG Y

3.2 × 106 3.0 × 106 5.7 × 106 2 × 106 5.4 × 106

0.3-1.2 kb 0.5-4.0 kb (1.8 kb) (>1.3) 0.7-3.5 kb (1.2 kb) 0.3-0.8 kb

Invitrogen Clontech OriGene S. Witte Invitrogen

JG JG JG

4.0 × 106 >106 1.5 × 106

0.4-4.0 kb (2.0 kb) 0.3-2.5 kb (>0.5 kb) 0.3-3.5 kb

Clontech R. Brent R. Brent

Vector

HeLa cells (human cervical carcinoma) WI-38 cells (human lung fibroblasts), serum-starved, cDNA Jurkat cells (human T cell leukemia), exponentially growing, cDNA Jurkat cells (human T cell leukemia) Jurkat cells (human T cell leukemia) Jurkat cells (human T cell leukemia) Jurkat cells (human T cell leukemia) Be Wo cells (human fetal placental choriocarcinoma) Human lymphocyte CD4+ T cell, murine, cDNA Chinese hamster ovary (CHO) cells, exponentially growing, cDNA A20 cells (mouse B cell lymphoma) Human B cell lymphoma Human 293 adenovirus–infected (early and late stages) SKOV3 human Y ovarian cancer MDBK cell, bovine kidney MDCK cells HepG2 cell line cDNA MCF7 breast cancer cells, untreated MCF7 breast cancer cells, estrogen-treated MCF7 cells, serum-grown LNCAP prostate cell line, untreated LNCAP prostate cell line, androgen-treated Mouse pachytene spermatocytes

Y JG JG

3.11 × 106 — —

0.3-1.2 kb — —

Invitrogen H. Niu K. Gustin

Y JG JG JG JG JG JG JG JG JG

5.0 × 106 5.8 × 106 — 2 × 106 1.0 × 107 1.0 × 107 1.0 ×107 2.9 × 106 4.6 × 106 —

(>1.4 kb) (>1.2 kb) — — (>1.5 kb) (>1.1 kb) 0.4-3.5 kb (>0.8 kb) (>0.9 kb) —

OriGene OriGene D. Chen M. Melegari OriGene OriGene OriGene OriGene OriGene C. Hoog

Tissues Human breast Human breast tumor Human liver Human liver

Y Y JG Y

9 × 106 8.84 × 106 >106 2.2 × 106

Invitrogen Invitrogen R. Brent Clontech

Human liver Human liver Human lung Human lung tumor Human brain Human brain Human testis Human testis Human ovary

JG JG Y Y JG Y Y JG Y

3.2 × 106 1.1 × 107 5.9 × 106 1.9 × 106 3.5 × 106 8.9 × 106 6.4 × 106 3.5 × 106 4.6 × 106

0.4-1.2 kb 0.4-1.2 kb 0.6-4.0 kb (>1 kb) 0.5-4 kb (1.3 kb) 0.3-1.2 kb (> 1 kb) 0.4-1.2 kb 0.4-1.2 0.5-4.5 kb (1.4 kb) 0.3-1.2 kb 0.3-1.2 kb 0.4-4.5 kb (1.6 kb) 0.3-1.2 kb

Invitrogen OriGene Invitrogen Invitrogen Clontech Invitrogen Invitrogen Clontech Invitrogen continued



Table 20.1.3 Libraries Compatible with the Interaction Trap Systema, continued

Source of RNA/DNA

Vector

Independent clones


Human ovary Human ovary Human heart Human placenta Human placenta Human mammary gland Human peripheral blood leucocyte Human kidney Human fetal kidney Human spleen Human prostate Human normal prostate Human prostate Human prostate cancer Human fetal prostate Human fetal liver Human fetal liver Human fetal liver Human fetal brain

JG JG JG Y JG JG JG JG JG Y Y JG JG JG JG JG Y JG JG

4.6 × 106 3.5 × 106 3.0 × 106 4.8 × 106 3.5 × 106 3.5 × 106 1.0 × 107 3.5 × 106 3.0 × 106 1.14 × 107 5.5 × 106 1.4 × 106 1.4 × 106 1.1 × 106 — 3.5 × 106 2.37 × 106 8.6 × 106 3.5 × 106

(>1.3 kb) 0.5-4.0 kb (1.8 kb) 0.3-3.5 kb (1.3 kb) 0.3-1.2 kb 0.3-4.0 kb (1.2 kb) 0.5-5 kb (1.6 kb) (>1.3 kb) 0.4-4.5 kb (1.6 kb) (>1 kb) 0.4-1.2 kb 0.4-1.2 kb 0.4-4.5 kb (1.7 kb) (>1 kb) (>0.9 kb) — 0.3-4.5 kb (1.3 kb) 0.3-1.2 kb (>1 kb) 0.5-1.2 kb (1.5 kb)

Mouse brain Mouse brain Mouse breast, lactating Mouse breast, involuting Mouse breast, virgin Mouse breast, 12 days pregnant Mouse skeletal muscle Rat adipocyte, 9-week-old Zucker rat Rat brain Rat brain (day 18) Rat testis Rat thymus Mouse liver Mouse spleen Mouse ovary Mouse prostate Mouse embryo, whole (19-day) Mouse embryo Drosophila melanogaster, adult, cDNA D. melanogaster, embryo, cDNA D. melanogaster, 0-12 hr embryos, cDNA D. melanogaster, ovary, cDNA D. melanogaster, disc, cDNA D. melanogaster, head

JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG

6.1 × 106 4.5 × 106 1.0 × 107 1.0 × 107 1.0 × 107 6.3 × 106 7.2 × 106 1.0 × 107 4.5 ×106 — 8.0 × 106 8.2 × 106 9.5 × 106 1.0 × 107 4.0 × 106 — 1.0 × 105 3.6 × 106 1.8 × 106 3.0 × 106 4.2 × 106

(>1 kb) 0.4-4.5 kb (1.2 kb) 0.4-3.1 kb 0.4-7.0 kb 0.4-5.5 kb 0.4-5.3 kb 0.4-3.5 kb 0.4-5.0 kb 0.3-3.4 kb — (>1.2 kb) (>1.3 kb) (>1.4 kb) (>1 kb) (>1.2 kb) — 0.2-2.5 kb 0.5-5 kb (1.7 kb) (>1.0 kb) 0.5-3.0 kb (1.4 kb) 0.5-2.5 kb (1.0 kb)

OriGene Clontech Clontech Invitrogen Clontech Clontech OriGene Clontech OriGene Invitrogen Invitrogen Clontech OriGene OriGene OriGene Clontech Invitrogen OriGene R. Brent, Clontech, Invitrogen, OriGene OriGene Clontech OriGene OriGene OriGene OriGene OriGene OriGene OriGene H. Niu OriGene OriGene OriGene OriGene OriGene OriGene OriGene Clontech OriGene Clontech R. Brent

JG JG JG

3.2 × 106 4.0 × 106 —

0.3-1.5 kb (800 bp) 0.3-2.1 kb (900 bp) —

R. Brent R. Brent M. Rosbash continued


Supplement 46

Table 20.1.3 Libraries Compatible with the Interaction Trap Systema, continued

Source of RNA/DNA

Vector

Miscellaneous Synthetic aptamers Saccharomyces cerevisiae, S288C, genomic S. cerevisiae, S288C, genomic Sea urchin ovary Caenorhabditis elegans Agrobacterium tumefaciens Arabidopsis thaliana, 7-day-old seedlings Tomato (Lycopersicon esculentum) Xenopus laevis embryo

PJM-1 JG JG JG JG JG JG JG JG

Independent clones

>1× 109 >3 × 106 4.0 × 106 3.5 × 106 3.8 × 106 — — 8 × 106 2.2 × 106


60 bp 0.8-4.0 kb 0.5-4.0 kb (1.7 kb) (>1.2 kb) — — — 0.3-4 kb (1.0 kb)

R. Brent R. Brent OriGene Clontech OriGene — H.M. Goodman G.B. Martin Clontech

aMost libraries are constructed in either the pJG4-5 vector or the pYESTrp vector (JG or Y in the Vector column); the peptide aptamer library is made in the pJM-1 vector. Libraries available from the public domain were constructed by the following individuals: (1) J. Gyuris; (3) C. Sardet and J. Gyuris; (4) W. Kolanus, J. Gyuris, and B. Seed; (39) D. Krainc; (50-52) R. Finley; (55) P. Watt; (54) P. Colas, B. Cohen, T. Jessen, I. Grishina, J. McCoy, and R. Brent (Colas et al., 1996). All libraries mentioned above were constructed in conjunction with and are available from the laboratory of Roger Brent, (510) 647-0690 or [email protected]. The following individual investigators must be contacted directly: (18) J. Pugh, Fox Chase Cancer Center, Philadelphia, Pa.; (8,9) Vinyaka Prasad, Albert Einstein Medical Center New York, N.Y.; (57, 58) Gregory B. Martin, [email protected]; (11) Huifeng Niu, [email protected]; (16) Christer Hoog, [email protected]; (12) Kurt Gustin, [email protected]; (6) Stephan Witte, [email protected]. bInsert size ranges for pJG4-5 based libraries originally constructed in the Brent laboratory, which are now commercially available from Clontech, were reestimated by the company.

Transform the library 1. Grow an ∼20-ml culture of EGY48 or EGY191 containing a LexA-operator-lacZ reporter plasmid and pBait in Glu/CM −Ura, −His liquid dropout medium overnight at 30°C. For best results, the pBait and lacZ reporter plasmids should have been transformed into the yeast within ∼7 to 10 days of commencing a screen.

2. In the morning, dilute culture into 300 ml Glu/CM −Ura, −His liquid dropout medium to 2 × 106 cell/ml (OD600 = ∼0.10). Incubate at 30°C until the culture contains ∼1 × 107 cells/ml (OD600 = ∼0.50). 3. Centrifuge 5 min at 1000 to 1500 × g in a low-speed centrifuge at room temperature to harvest cells. Resuspend in 30 ml sterile water and transfer to 50-ml conical tube. 4. Centrifuge 5 min at 1000 to 1500 × g. Decant supernatant and resuspend cells in 1.5 ml TE buffer/0.1 M lithium acetate. 5. Add 1 µg library DNA in pJG4-5 and 50 µg high-quality sheared salmon sperm carrier DNA to each of 30 sterile 1.5-ml microcentrifuge tubes. Add 50 µl of the resuspended yeast solution from step 4 to each tube. The total volume of library and salmon sperm DNA added should be 500/plate) within 24 to 48 hr after plating on selective medium. Some investigators omit use of a Gal/Raff/CM −Ura, −His, −Trp, −Leu master plate, restreaking directly to a Glu/CM −Ura, −His, −Trp master plate as in step 19.

Test for Gal dependence The following steps test for Gal dependence of the Leu+ insert and lacZ phenotypes to confirm that they are attributable to expression of the library-encoded proteins. The GAL1 promoter is turned off and −Leu selection eliminated before reinducing. 18. Restreak from the Gal/Raff/CM −Ura, −His, −Trp, −Leu master dropout plate to a 100-mm Glu/CM −Ura, −His, −Trp master dropout plate. Incubate overnight at 30°C until colonies form. 19. Restreak or replica plate from this plate to the following plates: Glu/Xgal/CM −Ura, −His, −Trp Gal/Raff/Xgal/CM −Ura, −His, −Trp Glu/CM −Ura, −His, −Trp, −Leu Gal/Raff/CM −Ura, −His, −Trp, −Leu. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

At this juncture, colonies and the library plasmids they contain are tentatively considered positive if they are blue on Gal/Raff/Xgal plates but not blue or only faintly blue on Glu/Xgal plates, and if they grow on Gal/Raff/CM −Leu plates but not on Glu/CM −Leu plates. The number of positives obtained will vary drastically from bait to bait. How they are processed subsequently will depend on the number initially obtained and on the preference



of the individual investigator. If none are obtained using EGY48 as reporter strain, it may be worth attempting to screen a library from an additional tissue source. If a relatively small number (≤30) are obtained, proceed to step 20. However, sometimes searches will yield large numbers of colonies (>30 to 300, or more). In this case, there are several options. The first option is to warehouse the majority of the positives and work up the first 30 that arise; those growing fastest are frequently the strongest interactors. These can be checked for specificity, and restriction digests can be used to establish whether they are all independent cDNAs or represent multiple isolates of the same, or a small number, of cDNAs. If the former is true, it may be advisable to repeat the screen in a less sensitive strain background, as obtaining many different interactors can be a sign of low-affinity nonspecific background. Alternatively, if initial indications are that a few cDNAs are dominating the positives obtained, it may be useful to perform a filter hybridization with yeast (see Support Protocol 3) using these cDNAs as a probe to establish the frequency of their identification and exclude future reisolation of these plasmids. The second major option is to work up large numbers of positives to get a complete profile of isolated interactors (see Support Protocol 4). A third option is to temporarily warehouse the entire results of this first screen, and repeat the screen with a less sensitive strain such as EGY191, on the theory that it is most important to get stronger interactors first and a complete profile of interactors later. Finally, some investigators prefer to work up the entire set of positives initially obtained, even if such positives number in the hundreds. Particularly in this latter case, it is most effective to use Alternate Protocol 1 as a means to identify unique versus common positives.

Isolate plasmid from positive colonies by transfer into E. coli 20a. Transfer yeast plasmids directly into E. coli by following the protocol for direct electroporation (UNIT 1.8, Alternate Protocol 2). Proceed to step 22. 20b. Isolate plasmid DNA from yeast by the rapid miniprep protocol (UNIT 13.11) with the following alteration: after obtaining aqueous phase, precipitate by adding sodium acetate to 0.3 M final and 2 vol ethanol, incubate 20 min on ice, microcentrifuge 15 min at maximum speed, wash pellet with 70% ethanol, dry, and resuspend in 5 µl TE buffer. Cultures can be grown prior to the miniprep using Glu/CM −Trp to select only for the library plasmid; this may increase the proportion of bacterial colonies that contain the desired plasmid.

21. Use 1 µl DNA to electroporate (UNIT 1.8) into competent KC8 bacteria, and plate on LB/ampicillin plates. Incubate overnight at 37°C. Electroporation must be used to obtain transformants with KC8 because the strain is generally refractory to transformation.

22. Restreak or replica plate colonies arising on LB/ampicillin plates to bacterial defined minimal A medium plates containing vitamin B1 and supplemented with Ura, His, and Leu but lacking Trp. Incubate overnight at 37°C. Colonies that grow under these conditions contain the library plasmid. The yeast TRP1 gene can successfully complement the bacterial trpC-9830 mutation, allowing the library plasmid to be easily distinguished from the other two plasmids contained in the yeast. It is helpful to first plate transformations on LB/ampicillin plates, which provides a less stringent selection, followed by restreaking to bacterial minimal medium to maximize the number of colonies obtained (E.G., unpub. observ.).

23. Purify library-containing plasmids using a bacterial miniprep procedure (UNIT 1.6). Some investigators are tempted to immediately sequence DNAs obtained at this stage. At this point, it is still possible that none of the isolated clones will express bona fide interactors, and it is suggested that the following specificity tests be completed before committing the effort to sequencing (also see annotation to step 28).



Supplement 46

Because multiple 2ìm plasmids with the same marker can be simultaneously tolerated in yeast, it sometimes happens that a single yeast will contain two or more different library plasmids, only one of which encodes an interacting protein. The frequency of this occurrence varies in the hands of different investigators and may in some cases account for disappearing positives if the wrong cDNA is picked. When choosing colonies to miniprep, it is generally useful to work up at least two individual bacterial transformants for each yeast positive. These minipreps can then be restriction digested (UNIT 3.1) with EcoRI + XhoI to release cDNA inserts, and the size of inserts determined on an agarose minigel (UNIT 2.5A) to confirm that both plasmids contain the same insert. An additional benefit of analyzing insert size is that it may provide some indication as to whether repeated isolation of the same cDNA is occurring, generally a good indication concerning the biological relevance of the interactor. See Background Information for further discussion.

Assess positive colonies with specificity tests Much spurious background will have been removed by the previous series of controls. Other classes of false positives can be eliminated by retransforming purified plasmids into “virgin” LexA-operator-LEU2/LexA-operator-lacZ/pBait–containing strains that have not been subjected to Leu selection and verifying that interaction-dependent phenotypes are still observed. Such false positives could include mutations in the initial EGY48 yeast that favor growth on Gal medium, library-encoded cDNAs that interact with the LexA DNA-binding domain, or proteins that are sticky and interact with multiple biologically unrelated fusion domains. 24. In separate transformations, use purified plasmids from step 23 to transform yeast that already contain the following plasmids and are growing on Glu/CM −Ura, −His plates: EGY48 containing pSH18-34 and pBait EGY48 containing pSH18-34 and pRFHM-1 EGY48 containing pSH18-34 and a nonspecific bait (optional). 25. Plate each transformation mix on Glu/CM −Ura, −His, −Trp dropout plates and incubate 2 to 3 days at 30°C until colonies appear. 26. Create a Glu/CM −Ura, −His, −Trp master dropout plate for each library plasmid being tested. Streak adjacently five or six independent colonies derived from each of the transformation plates. Incubate overnight at 30°C. 27. Restreak or replica plate from this master dropout plate to the same series of test plates used for the actual screen: Glu/Xgal/CM −Ura, −His, −Trp Gal/Raff/Xgal/CM −Ura, −His, −Trp Glu/CM −Ura, −His, −Trp, −Leu Gal/CM −Ura, −His, −Trp, −Leu. True positive cDNAs should make cells blue on Gal/Raff/Xgal but not on Glu/Xgal plates, and should make them grow on Gal/Raff/CM −Leu but not Glu/CM −Leu dropout plates only if the cells contain LexA-bait. cDNAs that meet such criteria are ready to be sequenced (see legend to Fig. 20.1.3 for primer sequence) or otherwise characterized. Those cDNAs that also encode proteins that interact with either RFHM-1 or another nonspecific bait should be discarded.


It may be helpful to cross-check the isolated cDNAs with a database of cDNAs thought to be false positives. This database is available on the World Wide Web as a work in progress at http://www.fccc.edu:80/research/labs/golemis/InteractionTrapInWork.html. cDNAs reported to this database are generally those isolated only once in a screen in which obviously true interactive partners were isolated multiple times, cDNAs that may interact with more than one bait, or cDNAs for which the interaction does not appear to make biological sense



in the context of the starting bait. Although some proteins in this database may ultimately turn out in fact to associate with the bait that isolated them, they are by default unlikely to possess a unique and interesting function in the context of that bait if they are well represented in the database.

28. If appropriate, conduct additional specificity tests (see Support Protocol 5). Analyze and sequence positive isolates. The primer sequence for use with pJG4-5 is provided in the legend to Figure 20.1.4. DNA prepared from KC8 is generally unsuitable for dideoxy or automated sequencing even after use of Qiagen columns and/or cesium chloride gradients. Library plasmids to be sequenced should be retransformed from the KC8 miniprep stock (step 23) to a more amenable strain, such as DH5α, before sequencing is attempted.

RAPID SCREEN FOR INTERACTION TRAP POSITIVES Under some circumstances, it may be desirable to attempt the analysis of a large number of positives resulting from a two-hybrid screen. One such hypothetical example would be a bait with a leucine zipper or coiled coil known to dimerize with partner “A” that is highly expressed. In order to identify the rare novel partner “B”, it is necessary to work through the high background of “A” reisolates. This protocol uses the polymerase chain reaction (PCR) in a strategy to sort positives into redundant (multiple isolates) and unique classes prior to plasmid rescue from yeast, thus greatly reducing the number of plasmid isolations that must be performed. An additional benefit is that this protocol preidentifies positive clones containing one or multiple library plasmids; for those containing only one library plasmid, only a single colony needs to be prepared through KC8/DH5α.


Additional Materials (also see Basic Protocol 2) Yeast plated on Glu/CM −Ura, −His, −Trp master plate (see Basic Protocol 2, step 19) Lysis solution (see recipe) 10 µM forward primer (FP1): 5′-CGT AGT GGA GAT GCC TCC-3′ 10 µM reverse primer (FP2): 5′-CTG GCA AGG TAG ACA AGC CG-3′ Toothpicks or bacterial inoculating loops (UNIT 1.1), sterile 96-well microtiter plate Sealing tape, e.g., wide transparent tape 150- to 212-µm glass beads, acid-washed (UNIT 13.13) Vortexer with flat plate Additional reagents and equipment for performing an interactor hunt (see Basic Protocol 2), PCR amplification of DNA (UNIT 15.1), agarose gel electrophoresis (UNITS 2.5A & 2.6), restriction endonuclease digestion (UNIT 3.1), electroporation (UNIT 1.8), and miniprep isolation of bacterial DNA (UNIT 1.6) 1. Perform an interactor hunt (see Basic Protocol 2, steps 1 to 19). 2. Use a sterile toothpick or bacterial inoculating loop to transfer yeast from the Glu/CM, −Ura, −His, −Trp master plate into 25 µl lysis solution in a 96-well microtiter plate. Seal the wells of the microtiter plate with sealing tape and incubate 1.5 to 3.5 hr at 37°C with shaking. The volume of yeast transferred should not exceed ∼2 to 3 ìl of packed pellet; larger quantities of yeast will reduce quality of the DNA. DNA can be efficiently recovered from master plates that have been stored up to 1 week at 4°C. If yeast have been previously gridded on master plates, transfer to microtiter plates can be facilitated by using a multicolony replicator.



Supplement 46

3. Remove tape from the plate, add ∼25 µl acid-washed glass beads to each well, and reseal with the same tape. Firmly attach the microtiter plate to a flat-top vortexer, and vortex 5 min at medium-high power. The microtiter plate can be attached to the vortexer using 0.25-in (0.64-cm) rubber bands.

4. Remove the tape and add ∼100 µl sterile water to each well. Swirl gently to mix, then remove sample for step 5. Press the tape back firmly to seal the microtiter plate and place in the freezer at −20°C for storage. 5. Amplify 0.8 to 2.0 µl of sample by standard PCR (UNIT 15.1) in a ∼30-µl volume using 3 µl each of the forward primer FP1 and the reverse primer FP2. Perform PCR using the following cycles: Initial step: 31 cycles:

2 min 45 sec 45 sec 45 sec

94°C 94°C 56°C 72°C.

These conditions have been used successfully to amplify fragments up to 1.8 kb in length; some modifications, such as extension of elongation time, are also effective.

6. Load 20 µl of the PCR reaction product on a 0.7% low melting temperature agarose gel (UNIT 2.6) to resolve PCR products. Based on insert sizes, group the obtained interactors in families, i.e., potential multiple independent isolates of identical cDNAs. Reserve gel until results of step 7 are obtained. No special precautions are needed for storing the gel. Since HaeIII digests typically yield rather small DNA fragments, running the second gel does not take a lot of time. Usually, the delay does not exceed 45 to 60 min, during which time the first gel may be stored in a gel box at room temperature or wrapped in plastic wrap at 4°C.

7. While the gel is running, use the remaining 10 µl of PCR reaction product for a restriction endonuclease digestion with HaeIII in a digestion volume of ∼20 µl (UNIT 3.1). Based on analysis of the sizes of undigested PCR products in the gel (step 6), rearrange the tubes with HaeIII digest samples so that those thought to represent a family are side by side. Resolve the digests on a 2% to 2.5% agarose gel (UNIT 2.5A). Most restriction fragments will be in the 200-bp to 1.0-kb size range so using a long gel run is advisable. This analysis should produce a distinct fingerprint of insert sizes and allow definition of library cDNAs as unique isolates or related groups. A single positive yeast will sometimes contain multiple library plasmids. An advantage of this protocol is the ready detection of multiple library plasmids in PCR reactions; thus, following subsequent bacterial transformations, only a single TRP1 colony would need to be analyzed unless multiple plasmids were already known to be present.

8. Isolate DNA fragments from the low melting temperature agarose gel (step 6). If inspection of the banding pattern on the two gels suggests that a great many reisolates of a small number of cDNAs are present, it may be worthwhile to immediately sequence PCR products representative of these clusters, but it is generally still advisable to continue through specificity tests before doing so. If the PCR products are sequenced, the FP1 forward primer works well in automated sequencing of PCR fragments, but the FP2 primer is only effective in sequencing from purified plasmid. In general, priming from the AT-rich ADH terminator downstream of the polylinker/cDNA in library plasmid is less efficient than from upstream of the cDNA, and it is hard to design effective primers in this region. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

9. Remove the microtiter plate of lysates from the freezer, thaw it, and remove 2 to 4 µl of lysed yeast for each desired positive. Electroporate DNA into either DH5α or KC8 E. coli as appropriate, depending on the choice of bait and reporter plasmids (see



Table 20.1.1 and see Background Information for further information). Refreeze the plate as a DNA reserve in case bacteria fail to transform on the first pass. KC8 E. coli should be used for electroporation when the original reagents pEG202/pJG45/pJK101 are used for the interaction trap. An additional strength of this protocol is that it identifies redundant clones before transfer of plasmids to bacteria, thus reducing the amount of work required in cases where plasmid identity can be unambiguously assigned. However, although restriction endonuclease digestion and PCR analysis are generally highly predictive, they are not 100% certain methods for estimating cDNA identity. Thus, if there is any doubt about whether two cDNAs are the same, investigators are urged to err on the side of caution.

10. Prepare a miniprep of plasmid DNA from the transformed bacteria (UNIT 1.6) and perform yeast transformation and specificity assessment (see Basic Protocol 2, steps 24 to 28). PERFORMING A HUNT BY INTERACTION MATING An alternative way of conducting an interactor hunt is to mate a strain that expresses the bait protein with a strain that has been pretransformed with the library DNA, and screen the resulting diploid cells for interactors (Bendixen et al., 1994; Finley and Brent, 1994). This “interaction mating” approach can be used for any interactor hunt, and is particularly useful in three special cases. The first case is when more than one bait will be used to screen a single library. Interaction mating allows several interactor hunts with different baits to be conducted using a single high-efficiency yeast transformation with library DNA. This can be a considerable savings, since the library transformation is one of the most challenging tasks in an interactor hunt. The second case is when a constitutively expressed bait interferes with yeast viability. For such baits, performing a hunt by interaction mating avoids the difficulty associated with achieving a high-efficiency library transformation of a strain expressing a toxic bait. Moreover, the actual selection for interactors will be conducted in diploid yeast, which are more vigorous than haploid yeast and can better tolerate expression of toxic proteins. The third case is when a bait cannot be used in a traditional interactor hunt using haploid yeast strains (see Basic Protocol 2) because it activates transcription of even the least sensitive reporters. In diploids the reporters are less sensitive to transcription activation than they are in haploids. Thus, the interaction mating hunt provides an additional method to reduce background from transactivating baits.


In the protocol described below, the library DNA is used to transform a strain with a LEU2 reporter (e.g., EGY48). This pretransformed library strain is then frozen in many aliquots, which can be thawed and used for individual interactor hunts. The bait is expressed in a strain of mating type opposite to that of the pretransformed library strain, and also bearing the lacZ reporter. A hunt is conducted by mixing the pretransformed library strain with the bait strain and allowing diploids to form on YPD medium overnight. The diploids are then induced for expression of the library-encoded proteins and screened for interactors as in Basic Protocol 2. NOTE: Strain combinations other than those described below can also be used in an interaction-mating hunt. The key to choosing the strains is to ensure that the bait and prey strains are of opposite mating types and that both have auxotrophies to allow selection for the appropriate plasmids and reporter genes. Also, once the bait plasmid and lacZ reporter plasmid have been introduced into the bait strain, and the library plasmids have been introduced into the library strain, the resulting bait strain and library strain must each have auxotrophies that can be complemented by the other, so that diploids can be selected.



Supplement 46

Additional Materials (also see Basic Protocols 1 and 2) Yeast strains: either RFY206 (Finley and Brent, 1994), YPH499 (Sikorski and Hieter, 1989; ATCC #6625), or an equivalent MATa strain with auxotrophic markers ura3, trp1, his3, and leu2 YPD liquid medium (UNIT 13.1) Glu/CM –Trp plates: CM dropout plates −Trp (UNIT 13.1) supplemented with 2% glucose pJG4-5 library vector (Fig. 20.1.6), empty 100-mm YPD plates (UNIT 13.1) Additional reagents and equipment for lithium acetate transformation of yeast (UNIT 13.7) Construct the bait strain The bait strain will be a MATa yeast strain (mating type opposite of EGY48) containing a lacZ reporter plasmid like pSH18-34 and the bait-expressing plasmid, pBait. 1. Perform construction of the bait plasmid (pBait; see Basic Protocol 1, step 1). 2. Cotransform the MATa yeast strain (e.g., either RFY206 or YPH499) with pBait and pSH18-34 using the lithium acetate method (UNIT 13.7). Select transformants on Glu/CM –Ura,–His plates by incubating plates at 30°C for 3 to 4 days until colonies form. Combine 3 colonies for all future tests and for the mating hunt. The bait strain (RFY206/pSH18-34/pBait or YPH499/pSH18-34/pBait) can be tested by immunoblotting to ensure that the bait protein is expressed (see Support Protocol 1). Synthesis and nuclear localization of the bait protein can also be tested by the repression assay (see Basic Protocol 1, steps 8 to 12).

3. Optional: Assay lacZ gene activation in the bait strain (see Basic Protocol 1, steps 4 to 7). If the bait activates the lacZ reporter, a less sensitive lacZ reporter plasmid (Table 20.1.1), or an integrated version of the lacZ reporter should be tried. A bait that strongly activates the lacZ reporters usually cannot be used in a hunt based on selection of interactors with the LEU2 reporter, because the LEU2 reporters are more sensitive than the lacZ reporters. However, both reporters are less sensitive to activation by a bait in diploid cells, as compared to haploid cells. Thus, a more important test of the transactivation potential of a bait is to test the leucine requirement of diploid cells expressing it, as described in steps 6 to 20, below.

Prepare the pretransformed library strain (EGY48 + library plasmids) 4. Perform a large-scale transformation of EGY48 with library DNA using the lithium acetate method (see Basic Protocol 2, steps 1 to 8, except start with EGY48 bearing no other plasmids). To prepare for transformation, grow EGY48 in YPD liquid medium. Select library transformants on Glu/CM –Trp plates by incubating 3 days at 30°C. 5. Collect primary transformants by scraping plates, washing yeast, and resuspending in 1 pellet vol glycerol solution (see Basic Protocol 2, steps 9 to 12). Freeze 0.2 to 1.0 ml aliquots at −70° to −80°C.


The cells will be stable for at least 1 year. Refreezing a thawed aliquot will result in loss of viability. Thus, many frozen aliquots should be made, so that each thawed aliquot can be discarded after use.



Prepare the pretransformed control strain (EGY48 + pJG4-5) 6. Transform EGY48 grown in YPD liquid medium with the empty library vector, pJG4-5, using the lithium acetate method (UNIT 13.7). Select transformants on Glu/CM –Trp plates by incubating 3 days at 30°C. 7. Pick and combine three transformant colonies and use them to inoculate 30 ml of Glu/CM –Trp medium. Incubate 15 to 24 hr at 30°C (to OD600 >3). 8. Centrifuge 5 min at 1000 to 1500 × g, room temperature, and remove supernatant. Resuspend in 10 ml sterile water to wash cells. 9. Centrifuge 5 min at 1000 to 1500 × g, room temperature, and remove supernatant. Resuspend in 1 pellet vol glycerol solution and freeze 100-µl aliquots at −70° to −80°C. Determine plating efficiency of pretransformed library and pretransformed control strains 10. After freezing (at least 1 hr) thaw an aliquot of each pretransformed strain (from step 5 and step 9) at room temperature. Make several serial dilutions in sterile water, including aliquots diluted 105-fold, 106-fold, and 107-fold. Plate 100 µl of each dilution on 100-mm Glu/CM –Trp plates and incubate 2 to 3 days at 30°C. 11. Count the colonies and determine the number of colony-forming units (cfu) per aliquot of transformed yeast. The plating efficiency for a typical library transformation and for the control strain will be ∼1 × 108 cfu per 100 ìl.

Mate the bait strain with the pretransformed library strain and the pretransformed control strain In steps 12 through 20, an interactor hunt is conducted concurrently with testing LEU2 reporter activation by the bait itself. For most baits, this approach will be the quickest way to isolate interactors. However, for some baits, such as those that have a high transactivation potential, or those that affect yeast mating or growth, steps 12 through 20 will serve as a pilot experiment to determine the optimal parameters for a subsequent hunt. 12. Grow a 30-ml culture of the bait strain in Glu/CM –Ura,–His liquid dropout medium to mid to late log phase (OD600 = 1.0 to 2.0, or 2 to 4 × 107cells/ml). A convenient way to grow the bait strain is to inoculate a 5-ml culture with approximately three colonies from a plate and grow it overnight at 30°C with shaking. In the morning, measure the OD600, dilute into a 30-ml culture to a final OD600 = 0.2, and grow at 30°C with shaking. The culture should reach mid to late log phase before the end of the day.

13. Centrifuge the culture 5 min at 1000 to 1500 × g, room temperature, to harvest cells. Resuspend the cell pellet in sterile water to make a final volume of 1 ml. This should correspond to ∼1 × 109 cells/ml.

14. Set up two matings. In one sterile microcentrifuge tube mix 200 µl of the bait strain with 200 µl of a thawed aliquot of the pretransformed control strain from step 9. In a second microcentrifuge tube mix 200 µl of the bait strain with ∼1 × 108 cfu (∼0.1 to 1 ml) of the pretransformed library strain from step 5. The library mating should be set up so that it contains a ∼2-fold excess of bait strain cfu over pretransformed library strain cfu. Because the bait strain was harvested in log phase, most of the cells will be viable (i.e., cells/ml = ∼cfu/ml), and the number of cfu can be sufficiently estimated from optical density (1 OD600 = ∼2 × 107 cells/ml). Under these conditions, ∼10% of the cfu in the pretransformed library strain will mate with the bait



Supplement 46

strain. Thus, a complete screen of 107 library transformants will require a single mating with at least 108 cfu of the pretransformed library strain and at least 2 × 108 cfu of the bait strain. To screen more library transformants, set up additional matings. The number of pretransformed library transformants to screen depends on the size of the library and the number of primary transformants obtained in step 5. If the size of the library is larger than the number of transformants obtained in step 5, the goal will be to screen all of the yeast transformants. In this case, complete screening of the library will require additional transformations of EGY48 and additional interactor hunts. If the size of the library is smaller than the number of transformants obtained in step 5, the goal will be to screen at least a number of transformants equivalent to the size of the library.

15. Centrifuge each cell mixture for 5 min at 1000 to 1500 × g, pour off medium, and resuspend cells in 200 µl YPD medium. Plate each suspension on a 100-mm YPD plate. Incubate 12 to 15 hr at 30°C. 16. Add ∼1 ml of Gal/Raff/CM –Ura, –His, –Trp to the lawns of mated yeast on each plate. Mix the cells into the medium using a sterile applicator stick. 17. Transfer each slurry of mated cells to a 500-ml flask containing 100 ml of Gal/Raff/CM –Ura, –His, –Trp dropout medium. Incubate with shaking 6 hr at room temperature to induce the GAL1 promoter, which drives expression of the cDNA library. 18. Centrifuge the cell suspensions 5 min at 1000 to 1500 × g, room temperature, to harvest the cells. Wash by resuspending in 30 ml of sterile water and centrifuging again. Resuspend each pellet in 5 ml sterile water. Measure OD600 and, if necessary, dilute to a final concentration of ∼1 × 108 cells/ml. This is a mixture consisting of haploid cells that have not mated and diploid cells. Under a microscope, the two cell types can be distinguished by size (diploids are ∼1.7× bigger than haploids) and shape (diploids are slightly oblong and haploids are spherical). Because diploids grow faster than haploids, this mixture will contain ∼10% to 50% diploid cells. The actual number of diploids will be determined by plating dilutions on –Ura, –His, –Trp medium, which will not support the growth of the parental haploids.

19. For each mating make a series of 1⁄10 dilutions in sterile water, at least 200 µl each, to cover a 106-fold concentration range. Plate 100 µl from each tube (undiluted, 10−1, 10−2, 10−3, 10−4, 10−5, and 10−6 dilution) on 100-mm Gal/Raff/CM –Ura, –His, –Trp, –Leu plates. Plate 100 µl from the 10−4, 10−5, and 10−6 tubes on 100-mm Gal/Raff/CM –Ura, –His, –Trp plates. Incubate plates at 30°C. Count the colonies on each plate after 2 to 5 days. 20. For the mating with the pretransformed library, prepare an additional 3 ml of a 10−1 dilution. Plate 100 µl of the 10−1 dilution on each of 20 100-mm Gal/Raff/CM –Ura, –His, –Trp, –Leu plates. Also plate 100 µl of the undiluted cells on each of 20 100-mm Gal/Raff/CM –Ura, –His, –Trp, –Leu plates. Incubate at 30°C. Pick Leu+ colonies after 2 to 5 days and characterize them beginning with step 17 of Basic Protocol 2.


The number of Leu+ colonies to pick to ensure that all of the pretransformed library has been screened depends on the transactivation potential of the bait protein itself. The transactivation potential is expressed as the number of Leu+ colonies that grow per cfu (Leu+/cfu) of the bait strain mated with the control strain, as determined in step 19 of this protocol. It can be calculated as the ratio of the number of colonies that grow on Gal/Raff/CM –Ura,–His, –Trp, –Leu to the number of colonies that grow on Gal/Raff/CM –Ura, –His, –Trp for a given dilution of the mating between the bait strain and the control strain. A bait with essentially no transactivation potential will produce less than 10−6 Leu+/cfu. For a bait to be useful in an interactor hunt it should not transactivate more than 10−4 Leu+/cfu.



To screen all of the pretransformed library, it will be necessary to pick a sufficient number of Leu+ colonies in addition to background colonies produced by the transactivation potential of the bait itself. Thus, the minimum number of Leu+ colonies that should be picked in step 20 of this protocol is given by: (transactivation potential, Leu+/cfu) × (# library transformants screened). For example, if 107 library transformants were obtained in step 2 (and at least 108 cfu of these transformants were mated with the bait strain in step 14, since only ∼10% will form diploids), and the transactivation potential of the bait is 10−4 Leu+/cfu, then at least 1000 Leu+ colonies must be picked and characterized. In other words, if the rarest interactor is present in the pretransformed library at a frequency of 10−7, to find it one needs to screen through at least 107 diploids from a mating of the library strain. However, at least 1000 of these 107 diploids would be expected to be Leu+ due to the bait background if the transactivation potential of the bait is 10−4. The true positives will be distinguished from the bait background in the next step by the galactose dependence of their Leu+ and lacZ+ phenotypes.

PREPARATION OF PROTEIN EXTRACTS FOR IMMUNOBLOT ANALYSIS To confirm that the bait fusion protein constructed in Basic Protocol 1 is synthesized properly, a crude lysate is prepared for SDS-PAGE and immunoblot analysis (UNITS 10.2 & 10.8). The presence of the target protein is detected by antibody to LexA or the fusion domain.

SUPPORT PROTOCOL 1

Materials Master plates with pBait-containing positive and control yeast on Glu/CM −Ura, −His dropout medium (see Basic Protocol 1, step 4) Glu/CM −Ura, −His dropout liquid medium: CM dropout plates −Ura, −His (UNIT 13.1) supplemented with 2% glucose 2× Laemmli sample buffer (see recipe) Antibody to fusion domain or LexA: monoclonal antibody to LexA (Clontech, Invitrogen) or polyclonal antibody to LexA (available by request from R. Brent or E. Golemis) 30°C incubator 100°C water bath Additional reagents and equipment for SDS-PAGE (UNIT 10.2) and immunoblotting and immunodetection (UNIT 10.8) 1. From the master plates, start a 5-ml culture in Glu/CM −Ura, −His liquid medium for each bait being tested and for a positive control for protein expression (i.e., RFHMI or SH17-4). Incubate overnight at 30°C. For each construct assayed, it is a good idea to grow colonies from at least two primary transformants, as levels of bait expression are sometimes heterogenous.

2. From each overnight culture, start a fresh 5-ml culture in Glu/CM −Ura, −His at OD600 = ∼0.15. Incubate again at 30°C. 3. When the culture has reached OD600 = 0.45 to 0.7 (∼4 to 6 hr), remove 1.5 ml to a microcentrifuge tube. For some LexA fusion proteins, levels of the protein drop off rapidly in cultures approaching stationary phase. This is due to a combination of the diminishing activity of the ADH1 promoter in late growth phases and the relative instability of particular fusion domains. Thus, it is not a good idea to let cultures become saturated in the hopes of obtaining a higher yield of protein.



Supplement 46

4. Microcentrifuge cells 3 min at 13,000 × g, room temperature. When the pellet is visible, remove the supernatant. Inspection of the tube should reveal a pellet ∼1 to 3 ìl in volume. If the pellet is not visible, microcentrifuge another 3 min.

5. Working rapidly, add 50 µl of 2× Laemmli sample buffer to the visible pellet in the tube, vortex, and place the tube on dry ice. Samples may be frozen at −70°C.

6. Transfer frozen sample directly to a boiling water bath or a PCR machine set to cycle at 100°C. Boil 5 min. 7. Microcentrifuge 5 sec at maximum speed to pellet large cellular debris. 8. Perform SDS-PAGE (UNIT 10.2) using 20 to 50 µl sample per lane. 9. To detect the protein, immunoblot and analyze (UNIT 10.8) using antibody to the fusion domain or LexA. SUPPORT PROTOCOL 2

PREPARATION OF SHEARED SALMON SPERM CARRIER DNA This protocol generates high-quality sheared salmon sperm DNA (sssDNA) for use as carrier in transformation (Basic Protocol 2). This DNA is also suitable for other applications where high-quality carrier DNA is needed (e.g., hybridization). This protocol is based on Schiestl and Gietz (1989). For more details of phenol extraction or other DNA purification methods, consult UNIT 2.1A. Materials High-quality salmon sperm DNA (e.g., sodium salt from salmon testes, Sigma or Boehringer Mannheim), desiccated TE buffer, pH 7.5 (APPENDIX 2), sterile TE-saturated buffered phenol (UNIT 2.1A) 1:1 (v/v) buffered phenol/chloroform Chloroform 3 M sodium acetate, pH 5.2 (APPENDIX 2) 100% and 70% ethanol, ice cold Magnetic stirring apparatus and stir-bar, 4°C Sonicator with probe 50-ml conical centrifuge tube High-speed centrifuge and appropriate tube 100°C and ice-water baths 1. Dissolve desiccated high-quality salmon sperm DNA in TE buffer, pH 7.5, at a concentration of 5 to 10 mg/ml by pipetting up and down in a 10-ml glass pipet. Place in a beaker with a stir-bar and stir overnight at 4°C to obtain a homogenous viscous solution. It is important to use high-quality salmon sperm DNA. Sigma Type III sodium salt from salmon testes has worked well, as has a comparable grade from Boehringer Mannheim. Generally it is convenient to prepare 20- to 40-ml batches at a time.

2. Shear the DNA by sonicating briefly using a large probe inserted into the beaker. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

The goal of this step is to generate sheared salmon sperm DNA (sssDNA) with an average size of 7 kb, but ranging from 2 to 15 kb. Oversonication (such that the average size is closer to 2 kb) drastically decreases the efficacy of carrier in enhancing transformation. The original version of this protocol (Schiestl and Gietz, 1989) called for two 30-sec pulses at



Table 20.1.4

System

Two-hybrid System Variantsa

DNA-binding Activation Selection domain domain

Two-hybrid Interaction trap “Improved two-hybrid” Modified two-hybrid KISS Contingent replication

GAL4 LexA GAL4 LexA GAL4 GAL4

GAL4 B42 GAL4 VP16 VP16 VP16

Activation of lacZ, HIS3 Activation of LEU2, lacZ Activation of HIS3, lacZ Activation of HIS3, lacZ Activation of CAT, hygr Activation of T-Ag, replication of plasmids

Reference Chien et al., 1991 Gyuris et al., 1993 Durfee at al., 1993 Vojtek at al., 1993 Fearon et al., 1992 Vasavada et al., 1991

aAbbreviations: CAT, chloramphenicol transferase gene; hygr, hygromycin resistance gene; T-Ag, viral large T antigen.

three-quarter power, but optimal conditions vary between sonicators. The first time this protocol is performed, it is worthwhile to sonicate briefly, then test the size of the DNA by running out a small aliquot alongside molecular weight markers on an agarose gel containing ethidium bromide. The DNA can be sonicated further if needed.

3. Once DNA of the appropriate size range has been obtained, extract the sssDNA solution with an equal volume of TE-saturated buffered phenol in a 50-ml conical tube, shaking vigorously to mix. 4. Centrifuge 5 to 10 min at 3000 × g, room temperature, or until clear separation of phases is obtained. Transfer the upper phase containing the DNA to a clean tube. 5. Repeat extraction using 1:1 (v/v) buffered phenol/chloroform, then chloroform alone. Transfer the DNA into a tube suitable for high-speed centrifugation. 6. Precipitate the DNA by adding 1⁄10 vol of 3 M sodium acetate and 2.5 vol of ice-cold 100% ethanol. Mix by inversion. Centrifuge 15 min at ∼12,000 × g, room temperature. 7. Wash the pellet with 70% ethanol. Briefly dry either by air drying, or by covering one end of the tube with Parafilm with a few holes poked in and placing the tube under vacuum. Resuspend the DNA in sterile TE buffer at 5 to 10 mg/ml. Do not overdry the pellet or it will be very difficult to resuspend.

8. Denature the DNA by boiling 20 min in a 100°C water bath. Then immediately transfer the tube to an ice-water bath. 9. Place aliquots of the DNA in microcentrifuge tubes and store frozen at −20°C. Thaw as needed. DNA should be boiled again briefly (5 min) immediately before addition to transformations. Before using a new batch of sssDNA in a large-scale library transformation, it is a good idea to perform a small-scale transformation using suitable plasmids to determine the transformation efficiency. Optimally, use of sssDNA prepared in the manner described will yield transformation frequencies of >105 colonies/ìg input plasmid DNA.



Supplement 46

SUPPORT PROTOCOL 3

YEAST COLONY HYBRIDIZATION This protocol is adapted from a modification of the classic protocol of Grunstein and Hogness (1975; Kaiser et al., 1994). It is primarily useful when a large number of putative interactors has been obtained, and initial minipreps and restriction digests have indicated that many of them derive from a small number of cDNAs; these cDNAs can then be used as probes to screen and eliminate identical cDNAs from the pool. Materials Glu/CM −Trp plates: CM dropout plates −Trp (UNIT 13.1) supplemental with 2% glucose Master dropout plate of yeast positive for Gal dependence (see Basic Protocol 2, step 18) 1 M sorbitol/20 mM EDTA/50 mM DTT (prepare fresh) 1 M sorbitol/20 mM EDTA 0.5 M NaOH 0.5 M Tris⋅Cl (pH 7.5)/6× SSC (APPENDIX 2) 2× SSC (APPENDIX 2) 100,000 U/ml β-glucuronidase (type HP-2 crude solution from Helix pomatia; Sigma) 82-mm circular nylon membrane, sterile Whatman 3MM paper 80°C vacuum oven or UV cross-linker Additional reagents and equipment for bacterial filter hybridization (UNITS 6.3 & 6.4) 1. Place a sterile nylon membrane onto a Glu/CM −Trp dropout plate. From the master dropout plate of Gal-dependent positives, gently restreak positives to be screened onto the membrane and mark the membrane to facilitate future identification of hybridizing colonies. Grow overnight (∼12 hr) at 30°C. Growth for extended periods of time (i.e., 24 hr) may result in difficulty in obtaining good lysis. It is a good idea to streak positive and negative controls for the cDNAs to be hybridized on the membrane.

2. Remove membrane from plate. Air dry briefly. Incubate ∼30 min on a sheet of Whatman 3MM paper saturated with 1 M sorbitol/20 mM EDTA/50 mM DTT. Optionally, before commencing chemical lysis, membranes can be placed at −70°C for 5 min, then thawed at room temperature for one or more cycles to enhance cell wall breakage.

3. Cut a piece of Whatman 3MM paper to fit inside a 100-mm petri dish. Place the paper disc in the dish and saturate with 100,000 U/ml β-glucuronidase diluted 1:500 in 1 M sorbitol/20 mM EDTA (2 µl glucuronidase per ml of sorbitol/EDTA to give 200 U/ml final). Layer nylon membrane on dish, cover dish, and incubate up to 6 hr at 37°C until >80% of the cells lack a cell wall. The extent of cell wall removal can be determined by removing a small quantity of cells from the filter to a drop of 1 M sorbitol/20 mM EDTA on a microscope slide and observing directly with a phase-contrast microscope at ≥60× magnification. Cells lacking cell wall are nonrefractile.

4. Place membrane on Whatman 3MM paper saturated with 0.5 M NaOH for ∼8 to 10 min. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins

5. Place membrane on Whatman 3MM paper saturated with 0.5 M Tris⋅Cl (pH 7.5)/6× SSC for 5 min. Repeat with a second sheet of Whatman 3MM paper.



6. Place membrane on Whatman 3MM paper saturated with 2× SSC for 5 min. Then place membrane on dry Whatman paper to air dry for 10 min. 7. Bake membrane 90 min at 80°C in vacuum oven or UV cross-link. 8. Process as for bacterial filter hybridization (UNITS 6.3 & 6.4), hybridizing the membrane with probes complementary to previously isolated cDNAs. When selecting probes, either random-primed cDNAs or oligonucleotides complementary to the cDNA sequence may be used. If the cDNA is a member of a protein family, it may be advantageous to use oligonucleotides to avoid inadvertently excluding genes related but not identical to those initially obtained.

MICROPLATE PLASMID RESCUE In some cases, it is desirable to isolate plasmids from a large number of positive colonies (Basic Protocol 2, steps 18 and 19). The protocol described below is a batch DNA preparation protocol developed by Steve Kron (University of Chicago, Chicago, Ill.) as a scale-up of a basic method developed by Manuel Claros (Laboratoire de Génétique Moleculaire, Paris, France).

SUPPORT PROTOCOL 4

Materials 2× Glu/CM −Trp liquid medium: 2× CM −Trp liquid medium (UNIT 13.1) supplemented with 4% glucose Master plate of Gal-dependent yeast colonies (see Basic Protocol 2, step 18) Rescue buffer: 50 mM Tris⋅Cl (pH 7.5)/10 mM EDTA/0.3% (v/v) 2-mercaptoethanol (prepare fresh) Lysis solution: 2 to 5 mg/ml Zymolyase 100T/rescue buffer or 100,000 U/ml β-glucuronidase (type HP-2 crude solution from Helix pomatia; Sigma) diluted 1:50 in rescue buffer 10% (w/v) SDS 7.5 M ammonium acetate (APPENDIX 2) Isopropanol 70% ethanol TE buffer, pH 8.0 (APPENDIX 2) 24-well microtiter plates Centrifuge with microplate holders, refrigerated Repeating micropipettor 37°C rotary shaker Grow yeast cultures 1. Aliquot 2 ml of 2× Glu/CM −Trp medium into each well of a 24-well microtiter plate. Into each well, pick a putative positive colony. Grow overnight with shaking at 30°C. The 2× minimal medium is used to maximize the yield of yeast. Four plates can generally be handled conveniently at once, based on the number that can be centrifuged simultaneously.

2. Centrifuge 5 min at 1500 × g, 4°C. Shake off supernatant with a snap and return the plate to upright. 3. Swirl or lightly vortex the plate to resuspend cell pellets in remaining liquid. Add 1 ml water to each well and swirl lightly. Cell pellets can most easily be resuspended in residual liquid before adding new solutions. Addition of liquid can be accomplished using a repeating pipettor.



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4. Centrifuge 5 min at 1500 × g, 4°C. Shake off supernatant and resuspend pellet. Add 1 ml rescue buffer. 5. Centrifuge 5 min at 1500 × g, 4°C. Shake off supernatant and resuspend pellet in the small volume of liquid remaining in the plate. Lyse cells 6. To each well, add 25 µl lysis solution. Swirl or vortex to mix. Incubate (with cover on) on a rotary shaker ∼1 hr at 37°C. Lysis solution need not be completely dissolved before use. By 1 hr, lysis should be obvious as coagulation of yeast into a white precipitate. Susceptibility of yeast strains to lytic enzymes varies. If lysis occurs rapidly, then less lytic enzyme should be used. If the lysis step is allowed to go too far, too much of the partially dissolved cell wall may contaminate the final material. Lysis can be judged by examining cells with a phase-contrast microscope. Living cells are white with a dark halo and dead cells are uniformly gray. Lysis leads to release of granular cell contents into the medium. Once cells are mostly gray and many are disrupted, much of the plasmid should have been released.

7. To each well, add 25 µl of 10% SDS. Mix gently by swirling to completely disperse the precipitates. Allow plates to sit 1 min at room temperature. At this point, the wells should contain a clear, somewhat viscous solution.

Purify plasmid 8. To each well, add 100 µl of 7.5 M ammonium acetate. Swirl gently, then incubate 15 min at −70°C or −20°C until frozen. Addition of acetate should result in the formation of a massive white precipitate of cell debris and SDS. The freezing step appears to improve removal of inhibitors of E. coli transformation.

9. Remove plate from freezer. Once it begins to thaw, centrifuge 15 min at 3000 × g, 4°C. Transfer 100 to 150 µl of the resulting clear supernatants to clean 24-well plates. In general, some contamination of the supernatant with pelleted material cannot be avoided. However, it is better to sacrifice yield in order to maintain purity.

10. To each well, add ∼0.7 vol isopropanol. Mix by swirling and allow to precipitate 2 min at room temperature. A cloudy fine precipitate should form immediately after isopropanol is added.

11. Centrifuge 15 min at 3000 × g, 4°C. Shake off supernatant with a snap. 12. To each well, add ∼1 ml cold 70% ethanol. Mix by swirling, centrifuge 5 min at 3000 × g, 4°C. Shake off supernatant with a snap, invert plates and blot well onto paper towel. Allow plates to air dry. 13. To each well, add 100 µl TE buffer. Swirl well and allow to rest on bench several minutes, until the pellets appear fully dissolved. Transfer preps to microcentrifuge tubes or 96-well plates for storage at −20°C.


One to five microliters of each of the resulting preparations can be used to transform competent E. coli: for KC8, electroporation should be used (see Basic Protocol 2, step 21). Sometimes, the yield of transformants is low if E. coli carrying plasmids are not permitted time to increase the plasmid copy number above a critical threshold before the cells are placed on selective medium. Allow plenty of time for cells to express antibiotic resistance or the TRP1 gene before plating. If insufficient numbers of colonies are obtained by this approach, the final plasmid preparation can be resuspended in 20 ìl instead of 100 ìl TE buffer to concentrate the DNA stock.



ADDITIONAL SPECIFICITY SCREENING The three test plasmids outlined (pSH18-34, pRFHM1, and pEG202; see Basic Protocol 2, step 24) represent a minimal test series. If other LexA-bait proteins that are related to the bait protein used in the initial library screen are available, substantial amounts of information can be gathered by additional specificity tests. For example, if the initial bait protein was LexA fused to the leucine zipper of c-Fos, specificity screening of interactor-hunt positives against the leucine zippers of c-Jun or GCN4 in addition to that of c-Fos might allow discrimination between proteins that are specific for fos versus those that generically associate with leucine zippers.

SUPPORT PROTOCOL 5


Glycerol solution 65% (v/v) glycerol, sterile 0.1 M MgSO4 25 mM Tris⋅Cl, pH 8.0 (APPENDIX 2) Store up to 1 year at room temperature Laemmli sample buffer, 2× 10% (v/v) 2-mercaptoethanol (2-ME) 6% (w/v) SDS 20% (v/v) glycerol 0.2 mg/ml bromphenol blue 0.025× Laemmli stacking buffer (see recipe; optional) Store up to 2 months at room temperature This reagent can conveniently be prepared 10 ml at a time.

Laemmli stacking buffer, 2.5× 0.3 M Tris⋅Cl, pH 6.8 0.25% (w/v) SDS Store up to 1 month at 4°C Lysis solution 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 10 mM EDTA 0.3% (v/v) 2-mercaptoethanol (2-ME), added just before use 2% (v/v) β-glucuronidase from Helix pomatia (Type HP-2; Sigma), added just before use COMMENTARY Background Information Interaction-based cloning is derived from three experimental observations. In the first, Brent and Ptashne (1985) demonstrated that it was possible to assemble a novel, functional transcriptional activator by fusing the DNAbinding domain from one protein, LexA, to the activation domain from a second protein, GAL4. This allowed the use of a single reporter system containing a single DNA-binding motif, the LexA operator, to study transcriptional ac-

tivation by any protein of interest. In the second, Ma and Ptashne (1988) built on this work to demonstrate that the activation domain could be brought to DNA by interaction with a DNAbinding domain. In the third, Fields and Song (1989), working independently of Ma and Ptashne, used two yeast proteins, SNF1 and SNF4, to make an SNF1 fusion to the DNAbinding domain of GAL4 and an SNF4 fusion to the GAL4 activation domain. They demonstrated that the strength of the SNF1-SNF4



Supplement 46


interaction was sufficient to allow activation through a GAL4 DNA-binding site. From this, they suggested the feasibility of selecting interacting proteins by performing screens of cDNA libraries made so that library-encoded proteins carried activating domains. Several groups have developed cDNA library strategies along these lines, with some systems using LexA and others using GAL4 as the DNA-binding domain (Table 20.1.4). LexA and GAL4 each have different properties that should be considered when selecting a system. LexA is derived from a heterologous organism, has no known effect on the growth of yeast, possesses no residual transcriptional activity, can be used in GAL4+ yeast, and can be used with a Gal-inducible promoter. Because GAL4 is an important yeast transcriptional activator, it has the disadvantage that experiments must be performed in gal4− yeast strains to avoid background due to activation of the reporter system by endogenous GAL4. Such gal4− strains are frequently less healthy and more difficult to transform than wild-type strains, and either libraries must be constitutively expressed or alternate inducible systems must be used. By contrast, the GAL4 DNA-binding domain may be more efficiently localized to the nucleus and may be preferred for some proteins (for a review of GAL4-based systems, see Bartel et al., 1993). Whichever system is used, it is important to remember that the bait protein constitutes a novel fusion protein whose properties may not exactly parallel those of the original unfused protein of interest. Although systems using the two-hybrid paradigm have been developed in mammalian cells (see Table 20.1.4), these have not been used effectively in library screens. It seems likely that the organism of choice for two-hybrid identification of novel partner proteins will remain yeast. cDNAs that pass specificity tests are referred to as positives, or “true positives.” In interactor hunts conducted to date, anywhere from zero to practically all isolated plasmids passed the final specificity test. If no positives are obtained, the tissue source for the library originally used may not be appropriate, and a different library may produce better results. However, there are some proteins for which no positives are found. Various explanations for this are provided below. Conversely, some library-encoded proteins are known to be isolated repeatedly using a series of unrelated baits, and these proteins demonstrate at least some specificity. One of these, heat shock protein 70, might be explained by positing that it

assists the folding of some LexA-fused bait proteins, or alternatively, that these bait proteins are not normally folded. This example illustrates the point that the physiological relevance of even quite specific interactions may sometimes be obscure. Because the screen involves plating multiple cells to Gal/CM −Ura, −His, −Trp, −Leu dropout medium for each primary transformant obtained, multiple reisolates of true positive cDNAs are frequently obtained. If a large number of specific positives are obtained, it is generally a good idea to attempt to sort them into classes—for example, digesting minipreps of positives with EcoRI, XhoI, and HaeIII will generate a fingerprint of sufficient resolution to determine whether multiple reisolates of a small number of clones or single isolates of many different clones have been obtained. The former situation is a good indication that the system is working well. An important issue that arises in an interactor hunt is the question of how biologically relevant interacting proteins that are isolated are likely to be. This leads directly to the question of what Kd of association two molecules must have to be detected by an interactor hunt. In fact, this is not at all a simple issue. For the system described here, most fusion proteins appear to be expressed at levels ranging from 50 nM to 1 µM (Golemis and Brent, 1992). Given the strength of the GAL promoter, it is likely that many library-encoded proteins are expressed at similarly high levels, ≥1 µM in the nucleus (Golemis and Brent, 1992). At this concentration, which is in considerable excess over the nuclear concentration of operatorbound bait protein, a cDNA library–encoded protein should half-maximally occupy the DNA-bound bait protein if it possesses a Kd of 10−6 M, making it theoretically possible that very-low-affinity interactions could be detected. Such interactions have been observed in some cases. In contrast, some interactions that have been previously established using other methods and are predicted by known Kd to be easily detected by these means, either are not detected or are detected only weakly (Finley and Brent, 1994; Estojak et al., 1995). Because of the conservation of many proteins between lower and higher eukaryotes, one explanation for this observation is that either one or both of the partners being tested is being sequestered from the desired interaction by fortuitous association with an endogenous yeast protein. A reasonably complete investigation of the degree of correlation between in vitro determina-



tions of interaction affinity and apparent strength of interaction in the interaction trap is included in Estojak et al. (1995). The result of this investigation suggests it is important to measure the affinity of detected interactions under different conditions, using a second assay system, rather than to draw conclusions about affinity based on detection in the interaction trap. A number of different plasmids can be used for conducting an interactor hunt. Their properties are summarized in Tables 20.1.1 and 20.1.2. Because of the generous and open scientific exchange between investigators using the system, the number of available plasmids and other components has greatly expanded since the appearance of the initial two-hybrid reagents, facilitating the study of proteins inaccessible by the original system. The original parent plasmid for generating LexA fusions, pEG202 is a derivative of 202 + PL (Ruden et al., 1991; see Fig. 20.1.3) that contains an expanded polylinker region. The available cloning sites in pEG202 include EcoRI, BamHI, SalI, NcoI, NotI, and XhoI, with the reading frame as described in the legend to Figure 20.1.3. Since the original presentation of this system, a number of groups have developed variants of this plasmid that address specialized research needs. Those currently available, as well as purposes for which they are suited, are listed in Table 20.1.1. pGilda, created by David A. Shaywitz, places the LexAfusion cassette under the control of the inducible GAL1 promoter, allowing expression of the bait protein for limited times during library screening, reducing the exposure of yeast to toxic baits. pJK202, created by Joanne Kamens, adds nuclear localization sequences to pEG202, facilitating assay of the function of proteins lacking internal nuclear localization sequences. pNLexA, created by Ian York, places LexA carboxy-terminal in the fusion domain, allowing assay of interactions that require an unblocked amino-terminus on the bait protein. pEE202I, created by Mike Watson and Rich Buckholz, allows chromosomal integration of a pEG202-like bait, thus reducing expression levels so they are more physiological for bait proteins normally present at low levels intracellularly. All of these have been extensively tested by numerous researchers. pGilda, pJK202, and pEE202I work with complete reliability. pNLexA works effectively with ∼50% of the fusion domains tried, but synthesizes only very low levels of protein (relative to expression of the same fusion domain as a

pEG202 fusion) with the remaining 50%. Attachment of fusion domains amino-terminal either to LexA or GAL4 has been generally problematic in the hands of many investigators; it may be that appending additional protein sequences to the amino termini of these proteins is destabilizing, although the problem has not been rigorously investigated. A series of lacZ reporters of differing sensitivity to transcriptional activation can be used to detect interactions of varying affinity (see Table 20.1.2). These plasmids are LexA operator–containing derivatives of the plasmid LR1∆1 (West et al., 1984). In LR1∆1, a minimal GAL1 promoter lacking the GAL1 upstream activating sequences (GALUAS) is located upstream of the bacterial lacZ gene. In pSH18-34, eight LexA operators have been cloned into an XhoI site located 167 bp upstream of the lacZ gene (S. Hanes, unpub. observ.). pJK103 and pRB1840 contain two and one operator, respectively. pJK101 is similar to pSH18-34, except that it contains the GAL1 upstream activating sequences (GALUAS) upstream of two LexA operator sites. A derivative of del20B (West et al., 1984), it is used in the repression assay (Brent and Ptashne, 1984; see Fig. 20.1.5) to assess LexA fusion binding to operator. pSH17-4 is a HIS3 2µm plasmid that encodes LexA fused to the activation domain of the yeast activator GAL4. EGY48 cells bearing this plasmid will produce colonies in overnight growth on medium lacking Leu, and yeast that additionally contain pSH18-34 will turn deep blue on plates containing Xgal. This plasmid serves as a positive control for the activation of transcription. pRFHM1 is a HIS3 2µm plasmid that encodes LexA fused to the N-terminus of the Drosophila protein bicoid. The plasmid has no ability to activate transcription, so EGY48 cells that contain pRFHM1 and pSH18-34 do not grow on −Leu medium and remain white on plates containing Xgal. pRFHM1 is a good control for specificity testing, because it has been demonstrated to be sticky—that is, to associate with a number of library-encoded proteins that are clearly nonphysiological interactors (R. Finley, Wayne State University, Detroit, Mich., unpub. observ.). This protocol uses interaction libraries (Table 20.1.3) made in pJG4-5 or its derivatives (see Fig. 20.1.6). pJG4-5 was developed to facilitate isolation and characterization of novel proteins in interactor hunts (Gyuris et al., 1993). The pJG4-5 cDNA library expression



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cassette is under control of the GAL1 promoter, so library proteins are expressed in the presence of galactose (Gal) but not glucose (Glu). This conditional expression has a number of advantages, the most important of which is that many false-positives obtained in screens can be easily eliminated because they do not demonstrate a Gal-dependent phenotype. The expression cassette consists of an ATG to start translation, a nuclear localization signal to extend the interaction trap’s range to include proteins that are normally predominantly localized in the cytoplasm, an activation domain (acid blob; Ma and Ptashne, 1987), the hemagglutinin epitope tag to permit rapid assessment of the size of encoded proteins, EcoRI-XhoI sites designed to receive directionally synthesized cDNAs, and the alcohol dehydrogenase (ADH) termination sequences to enhance the production of high levels of library protein. The plasmid also contains the TRP1 auxotrophy marker and 2µm origin for propagation in yeast. A derivative plasmid, pJG4-5I, was created by Mike Watson and Richard Buckholz to facilitate chromosomal integration of the activation domain fusion expression plasmid. A series of recently developed derivatives of pEG202, pJG4-5, and lacZ reporter plasmids (MW101 to MW112) alter the antibiotic resistance markers on these plasmids from ampicillin (Apr) to either kanamycin (Kmr) or chloramphenicol (Cmr; Watson et al., 1996). Judiciously mixing and matching these plasmids in conjunction with Apr libraries would considerably reduce work subsequent to library screening, because the KC8 transformation, which involves trpC complementation in bacteria, could be omitted. EGY48 and EGY191 (see Table 20.1.2) are both derivatives of the strain U457 (a gift of Rodney Rothstein, Columbia University, New York, N.Y.) in which the endogenous LEU2 gene has been replaced by homologous recombination with LEU2 reporters carrying varying numbers of LexA operators, using a procedure detailed in Estojak et al. (1995). Interaction Trap–compatible reagents have recently become commercially available; Clontech and Invitrogen were the first to market such reagents and have recently been joined by OriGene. All suppliers use systems with the most sensitive reporters (EGY48 and pSH1834), and provide their own positive and negative controls for testing activation or interaction between defined proteins. For expression of bait and library proteins, the Clontech Matchmaker LexA two-hybrid system and the

OriGene Duplex-A system use some of the basic set of plasmids described here (see Table 20.1.1 for availability). Forward sequencing primers for bait and library plasmids are included in the Clontech kit, and Insert Screening Amplimer Sets for both plasmids can be acquired separately. Additional related products from Clontech include KC8 competent cells, anti-LexA monoclonal antibodies, a yeast transformation system, a yeast plasmid isolation kit, and an EGY48 partner strain for yeast mating to facilitate the analysis of interaction specificity. OriGene has a generally similar product line to Clontech. In contrast, Invitrogen has substantially modified the Interaction Trap core reagents to develop its own bait and library plasmids. pHybLex/Zeo, a novel bait plasmid, is ∼50% smaller than the original pEG202 (making it easier to clone into), and it has an enriched polylinker. Significantly, it replaces both the Apr and HIS3 genes with a novel gene that confers resistance to the antibiotic Zeocin (supplied with the kit), which provides selection in both bacteria and yeast. This elimination of auxotrophic selection for the bait plasmid renders the LexA-fusion construct usable with libraries and strains from all existing two-hybrid systems and additionally facilitates the direct selection of library plasmid in strains other than KC8. Some changes, which are designed to make the vector easier to use, have also been introduced in the library vector pYESTrp (e.g., it uses a V5 epitope tag for protein detection). The Invitrogen kit, termed Hybrid Hunter, includes the bait/library/reporter plasmids and EGY48 yeast strain as noted, and additionally includes primer sets for bait and library plasmids and the L40 yeast strain, should an investigator wish to use a HIS3 auxotrophy selection. Additional related products from Invitrogen include antibodies for detection of bait and prey fusion proteins (antiLexA and anti-V5), pJG4-5 library vector primers, and a Transformation Kit. A significant advantage of the entry of commercial entities into the Interaction Trap field is the rapid increase in the number of compatible cDNA libraries. A list of currently available premade libraries available from these companies is presented in Table 20.1.3, and custommade libraries are also available upon request. Because new libraries and other related reagents are being constantly added to the line of two-hybrid related products, it is advisable to contact the companies or visit their Web sites (www.clontech.com, www.invitrogen.com, and www.origene.com) for the latest information.



Finally, over the last several years, a number of groups have adapted basic two-hybrid strategies to more specialized applications, and they have devised strategies to broaden their basic functionality. Interaction Mating (Finley and Brent, 1994) has been used to establish extended networks of targeted protein-protein interaction. In this approach, a panel of LexAfused proteins are transformed into a MATa haploid selective strain (such as RFY206), a panel of activation-domain fused proteins are transformed into a suitable MATα haploid (such as EG448), and the two panels are crossgridded against each other for mating. Selected diploids are then screened by replica plating to selective medium. This approach complements library screening in large-scale applications, such as proposed definition of interaction maps for entire genomes (Bartel et al., 1996). Interaction mating has also provided the basis for an alternative two-hybrid hunt protocol (see Alternate Protocol 2), useful in cases when a single library will be screened with different baits. In this approach (Bendixen et al., 1994; Finley and Brent, 1994: Kolonin and Finley, 1998), a library is introduced into a single strain, like EGY48, and aliquots are stored frozen. To conduct a hunt, an aliquot is thawed and mated with a strain expressing a bait. This allows one to avoid repeated high-efficiency transformations, since a single library transformation can provide enough pretransformed yeast to conduct dozens of interactor hunts. Moreover, some yeast strains pretransformed with libraries are becoming commercially available, which may eliminate altogether the need to conduct a high-efficiency library transformation for some researchers. Two-hybrid approaches have been shown to be effective in identifying small peptides with biological activities on selected baits (Yang et al., 1995; Colas et al., 1996), which may prove to be useful as a guide to targeted drug design. Rapid screening protocols have been devised using custom-synthesized libraries expressing sheared plasmid DNA to facilitate rapid mapping of interaction interfaces (Stagljar et al., 1996). Osborne and coworkers have demonstrated the effectiveness of a tribrid (or tri-hybrid) approach, in which an additional plasmid expresses a tyrosine kinase to specifically modify a bait protein, allowing detection of SH2domain-containing partner proteins that recognize specific phosphotyrosine residues (Osborne et al., 1995). A variety of more elaborate tribrid approaches, in which a DNA-binding domain fused protein is used to present an

intermediate nonprotein compound for interaction with a library, have been developed and proven effective. These approaches have allowed the identification of proteins binding specific drug ligands (Chiu et al., 1994; Licitra and Liu, 1996), as well as the identification of proteins binding to RNA sequences (SenGupta et al., 1996; Wang et al., 1996). It is expected that the range of utility of these systems will continue to expand.

Critical Parameters and Troubleshooting To maximize chances of a successful interactor hunt, a number of parameters should be taken into account. Before attempting a screen, bait proteins should be carefully tested to ensure that they have little or no intrinsic ability to activate transcription. Bait proteins must be expressed at reasonably high levels and must be able to enter the yeast nucleus and bind DNA (as confirmed by the repression assay). Optimally, integrity and levels of bait proteins should be confirmed by immunoblot analysis, using an antibody to either LexA or the fused domain. In particular, at this time, bait proteins that have extensive transmembrane domains or are normally excluded from the nucleus are not likely to be productively used in a library screen. Proteins that are moderate to strong activators will need to be truncated to remove activating domains before they can be used. If a protein neither activates nor represses, the most likely reason is that it is not being made. This can be determined by immunoblot analysis of a crude lysate protein extract of EGY48 (UNIT 10.8; Samson et al., 1989) containing the plasmid, using anti-LexA antibodies as primary antiserum. If the full protein is not made, it may be possible to express truncated derivatives of the protein. If the protein is made, but still does not repress, it may not enter the yeast nucleus effectively, although this appears to be a relatively rare problem. In this case, introducing the coding sequence for the fused moieity into a LexA fusion vector containing a nuclear localization motif (e.g., pJK202; J. Kamens, BASF, Worcester, Mass., unpub. observ.) may solve the problem. The test for the leucine (Leu) requirement is extremely important to determine whether the bait protein is likely to yield an unworkably high background. The LEU2 reporter in EGY48 is more sensitive than the pSH18-34 reporter for some baits (Estojak et al., 1995). Therefore, it is possible that a bait protein demonstrating little or no signal in a β-galac-



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tosidase assay may nevertheless permit some growth on −Leu medium. If this occurs, there are several options. First, a less sensitive strain can be used, as described in the text. Second, background can sometimes be reduced further by making the EGY strain diploid (e.g., D. Krainc, Harvard Medical School, Boston, Mass.; R. Finley and R. Brent, unpub. observ.) or by performing the hunt by interaction mating as described in Alternate Protocol 2. A third option is to attempt to truncate the bait protein to remove activating function. In general, it is useful to extrapolate from the number of cells that grow on −Leu medium to the number that would be obtained in an actual library screen, and determine if this is a background level that can be tolerated. For example, if two colonies arise from 100,000 plated cells on −Leu medium, 200 to 400 would be expected in an actual screen of 106 cDNAs. Although this is a high initial number of positives, the vast majority should be eliminated immediately through easily performed controls. This is a judgment call. Finally, very rarely it happens that a bait that appears to be well behaved and negative for transcriptional activation through all characterization steps will suddenly develop a very high background of transcriptional activation following library transformation. The reason for this is currently obscure, and no means of addressing this problem has as yet been found: such baits are hence inappropriate for use in screens. The protocols described in this unit use initial screening with the most sensitive reporters followed by substitution with less sensitive reporters if activation is detected. An obvious question is, why not start out working with extremely stringent reporters and know immediately whether the system is workable? In fact, some researchers routinely use a combination of pJK103 or pRB1840 with EGY191, and obtain proteins that to date appear to be biologically relevant partners from library screens. However, extensive comparison studies using interactors of defined in vitro affinity with different combinations of LacZ and LEU2 reporters (Estojak et al., 1995) have indicated that although the most sensitive reporters (pSH1834) may in some cases be prone to background problems, the most stringent reporters (EGY191, pRB1840) may miss some interactions that certainly are biologically relevant and occur inside cells. In the end, the choice of reporters devolves to the preference of individual investigators: the bias of the authors is to cast a broad net in the early stages of a screen,

and hence to use more sensitive reporters when practicable. It is important to move expeditiously through characterization steps and to handle yeast transformed with bait plasmids with care. In cases where yeasts have been maintained on plates for extended periods (e.g., 4 days at room temperature or >2 to 3 weeks at 4°C), unexpected problems may crop up in subsequent library screens. The transformation protocol is a version of the lithium acetate transformation protocol described by Schiestl and Gietz (1989) and Gietz et al. (1992; see UNIT 13.7) that maximizes transformation efficiency in Saccharomyces cerevisiae and produces up to 105 colonies/µg plasmid DNA. In contrast to Escherichia coli, the maximum efficiency of transformation for S. cerevisiae is ∼104 to 105/µg input DNA. It is extremely important to optimize transformation conditions before attempting an interactor hunt. Perform small-scale pilot transformations to ensure this efficiency is attained and to avoid having to use prohibitive quantities of library DNA. In addition, as for any effort of this type, it is a good idea to obtain or construct a library from a tissue source in which the bait protein is known to be biologically relevant. In practice, the majority of proteins isolated by interaction with a LexA fusion turn out to be specific for the fused domain; a smaller number are nonspecifically sticky, and to date there appears to have been only one isolation from a eukaryotic library of a protein specific for LexA. However, it is generally informative to retest positive clones on more than one LexA bait protein; ideally, library-derived clones should be tested against the LexA fusion used for their isolation, several LexA fusions to proteins that are clearly unrelated to the original fusion, and if possible, several LexA fusions that there is reason to believe are related to the initial protein (e.g., if the initial probe was LexA-Fos, a good related set would include LexA-Jun and LexA-GCN4). Colony selection for master plate production is one of the more variable parts of the procedure. For strong interactors, colonies will grow up in 2 days. However, if plates are left at 30°C, new colonies will continue to appear every day. Those that appear rapidly are most likely to reflect interactors that are biologically relevant to the bait protein. Those that appear later may or may not be relevant. However, many parameters can delay the time of colony formation of cells that contain valid interactions, including the strength of the interaction



and the level of expression of the library-encoded protein.

Anticipated Results Depending on the protein used as bait, anywhere from zero to hundreds of specific interactors will be obtained from 106 primary transformants.

Time Considerations If all goes well, once the required constructions have been made it will take ∼1 week to perform yeast transformations, obtain colonies, and determine whether bait proteins are appropriate. It will take a second week to perform library transformations, replate to selective medium, and obtain putative positives. A third week will be required to rescue the plasmid from the yeast, passage it through E. coli, transform fresh yeast, and confirm specificity.

Literature Cited Bartel, P.L., Chien, C.-T., Sternglanz, R., and Fields, S. 1993. Using the two-hybrid system to detect protein-protein interactions. In Cellular Interactions in Development: A Practical Approach (D.A. Hartley, ed.) pp. 153-179. Oxford University Press, Oxford. Bartel, P.L., Roecklein, J.A., SenGupta, D., and Fields, S. 1996. A protein linkage map of Escherichia coli bacteriophage T7. Nature Genet. 12:72-77. Bendixen, C., Gangloff, S., and Rothstein, R. 1994. A yeast mating-selection scheme for detection of protein-protein interactions. Nucl. Acids Res. 22:1778-1779. Breeden, L. and Nasmyth, K. 1985. Regulation of the yeast HO gene. Cold Spring Harbor Symp. Quant. Biol. 50:643-650. Brent, R. and Ptashne, M. 1984. A bacterial repressor protein or a yeast transcriptional terminator can block upstream activation of a yeast gene. Nature 312:612-615. Brent, R. and Ptashne, M. 1985. A eukaryotic transcriptional activator bearing the DNA specificity of a prokaryotic repressor. Cell 43:729-736. Chien, C.-T., Bartel, P.L., Sternglanz, R., and Fields, S. 1991. The two-hybrid system: A method to identify and clone genes for proteins that interact with a protein of interest. Proc. Natl. Acad. Sci. U.S.A. 88:9578-9582. Chiu, M.I., Katz, H., and Berlin, V. 1994. RAPT1, a mammalian homolog of yeast Tor, interacts with the FKBP12/rapamycin complex. Proc. Nat. Acad. Sci. U.S.A. 91:12574-12578. Colas, P., Cohen, B., Jessen, T., Grishina, I., McCoy, J., and Brent, R. 1996. Genetic selection of peptide aptamers that recognize and inhibit cyclindependent kinase 2. Nature 380:548-550. Durfee, T., Becherer, K., Chen, P.L., Yeh, S.H., Yang, Y., Kilburn, A.E., Lee, W.H., and Elledge, S.J.

1993. The retinoblastoma protein associates with the protein phosphatase type 1 catalytic subunit. Genes & Dev. 7:555-569. Estojak, J., Brent, R., and Golemis, E.A. 1995. Correlation of two-hybrid affinity data with in vitro measurements. Mol. Cell. Biol. 15:58205829. Fearon, E.R., Finkel, T., Gillison, M.L., Kennedy, S.P., Casella, J.F., Tomaselli, G.F., Morrow, J.S., and Dang, C.V. 1992. Karyoplasmic interaction selection strategy: A general strategy to detect protein-protein interactions in mammalian cells. Proc. Nat. Acad. Sci. U.S.A. 89:7958-7962. Fields, S. and Song, O. 1989. A novel genetic system to detect protein-protein interaction. Nature 340:245-246. Finley, R.L., Jr., and Brent, R. 1994. Interaction mating reveals binary and ternary connections between Drosophila cell cycle regulators. Proc. Natl. Acad. Sci. U.S.A. 91:12980-12984. Gietz, D., St. Jean, A., Woods, R.A., and Schiestl, R.H. 1992. Improved method for high-efficiency transformation of intact yeast cells. Nucl. Acids Res. 20:1425. Golemis, E.A. and Brent, R. 1992. Fused protein domains inhibit DNA binding by LexA. Mol. Cell Biol. 12:3006-3014. Grunstein, M., and Hogness, D.S. 1975. Colony hybridization: A method for the isolation of cloned DNAs that contain a specific gene. Proc. Natl. Acad. Sci. U.S.A. 72:3961-3965. Gyuris, J., Golemis, E.A., Chertkov, H., and Brent, R. 1993. Cdi1, a human G1- and S-phase protein phosphatase that associates with Cdk2. Cell 75:791-803. Kaiser, C., Michaelis, S., and Mitchell, A. 1994. Methods in Yeast Genetics, a Cold Spring Harbor Laboratory Course Manual, pp.135-136. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Kolonin, M.G. and Finley, R.L., Jr. 1998. Targeting cyclin-dependent kinases in Drosophilia with peptide aptamers. Proc. Natl. Acad. Sci. U.S.A. In press. Licitra, E.J. and Liu, J.O. 1996. A three-hybrid system for detecting small ligand-protein receptor interactions. Proc. Nat. Acad. Sci. U.S.A. 93:12817-12821. Ma, J. and Ptashne, M. 1987. A new class of yeast transcriptional activators. Cell 51:113-119. Ma, J. and Ptashne, M. 1988. Converting an eukaryotic transcriptional inhibitor into an activator. Cell 55:443-446. Osborne, M., Dalton, S., and Kochan, J.P. 1995. The yeast tribrid system: Genetic detection of transphosphorylated ITAM-SH2 interactions. Bio/Technology 13:1474-1478. Ruden, D.M., Ma, J., Li, Y., Wood, K., and Ptashne, M. 1991. Generating yeast transcriptional activators containing no yeast protein sequences. Nature 350:426-430. Samson, M.-L., Jackson-Grusby, L., and Brent, R. 1989. Gene activation and DNA binding by



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Drosophila Ubx and abd-A proteins. Cell 57:1045-1052. Schiestl, R.H. and Gietz, R.D. 1989. High-efficiency transformation of intact yeast cells using single-stranded nucleic acids as a carrier. Curr. Genet. 16:339-346. SenGupta, D.J., Zhang, B., Kraemer, B., Pochart, P., Fields, S., and Wickens, M. 1996. A three-hybrid system to detect RNA-protein interactions in vivo. Proc. Nat. Acad. Sci. U.S.A. 93:8496-8501. Sikorski, R.S. and Hieter, P. 1989. A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122:19-27. Stagljar, I., Bourquin, J.-P., and Schaffner, W. 1996. Use of the two-hybrid system and random sonicated DNA to identify the interaction domain of a protein. BioTechniques 21:430-432. Vasavada, H.A., Ganguly, S., Germino, F.J., Wang, Z.X., and Weissman, S.M. 1991. A contingent replication assay for the detection of protein-protein interactions in animal cells. Proc. Nat. Acad. Sci. U.S.A. 88:10686-10690. Vojtek, A.B., Hollenberg, S.M., and Cooper, J.A. 1993. Mammalian Ras interacts directly with the serine/threonine kinase Raf. Cell 74:205-214. Wang, Z.F., Whitfield, M.L., Ingledue, T.C.3, Dominski, A., and Marzluff, W.F. 1996. The protein that binds the 3′ end of histone mRNA: A novel RNA-binding protein required for histone pre-mRNA processing. Genes & Dev. 10:3028-3040. Watson, M.A., Buckholz, R., and Weiner, M.P. 1996. Vectors encoding alternative antibiotic resistance for use in the yeast two-hybrid system. BioTechniques 21:255-259. West, R.W.J., Yocum, R.R., and Ptashne, M. 1984. Saccharomyces cerevisiae GAL1-GAL10 divergent promoter region: Location and function of the upstream activator sequence UASG. Mol. Cell Biol. 4:2467-2478. Yang, M., Wu, Z., and Fields, S. 1995. Protein-peptide interactions analyzed with the yeast two-hybrid system. Nucl. Acids Res. 23:1152-1156.

Key Reference

Internet Resources http://www.clontech.com http://cmmg.biosci.wayne.edu/rfinley/lab.html Source of two-hybrid information, protocols, and links. http://www.invitrogen.com http://www.origene.com Commercial sources for basic plasmids, strains, and libraries for interaction trap experiments. [email protected] [email protected] Sources of interaction trap plasmids for specialized interactions. http://www.fccc.edu:80/research/labs/golemis/ InteractionTrapInWork/html Database for false positive proteins detected in interaction trap experiments; analysis of two-hybrid usage. http://xanadu.mgh.harvard.edu/brentlabhome page4.html Database of interaction trap protocols and related issues.

Contributed by Erica A. Golemis and Ilya Serebriiskii Fox Chase Cancer Center Philadelphia, Pennsylvania Russell L. Finley, Jr. and Mikhail G. Kolonin (hunt by interaction mating) Wayne State University School of Medicine Detroit, Michigan Jeno Gyuris Mitotix, Inc. Cambridge, Massachusetts Roger Brent The Molecular Sciences Institute Berkeley, California

Gyuris et al., 1993. See above. Initial description of interaction trap system.




Affinity Purification of Proteins Binding to GST Fusion Proteins

UNIT 20.2

This unit describes the use of proteins fused to glutathione-S-transferase (GST fusion proteins) to affinity purify other proteins, a technique also known as GST pulldown purification. The GST fusion protein is first purified on glutathione-agarose beads as described in UNIT 16.7. The bead-bound fusion protein is then used as “bait” to test for binding to a known or suspected “test” protein which may be either purified or labeled by in vitro translation (UNIT 10.17). Beads with bound protein are washed, and the amount of test protein retained is determined by elution with glutathione or salt or simply by applying the beads and bound proteins to an SDS-polyacrylamide gel (UNIT 10.2; Fig. 20.2.1). The bead-bound fusion protein can also be incubated with a complex mixture of proteins. If performed on a large scale, this technique can be used to purify known or unknown proteins that interact directly or indirectly with the bait protein (Fig. 20.2.2). When a method (e.g., antibody binding) exists for detecting a particular protein, it is possible to determine if a test protein is retained from a complex mixture of proteins such as a crude cellular lysate. This GST pulldown technique can complement other methods for assessing protein-protein interactions—in vitro assays such as electrophoretic mobility shift assays (EMSA; see UNIT 12.2) or coimmunoprecipitation (see UNIT 10.16), or in vivo assays such as the two-hybrid interaction trap (UNIT 20.1) or ubiquitin-based split-protein sensor (Johnsson and Varshavsky, 1994). The GST pulldown affinity technique is simple to perform and quite powerful. However, it is prone to artifacts due to mass-action effects, and some care should be used in interpreting positive results. The Basic Protocol describes a strategy that can be used to isolate proteins that display affinity for (bind to) proteins fused to GST. GST fusion protein bound to agarose affinity beads is used to assay the binding of a specific test protein that has been labeled with [35S]methionine by in vitro translation (UNIT 10.17). However, this method can be adapted for use with other types of fusion proteins—for example, His6 (UNIT 10.11B), biotin tags, or maltose-binding protein fusions (MBP; UNIT 16.6), and these may offer particular advantages. A Support Protocol describes preparation of an Escherichia coli extract that is added to the reaction mixture with purified test protein to reduce nonspecific binding. STRATEGIC PLANNING There are three choices to be made in planning a GST fusion protein affinity purification: the selection and design of the affinity fusion to the bait protein, the assay used to detect the test protein(s), and the controls used to check for specificity. A GST fusion protein serves well for most applications; the features of this fusion and the methods involved are described in UNIT 16.7. If either glutathione-S-transferase or the agarose matrix retains an interfering contaminating protein, changing either the affinity moiety (e.g., nickel, maltose, or streptavidin) or the matrix (e.g., Streptavidin Dynabeads, Dynal) may avoid the problem. If the purpose of the experiment is to determine if the bait and test proteins multimerize, it may be better to use monomeric MBP or His6 to construct the fusion protein because GST is known to form dimers. Each system has its own advantages and disadvantages. One advantage of GST or MBP fusions is that they can increase the solubility of certain insoluble or semi-soluble proteins. A second advantage is that peptides are more readily fused to a protein (GST, MBP) than to His6. A third advantage is the low background compared to His6/Ni agarose. One disadvantage, however, is that the capacity of glutathione agarose beads for GST fusion proteins falls steeply with increasing size of the fusion.


Contributed by Jonathan C. Swaffield and Stephen Albert Johnston

20.2.1


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binding and washing

SDS-PAGE

radiolabeled test protein

GST-bait fusion

autoradiogram

GST-bait fusion protein

GST-bait bound to glutathione-agarose

Coomassie-stained gel

radiolabeled test protein

complex competitor proteins

Figure 20.2.1 Schematic representation of GST pulldown purification of proteins.




The two methods commonly used to detect binding of the test protein are immunoblotting with a protein-specific antibody (UNIT 10.8) or measuring bound radioactivity of an in vitro–translated test protein (UNIT 10.17) that has been labeled with [35S]methionine. Radioactive detection is preferable because it is possible to accurately quantitate the protein by phosphorimaging or scanning densitometry, and confers greater sensitivity of detection. Furthermore, after labeled proteins are separated on SDS-PAGE analytical gels, the gels can be fixed, stained for protein, and scanned as a control for loss of beads and bait protein during washing steps. With immunoblotting, accurate quantitation of the signal on a sheet of film (for chemiluminescent detection) or membrane (for the insoluble dye method) is difficult. In addition, the relatively large amount of GST fusion protein bait may exhibit nonspecific cross-reactivity with the antibodies employed, making detection difficult if the GST-bait and test proteins are similar in size. This problem can be circumvented by constructing a GST fusion protein having a protease site at the junction between GST and the bait protein sequence (see UNIT 16.4B). After binding and washing, the bait protein and any bound test protein can be released from the beads by treatment with the appropriate protease. Should nonspecific binding of the test protein to the agarose beads be a problem, protease or glutathione can be employed to elute only that portion of the test protein that is specifically bound to the bait protein or to glutathione, as opposed to the matrix. A third method to detect binding of the target protein to the bait protein is an activity assay of the washed beads after the binding reaction. When employing an activity assay, however, it may be advantageous to elute the bait/target protein complex first. This will allow the protein to be assayed in a soluble form in the buffer of choice. Glutathione-agarose beads with bound GST alone are often used as a negative control for nonspecific retention. Two better controls, however, are competition with a large excess of cold target protein or use of a variant of the bait protein which is known not to bind the test protein in vivo (Fig. 20.2.3). When using His6 or biotin tags on the bait protein, a negative control with an unrelated protein may be used, but, again, it is best to use a nonbinding variant of the bait protein. It should be noted, however, that a number of naturally biotinylated proteins occur in cells—one in E. coli (Agarana et al., 1986), four or five in Saccharomyces cerevisiae, depending on growth conditions (Lim et al., 1987), and four in mammalian cells (Chandler and Ballard, 1988) and in plants (Nikolau et al., 1985). These may cause high background by binding to streptavidin beads used to immobilize biotinylated bait protein and give nonspecific bands. GST FUSION PROTEIN–AFFINITY PURIFICATION Bait protein fused to GST and bound to glutathione-agarose beads is incubated with radiolabeled test protein in the presence of an Escherichia coli protein extract. The beads and associated protein complexes are removed from the incubation mixture, washed, eluted in loading buffer, and analyzed on an SDS-PAGE gel. The resulting gel is quantitated using scanning densitometry or phosphorimaging. Materials E. coli extract in bead binding buffer (see Support Protocol) Test protein labeled with [35S]methionine (UNIT 10.17) 100 to 250 µg/ml GST and GST fusion protein bound to agarose beads (UNIT 16.7), freshly prepared 1× SDS sample buffer (UNIT 10.2) Gel fixative: 10% (v/v) glacial acetic acid/45% (v/v) methanol Centrifuge and Beckman TLA-100 rotor or equivalent

BASIC PROTOCOL



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Microcentrifuge, 4°C X-ray film or Storage Phosphor screen (Molecular Dynamics) Scanning densitometer or phosphorimager Additional reagents and equipment for SDS-PAGE (UNIT 10.2), Coomassie blue staining (UNIT 10.6; optional), and autoradiography (APPENDIX 3) Prepare the extract 1. Thaw soluble E. coli protein extract on ice. 2. For each reaction, centrifuge 500 µl thawed extract 30 min at 175,000 × g (70,000 rpm in a Beckman TLA-100 rotor), 4°C Centrifugation removes any insoluble protein aggregates that form during storage or thawing.

3. Remove the supernatant (≥400 µl). Transfer 200 µl supernatant into each of two tubes, and store on ice. Prepare test protein 4. Add 1 to 5 µl [35S]methionine-labeled test protein to one tube containing 200 µl of the supernatant from step 3. Incubate 15 min on ice. The specific activity of the [35S]methionine-labeled test protein should be high enough to permit detection within a reasonable amount of time.

5. Microcentrifuge 15 min at maximum speed, 4°C. This step removes insoluble protein aggregates from the in vitro translation mix.

6. Transfer 200 µl of the supernatant (containing labeled test protein + extract) to a clean microcentrifuge tube. Prepare the bait protein 7. Add 20 µl GST or GST fusion protein (bait) bound to agarose beads to the other tube containing 200 µl of supernatant from the E. coli extract (step 3) and mix thoroughly. From 2 to 5 ìg GST bait protein is required for each reaction. It is best to include ∼20 ìl beads in each reaction so the bead pellet will be visible during the washes. However, if 20 ìl of beads would add more than 2 to 5 ìg protein, add unbound glutathione beads to provide the necessary volume. It may be necessary to minimize the amount of bait protein to decrease nonspecific interactions. Glutathione beads bound with GST alone may be stored a number of months in PBS at 4°C, but fusion proteins are likely to be less stable and preparations should be checked for degradation before use. Excessive degradation will alter the concentration of the bait protein (see Critical Parameters), and released protein fragments may bind test protein and may not be retained on the glutathione beads during the washes, thus interfering with subsequent analysis. Although such degradation usually results in complete loss of the bait component of the fusion protein, partial degradation may leave N-terminal fragments that may or may not bind test protein. SDS-PAGE can be used to check for degradation. Proteolysis may be limited by including protease inhibitors (UNIT 10.11B) in all buffers. Unless a specific GST fusion protein is known to be stable, the binding assay should use freshly prepared beads bound with purified full-length fusion protein. Stored beads should be washed to remove any degradation products.


Perform binding assay 8. Add 200 µl well-mixed bead + extract suspension (step 7) to the tube containing test protein + extract (step 6). Incubate 1 to 2 hr at 4°C with gentle mixing.



9. Microcentrifuge the reaction 1 min at maximum speed, 4°C, to pellet the beads. 10. Remove supernatant and wash the beads three times with 1 ml bead binding buffer each time. Microcentrifuge 1 min at maximum speed, 4°C, between washes. 11. After the final wash, carefully remove all liquid. It is important to remove all liquid to ensure that equal volumes are loaded on the analytical SDS-PAGE gel. A twisted Kimwipe or a very thin gel-loading pipet tip is useful for removing the final few microliters.

12. Add 25 µl of 1× SDS sample buffer directly to each tube and boil 5 min. A reduction in background and an increase in the specificity of the reaction can be obtained by eluting only the GST-bait protein and any complexed target from the beads with either glutathione or protease. The eluted material is then mixed with SDS sample buffer and analyzed. Elution may be particularly advantageous when monitoring binding of the target protein by an activity assay. Bait/target complexes can be eluted into the buffer of choice and assayed away from potential interfering activities nonspecifically bound to the beads. One disadvantage of elution, however, is that elution is rarely 100% efficient, which makes quantitation of binding difficult.

Run and analyze an analytical gel 13. Run an analytical SDS-PAGE gel (see UNIT 10.2). The amount of sample required varies with different bait/target protein pairs and the manner of detection. Loading all the sample avoids the question of loading volumes because there is no uncertainty about the ratio of loaded to unloaded sample.

14. Optional: Stain the gel for protein with Coomassie blue (UNIT 10.6). The stained gel provides a visual confirmation that beads and the bait protein complexes were not lost during the washes and that all reactions contain equivalent amounts of bait protein. To verify that no material was lost during the washes, a parallel gel loaded with one-tenth the sample can be run and stained for protein to more accurately compare the GST fusion protein bands. Also, degradation of the fusion protein may lead to a decrease in the amount of bait protein, especially when using crude extracts (see Fig. 20.2.1) instead of purified labeled test protein. Such degradation must be taken into account in quantifying bound test protein.

15. Fix the gel in gel fixative 30 min at room temperature with gentle shaking. 16. Dry the gel and autoradiograph it or place it on a Storage Phosphor screen. 17. Quantitate the film using a scanning densitometer or phosphorimager. PREPARATION OF E. COLI EXTRACT It is necessary to perform the binding reactions in the presence of a complex competitor to reduce nonspecific background binding. This is best done by preparing a soluble protein extract from Escherichia coli in the buffer that will be used in the GST fusion protein binding reaction (see Basic Protocol).

SUPPORT PROTOCOL

Materials E. coli overnight culture grown in LB medium (UNIT 1.1) Bead binding buffer (see recipe) with and without Triton X-100 and glycerol, 4°C Tween 20 Glycerol Sonicator with microprobe Centrifuge and Sorvall SS-34 rotor or equivalent



Supplement 37

Additional reagents and equipment for growing E. coli (UNIT 1.2) and quantitating proteins in solution (UNIT 10.1) 1. Harvest an overnight culture of E. coli grown in LB medium. Wash the cells in cold bead binding buffer. If proteolysis of either the target or the GST-bait fusion protein proves to be a problem during the incubation period, a protease-deficient strain of E. coli may be used to prepare the complex competitor extract. However, it should be noted that such strains are often fragile and may not grow well when they contain the pLysS plasmid described in the annotation to step 3. Additional protease inhibitors also may be added to the bead binding buffer. If proteolysis continues to be a problem, alternative blocking/competing reagents— such as BLOTTO or UNI-BLOCK (Analytical Genetic Testing Center)—may be tried. The size of the culture is dictated by the amount of extract required for the binding assays. A 1-liter culture will yield at least 10 ml of a 10 mg/ml extract.

2. Pellet the cells. Resuspend the pellet in 10 ml per liter of culture of cold bead binding buffer without Triton X-100 and glycerol. Glycerol and Triton X-100 tend to cause foaming during sonication, thus reducing its effectiveness.

3. Sonicate five times, 1 min each, with a 2-min pause on ice between each cycle. If a sonicator is unavailable, use E. coli strain BL21 (DE3) containing the pLysS plasmid (Novagen), harvest as in steps 1 and 2 but include 1% (v/v) Triton X-100, and subject the resuspended pellet to a single freeze-thaw cycle, which is sufficient to lyse the cells of this strain. DNaseI treatment in the presence of 10 mM Mg2+ can be used to reduce the viscosity caused by released DNA.

4. Centrifuge the sonicate 30 min at 47,800 × g avg (20,000 rpm in a Sorvall SS-34 rotor), 4°C. 5. Remove the supernatant and store on ice. 6. Add Triton X-100 to 1% (v/v) and glycerol to 10% (v/v) and mix thoroughly. 7. Assay the protein concentration (UNIT 10.1). Dilute the extract with cold bead binding buffer to give a final concentration of 10 mg/ml protein and use. Undiluted extracts may also be stored in 5-ml (10 reactions) or 10-ml (20 reactions) aliquots at −80° or −20°C at least three months. After the aliquot is thawed, repeat step 4 before assaying the protein concentration and diluting to 10 mg/ml.


Bead binding buffer 50 mM potassium phosphate, pH 7.5 150 mM KCl 1 mM MgCl2 10% (v/v) glycerol 1% (v/v) Triton X-100 Protease inhibitors (UNIT 10.11B or Protease Cocktail, Boehringer Mannheim) Store buffer at room temperature and add protease inhibitors just before use Affinity Purification of Proteins Binding to GST Fusion Proteins

Also, prepare the buffer without Triton X-100 and glycerol to be used for resuspending extracts for sonication.



COMMENTARY Background Information The glutathione-S-transferase (GST) fusion–affinity purification is a simple method to present a native bait protein (as a GST fusion protein) to a test protein or complex of proteins. The assay does not involve denaturing either bait or test proteins, nor does it rely on the more restrictive assay of coimmunoprecipitation with an antibody. It is faster than using the in vivo two-hybrid interaction trap system (UNIT 20.1) to detect protein-protein interaction and can be performed in a defined medium. It is also quite sensitive—capable of detecting interactions with Kds in the micromolar range. The GST pulldown assay has usually been used for qualitative assessment of whether two proteins interact, but demonstration of binding in vitro does not necessarily indicate that the binding has in vivo biological relevance. Once a binding reaction is identified, variations on the basic assay can be used to further characterize the reaction. The actual bait sequence sufficient for interaction with the test protein can be identified by constructing GST fusions to different regions or fragments of the bait protein and testing them in the binding assay. If it is possible to quantitate the amount of bait protein, the binding constant can then be determined. The half-life of the binding reaction can be determined by binding the labeled test protein in an excess of bait protein: after equilibrium is reached, a large excess of cold test protein is added, and the amount of labeled test protein retained on the beads is assayed at various time points. Because it is not generally feasible to produce large amounts of labeled protein by in vitro translation, the test protein can alternatively be labeled with 32P in vitro by incorporating a five-amino-acid recognition element for the catalytic domain of heart muscle kinase into the amino-terminal sequence of the protein (Blanar and Rutter, 1992). Competition assays can indicate whether two proteins use exclusive binding sites. With a limiting amount of bait protein, equimolar amounts of two test proteins are added, and the relative amounts of these proteins bound to the bait protein at equilibrium are assayed. The system can also be used to construct a binding pathway for a complex. For example, it may be determined that test protein A may only bind the bait in the presence of test protein B, whereas protein B binds the bait independently of A. This approach has been elegantly used to reconstruct the TFIID complex (Chen et al., 1994).

Many of the potential applications of the pulldown technique rely on accurate quantitation of the reactants and products. However, the sensitivity of this assay rests in the mass-action effects of using high concentrations of bait which allows detection of even weak interactions but also creates the possibility of artifacts. Paying attention to the quantitative features of the pulldown assays leads to an appreciation of both the limitations and powers of this technique.

Critical Parameters Because of the potential for artifacts, merely showing that a test protein binds the bait protein but not GST is not sufficient to establish that this interaction takes place in vivo. Support for a positive binding result can come either from independent assays, such as the two-hybrid interaction trap or immunoprecipitation, or from other control affinity experiments. The most convincing demonstrations are those in which a control variant bait protein that has lost function in vivo also does not bind in vitro (see Fig. 20.2.3) and those in which the bait–test protein complex reconstitutes an in vivo activity. Quantitation of the amounts of bait and test proteins relative to the question being posed is critical. If the bait is present at a high concentration, nonspecific interactions can be enhanced. If the test protein is in excess, no estimate of relative binding to the bait protein can be made. If substantially more test protein binds to the beads than to bait protein, this probably indicates a nonspecific interaction with the beads; however, test proteins may, in some cases, exhibit a genuine interaction in molar excess over bait (Fig. 20.2.2). For quantitative purification of a test protein, however, the bait should be in a large excess and the amount of bait bound to the beads should be directly determined. Some GST fusion proteins are prone to degradation, so the beads will contain partial fragments in addition to full-length fusions. Usually, proteolysis leaves an intact GST with little or no bait protein attached, and the amount of GST verses GST-fusion protein can be directly estimated by Coomassie stain on an SDS gel. However, some fusion proteins are prone to small C-terminal deletions, and if estimation of the amount of full-length protein is critical, it should be done carefully. Degradation can result in the accumulation of N-terminal bait fragments on



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retained –80

+80

extract –80

+80

GAL80

GST-34 GST breakdown product

Figure 20.2.2 Purification of an interacting test protein from a crude E. coli extract (−80)/(+80). Crude E. coli extracts from control cells (−80) or cells expressing the yeast negative regulator Gal80p (+80) were incubated with a GST fusion protein containing a 34-amino-acid fragment that included the activation domain of the yeast transactivator Gal4p (GST-34). After extensive washing, the beads were boiled in 2× SDS sample buffer and analyzed by SDS-PAGE. Proteins were visualized by Coomassie blue staining. This gel shows that the retained fraction includes GST that has lost the 34-mer bait protein. Gal80p binds to GST-34 as a dimer, hence the molar excess seen in the +80 retained fraction. Photograph provided by Karsten Melcher.

the agarose beads, because these are still able to bind via the GST moiety. If the point of fusion between GST and the bait protein is attacked, GST alone is bound to the agarose beads. If GST is degraded, the fusion protein is unable to bind to the beads. If there are a substantial number of GST fusions with N-terminal bait protein fragments, quantitation of active protein will require limiting proteolysis or purifying the full-length fusion protein.

Troubleshooting


If an interaction occurs between two proteins in vivo, a GST pulldown assay will usually detect it. If it is suspected that two proteins interact in vivo but the interaction is not detected in the pulldown assay, there are two possible explanations. One is that there is no interaction. The other is that something is suboptimal or missing in the assay. The bait protein may be sterically inhibited as a fusion protein. In this case, the bait protein can be switched to the N-terminus of GST or con-

structed using another affinity tag. Another possibility is that the bait protein is not properly modified when produced as a fusion protein in E. coli. In this case, the fusion protein can be produced in yeast (Higgins, 1995; Romanos et al., 1995; Strausberg and Strausberg, 1995), baculovirus (see UNITS 16.9, 16.10 & 16.11), or mammalian cells (see UNITS 16.12, 16.13 & 16.14). However, if one of these alternative systems is chosen, it is better to use an affinity tag other than GST (such as maltose or biotin) because the level of glutathione in eukaryotic cells can inhibit binding to the agarose beads. Finally, the binding reaction may require one or more additional factors. Investigating this possibility will require binding the test protein out of a cellular extract or adding cellular extract to the labeled, in vitro–translated test protein. If the test protein is specifically retained on the bait protein agarose but not on the GST or negative control agarose, it does not necessarily mean that the bait and test proteins interact in vivo. Artifacts may result from mass-action



input

–

+

–

(+ –)

+

–

+

retained Gal80p

1

2

3

4

5

6

7

Figure 20.2.3 Replication in vitro of in vivo Gal4p/Gal80p interactions. In the presence of 10 to 20 µg bait protein and in the absence of complex competitor protein, Gal80p binds to all the mutant Gal4p activation domains tested, even those it does not repress in vivo (data not shown). However, this figure shows that when the amount of bait protein is reduced to 4 µg and 4 mg complex competitor is included, the in vitro pulldown assay parallels the in vivo interaction seen between Gal80p and several different Gal4p activation domain mutants. Lane 1, GST alone; Lane 2, GST + wild-type Gal4p activation domain; Lanes 3 to 7, GST + mutant Gal4p activation domains. A + indicates mutants with a positive in vivo Gal4p/Gal80p interaction; a − indicates mutants with negative in vivo Gal4p/Gal80p interaction; and ± indicates mutants with low-level in vivo Gal4p/Gal80p interaction. Abbreviation: GST, glutathione-S-transferase. Photograph provided by Karsten Melcher.

effects of large amounts of bait or target protein that drive nonphysiological interactions. Artifacts may also be the consequence of taking a protein out of its normal context. A protein that normally exists in a complex may, when expressed as a GST fusion protein, expose many surfaces that are not normally exposed in the cell. These surfaces may be “sticky” and lead to a general affinity for other proteins. Artifactual interactions can be minimized by reducing the amount of bait protein. Initial GST pulldown assays used tens or even hundreds of micrograms of bait protein. Such high concentrations can produce mass-action effects that drive any interactions with the test protein. While developing conditions to replicate in vitro the known in vivo interactions between the activation domain of the yeast transactivator Gal4p and its repressor Gal80p (Fig. 20.2.3), the authors found that the amount of bait protein needed to be reduced to 1 to 5 µg per reaction. Inclusion of a complex protein competitor, such as an extract of soluble E. coli proteins, in the reaction mix also reduces artifactual interactions. This is very important for the reduction of nonspecific binding, for which a complex competitor is much more effective than a single competing protein like BSA. In the standard assay, the complex protein competitor is pre-

sent in ≥100-fold excess over the GST fusion protein.

Anticipated Results In the protocol described, with E. coli competitor, typically 80% of the input test protein binds to the bait if the affinity is ≤10−8 M. For proteins with binding constants of 10−6 to 10−8 M, typically 10% to 40% of input test protein is retained. Of course, with large amounts of bait and no competitor even weakly binding proteins can be detected. If the interaction assay identifies a novel or unknown protein, the system can be scaled up to obtain biochemical amounts of the novel protein for further analysis and identification. After elution with protease or glutathione, the target protein should be highly enriched. Further purification can then be performed following standard protocols (see Chapter 10).

Time Considerations One of the attractive features of this protocol is how quickly it can be performed. Using the incubation times outlined in the protocol description and labeled test protein, it is easy to determine whether the test protein binds to the GST fusion protein within 1 day. Analysis of the bound test protein by immunoblotting (UNIT 10.8) requires an additional day. Construction,



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expression, and purification of the GST fusion protein and binding to beads (UNIT 16.7) requires additional time.

Literature Cited Agarana, C.E., Kuntz, I.D., Birkin, S., Axel, R., and Cantor, C.R. 1986. Molecular cloning and nucleotide sequence of the streptavidin gene. Nucl. Acids Res. 14:871-882. Blanar, M.A. and Rutter, W.J. 1992. Interaction cloning: Identification of a helix-loop-helix zipper protein that interacts with c-fos. Science 256:1014-1018. Chandler, C.S. and Ballard, F.J. 1988. Regulation of the breakdown rates of biotin-containing proteins in Swiss 3T3-L1 cells. Biochem. J. 251:749-755. Chen, J.-L., Attardi, L.D., Verrijzer, C.P., Yokomori, K., and Tjian, R. 1994. Assembly of recombinant TFIID reveals differential coactivator requirements for distinct transcriptional activators. Cell 79:93-105. Higgins, D.R. 1995. Overview of protein expression in Pichia pastoris. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, H.L. Ploegh, D.W. Speicher, and P.T. Wingfield, eds.) pp. 5.7.1-5.7.16. John Wiley & Sons, New York. Johnsson, N. and Varshavsky, A. 1994. Split ubiquitin as a sensor of protein interactions in vivo. Proc. Natl. Acad. Sci. U.S.A. 91:1034010344.

Nikolau, B.J., Wurtele, E.S., and Stumpf, P.K. 1985. Use of streptavidin to detect biotin-containing proteins in plants. Anal. Biochem. 149:448-453. Romanos, M.A., Clare, J.I., and Brown, C. 1995. Culture of yeast for the production of heterologous proteins. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, H.L. Ploegh, D.W. Speicher, and P.T. Wingfield, eds.) pp. 5.8.1-5.8.17. John Wiley & Sons, New York. Strausberg, R.L. and Strausberg, S.I. 1995. Overview of protein expression in Saccharomyces cerevisiae. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, H.L. Ploegh, D.W. Speicher, and P.T. Wingfield, eds.) pp. 5.6.1-5.6.7 John Wiley & Sons, New York.

Key Reference Melcher, K. and Johnston, S.A. 1995. Gal4 interacts with TATA-binding protein and co-activators. Mol. Cell Biol. 15:2839-2848. Describes binding from extracts, binding of in vitrotranslated and labeled protein, binding-deficient mutants, correlation of in vitro and in vivo results, and determination of binding constant.

Contributed by Jonathan C. Swaffield and Stephen Albert Johnston University of Texas Southwestern Medical Center Dallas, Texas

Lim, P., Rhode, M., Morris, C.P., and Wallace, J.C. 1987. Pyruvate carboxylase in the yeast pyc mutant. Arch. Biochem. Biophys. 258:219-264.




Phage-Based Expression Cloning to Identify Interacting Proteins

UNIT 20.3

Interaction cloning (also known as expression cloning) is a technique to identify and clone genes which encode proteins that interact with a protein of interest, or “bait” protein. Phage-based interaction cloning requires a gene encoding the bait protein and an appropriate expression library constructed in a bacteriophage expression vector, such as λgt11. The gene encoding the bait protein is used to produce recombinant fusion protein in E. coli. The cDNA is radioactively labeled with 32P. A recognition site for cyclic adenosine 3′, 5′-phosphate (cAMP)–dependent protein kinase (protein kinase A; PKA) is introduced into the recombinant fusion protein to allow its enzymatic phosphorylation by PKA and [γ-32P]ATP. The procedure presented here (see Basic Protocol) involves a fusion protein consisting of bait protein and glutathione-S-transferase (GST) with a PKA site at the junction between them (the protocol can, however, be adapted to use other PKA-containing recombinant proteins). The labeled protein is subsequently used as a probe to screen a λ bacteriophage-derived cDNA expression library, which expresses β-galactosidase fusion proteins that contain in-frame gene fusions. The phages lyse cells, form plaques, and release fusion proteins that are adsorbed onto nitrocellulose membrane filters. The filters are blocked with excess nonspecific protein to eliminate nonspecific binding and probed with the radiolabeled bait protein (see Fig. 20.3.1). This procedure leads directly to the isolation of genes encoding the interacting protein, bypassing the need for purification and microsequencing or for antibody production. NOTE: Radioactive label is used in this protocol, and appropriate precautions and shielding should be used (APPENDIX 1F). STRATEGIC PLANNING There are two important choices one must make to begin this procedure: (1) how to design the bait protein and (2) how to construct or acquire an appropriate phage-derived expression library. Vectors for recombinant fusion protein expression that contain a PKA recognition site can be obtained commercially. Several companies now sell these vectors with various affinity tags such as GST (Pharmacia Biotech), histidine (Novagen), or calmodulin-binding protein (Stratagene). Alternatively, one can engineer the PKA recognition site (the five–amino acid sequence RRASV) into existing vectors by using synthetic DNA that encodes it. Lambda-derived expression libraries that direct the expression of cDNAs made from many different mRNA sources are widely available. Alternatively, a library may be constructed for a particular experimental purpose; when designing a library to be used in expression cloning, several points should be considered. Libraries made from cDNA synthesized with random primers or an oligo dT-primer during first-strand synthesis (UNITS 5.5 & 5.6) can often be advantageous in that multiple clones representing different portions of the same protein can be identified in the screening. Analysis of coding regions present in these multiple clones could provide useful information about what region of the protein is responsible for the observed interaction. Full-length cDNA clones can subsequently be obtained from other available libraries. Many suitable λ vectors are available for constructing cDNA expression libraries. The most widely used cDNA expression vector has been λgt11 (Huynh et al., 1985). There are modifications of these λ vectors, however, that facilitate the recovery of the cDNA Contributed by Julie M. Stone Current Protocols in Molecular Biology (1997) 20.3.1-20.3.9 Copyright © 1997 by John Wiley & Sons, Inc.



inserts by avoiding the necessity of time-consuming λ phage DNA preparations. These modified vectors include those that make use of the cre-lox recombination for in vivo conversion of recombinant phages into plasmid DNA in cre recombinase-expressing host strains (e.g., λZipLox, Life Technologies) as well as those that employ helper phage for in vivo excision of a phagemid vector (e.g., the λZAP series of vectors, Stratagene). Most λ vectors available produce fusions of the cDNA inserts to β-galactosidase, because they are cloned into the lacZ gene. However, some λ-derived expression vectors (e.g., λSCREEN-1, Novagen) direct expression of proteins fused to the T7 DNA polymerase promoter (gene 10 under control of the T7 promoter) and may yield higher expression of the library proteins (Margolis et al., 1992). BASIC PROTOCOL

INTERACTION CLONING Phage-based interaction expression cloning is a simple, rapid, and powerful technique to identify interacting proteins. A protein of interest is expressed as a recombinant fusion protein and labeled with 32P at a serine residue in an engineered PKA recognition site to facilitate detection. β-galactosidase proteins that are fused in-frame to cDNA inserts in a bacteriophage λ-derived expression library are produced by the phage and adsorbed onto nitrocellulose filters. The filters are then screened with the radiolabeled protein probe to identify phage clones that express an interacting protein.

A G ST

PKA ba it

B probed with GST-bait probed with GST

Phage-Based Expression Cloning

Figure 20.3.1 Schematic representation of the interaction cloning technique used to identify proteins that associate with a protein of interest (bait protein), and the expected results of a successful screen. (A) Expression of β-galactosidase fusion proteins from in-frame cDNA inserts of a λgt11 Arabidopsis library is induced with IPTG-impregnated nitrocellulose membrane filters (indicated by an oval). Filters are probed with GST-bait fusion protein labeled with 32P (•) at the PKA recognition site located at the junction of the fusion. Interacting clones are detected by autoradiography. (B) Representative autoradiogram of a tertiary screen of a positive plaque after the control experiment to determine whether the interaction is specific for the bait portion of the probe. The top half of the filter is probed with GST-bait while the bottom half is probed with GST alone as a control.



Materials cAMP-dependent protein kinase (PKA; e.g., 250-U lots from Sigma) 40 mM DTT, prepared fresh 10× PKA buffer (see recipe) 10 mCi/ml [γ-32P]ATP (6000 mCi/mmol) Purified glutathione-S-transferase (GST)–bait protein fusion protein with a PKA recognition site (UNIT 16.7), at ∼0.1 to 1 µg/µl concentration Z′-KCl (see recipe), ice cold Sephadex G-50 equilibrated in Z′-KCl E. coli Y1090r− or other appropriate host strain LB medium containing appropriate selective antibiotic (see UNIT 1.1, APPENDIX 3F, and Table 1.4.1), 10 mM MgSO4, and 0.2% maltose 10 mM MgSO4 10 mM IPTG (Table 1.4.2) 150- or 100-mm LB plates (with antibiotic, if necessary; UNIT 1.1) 0.7% top agarose (UNIT 1.1), 47°C Tris-buffered saline with Triton X-100 (TBS-T; see recipe) India ink HEPES blocking buffer (HBB; see recipe) Binding buffer (BB; see recipe) Suspension medium (SM; UNIT 6.12) Chloroform 3-ml disposable plastic columns or disposable syringe and glass wool Scintillation counter and fluid Tabletop centrifuge or equivalent Nitrocellulose membrane filters (137- and 82-mm disks) 22-G needle Additional reagents and equipment for preparation and purification of recombinant glutathione-S-transferase fusion protein (UNIT 16.7), SDS-PAGE (optional; UNIT 10.2), autoradiography (APPENDIX 3A), titering and plating λ phage to generate plaques (UNIT 1.11 & UNIT 6.1), and purification of bacteriophage clones (UNIT 6.5) Prepare the 32P-labeled protein probe 1. Resuspend 250 U PKA in 25 µl freshly prepared 40 mM DTT. Let the reconstituted enzyme stand at room temperature ∼10 min before use. It is important to use freshly prepared 40 mM DTT. PKA is extremely unstable after reconstitution; the enzyme stock solution can be stored temporarily at 4°C but retains activity for only 2 to 3 days.

2. Prepare a phosphorylation reaction mixture containing: 1 µl 10 U/µl PKA (from step 1) 3 µl 10× PKA buffer 5 µl 10 mCi/ml (6000 mCi/mmol) [γ-32P]ATP 1 to 10 µl (∼1 µg) purified GST-bait fusion protein H2O to 30 µl. Incubate 1 hr at room temperature. A fusion protein unrelated to the bait protein, but containing the PKA recognition site, should also be expressed for use as a control to determine whether the observed interaction is specific for the bait moiety. Due to the instability of PKA, it is advisable to label both the bait protein and the unrelated control fusion protein simultaneously (in separate reactions).



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3. Add 170 µl ice-cold Z′-KCl (to stop reaction) and store on ice until use. 4. Prepare a gel filtration column by pouring Sephadex G-50 equilibrated in Z′-KCl into a 3-ml disposable plastic column (or a 3-ml syringe with a glass wool plug) for a final bed volume of ∼3 ml. Allow the column to drain until the level of the buffer is at the top of the column bed. The Sephadex G-50 can be swelled in water or buffer and stored at 4°C for months. In this case, five to ten column volumes of Z′-KCl can be used to equilibrate the column after it is poured.

5. Load the entire phosphorylation reaction (from step 3) onto the column and collect the effluent in a 1.5-ml microcentrifuge tube. Place a second tube under the column, add a 200-µl aliquot of Z′-KCl, and collect the effluent again. Repeat the above loading and collecting steps an additional ten times to collect a total of twelve 200-µl fractions. 6. Measure the Cerenkov counts with a scintillation counter to identify the fractions with the highest specific activities, and calculate cpm/µl. Typically two peaks of radioactivity are observed. The first peak elutes in fractions five to nine and corresponds to labeled protein. The second peak, usually found in the last few fractions collected, corresponds to unincorporated ATP and should be discarded. The hottest fraction(s), which elute first, should be used. Fractions can be stored at 4°C for several weeks.

7. Optional: Analyze a small amount (1 to 2 µl) of the 32P-labeled protein probe by SDS-PAGE and autoradiography (UNIT 10.2 & APPENDIX 3A). Typically two strong signals are observed: one at the predicted size of the GST-bait fusion protein and the other at the predicted molecular weight of the GST alone (28 kDa). This is due to the fusion protein’s inherent susceptibility to protease cleavage at the GST-bait protein junction. The presence of labeled GST protein in the probe should not interfere with the screen; a control experiment with labeled GST will be performed at the later stages of screening.

Prepare host strain cells and dilution of bacteriophage cDNA library 8. Using serial dilution as described in UNIT 1.11, determine the titer of the bacteriophage cDNA library. 9. Grow an overnight 50-ml culture of E. coli Y1090r− (or other appropriate host strain; UNIT 1.4) in LB medium containing an appropriate selective antibiotic, 10 mM MgSO4, and 0.2% maltose. 10. Centrifuge cells 10 min at 2000 × g, room temperature, and resuspend in 25 ml of 10 mM MgSO4. The resuspended cells can be stored up to 1 week at 4°C before use.

Prepare the filters to screen the bacteriophage cDNA expression library 11. Soak nitrocellulose filters in 10 mM IPTG for 15 min at room temperature and air dry. IPTG-impregnated filters can be stored in a petri plate until use.

12. Prepare eight 1.5-ml microcentrifuge tubes each containing 0.6 ml Y1090r− cells (from step 10) and ∼40,000 pfu bacteriophage (from step 8), and incubate 15 min at 37°C. Phage-Based Expression Cloning



13. Add contents of each tube to 7 ml of 0.7% top agarose at 47°C and pour onto 150-mm LB plates (with antibiotic, if necessary). Incubate plates ∼3 hr at 42°C, until small plaques are visible. 14. Overlay the plates with IPTG-impregnated filters and incubate an additional 6 to 8 hr at 37°C. Formation of plaques in the absence of induction of the lacZ gene promoter ensures that any library-encoded proteins that are deleterious to phage growth will not be expressed while the phage are forming a plaque. Incubation in the presence of IPTG-impregnated filters is usually performed for a 6- to 8-hr period, but can proceed overnight for convenience.

15. Chill plates 15 min (or overnight) at 4°C. 16. Pierce each filter in several locations with a 22-G needle dipped in India ink to mark orientation. Remove filters from plates and wash 15 min in TBS-T at room temperature with shaking. Screen bacteriophage cDNA expression library 17. Incubate filters 1 to 4 hr with rocking at 4°C in 100 ml HBB. This blocking step (to reduce nonspecific binding) can also be performed overnight for convenience.

18. Incubate overnight with rocking at 4°C in BB containing 2.5–5 × 105 cpm/ml of radiolabeled fusion protein (from step 6). All eight filters can be placed in one 150-mm plate and probed with 30 to 40 ml of solution. The plate is wrapped in Parafilm and placed in a Plexiglas box for shielding. Alternatively, the filters can be placed with solution in a heat-sealable bag. The probe solution can be stored at 4°C and used for the subsequent secondary and tertiary screenings.

19. Wash membrane filters three times, 10 min each, in 100 ml BB with shaking at room temperature. Air dry and expose to film (see APPENDIX 3). 20. Using the large end of a Pasteur pipet, take an agarose plug at the position of the positive clone. Place into a 1.5-ml microcentrifuge tube and add 1 ml SM and 1 drop of chloroform. Agarose plugs can be stored at 4°C for months. Therefore, if many putative positive clones are identified in the primary screen, they can all be stored for future analysis if necessary.

21. Determine the titer by serial dilution (UNIT 1.11) and perform successive screening procedures to obtain purified clones as described in UNIT 6.5. Subsequent screens are performed on 100-mm LB plates with ∼2000 pfu/plate for secondary screens and ∼300 to 500 pfu/plate for tertiary screens. Before purified clones are obtained (usually during the tertiary screen), a control to eliminate clones that might interact with the GST fusion portion of the probe should be performed. To do this, cut the filters in half and probe one half with the labeled GST fusion to the protein of interest and the other with labeled GST alone or an unrelated control GST fusion protein (see Fig. 20.3.1).



Supplement 39

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. All solutions should be prepared from sterile, autoclaved stock solutions, except Z′-KCl, which should be filter sterilized. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Binding buffer (BB) 20 mM HEPES⋅OH, pH 7.4 7.5 mM KCl 0.1 mM EDTA 2.5 mM MgCl2 1% (w/v) nonfat dry milk The solution can be prepared without milk and stored indefinitely at room temperature (add milk prior to use).

HEPES blocking buffer (HBB) 20 mM HEPES⋅OH, pH 7.4 5 mM MgCl2 1 mM KCl 5% (w/v) nonfat dry milk The solution can be prepared without milk and stored indefinitely at room temperature (add milk prior to use).

PKA buffer, 10× 200 mM Tris⋅Cl, pH 7.5 10 mM DTT 1 M NaCl 120 mM MgCl2 Store up to 6 months at room temperature Tris-buffered saline with Triton X-100 (TBS-T) 10 mM Tris⋅Cl, pH 8.0 150 mM NaCl 0.05% (v/v) Triton X-100 Store up to 6 months at room temperature Z′-KCl 25 mM HEPES⋅OH, pH 7.4 12.5 mM MgCl2 20% (w/v) glycerol 100 mM KCl 1 mg/ml BSA 1 mM DTT Filter sterilize Store up to 6 months at 4°C COMMENTARY Background Information


Historically, interacting proteins have been isolated by biochemical approaches, which required purification of the interacting protein for antibody production or microsequencing before a clone encoding the protein could be identified. Two of the molecular biological approaches described in this chapter, the yeast two-hybrid system (UNIT 20.1) and phage-based

interaction expression cloning (this unit), however, directly yield a clone encoding the interacting protein. Bacteriophage cDNA expression libraries are commonly screened using antibodies (see UNIT 6.7 and references therein) or radiolabeled DNA probes to identify DNA-binding proteins (see UNIT 12.7 and references therein). Modifications to identify interacting proteins include



screening with the protein of interest (the bait protein) and detecting that protein with antibodies (Chapline et al., 1993). However, by screening with a radiolabeled protein probe, one avoids the additional incubations and washes which are necessary for immunodetection but that increase the likelihood of disrupting weak proteinprotein interactions. 125I-labeled protein probes have been used successfully to screen for interacting proteins (Hoeffler et al., 1991), although the technique described in this unit avoids the need to label the proteins with 125I, and the complications of handling this isotope. Protein probes autophosphorylated with 32P were originally used to screen cDNA expression libraries in the isolation of proteins that interact with receptor protein kinases, a technique referred to as CORT (cloning of receptor targets; Skolnik et al., 1991; Lowenstein et al., 1992; Margolis et al., 1992). By introducing a PKA recognition site into the protein probe, the technique was made suitable for proteins that were not themselves protein kinases (Blanar and Rutter, 1992; Kaelin et al., 1992). Phage-based interaction expression cloning as described in this unit has been used successfully to identify many interacting proteins, but may not be successful for all types of interactions. For example, many related proteins (known as A-kinase anchoring proteins, or AKAPs) have been identified by their ability to interact with the type II cAMP-dependent kinase regulatory subunit, RII (Carr and Scott, 1992). All the AKAPs can bind RII under denaturing conditions (Lester et al., 1996), illustrating that the success of interaction cloning is often dependent on the nature of the interactions and that interactions less dependent on three-dimensional structure may be favored. However, there are many examples of proteins identified by this technique that cannot interact under the denaturing conditions often used to confirm the binding (commonly referred to as overlay assays or Far Western analysis). For example, a protein domain identified by its interaction with a plant receptor–like protein kinase is capable of interacting with a denatured form of the bait protein, but is unable to interact when the interacting protein is denatured (Stone et al., 1994). This fact is consistent with the idea that immobilization on membranes denatures some proteins but not others.

Critical Parameters and Troubleshooting The success of phage-based interaction expression cloning is inherently dependent on the

quality of the protein probe used and the extent of representation of the bacteriophage cDNA library. GST fusion proteins are often obtained in high quantity in a soluble form, avoiding the necessity of solubilizing and refolding during protein purification from E. coli extracts. In most cases the PKA recognition site is readily accessible, and allows production of radiolabeled protein with a high specific activity. The strength of the protein-protein interaction is also critical. Depending on the nature of the interaction, a filter-binding technique might not be suitable. This technique may not detect weak or transient interactions, such as an enzyme/substrate interaction, and techniques such as the yeast two-hybrid system (UNIT 20.1) might be more appropriate. If no positive clones are identified or if background is high, it may be helpful to add reducing agents or detergents or to alter the salt conditions of the binding and wash solutions (Vinson et al., 1988). Moreover, denaturation and renaturation of the proteins on the filters using 6 M guanidine⋅HCl, as described in UNIT 12.7 (Alternate Protocol), may facilitate the recovery of clones expressing interacting proteins, because adsorption of the β-galactosidase fusion proteins to nitrocellulose filters may alter the conformation of the proteins. For a discussion of common problems with this procedure and their diagnosis and possible solutions, see Table 20.3.1.

Anticipated Results Depending on the nature of the interaction being sought by this technique, many interacting clones, none, or just a few may be identified. However, the technique is as simple as screening a library by hybridization and often well worth the effort. In any case, observed interactions should be confirmed by other means.

Time Considerations The interaction cloning technique described in this unit is extremely rapid in comparison with other techniques to identify interacting proteins, such as the yeast two-hybrid system (UNIT 20.1). The technique does not require any unusual reagents that would not be readily available in any laboratory routinely engaged in molecular biology, other than the cAMP-dependent protein kinase. Once the appropriate recombinant fusion protein and cDNA expression library are obtained, the primary screen can be completed in a few days. Once the appropriate construct for expression of the GST-bait is obtained, purification of the recom-



Supplement 39

Table 20.3.1

General Troubleshooting Guide for Interaction Cloning

Problem

Possible cause

Solution

High background

Insufficient washing

Use more BB wash solution

Poor choice of nitrocellulose membrane filters Insufficient blocking

Try a different supplier for membranes Block overnight in HBB

Poor-quality protein probe

Check the probe by SDS-PAGE and autoradiography

Wash conditions too stringent

Reduce wash times Vary salt and/or detergent concentration

Poorly folded library fusion proteins

Perform 6 M guanidine⋅HCl denaturation/renaturation (UNIT 12.7)

Wash and binding conditions insufficiently stringent Library biased to proteins that interact with the affinity tag

Increase salt and/or detergent in BB

No positive plaques

Too many positive plaques

binant GST-bait protein can be achieved in 1 day. 32P labeling of the bait protein requires 2 hr (several additional hours if the optional SDSPAGE and autoradiography are performed). The initial screening of the bacteriophage cDNA library takes 3 days; day 1 to prepare the filters and induce expression of library-encoded proteins, day 2 for blocking and probing overnight with 32 P-labeled bait protein, and day 3 for washing and autoradiography. Subsequent purification of the cDNA clones should only take an additional week. There are a number of steps that can be performed either rapidly or overnight, providing a great deal of convenience and flexibility.

Literature Cited Blanar, M.A. and Rutter, W.J. 1992. Interaction cloning: Identification of a helix-loop-helix zipper protein that interacts with c-Fos. Science 256:1014-1018. Carr, D.W. and Scott, J.D. 1992. Blotting and bandshifting: Techniques for studying protein-protein interactions. Trends Biochem. Sci. 17:246-249. Chapline, C., Ramsay, K., Klauck, T., and Jaken, S. 1993. Interaction cloning of protein kinase C substrates. J. Biol. Chem. 268:6858-6861.


Hoeffler, J.B., Lustbader, J.W., and Chen, C.Y. 1991. Identification of multiple nuclear factors that interact with cyclic AMP response element-binding protein and activation transcription factor-2 by protein interactions. Mol. Endocrinol. 5:256266.

Include unlabeled affinity tag in blocking, binding, and wash solutions

Huynh, T.V., Young, R.A., and Davis, R.W. 1985. Constructing and screening cDNA libraries in λgt10 and λgt11. In DNA Cloning: A Practical Approach (D.M. Glover, ed.) pp. 49-78. IRL Press, Oxford. Kaelin, W.G.J., Krek, W., Sellers, W.R., DeCaprio, J.A., Ajchenbaum, F., Fuchs, C.S., Chittenden, T., Li, Y., Farnham, P.J., Blanar, M.A., Livingston, D.M., and Flemington, E.K. 1992. Expression cloning of a cDNA encoding a retinoblastoma-binding protein with E2F-like properties. Cell 70:351-364. Lester, L.B., Coghlan, V.M., Nauert, B., and Scott, J.D. 1996. Cloning and characterization of a novel A-kinase anchoring protein: AKAP220, association with testicular peroxisomes. J. Biol. Chem. 271:9460-9465. Lowenstein, E.J., Daly, R.J., Batzer, A.G., Li, W., Margolis, B., Lammers, R., Ullrich, A., Skolnik, E.Y., Bar-Sagi, D., and Schlessinger, J. 1992. The SH2 and SH3 domain-containing protein GRB2 links receptor tyrosine kinases to ras signaling. Cell 70:431-442. Margolis, B., Silvennoinen, O., Comoglio, F., Roonprapunt, C., Skolnik, E., Ullrich, A., and Schlessinger, J. 1992. High-efficiency expression/cloning of epidermal growth factor–receptor-binding proteins with src homology 2 domains. Proc. Natl. Acad. Sci. U.S.A 89:8894-8898. Skolnik, E.Y., Margolis, B., Mohammadi, M., Lowenstein, E., Fischer, R., Drepps, A., Ullrich, A., and Schlessinger, J. 1991. Cloning of PI3 kinase–associated p85 utilizing a novel method of expression/cloning of target proteins for receptor tyrosine kinases. Cell 65:83-90.



Stone, J.M., Collinge, M.A., Smith, R.D., Horn, M.A. and Walker, J.C. 1994. Interaction of a protein phosphatase with an Arabidopsis serinethreonine receptor kinase. Science 266:793-795. Vinson, C.R., LaMarco, K.L., Johnson, P.F., Landschulz, W.H., and McKnight, S.L. 1988. In situ detection of sequence-specific DNA binding activity specified by a recombinant bacteriophage. Genes & Dev. 2:801-806.

Key References Blanar and Rutter, 1992. See above. The basic protocol described in this unit is modified directly from the Blanar and Rutter protocol. Huynh et al., 1985. See above. Provides an excellent description of constructing and screening λgt11 cDNA expression libraries.

Contributed by Julie M. Stone University of Missouri, Columbia Columbia, Missouri



Supplement 39

Surface Plasmon Resonance for Measurements of Biological Interest

UNIT 20.4

Genetic manipulations, including gene knockouts and mutant screens, provide an initial hint as to the function of a gene product and indicate possible associated factors. To unravel complicated biological processes that control the development of organisms one must identify the interacting components. Yeast two-hybrid methods (UNIT 20.1; Gyuris et al., 1993; Zervos et al., 1993; Bartel and Fields, 1995) provide an in vivo means of identifying species that directly associate with each other. However, many biological systems are notoriously redundant and consist of networks of interacting components. For these and other reasons, it is advisable to verify the in vivo observation by an in vitro detection method where there are fewer variables and where reaction conditions can be controlled. Few techniques measure equilibrium binding because routine washing steps can cause the dissociation of the interaction under study. Moreover, most in vitro techniques used for the detection of intermolecular interactions require the use of antibodies or protein-labeling and subject the interaction under study to harsh and potentially denaturing conditions. These techniques only provide static “snap-shots” of a dynamic process, and kinetic parameters must be inferred from successive experiments. An in vitro technique based on an optical phenomenon, called surface plasmon resonance (SPR; Liedberg et al., 1983; Daniels et al., 1988), can simultaneously detect interactions between unmodified proteins and directly measure kinetic parameters of the interaction. The use of SPR to study macromolecular interactions has recently become popular with the availability of user-friendly instruments. The most popular commercially available SPR device is the BIAcore instrument (BIAcore). This instrument consists of sensing optics, an automated sample delivery system, and a computer for instrument control, data collection, and data processing. Experiments are performed on disposable chips (Löfås and Johnsson, 1990). In practice, a ligand protein is immobilized on the chip while buffer continuously flows over the surface. The buffer flow is interrupted when sample “plugs” containing putative binding partners (analyte or target proteins) are sequentially flowed over the protein surface. The sensing apparatus monitors changes in the angle of minimum reflectance from the interface that result when a target protein associates with the ligand protein. Molecular interactions can be directly visualized (on the computer monitor) in real time as the optical response is plotted against time. This response is measured in resonance units (RUs, where 1000 RUs = 1 ng protein/mm2). The user handbook with the BIAcore SPR apparatus contains detailed information on the use of the equipment, as well as a number of protocols and information for experimental design. Basic Protocol 1, presented here, is a modification of the standard protocol in the BIAcore user handbook, and is used with the CM-5 BIAcore carboxylated dextran chip. Basic Protocol 2 describes the particular details necessary when using NTA-SAM chips. A detailed discussion on choosing between these two kinds of chips can be found in Critical Parameters and Anticipated Results. The use of SPR provides an accurate measurement of kinetic rate constants for a ligand-target binding reaction. If a ball-park estimate of the range of the equilibrium constants is desired, or if an absolute kinetic rate constant is desired, refer to the discussion in Anticipated Results.

Analysis of Protein Interactions Contributed by Cynthia Bamdad Current Protocols in Molecular Biology (1997) 20.4.1-20.4.12 Copyright © 1997 by John Wiley & Sons, Inc.


BASIC PROTOCOL 1

SPR USING BIAcore CHIPS The following protocol is used when performing an SPR experiment on a BIAcore device with the BIAcore CM-5 chip. This is the most commonly used SPR chip and is made with carboxylated dextran. Two particular deviations from the standard BIAcore protocol are discussed within annotations (steps 3 and 6); both deviations concern ligand binding to the chip, and were designed to avoid artifactual pitfalls that are peculiar to dextran-based chips. For detailed steps in the performance of an SPR experiment, and for additional reagents and equipment information, refer to the BIAcore user handbook. Materials CM-5 dextran chips (BIAcore) PBS (APPENDIX 2) Ligand protein Target protein Amine-coupling kit (BIAcore), containing: N-ethyl-N′-[(dimethylamino) propyl] carbodiimide hydrochloride (EDC) N-hydroxysuccinimide (NHS) Sodium acetate buffer, low pH Ethanolamine BIAcore SPR equipment BIA evaluation point-and-click software Determine binding conditions 1. Insert a new BIAcore CM-5 dextran chip. 2. Allow the system to equilibrate with PBS as the running buffer in the constant flow system. 3. Experimentally determine buffer conditions and concentrations that minimize nonspecific binding of target and ligand proteins to the carboxylated dextran. With dextran chips, conditions must be determined for the ligand because a ligand which has nonspecifically adsorbed to the highly negatively charged dextran is likely to be denatured and can nonspecifically bind the target protein in solution. This process can easily use up all four flow channels of two chips.

Determine regeneration conditions 4. Insert new chip. 5. Activate the carboxylates of the dextran support by injecting N-ethyl-N′-[(dimethylamino) propyl] carbodiimide hydrochloride (EDC) in the presence of N-hydroxysuccinimide (NHS). These reagents are included in an amine-coupling kit from BIAcore.

6. Inject ligand protein in low-pH sodium acetate buffer such that an adequate amount of the protein is presented on the dextran. Perform several parallel experiments in which the density of ligand presented by the dextran is varied.


This can be accomplished by altering contact times and protein concentration. Proceed with steps 7 through 14, and then measure the rates of dissociation of the target protein from each of the ligand densities (step 15). The rate of dissociation should be constant and independent of the concentration of the surface density of the ligand (see Anticipated Results). The ligand density should be optimized to produce a surface presenting enough molecules to elicit a clear binding response yet dilute enough to measure a dissociation rate constant that is constant.



7. Cap any unreacted activated carboxylates by treatment with ethanolamine. 8. Inject a solution containing a known concentration of the putative target protein under buffer conditions and within the concentration range determined in step 3. Vary the flow rate and the concentration of the analyte to minimize diffusion or “mass transport” effects. The association rate constant for a particular reaction should be just that: constant (see Anticipated Results discussion on Rate and equilibrium constants).

9. Experimentally determine regeneration conditions, i.e., conditions under which the target protein is completely dissociated from the covalently coupled ligand. One must be careful to determine conditions for the regeneration of the surface that preserve the activity of the immobilized species. This procedure can easily consume another chip.

Perform experimental measurements 10. Insert new chip, if necessary. 11. Perform steps 2, 5, 6, 7, and 8 (i.e., activate the dextran carboxylates, covalently attach the protein of interest, cap unreacted sites, and introduce a putative binding partner in solution). Use the optimal buffer conditions and ligand density determined in steps 3 and 6, respectively.

12. Regenerate using reagents determined in step 9. 13. Repeat steps 8 and 12 as necessary to obtain data using a total of at least three different concentrations of the target protein. 14. Using BIA evaluation point-and-click software, measure dissociation and association rate constants. Calculate an equilibrium constant using average values of the association and dissociation rates that resulted from the three different target protein concentrations. The kinetic parameters measured for the various concentrations of analyte in solution should be within reasonable experimental error limits, similar to the error limits obtained from one “identical” experiment to another.

15. Reverse the experimental configuration: confirm that the same kinetic parameters are measured when the previous analyte is immobilized and the previous ligand is in solution. SPR USING NTA-SAM CHIPS The following protocol was developed for SPR experiments using nitrilotri-acetic acid self-assembled monolayer (NTA-SAM; Bamdad et al., 1994; Sigal et al., 1996) chips. This protocol is simpler than Basic Protocol 1, as NTA-SAM chips do not require several of the control measures necessitated by dextran-based chips (see Critical Parameters discussion on NTA-SAM chips). For detailed steps in the performance of an SPR experiment, and for additional reagents and equipment information, refer to the BIAcore user handbook. Materials NTA-SAM chips (3% to 5% NTA relative to an inert ethylene glycol–terminated thiol) PBS or HeBS (APPENDIX 2) 1 mM NaOH 1% (w/v) Ni(II)SO4

BASIC PROTOCOL 2



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Histidine-tagged ligand protein Target protein BIAcore SPR equipment BIA evaluation point-and-click software 1. Insert an NTA-SAM chip. 2. Use PBS or HeBS as the running buffer. 3. Inject 10 µl of 1 mM NaOH. 4. Inject 25 µl of aqueous 1% Ni(II)SO4. 5. Inject a histidine-tagged ligand protein. 6. Inject a putative target protein. 7. Repeat steps 3 through 6 with a total of at least three concentrations of the target protein. Histidine-tagged proteins can be removed from the surface by treatment with 200-mM imidazole. Alternatively, a fresh flow cell can be used for each experiment (each chip has four flow cells).

8. Using BIA evaluation point-and-click software, measure dissociation and association rate constants. Calculate an equilibrium constant using average values of the association and dissociation rates that resulted from the three different target protein concentrations. The kinetic parameters measured for the various concentrations of analyte in solution should be within reasonable experimental error limits, similar to the error limits obtained from one “identical” experiment to another.



Use of SPR to analyze biomolecular interactions Optical sensing devices for detecting and kinetically characterizing interactions among biomolecules are now in mainstream use. Complicated, multicomponent interactions can be quickly “dissected” by sequential determination of pair-wise interactions. The BIAcore SPR instrument can tolerate a wide range of buffer conditions. A broad range of affinities are measurable by SPR: from ten millimolar to picomolar. Because binding events are viewed in real time, one can alter experiments immediately in response to initial results. Researchers have eagerly adopted the new methods because they don’t require time-consuming and hazardous labeling steps. Perhaps the most important advantage is that because the mass of a molecular species is detected by an optical response, no protein labeling is required and only microliter sample volumes are consumed. The microfluidic technology of these devices satisfies a long-standing research need—allow-

ing for both minimal sample consumption and rapid measurements. The commercial availability of user-friendly optical sensors is a fairly new development. Until recently, the design of the devices has received little input from the bioscientific community. We are now entering a new and exciting phase in the development of this technology that is being driven by the needs of researchers to not only carry out routine experiments faster but to venture into unexplored areas. Derivatized monolayers will play an increasingly important role in state-of-the-art detection devices, as they have demonstrated clear advantages over disordered polymer-based supports (see Critical Parameters discussion on NTASAM chips). With optical sensing, it is now possible to design experiments to measure the difference between monovalent and bivalent binding, detect very weak interactions between proteins and protein complexes, and characterize interactions between factors that facilitate the assembly (or catalyze the disruption) of large megadalton complexes. The future evolution of these devices is only limited by the



imagination and needs of researchers in their desire to unravel the complex processes of biological systems. There is an intrinsic danger in using any instrument as a black box. The fact that many biologists, largely untrained in theoretical physical chemistry, use the instrument to perform kinetic characterization of interactions has led to considerable skepticism among researchers regarding much of the BIAcore-generated data in the literature. An understanding of the physics of the instrument is essential for experimental design and data interpretation. Thus, this commentary treats (1) the theory behind SPR devices, (2) factors that affect the accuracy of an SPR experiment, (3) choice of SPR chips, (4) caveats for experimentation and additional control experiments, and (5) kinetic analysis and higher-order kinetics. SPR theory Optical phenomena involving electron oscillations in the surface of a metal have been studied for some time. Electrons move freely through a metal. However, at a metal-medium interface, dipole excitations cause the propagation of electron waves called surface plasma or plasmon waves (SPWs). Because the wave vectors of the SPWs are long, they cannot be excited by simple incident light, so direct coupling of the two waves is not possible. X-rays were originally used to excite SPWs and for this reason interest in the phenomenon was limited to physicists studying how electromagnetic energy behaves at surfaces. A major breakthrough came in the late 1960s, when researchers succeeded in exciting SPWs using simple incident light. Otto (1968a,b) and the team of Kretschmann and Raether (1968), building on the earlier work of Turbadar (1959), exploited the phenomenon of total internal reflection (TIR) to couple incident light to SPWs. This achievement made simple SPR devices possible. The basic configuration of modern SPR instruments has not deviated significantly from the original experimental setup (see Fig. 20.4.1). Total internal reflection occurs at an interface, when light propagates from a medium of higher refractive index to one of lower refractive index at an angle of incidence greater than the critical angle. The light energy is then totally reflected off of the interface, except for an evanescent field that extends a distance of one wavelength (of the incident light) into the second medium. The magnitude and direction

of the reflected light can be monitored by an array detector. If a very thin layer of a third dielectric material, of lower refractive index than either of the others, is sandwiched between the two different dielectrics within the one wavelength distance, the incident wave will be transmitted through the thin layer and into the second medium by photon tunneling. If the sandwiched material is a metal and the wave vector of the incident light can somehow be increased (achieved by passing it through a prism) to match that of an SPW propagating through the metal, then coupling of the incident light and the SPW will occur. Energy from the incident light will be fed into the SPW and not reflected off the interface. This is important because the coupling of the two waves will produce a minimum or “dip” in the reflected light; the angle at which this occurs is a parameter exquisitely dependent upon the second medium’s dielectric properties (Wahling et al., 1979). Recall that a small portion of incident light extends into the second medium, so that the angle at which the incident wave couples to the SPW is determined by the refractive index of the second medium by Snell’s Law (assuming the first dielectric is held constant). Small changes in the refractive index of the second material cause large shifts in the angle at which coupling and the corresponding dip in reflected light occur. In the study of protein-protein interactions, increasing the protein concentration of a solution will result in a more dense medium with an altered refractive index. One can infer how the protein content of the second material is changing over time by monitoring the change in the angle of minimum light intensity. If ligand is immobilized at the interface of the thin metal film and the second medium, then subsequent interaction with proteins injected into the second medium will cause a change in the refractive index of the interfacial region, again changing the minimum reflectance angle. It is important to appreciate that the attenuation of the “sensing wave” which projects into the second medium is severe. This allows kinetic information to be extracted from the optical response as a function of time because the sensing wave only detects proteins that are very close to the interface (where the first species is immobilized), as opposed to the total mass of protein in the solution sample. It is assumed that proteins that are so close to the immobilized species are associating.



Supplement 40

should have equal access to the immobilized ligands.

Critical Parameters Factors that affect the accuracy of an SPR experiment The physics of the optical phenomenon of SPR govern the accuracy of the device. Because the sensing wave falls off very rapidly, the change in optical response is only a linear function of the net change in protein mass up to a maximum distance of 150 nm from the interfacial region (Stenberg et al., 1991).To reliably measure the kinetics of an interaction, the sensing apparatus should be as inert as possible. Additionally, immobilized ligands should be oriented to provide equal accessibility of binding sites to the binding partners in solution (analytes), and analytes in solution

Dextran-based chips Virtually all of the commercially available chips that are suitable for the study of proteinprotein interactions using a BIAcore device have a functionalized dextran base that is ∼150 nm thick. This presents a problem, as it means that the processes under study are occurring at the very limit of accurate detection. Several other problems arise with the use of dextranbased SPR chips. The most commonly used chip is a carboxylated dextran chip, CM-5, that is available from BIAcore. Proteins are nonspecifically coupled to the carboxylates of the support by standard EDC/NHS chemistry,

where n1 > n2 > n3

surface plasmon

Θ2 d ≈λ

metal

Θ1 Θ3

normal incident light


n2

n3 prism n1

array detector

Figure 20.4.1 The physical configuration of modern SPR devices. Light passing through a dense medium (n1) incident at an angle (Θ1) greater than the critical angle, to a second medium of lower refractive index (n2) is totally reflected off the interface, except for an evanescent field that extends into the second medium for a distance of one wavelength. At a certain angle (Θ2) exquisitely dependent on the refractive index of the second medium, the incident light wave can couple to surface plasmon waves (electron oscillations) propagating through a thin metal film (n3) sandwiched between the two media. The incident light energy is fed into the surface plasmon wave and a corresponding minimum in reflected light is observed. Proteins immobilized at (or recruited to) the metal-medium interface alter the dielectric of the region, which causes a shift in the angle of minimum reflectivity (Θ3).



which targets primary amines on the protein surface. The carboxylated dextran carries a high net negative charge (Johnsson et al., 1991); it is thus not inert and may alter the kinetics of the system under study. The dextran support is also problematic with respect to mutual accessibility of ligand and target proteins. Experiments indicate that very large proteins may be completely prevented from entering the dextran matrix (see Critical Parameters discussion on NTA-SAM chips). The nonspecific coupling of proteins to the support renders the ligand’s active site randomly oriented and often inaccessible. Furthermore, flow over the nonuniform dextran polymer is turbulent. A large static layer separates the analyte-containing solution from the ligand-presenting dextran and hinders the diffusion of molecules into the matrix. Increasing the flow rate will help speed diffusion, but if the diffusion rate is slower than the association rate, then the diffusion rate will be the measured parameter. Although analytical methods have been proposed to model the diffusion or “mass transport” factor (Fisher and Fivash, 1994), it would be preferable to eliminate the problem rather than to model its effect into data evaluation software. NTA-SAM chips In 1994, a self-assembled monolayer (SAM; Nuzzo et al., 1987; Bain et al., 1989) SPR chip was developed that presents nitrilotri-acetic acid (NTA) ligands for the specific capture of histidine-tagged proteins to overcome the problems intrinsic to the dextran chip (Bamdad et al., 1994; Sigal et al., 1996). The thickness of the monolayer is 2 nm, as opposed to the 150nm thickness of the dextran chip. It has been possible to detect and characterize interactions on the NTA-SAM that were undetectable on the dextran chip because (1) processes under study are much closer to the emanation of the sensing wave, (2) the active sites are uniformly presented to analytes in solution, (3) noncovalent coupling to the support retains greater protein viability, and (4) flow over the monolayer is near laminar with an insignificant static layer. The first demonstration of the enhanced sensitivity of the NTA-SAM chip involved the interaction between a mutant form of a yeast holoenzyme component, Gal11p, and an inert DNA-binding domain, Gal4(58 to 100). Although there was considerable genetic evidence that the two proteins were associated, direct detection of the interaction by standard techniques was not demonstrable for a number of

years. The interaction between these two proteins could not be detected by SPR on dextranbased chips, but was easily detected and quantitated using the NTA-SAM chip (Barberis et al., 1995). Subsequently, a fusion protein of Gal11p, which could be prepared at much higher concentrations, was observed to interact with Gal4(58 to 100) using both dextran SPR chips and gel mobility shift assays (Farrell et al., 1996). It is reassuring that the kd measured on the NTA chip, where protein concentration was low, was virtually the same as that measured on the dextran chip using the fusion protein at a much higher concentration. The second demonstration of the enhanced sensing capabilities of the NTA-SAM chip was in detecting the interaction of yeast polymerase II with TFIIF (Bushnell et al., 1996), which had not been detectable by SPR with standard dextran chips. A third example of the superiority of monolayer-based SPR chips was the detection of the interaction between TFIIE and polymerase II using the NTA-SAM (Bushnell et al., 1996). The interaction could not be detected using standard dextran chips, presumably because of the size of the protein complexes and related diffusion problems. To measure the sensitivity of this method, side-by-side experiments were performed comparing dextran chips to NTA-SAMs; a chimeric T-cell receptor (TCR) was probed with two monoclonal antibodies (Sigal et al., 1996). One antibody recognized a particular conformation of the protein; the other recognized a linear epitope. Although six-times more protein was attached to the dextran surface (because of the increased surface area) than to the two-dimensional NTA array, only an effective monolayer of protein on the dextran was accessible to incoming ligand; thus, the total amount of antibody that bound to each surface was identical. The ratio of the conformationally sensitive antibody to the linear epitope–antibody that bound to each TCR-derivatized surface showed that five-times more protein remained in the active conformation when it was attached to the NTA-SAM than when attached to dextran. In addition to increased sensitivity, NTASAMs offer increased experimental simplicity. Because there are a finite and approximately constant number of NTA ligands on every NTA-SAM, the determination of optimal immobilization conditions (e.g., protein pH or concentration) or regeneration conditions is not necessary. For example, maximal NTA occupancy on 5% NTA-SAMs is virtually constant



Supplement 40

and is easily achieved by injecting 25 µl of a 25-kD protein at a concentration of 3 mg/ml; higher injection concentrations will not alter the amount of protein presented. At 3% to 5% NTA, rebinding events are minimal so there is no need to experimentally determine the ligand concentration that will minimize rebinding in order to reflect a true off-rate. It is not necessary to check for artifacts stemming from an attraction to the chip surface by performing reciprocal experiments (reverse configuration) since the NTA-SAM has been shown to be highly resistant to nonspecific binding of all proteins tested including TBP. An NTA SPR chip for the attachment of histidine-tagged proteins is now commercially available from BIAcore, but importantly the NTA ligands are attached to dextran (Gershon and Khilko, 1995), not incorporated into a monolayer. Noncovalent coupling of the protein of interest to the dextran support is an advantage because the chemical treatment used to promote covalent coupling denatures a significant portion of the protein. However, the chip still suffers from other problems intrinsic to dextran: high background charge, random orientation of the active site relative to analyte flow, and turbulent flow dynamics with a large static layer. Moreover, this chip may introduce a new experimental artifact, which was encountered when this approach was experimented with in 1994 (Bamdad and colleagues; unpub. observ.). The NTA group chelates a Ni(II) atom in a very-high-affinity interaction. However, the disordered nature of the dextran may position two NTA groups such that two ligands simultaneously coordinate one Ni(II) atom, which is much more unstable: subtle changes in pH and protein addition could trigger rearrangements that would act to crosslink the dextran and alter its optical response, thus mimicking protein binding.


Expanded applications with NTA-SAMs It is likely that monolayer chips will find wide use. This is because biological systems often consist of networks of proteins that interact with each other. For example, it now appears that the active transcription complex may consist of some thirty proteins (Kim et al., 1994; Koleske and Young, 1994). Future research may be increasingly focused on the study of proteins interacting with complexes of proteins.The nonuniform surface of dextran can interfere with the entry of large protein complexes into the matrix, whereas monolayers provide near laminar flow dynamics that allow

for the characterization of interactions between large protein complexes. On dextran, the stoichiometry of interactions between a protein in solution and an evolving protein complex assembling on the chip may not be possible, as the interstices of dextran would render variable amounts of the complex accessible at each stage. Another problem of great interest to biologists is the study of interactions where one or more of the partners is a membrane-associated protein. Plant et al. (1995) reported the development of a lipid bilayer chip for use in SPR instruments. Hydrophobic forces caused the spontaneous attachment of a phospholipid layer to a SAM comprised of hydrophobic alkyl thiolates. The work demonstrated the feasibility of using this approach to study interactions between components that are not water soluble: cell-surface receptor-ligand binding, proteinmembrane interactions, and cell-cell associations. Unfortunately, this chip has not yet been made commercially available. Many protein-protein interactions take place when one or both proteins are bound to DNA. The only method commercially available for the study of these interactions by SPR derivatizes dextran with streptavidin and then attaches it to biotinylated DNA. DNA-binding proteins can then be bound to the presented oligonucleotides for subsequent interaction studies with analytes in solution. These studies are subject to artifacts intrinsic to the dextranbased immobilization methods. A DNA-SAM was recently developed (Bamdad, 1997a) that presents single-stranded DNA above a background of inert alkane thiolates. Doublestranded DNA was hybridized to this surface via a complimentary single stranded “tail”. The method was specific for the desired DNA, accepted unmodified bacterially produced DNA, and can easily be extended to present mass arrays of heterogeneous oligonucleotides. The sense strand of the hybridized double-stranded DNA was attached to the surface by an enzyme reaction leaving the antisense strand dissociable by heat treatment. Endonuclease digestion of hybridized DNA would generate a universal acceptor surface for ligation of sample DNA cut with the same enzyme. Caveats and additional control experiments Obtaining reliable results from SPR experiments requires careful attention to experimental design. To optimize the veracity of SPR-obtained results, experimental techniques must be designed to minimize systematic artifacts. The



BIAcore handbook is instructive for designing a preliminary set of SPR experiments, but the following additional experiments are well advised. Once a set of experiments has been analyzed and kinetic rate constants obtained, the user should verify that the calculated rates are approximately correct using some simple experiments. For example, analyte injected at a concentration equal to the BIAcore-calculated kd should give rise to approximately half-maximal binding, in RUs. Similarly, if the protein in solution (at a concentration capable of eliciting maximal binding) is incubated (competitive inhibition) with ligand at a concentration equal to the calculated kd, and is then injected over immobilized ligand, approximately half-maximal binding should result. The kinetics of an interaction should be independent of the experimental configuration. To identify artifacts in the apparent kinetics of an interaction, a reciprocal experiment should be performed in which the protein in solution (the analyte) is immobilized as the ligand, and the previously immobilized protein becomes the analyte in solution. Performing the experiment in reverse configuration should result in comparable rate measurements. On dextran chips, discrepancies in reciprocal rates may be an indication of an interaction in which diffusion into the dextran, rather than association, is the rate-limiting step. Since diffusion into the dextran matrix is a function of the protein’s size and charge, use the rate that is obtained when the smaller or less-charged of the two proteins is used as the analyte. Verification must be done using another method. If a higher-order interaction is suspected, additional experiments should be designed to directly prove or disprove higher-order binding models, such as dimerization dependence (see Anticipated Results discussion on Kinetics analysis and higher-order kinetics). It is also useful to repeat binding experiments using genetic variants of one or both of the interacting proteins, particularly variants whose in vivo activities are known. Reducedactivity mutants often display differential affinities for functionally relevant target molecules. A panel of mutants possessing a range of in vivo activities would provide for excellent control experiments. Recall that SPR only detects the mass of a species near the interfacial region of the instrument, so the nonspecific binding of a few protein aggregates cannot be distinguished from multiple 1:1 binding events. One must there-

fore be rigorous about protein homogeneity. HPLC purification or centrifugation just prior to the experiment is recommended to rid the preparations of aggregates. Inaccuracies in measured binding constants can arise from discrepancies in the assessment of the active concentration of protein components. If the active concentration of one of the binding partners can be adequately assessed, that protein should be used as the analyte, since only the concentration of the analyte enters into the BIAcore kinetic equations. This minimizes problems associated with protein preparation heterogeneity.

Anticipated Results Rate and equilibrium constants The measured rate of dissociation should be a constant and independent of the concentration of either the surface density of the ligand or the analyte in solution. A dissociation rate that decreases as the ligand density increases indicates that dissociating molecules are rebinding to nearby sites. The ligand density should be optimized to produce a surface presenting enough molecules to elicit a clear binding response yet dilute enough to measure a dissociation rate constant that is constant. When extracting a dissociation rate from sensorgrams, choose time periods early in the dissociation phase to help eliminate rebinding events. The measured rate of association for a particular reaction should also be constant. However, if the rate of diffusion into the disordered dextran polymer is slower than the rate of association with the immobilized ligand, then the measurable parameter will be the diffusion rate, not the association rate. Because the diffusion rate is a function of the size of the protein in solution, the flow rate, and the size of the surface irregularities, it may not be possible to measure the true association rate between large analyte proteins (or protein complexes) and ligands immobilized on dextran. The accuracy of SPR measurements critically hinges on the stringency of the experimental design. For a ball-park estimate of the range of the equilibrium constant, determine the analyte concentrations that give rise to both maximal and minimal binding to a ligand surface. To a first approximation, the equilibrium constant should be the mean of the two concentrations. If an absolute kinetic rate is desired (i.e, one that is comparable to a solution measurement rather than a rank ordering of affinities), obtain an IC50 by performing a series of competitive inhibition experiments.



Supplement 40

Response (RUs)

Kinetics analysis and higher-order kinetics The BIAcore instrument comes with userfriendly kinetic evaluation software (modeled after the analysis described in O’Shannessy et al., 1993) that is now adequate to address simple first-order binding reactions. However, it may not be accurate for higher-order reactions. Others have developed more rigorous software for the kinetic analysis of BIAcore-derived data that uses numerical methods analysis and global fit programs (Fisher et al., 1994; Myszka, 1997). Fisher and Fivash (1994) have done considerable mathematical modeling of higher-order reactions that are then compared to experimental binding curves using a Cray supercomputer and their global fit analysis software. Less rigorous inferences of higher-order reactions based on curve-matching analysis of single experiments is risky. Additionally, overanalyzing data is less reliable than performing experiments designed to test a specific higherorder binding model. Experimental confirmation of a theorized higher-order binding model was recently accomplished using variable-density NTA-SAMs

21000 20200 19400 18600 17800 17000 16200 15400 14600 13800 13000

with SPR (Bamdad, 1997b). The concentration of NTA-terminated thiol was varied from 3% to 11% relative to an inert alkane thiol (the major component), producing a panel of SPR chips capable of presenting His-tagged peptides in variable-density arrays. It had previously been shown that four or more tandem repeats of an eight–amino acid motif could stimulate transcription in vivo and bind to the transcription factor TATA box–binding protein (TBP) in vitro (Tanaka, 1996). His-tagged peptides comprised of only two tandem repeats of the same motif were separately immobilized on NTA-SAMs of various density. BIAcore experiments showed that the peptides immobilized on low-density monolayers (3.8% NTA) were not able to bind TBP from solution. However, when the average distance between the peptides was decreased by immobilization on higher-density monolayers (5.7% NTA), a high-affinity interaction (20 nM) resulted (see Fig. 20.4.2). The complete study indicated that this observation resulted from the significant kinetic difference between monovalent and bivalent binding of the target protein, TBP. Com-

∆ 550 RUs

∆ 2080 RUs His-tagged peptide

hTBPc

∆ 5 RUs

∆ 1186 RUs 984

1184

1384

1584

1784

1984

2184

Time (sec)


Figure 20.4.2 BIAcore sensorgram showing that binding of hTBPc in solution to peptide motif surfaces is dependent upon density of ligand immobilization. The BIAcore SPR instrument plots changes in the angle of minimum reflectance in resonance units (RUs) as a function of time. The “square waves” represent injections of protein “plugs” that interrupt the constant buffer flow. The net change in measured RUs following a protein plug injection (arrows) indicates binding and can be correlated to a mass of protein recruited to the surface. An association constant can be derived from analysis of the initial phase of the injection, and a dissociation rate can be extracted from analysis of the system as it returns to buffer flow. Histidine-tagged fusion proteins terminated with the desired recognition motif were separately immobilized on NTA-SAMs presenting 3.8% (dashed line) or 5.7% NTA (solid line). hTBPc in solution was injected over the surfaces in two separate experiments. An overlay of the two resultant SPR sensorgrams shows that hTBPc cannot bind (∆5 RUs; arrowhead) to the repeats when they are immobilized at low density (3.8% NTA; average distance = 29 Å), but bind very tightly (∆550 RUs; arrowhead) when the peptides are positioned closer together at a higher density (5.7% NTA; average distance = 23 Å).



petitive inhibition experiments involving peptides either two or four repeats long demonstrated that there was a 250-fold difference in affinity between the two binding modes. The average distance between peptides on the chip surface, which presumably represents the distance between binding sites on TBP, was extrapolated from a statistical analysis of the lowest peptide density that elicited a large increase in target affinity. This analysis enabled the discrimination between three possible mechanistic models describing how reiterated repeats could synergistically effect the general transcription factor. Despite the advantages presented by the ability to perform SPR experiments on a monolayer surface, it is still possible that experiments on these surfaces reflect interactions that differ significantly from those in solution due to local concentration effects and/or differences between two- and three-dimensional diffusion. A solution measurement, the IC50, can be generated from a series of competitive inhibition experiments where the analyte is preincubated with varying concentrations of the ligand prior to injection of the mix over ligand-immobilized surfaces. In the above experiments, such an analysis revealed that the surface interaction was 20 times tighter than the comparable solution interaction.

Time Considerations The major time factor in performing SPR experiments is in working out the appropriate conditions and experimental design, which can take weeks or even months. A single binding assay can be carried out in 5 to 10 min per injection. Using NTA-SAM chips, a set of experiments using three protein concentrations can be performed in 2 to 3 hr. However, to perform the same experiments with a dextran chip, it is likely to take several days to work out parameters such as the appropriate binding conditions and regeneration conditions.

Literature Cited Bain, C.D., Evall, J., and Whitesides, G.M. 1989. Formation of monolayers by the coadsorption of thiols on gold: Variation in the head group, tail group and solvent. J. Am. Chem. Soc. 111:71557164. Bamdad, C. 1997a. Submitted for publication. Bamdad, C. 1997b. Submitted for publication. Bamdad, C., Sigal, G., Strominger, J., and Whitesides, G. 1994. A mixed monolayer for the immobilization of histidine-tagged proteins. Appendage: A self-assembled monolayer for the

presentation of oligonucleotides. U.S. patent application PCL01016-94. Barberis, A., Pearlberg, J., Simkovich, N., Farrell, S., Reinagel, P., Bamdad, C., Sigal, G., and Ptashne, M. 1995. Contact with a component of the polymerase II holoenzyme suffices for gene activation. Cell 81:359-368. Bartel, P.L. and Fields, S. 1995. Analyzing proteinprotein interactions using a two-hybrid system. Methods Enzymol. 254:241-263. Bushnell, D.A, Bamdad, C., and Kornberg, R. 1996. A minimal set of RNA poymerase II transcription protein interactions. J. Biol. Chem. 271:20170-20174. Daniels, P.B., Deacon, J.K., Eddowes, M.J., and Pedley, D.G. 1988. Surface plasmon resonance applied to immunosensing. Sens. Actuators 16:11-18. Farrell, S., Simkovich, N., Wu, Y., Barberis, A., and Ptashne, M. 1996. Gene activation by recruitment of the RNA polymerase II holoenzyme. Genes & Dev. 10:2359-2367. Fisher, R.J. and Fivash, M. 1994. Surface plasmon resonance based methods for measuring the kinetics and binding affinities of biomolecular interactions. Curr. Opin. Biotechnol. 5:389-395. Fisher, R.J., Fivash, M., Casas-Finet, J., Bladen, S., and Larson McNitt, K. 1994. Real-time BIAcore measurements of Escherichia coli singlestranded DNA binding (SSB) protein to polydeoxythymidylic acid reveal single-state kinetics with steric cooperativity. Methods 6:121-133. Gershon, P.D. and Khilko, S. 1995. Stable chelating linkage for reversible immobilization of oligohistidine tagged proteins in the BIAcore surface plasmon resonance detector. J. Immunol. Methods 183:65-76. Gyuris, J., Golemis, E., Chertkov, H., and Brent, R. 1993. Cdil, a human G1 and S-phase protein phosphotase that associates with Cdk2. Cell 75:791-803. Johnsson, B., Löfås, S., and Lindquist, G. 1991. I mm obiliza tion of proteins to a carboxymethyldextran-modified gold surface for biospecific interaction analysis in surface plasmo n resonance sensors. Anal. Biochem. 198:268-277. Kim, Y.J., Bjorklund, S., Li, Y., Sayre, M.H., and Kornberg, R.D. 1994. A multiprotein mediator of transcriptional activation and its interaction with the C-terminal repeat domain of RNA polymerase II. Cell 77:599-608. Koleske, A. and Young, R.A. 1994. An RNA polymerase II holoenzyme responsive to activators. Nature 368:466-469. Kretschmann, E. and Raether, H. 1968. Z. Naturf. 230:2135. Liedberg, B., Nylander, C., and Lundström, I. 1983. Surface plasmon resonance for gas detection and biosensing. Sens. Actuators 4:299-304.



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Löfås, S. and Johnsson, B. 1990. A novel hydrogel matrix on gold surfaces in surface plasmon resonance sensors for fast and efficient covalent immobilization of ligands. J. Chem. Soc. Chem. Commun. 1526-1528.

monolayer for the binding and study of histidinetagged proteins by surface plasmon resonance. Anal. Chem. 68:490-497.

Myszka, D. 1997. Kinetic analysis of macromolecular interactions using surface plasmon resonance biosensors. Curr. Opin. Biotechnol. 8:50-57.

Stenberg, E., Persson, B., and Roos, H. 1991. Quantitative determination of surface concentration of protein with surface plasmon resonance using radiolabeled proteins. J. Colloid Interface Sci. 143:513-526.

Nuzzo, R.G., Fusco, F.A., and Allara, D.L. 1987. Spontaneously organized molecular assemblies. 3. Preparation and properties of solution adsorbed monolayers of organic disulfides on gold surfaces. J. Am. Chem. Soc. 109:2358-2368.

Tanaka, M. 1996. Modulation of promoter occupancy by cooperative DNA binding and activation-function is a major determinant of transcriptional regulation by activators in vivo. Proc. Natl. Acad. Sci. U.S.A. 93:4311-4315.

O’Shannessy, D., Brigham-Burke, M., Soneson, K., Hensley, P., and Brooks, I. 1993. Determination of rate and equilibrium binding constants for macromolecular interactions using surface plasmon resonance: Use of nonlinear least squares analysis methods. Anal. Biochem. 212:457-468.

Wahling, G., Raether, H., and Mobius, D. 1979. Studies of organic monolayers on thin silver films using the attenuated total reflection method. Thin Solid Films 58: 391-395.

Otto, A. 1968a. Z. Phys. 216:398. Otto, A. 1968b. Phys. Stat. Solidi. 26:199. Plant, A., Brigham-Burke, M., Petrella, E., and O’Shannessy, D. 1995. Phospholipid/alkanethiol bilayers for cell-surface receptor studies by surface plasmon resonance. Anal. Biochem. 226:342-348. Sigal, G.B., Bamdad, C., Barberis, A., Strominger, J., and Whitesides, G.M. 1996. A self-assembled

Turbadar, T. 1959. Proc. Phys. Soc. London 73:40.

Zervos, A., Gyuris, J., and Brent, R. 1993. Mxi1, a protein that specifically interacts with Max to bind Myc-Max recognition sites. Cell 72:223232.

Contributed by Cynthia Bamdad Clinical Micro Sensors Pasadena, CA




Detection of Protein-Protein Interactions by Coprecipitation

UNIT 20.5

Coprecipitation of proteins from whole-cell extracts is a valuable approach to test for physical interactions between proteins of interest. When a precipitating antibody is used, this method is referred to as co-immunoprecipitation. Coprecipitation can be used to study interactions between known proteins under a variety of conditions and as a means of identifying components of a complex. Coprecipitation may be the single method of choice, or may be used in combination with other methods that detect protein-protein interactions, such as two-hybrid analysis and copurification schemes (UNIT 20.1), and tests of physical associations using purified proteins. This unit describes basic approaches to immunoprecipitating tagged proteins from whole-cell extracts. The approaches described can be adapted for other systems. In a typical experiment, as described here, cells are lysed and a whole-cell extract is prepared under nondenaturing conditions (see Strategic Planning). The protein is precipitated from the lysate with a solid-phase affinity matrix and the precipitate is tested for the presence of a second specifically associated protein (see Basic Protocol and Alternate Protocol). The approach can be used for native or epitope-tagged proteins for which antibodies are available, or for recombinant proteins that have been engineered to bind with high affinity to a molecule that can be coupled to a solid-phase matrix (see Strategic Planning). The presence of an associated protein is detected by separating the precipitated proteins by SDS-PAGE (UNIT 10.2) and immunoblotting (UNIT 10.8) with a second antibody that recognizes the putative associated protein. Controls to test specificity of interaction are crucial (see Strategic Planning). For additional background reading, the user should consult UNIT 10.16 for a theoretical discussion of immunoprecipitation; see Chapter 11 for principles of antibody production and immunoassays; see UNITS 10.15, 20.2 & 20.3 for approaches to tagging proteins; and see UNITS 13.7-13.10 for transformation and propogation of S. cerevisiae. For an in-depth review of immunoprecipitation techniqes, see chapter 11 of Harlow and Lane (1988). For an in-depth review of coprecipitation and other approaches to detect protein-protein interactions, see Phizicky and Fields (1995). STRATEGIC PLANNING Detecting the Proteins in Question The first step is to generate reagents that detect the two proteins in the coprecipitate under nondenaturing conditions. If antibodies are available that can immunoprecipitate the proteins under nondenaturing conditions, then they can be used. Alternatively, the proteins can be differentially tagged in a variety of ways to allow their detection with commercially available antibodies or other affinity reagents. The tagged proteins are then introduced into the host organism using expression vectors. All tagged proteins must be assessed for function in vivo. A frequently used option is to add a short peptide or epitope that is recognized by a commercially available high-affinity monoclonal antibody (mAb; UNIT 10.15). The epitope is typically added at the amino or carboxyl terminus, although internal positions that do not disrupt function can also be used. Two frequently used epitopes are derived from influenza hemagglutinin protein (HA) and human c-Myc and are recognized by highaffinity mAbs (12CA5 and 9E10, respectively; Kolodziej and Young, 1991). Others such Contributed by Elaine A. Elion Current Protocols in Molecular Biology (1999) 20.5.1-20.5.9 Copyright © 1999 by John Wiley & Sons, Inc.



as FLAG are also available (UNIT 10.15; BioSupplyNet Source Book, 1999). The choice of the epitope may be dictated by its amino acid composition. It is often useful to insert tandem copies of the epitope in order to increase sensitivity. The number of additional tandem copies can range widely from one (Field et al., 1988) to several (e.g., three; Tyers et al., 1993) to many (e.g., nine; Feng et al.,1998). Proteins can also be fused to small proteins or peptides that have high affinity to small molecules that can be attached to a solid support. This is a particularly valuable approach when the protein to be precipitated comigrates with immunoglobulin heavy or light chains in an SDS-polyacrylamide gel. Such alternative tagging methods include fusion to glutathione-S-transferase (to allow purification by a glutathione affinity matrix; UNIT 16.7) or maltose-binding protein (to allow purification by a maltose affinity matrix; UNIT 16.6). An excellent reference for identifying sources of commercially available antibodies and approaches to tagging proteins can be found in the BioSupplyNet Source Book (1999). See Chapter 11 and Harlow and Lane (1988) for the generation and purification of specific antibodies. See Chapter 16 for a discussion of tagging and expressing proteins. Preparing Whole-Cell Extracts The second step to a successful coprecipitation is generating whole-cell extracts that optimize the yield and activity of the proteins to be analyzed, using lysis buffer conditions that permit recognition of the proteins by the affinity matrix. The yield of total protein in a whole-cell extract is not always a reliable indicator of the relative yield and activity of specific proteins, so it is wise to verify both parameters at the onset of an experiment before proceeding with the coprecipitation. Yield and activity can be affected by a number of factors (see Chapter 10; see Harlow and Lane, 1988). Small variations in the relative amounts of salt and detergents in the lysis buffer can have large effects on yield and activity, as can the speed and efficiency of cell breakage. Both factors are particularly important for less soluble proteins that associate with macromolecular structures such as membranes or cytoskeleton. In addition, global inhibition of proteolysis through the inclusion of multiple classes of protease inhibitors may be essential. Methods for preparing whole-cell extracts from yeast (UNIT 13.13), E. coli (UNITS 16.1-16.8), insect cells (UNIT 16.11), and mammalian cells (UNITS 16.12-16.18) can be found elsewhere in this manual, and specifics will not be discussed here. In general, the lysis buffer conditions are not very different from the coprecipitation conditions. It is recommended that the investigator begin by comparing small-scale extract preparations that vary the amount of salt and nonionic detergent. As a starting point, a basic lysis buffer might contain the following components. Basic components. Basic components include a buffering agent (such as 50 mM Tris⋅Cl, pH 7.5), a small amount of nonionic detergent (such as 0.1% [v/v] Triton X-100), salt (such as 100 mM NaCl), a reducing agent (such as 1 mM DTT), and 10% (v/v) glycerol as stabilizer. Protease inhibitors. Protease inhibitor cocktails are described in UNIT 13.13 and are also commercially available. A reasonable starting point would be to include 5 µg/ml each chymostatin, pepstatin A, leupeptin, and antipain, as well as 1 mM phenylmethysulfonyl fluoride and 1 mM benzamidine.


Chelating agents. EGTA (∼15 mM) is commonly included to chelate divalent metal ions that are essential for metalloproteases. Because EGTA also inhibits other metal-dependent enzymes, it may be omitted, combined with the addition of a needed metal ion, and/or substituted with EDTA.



Phosphatase inhibitors. If the phosphorylation state of the proteins in question is important, a mixture of phosphatase inhibitors should also be included in the lysis buffer. A starting mixture could contain 2.5 mM each meta- and ortho-vanadate, 10 mM NaF, and 10 mM β-glycerol phosphate. Simple modifications of this initial buffer include varying the amount of NaCl (from 0 to 500 mM) and of Triton X-100 (from 0% to 1%). The investigator may choose to compare different means of breaking the cells (for example, glass-bead breakage versus liquid nitrogen/grinding methods for yeast cells; UNIT 13.13).

1. Generate antibodies to protein 1 and protein 2; or differentially tag them ( ) and introduce genes encoding tagged proteins into host cell

protein 1

protein 2

2. Prepare whole-cell extract

3. Incubate extract with antibody to protein 1 4. Incubate extract with protein A–Sepharose, which binds antibody

5. Collect Sepharose beads by centrifugation

discard

P2 P1

supernatant

pellet

P2-NT P2 P1 P1-NT

protein 2 protein 1 Ig h Ig I

6. Wash pellet several times to remove proteins not bound to protein A–Sepharose. 7. Dissociate proteins from protein A– Sepharose. Separate proteins by SDS - PAGE. Immunoblot with antibody to protein 2 (P2). Reprobe with antibody to protein 1 (P1).

Figure 20.5.1 Flowchart for the coprecipitation of two proteins that have been differentially tagged and introduced into the host organism. Ig h and Ig l, immunoglobulin heavy and light chains; NT, no tag.



Supplement 46

Total protein concentration in the whole-cell extract is generally assayed by using the Bio-Rad protein assay and calculating protein concentration (UNIT 10.1A). Extracts should be tested for the amount of each specific protein by immunoblot analysis (UNIT 10.8), analyzing 25 to 75 µg of total protein. In general, it is best to test for the presence of a second established protein (such as a housekeeping enzyme, cytoskeletal or ribosomal protein, or a previously defined component in the pathway being studied) as an internal control for normalization and as a positive control for the immunoblot. The amount of specific protein in the whole-cell extract is compared to the amount that is recovered by precipitation with an affinity matrix. Control Tests for Specificity of Interaction Controls are essential to verify that the antibodies and protein-protein interactions are specific. Proper controls are simplest to set up when the proteins are differentially tagged. In this instance, two parallel extracts are prepared from strains that contain each protein lacking the tag in the presence of the second tagged protein. An example is shown in the idealized gel in Figure 20.5.1, which includes lanes containing untagged protein 1 + tagged protein 2 and tagged protein 1 + untagged protein 2. If the antibodies are specific, untagged protein 1 will not immunoprecipitate. The presence of untagged protein 1 in the immunoprecipitate will indicate that it binds the affinity matrix nonspecifically. If the interaction between proteins 1 and 2 is specific, then tagged protein 2 will be present in the immunoprecipitate of tagged protein 1, but not in its absence. If antibodies to native proteins are used, it is necessary to compare extracts made from strains harboring deletions of the proteins in question to test for the specificity of the antibody and the interaction. However, this is obviously possible only if the deletions do not cause inviability. If deletion mutations are not possible, a commonly used approach is to show that the preimmune serum or an antibody not known to be specific to either of the proteins in question does not coprecipitate them in a parallel experiment. However, the latter two controls do not rule out the possibility that the antibody is precipitating the protein in question through an indirect association. It is also essential to compare the amount of coprecipitated protein with the amounts of the two proteins in question in the whole-cell extract. This allows one to determine whether apparent differences in the ability of the two proteins to coprecipitate is a secondary consequence of the relative abundance of the proteins. This control is particularly important when an interaction has been established and the investigator wishes to search for regulatory changes in association apart from changes in abundance. BASIC PROTOCOL


COPRECIPITATING PROTEINS WITH PROTEIN A/G–SEPHAROSE Once the conditions of extract preparation have been established (see Strategic Planning), the next step is to test for coprecipitation of the specific proteins. This protocol describes a standard coprecipitation procedure that uses an antibody coupled to protein A– Sepharose or protein G–Sepharose. An alternative coprecipitation method that uses GST coupled to glutathione-agarose is also provided (see Alternate Protocol). It is essential to keep all buffers and tubes cold by using an ice bath and a refrigerated centrifuge. The conditions of coprecipitation match the conditions of the lysis buffer described above. Materials Whole-cell extract (see Strategic Planning) Antibody Co-immunoprecipitation buffer (see recipe) 5 M NaCl Protein A/G–Sepharose slurry (see recipe)



2× sample buffer for SDS-PAGE (UNIT 10.2) 20-ml syringe and 18-G needle Hamilton syringe Additional reagents and equipment for SDS-PAGE (UNIT 10.2) and immunoblotting (UNIT 10.8) 1. Prepare duplicate samples in microcentrifuge tubes on ice: 0.5 to 1 mg whole-cell extract 1 µg antibody 5 M NaCl to equalize at 100 mM NaCl Co-immunoprecipitation buffer to 0.5 ml final volume. Adjust buffer by adding a divalent cation if necessary for activity of the protein in question.

2. Invert tube gently several times and incubate on ice for 90 min with occasional tube inversion. It is recommended that the investigator begin with a 90-min incubation. However, this incubation step can be shortened or lengthened.

3. Microcentrifuge 10 min at maximum speed, 4°C, to pellet nonspecific aggregates. Transfer supernatant to a new microcentrifuge tube. 4. Add 50 µl of protein A– or protein G–Sepharose slurry (25 to 30 µl bead volume). Be sure to evenly suspend the slurry before distributing it to the samples. Protein A has been used more frequently for historical reasons; however, protein G binds a broader range of Ig subtypes at higher efficiency. See UNIT 11.11 for a description of their relative binding capacities.

5. Rotate tube gently at 4°C for 30 to 60 min. Rocking is much less efficient and should be avoided.

6. Gently pellet protein A/G–Sepharose by centrifuging 30 sec at 1000 rpm in a tabletop centrifuge, 4°C. 7. Wash pellet three times with 1 ml co-immunoprecipitation buffer. For each wash, gently invert tube three times before pelleting. After each pelleting, use a 20-ml syringe with an 18-G needle to aspirate and remove supernatant. It may be possible to omit the costly protease inhibitors from the buffer at this stage, but this has not been attempted to date.

8. Aspirate as much liquid as possible from the final without touching the beads and add 25 µl of 2× sample buffer. If desired, samples containing sample buffer can be frozen up to several months at −80°C prior to SDS-PAGE. In this case the buffer should be prepared with sterile stock solutions and made with 1 mM sodium azide included.

9. Prepare for SDS-PAGE analysis by boiling for 5 min, vortexing, and microcentrifuging briefly to pellet beads. Use a Hamilton syringe to load eluates onto an SDS-polyacrylamide gel, arranging duplicate samples to allow preparation of duplicate blots. Separate by electrophoresis (UNIT 10.2). A Hamilton syringe works well to remove the eluate from the beads during loading.

10. Immunoblot duplicate samples separately with antibodies for each of the two proteins (UNIT 10.8). Be sure to include aliquots of the whole-cell extract for comparison and as a positive control for the immunoblot. Each immunoblot can be reprobed with the antibody to the other protein.



Supplement 46

ALTERNATE PROTOCOL

COPRECIPITATING A GST FUSION PROTEIN GST fusion proteins may be coprecipitated by following the co-immunoprecipitation procedure (see Basic Protocol) with the modifications outlined below. This procedure might be used when the protein in question comigrates with immunoglobulin heavy or light chain in an SDS-PAGE gel or if the antibodies being used precipitate too many cross-reacting proteins, such as the protein being tested for association. Furthermore, it is possible to dissociate the purified GST fusion from the solid-state glutathione resin under gentle conditions through the use of imidazole. The approach is also useful in that it will increase the size of the protein sufficiently that this size increase can be used as a diagnostic feature in analyzing complexes. Additional Materials (also see Basic Protocol) Glutathione-agarose or glutathione-Sepharose slurry (see recipe) 1. Prepare duplicate samples as described for protein A/G–Sepharose (see Basic Protocol, step 1), omitting antibody. 2. Microcentrifuge 10 min at maximum speed, 4°C, to pellet nonspecific aggregates. Transfer supernatant to new microcentrifuge tube. 3. Add 30 µl glutathione-agarose or glutathione-Sepharose slurry (25- to 30-µl bead volume). Be sure to evenly suspend the slurry before distributing it to the samples. 4. Rotate the sample, pellet and wash glutathione-agarose/Sepharose, and perform SDS-PAGE and immunoblot analysis (see Basic Protocol, steps 5 to 10). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Co-immunoprecipitation buffer 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 15 mM EGTA 100 mM NaCl 0.1% (w/v) Triton X-100 Store at 4°C Immediately before use add: 1× protease inhibitor mix (see recipe) 1 mM dithiothreitol (DTT) 1 mM phenylmethylsulfonyl fluoride (PMSF; from fresh 250 mM solution in 95% ethanol) The protease inhibitor mix, PMSF, and DTT should be added fresh at the time of experimentation. The mixture without those components can be stored for months at 4°C with the addition of 1 mM sodium azide. PMSF is labile in aqueous buffer and should be added at the last minute.


Glutathione-agarose or glutathione-Sepharose slurry Swell 1.5 g glutathione-agarose or glutathione-Sepharose beads (e.g., Pierce, Sigma) in 30 ml of 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2), for 1 to 2 hr on ice. Pellet beads by gravity or very gentle centrifugation (1 min at 1000 rpm in a tabletop centrifuge) and then wash four times with co-immunoprecipitation buffer (see recipe) that lacks protease inhibitor mix and contains 1 mM sodium azide. Resuspend beads in 15 ml of this buffer to yield a final slurry concentration of ∼100 mg/ml. Store at 4°C (stable for months).



Protease inhibitor mix, 1000× Dissolve in DMSO: 5 mg/ml chymostatin 5 mg/ml pepstatin A 5 mg/ml leupeptin 5 mg/ml antipain Store in aliquots up to 1 year at −20°C Protein A/G–Sepharose slurry Swell 1.5 g protein A– or protein G–Sepharose beads (e.g., Pierce, Sigma) in 30 ml of 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2), for 1 to 2 hr on ice. Pellet beads by gravity or very gentle centrifugation (1 min at 1000 rpm in a tabletop centrifuge) and then wash four times with co-immunoprecipitation buffer (see recipe) that lacks protease inhibitor mix and contains 1 mM sodium azide. Resuspend beads in 15 ml of this buffer to yield a final slurry concentration of ∼100 mg/ml. Store at 4°C (stable for months). The recipe can be scaled up or down.

COMMENTARY Background Information Coprecipitation is a powerful and simple approach to test for a physical interaction between proteins. There are many reasons to incorporate coprecipitation into a study. First, as a form of protein affinity chromatography, the method may be sensitive enough to detect weak associations that do not withstand the rigors of standard purification methods involving substantial dilution of the initial cell extract. Second, coprecipitation tests for associations between proteins within the milieu of a whole-cell extract, where the proteins are present at native concentration in a complex mixture of other cellular components. This feature makes it an important partner to two-hybrid methods and direct tests of interactions using purified proteins, for it provides a way to verify that a positive interaction reflects a true in vivo association. For example, nonphysiological interactions can be detected when purified proteins are present at too elevated a concentration. A falsely positive interaction between two proteins can also arise in a two-hybrid test when protein domains are inappropriately exposed due to altered folding. In addition, not all proteins are amenable to two-hybrid analysis; a negative result may mask a true association. Nevertheless, a word of caution is in order. The ability to coprecipitate two proteins from a cellular extract is not proof that a particular interaction normally takes place in vivo. Additional experiments are needed to argue that a given interaction is not the result of mixing cell contents during extract preparation. Such evi-

dence could include colocalization of the proteins or demonstration of functional relatedness. When performing coprecipitation, it is important to precipitate from both directions (i.e., individually precipitating protein 1 and protein 2, and testing for the presence of protein 2 and protein 1, respectively). This is important in that it can provide further verification of an interaction between the proteins. It is also important because it is possible the interaction will only be detected in one direction. An inability to detect an interaction in one direction could be due to a variety of factors including obstruction of an interaction by the binding of the antibody or other affinity agent, or differences in pool size representation of each protein. For example, protein 1 may bind to many proteins besides protein 2, while most of protein 2 binds to protein 1. In this scenario, it would be anticipated that detection of their association will be most efficient when protein 2 is precipitated.

Critical Parameters and Troubleshooting It is important to vary conditions of both the extract preparation and the coprecipitation to determine what is optimal. When starting from scratch, it is most prudent to use a range of lysis and precipitation conditions from less to more stringent in terms of the amount of salt and nonionic detergent. When no interaction is detected, it is worthwhile to use less stringent conditions (reduced salt with little or no non-



Supplement 46


ionic detergent). In addition, it may be necessary to avoid any dilution of the whole-cell extract. This can be done by adding the protein A/G–Sepharose directly to the extract after an initial clarification centrifugation and by using smaller wash volumes. Depending on the strength and nature of the interaction, the precipitation can be done in the presence of a mixture of detergents that includes ionic detergents (for example, 1% Triton X-100, 0.5% deoxycholate, 0.1% SDS), similar to that described in RIPA buffer (UNIT 10.16). In addition, it may be necessary to increase the expression levels of the proteins in question to be able to readily detect them by coprecipitation. A range of expression levels is recommended, because a level that is too high can lead to unregulated interactions (Feng et al., 1998). Alternatively, one can scale up the coprecipitation and use more than 0.5 to 1 mg of whole-cell extract (Feng et al., 1998). Here, the limiting factor is the concentration of the extracts, which must be high enough to allow the volume of the coprecipitation mixture to remain low. Larger-scale extract preparations may be necessary to generate more concentrated extracts. Finally, in cases of failure due to low abundance of the proteins in the host organism, one can overexpress a tagged version of one of the two proteins in the same or another host (such as E. coli), concentrate this protein by pre-immobilization on an appropriate affinity matrix, and then incubate the affixed protein with extracts from the host organism. The most important objective in these experiments is to generate as great a signal-tonoise ratio as possible and avoid problems of background. A variety of parameters can be changed to enhance the co-immunoprecipitation. Optimization of the precipitating antibody is one possibility. Protein A–Sepharose and protein G–Sepharose should give results comparable to anti-Ig serum. However, direct coupling of the antibody to Sepharose may lead to reduced background and more quantitative precipitation. In addition, varying the ratio of antibody to whole-cell extract and the total amount of whole-cell extract is strongly suggested to determine the optimal amount of antibody that gives the most precipitation with the least amount of background. Affinity purification of the antibody may be necessary if the antibody immunoprecipitates additional crossreacting proteins. Additional approaches can be taken to minimize background. First, better clarification of the cell extract can be done by precentrifugation

at 100,000 × g. These extracts can be directly used for coprecipitation without an intervening freezing step, which can increase the amount of protein precipitation. Second, both the lysis buffer and the coprecipitation buffer can be supplemented with 1% BSA to reduce the amount of nonspecific binding to the affinity matrix. Third, the whole-cell extract can be preincubated with protein A/G–Sepharose to remove nonspecific proteins that bind to the solid support. Fourth, the amount of salt and detergent can be increased in both the coprecipitation and the washes to reduce nonspecific binding. Fifth, increasing the number of washes may also help, although it may reduce the amount of specific protein that remains associated. Sixth, one can increase the expression levels of the proteins in question to generate a stronger signal that is above background binding. Alternatively, it may be possible to produce a whole-cell extract that is enriched for the proteins in question (e.g., by preparing a nuclear extract if the proteins are known to be in the nucleus). In instances where one of the proteins binds nonspecifically to Sepharose, the substitution of an agarose-based affinity matrix may help solve the problem. Finally, it may be necessary to generate a different set of reagents to precipitate the proteins in question (i.e., different antibodies and/or protein tag).

Anticipated Results Provided suitable antibodies are available to the proteins in question and the physical interaction is stable to the coprecipitation conditions, it should be possible to detect an interaction between two proteins.

Time Considerations Once the extracts are prepared, coprecipitation can be done within 3 to 4 hr, yielding samples ready to load on a gel for SDS-PAGE and immunoblot analysis.

Literature Cited BioSupplyNet Source Book. 1999. BioSupplyNet, Plainview, N.Y., and Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Feng, Y., Song, L.-Y., Kincaid, E., Mahanty, S.K., and Elion, E.A. 1998. Functional binding between Gβ and the LIM domain of Ste5 is required to activate the MEKK Ste11. Cur. Biol. 8:267-278. Field, J., Nikawa, J., Broek, D., MacDonald, B., Rodgers, L., Wilson, I.A., Lerner, R.A., and Wigler, M. 1988. Purification of RAS-responsive adenylyl cyclase complex from Sacchar-



omyces cerevisiae by use of an epitope addition method. Mol. Cell. Biol. 8:2159-2165. Harlow, E. and Lane, D. 1988. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Kolodziej, P.A. and Young, R.A. 1991. Epitope tagging and protein surveillance. Methods Enzymol. 194:508-519. Phizicky, E.M. and Fields, S. 1995. Protein-protein interactions: Methods for detection and analysis. Microbiol. Rev. 59:94-123. Tyers, M., Tokiwa, G., and Futcher, A.B. 1993. Comparison of the Saccharomyces cerevisiae G1 cyclins: Cln3 may be an upstream activator of Cln1, Cln2, and other cyclins. EMBO J. 11:17731784.

Key References BioSupplyNet Source Book. 1999. See above. Published yearly. Instant access is available on the WWW, at http://www.biosupplynet.com; hard copy may be requested by fax at (609) 786-4415. Information on its contents may be obtained by telephone at (516) 349-5595, fax at (516) 349-5598, or email at [email protected]. Phizicky and Fields, 1995. See above. General discussion of methodologies for detecting protein-protein interactions as well as their merits and drawbacks.

Contributed by Elaine A. Elion Harvard Medical School Boston, Massachusetts



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Identification of Protein Interactions by Far Western Analysis

UNIT 20.6

This unit describes far western blotting, a method of identifying protein-protein interactions. In a far western blot, one protein of interest is immobilized on a solid support membrane, then probed with a non-antibody protein. Far western blots can be used to identify specific interacting proteins in a complex mixture of proteins (see Basic Protocol). They are particularly useful for examining interactions between proteins that are difficult to analyze by other methods due to solubility problems or because they are difficult to express in cells. This method is performed totally in vitro, and the proteins of interest can be prepared in a variety of ways. Peptides can be used to determine the effects of specific residues or post-translational modifications on protein-protein interactions (see Alternate Protocol 2). In addition, many different detection techniques, either radioactive or nonradioactive, can be used. For example, the protein probe may be detected indirectly with an antibody, rather than being labeled radioactively (see Alternate Protocol 1). Thus, techniques and reagents already in hand can frequently be adapted for use with this assay. CAUTION: Appropriate safety precautions must be taken when working with radioactive materials. Information on proper handling and disposal of radioactive compounds can be found in APPENDIX 3A and may be obtained from local radiation safety officials. Specific information on handling 35 S-labeled compounds can be found in UNIT 10.18. FAR WESTERN ANALYSIS OF A PROTEIN MIXTURE The following is a basic method for detecting protein-protein interactions by far western blotting when one protein is contained within a simple or complex mixture of proteins. First, the protein sample is fractionated on an SDS-PAGE gel (UNIT 10.2A). After electrophoresis, the proteins are transferred from the gels onto a solid support membrane by electroblotting (UNIT 10.8). Transferred membranes may be stained with Ponceau S to facilitate location and identification of specific proteins. Nonspecific sites on the membranes are blocked with standard blocking reagents, and the membranes are then incubated with a radiolabeled non-antibody protein probe. After washing, proteins that bind to the probe are detected by autoradiography (APPENDIX 3A).

BASIC PROTOCOL

Materials Samples to be analyzed 1× SDS sample buffer (UNIT 10.2A) Ponceau S staining solution (see recipe) Blocking buffer I: 0.05% (w/v) Tween 20 in 1× PBS (see recipe for PBS); prepare fresh Blocking buffer II: dissolve 1 g bovine serum albumin (BSA; fraction V) in 100 ml 1× PBS (see recipe for PBS); prepare fresh Phosphate-buffered saline (PBS; see recipe), pH 7.9 cDNA encoding protein of interest cloned into an in vitro expression vector In vitro transcription/translation kit (Promega) 10 mCi/ml 35S-methionine (1000 Ci/mmol) Probe purification buffer (see recipe) Probe dilution buffer (see recipe) Analysis of Protein Interactions Contributed by Diane G. Edmondson and Sharon Y. Roth Current Protocols in Molecular Biology (2001) 20.6.1-20.6.10 Copyright © 2001 by John Wiley & Sons, Inc.


Polyvinyldifluoridine (PVDF) or nitrocellulose membrane for protein transfer Microfiltration centrifuge columns (e.g., Gelman Nanosep, Pall Filtron, or Millipore Microcon) Additional reagents and equipment for SDS-PAGE (UNIT 10.2A), electrophoretic transfer of proteins to a support membrane (UNIT 10.8), in vitro translation (UNIT 10.17), and autoradiography (APPENDIX 3A) NOTE: Always handle support membranes with gloves or membrane forceps. Prepare protein blot 1. Prepare the protein sample to be analyzed by resuspending it in 1× SDS sample buffer. UNIT 10.2A gives instructions on preparation of samples and amount of samples to load. In general, ∼50 to 100 ìg can be loaded in each lane for a complex mixture of proteins. A smaller amount, i.e., 10 to 20 ìg, is loaded for less complex protein samples. The amount loaded may also need to be adjusted for the size of gel. (Usually 30 ìg/mm2 loading surface can be resolved without smearing.)

2. Separate the samples on an SDS-polyacrylamide gel (UNIT 10.2A). 3. Transfer the proteins from the gel to a solid support membrane (e.g., PVDF or nitrocellulose) by semidry electroblotting (UNIT 10.8). Either nitrocellulose or PVDF membranes can be used with good results. PVDF membranes are easier to handle and tend to give a slightly higher signal-to-noise ratio, probably due to increased protein retention by the membrane.

Stain with Ponceau S 4. After transfer, stain the membrane for 5 min in ∼100 ml freshly diluted 1× Ponceau S staining solution. Stain the membrane in a plastic container large enough to hold the blot and use sufficient Ponceau S to cover the membrane completely. This step is optional. When the protein samples consist of a few proteins, or when there are clearly visible bands that facilitate orientation of the blot, staining with Ponceau S can provide helpful landmarks. One can unequivocally identify interacting bands, mark the position of molecular weight standards, and trim away excess membrane more exactly.

5. Destain the membrane washing in several changes of deionized water until the proteins are clearly visible. Place light pencil marks adjacent to important protein bands to mark them for future reference. Trim away excess membrane. The stain fades quickly so the marks must be placed immediately.

6. Destain an additional 5 min in water until the red staining fades. Block membrane 7. Block blot for 2 hr in 200 ml blocking buffer I at room temperature with gentle agitation. 8. Decant and add 200 ml blocking buffer II. Incubate as in step 7. 9. Decant blocking buffer II and rinse the membrane briefly in 100 ml of 1× PBS. At this point, the blot may be probed immediately or may be wrapped in plastic wrap and stored for up to 2 weeks at 4°C. Identification of Protein Interactions by Far Western Analysis



Prepare the probe 10. Following manufacturer’s procedures, prepare a radiolabeled in vitro–translated probe of the protein of interest using 35S methionine (also see UNIT 10.18). The probe can be conveniently prepared during the blocking steps. The authors routinely use the Promega TnT quick-coupled transcription/translation system for producing probes. For a small blot (e.g., ≤9 × 9–cm), one half of a standard in vitro transcription/translation reaction (i.e., 25 ìl) is sufficient for the probe. For larger blots, an entire 50-ìl reaction may be used. In the authors’ laboratory it is considered essential that the transcription/translation lysate not be repeatedly frozen and thawed.

11. After translation, dilute the probe with 500 µl probe purification buffer, and purify by microcentrifuging 15 to 30 min at 10,000 × g, room temperature, in a microfiltration column. Save aliquots of the purified probe for analysis by SDS-PAGE (i.e., 2 µl), and for scintillation counting (i.e., 2 to 5 µl). Check microfiltration column manufacturer’s procedure for exact centrifugation times required by different columns. In practice, it is not always necessary to purify the probe through a microfiltration column; many probes give good signals without purification. However, if signal-to-noise ratio is low, probe purification may improve results. In addition, it is possible to quantitate the proportion of probe bound if the probe has been purified.

Bind probe 12. Preincubate blot for 10 min in 50 ml of 1× probe dilution buffer (without probe) by gently agitating at room temperature. 13. Dilute the translated probe with 1× probe dilution buffer in a volume sufficient to cover the membrane to be probed (typically 3 ml). For small blots, i.e., ≤9 × 9–cm, a 50-ml conical tube makes a convenient incubation chamber. A volume of 3 ml is enough solution to cover the blot and tubes can be rotated on a mechanical rotator. In addition, the conical tube makes the radiolabeled probe easy to contain and dispose of. Larger blots can be incubated on a Nutator or orbital shaker in a heat-sealable bag, or rotated in a hybridization oven adjusted to room temperature.

14. Add the probe to the membrane and incubate 2 hr at room temperature. Rotate the tubes or agitate bags throughout the binding reaction. Wash the membrane 15. Transfer the membrane to a plastic dish and wash the membrane with 200 ml 1× PBS for 5 min, room temperature. Repeat for a total of four washes. Background is generally quite low and extended washing does not substantially reduce background.

16. Air dry the membrane and expose to X-ray film (autoradiography) or phosphor imager screen (see APPENDIX 3A for both techniques). Do not cover the blots with plastic wrap as this will quench the 35S signal. Overnight exposure to X-ray film is usually sufficient to detect positive interactions.

DETECTING INTERACTING PROTEINS BY IMMUNOBLOTTING In vitro–translated probes have the advantages of being quickly produced, easily detected, and quantitated to give an estimate of relative binding. In addition, mutations in the protein probe can be generated by simple cloning procedures and can provide information on binding domains and their critical residues. A disadvantage of in vitro–translated probes is the need for multiple methionine or cysteine residues to obtain a well labeled probe. For the same reasons, small peptide fragments are often not suitable for use as in




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vitro–translated probes. [14C]leucine and [3H]leucine can also be used for in vitro translation of proteins; however, in the authors’ laboratory [14C]leucine has not yielded probes suitable for use as far western probes. There are many other ways to generate probes for far western blots. The protein probe may be labeled in vitro with 125I (Schumacher and Tsomides, 1995) or enzymatically with 32 P (Kimball, 1998). Biotin-labeled probes may be detected with streptavidin-biotin detection schemes (Luna, 1996; Grulich-Henn et al., 1998; Kimball et al., 1998). Protein binding may be detected indirectly as well. If an antibody to the interacting protein is available, then an unlabeled protein probe can be bound to the blots as usual and then detected by western (immunoblot) analysis. This is especially useful when a tagged recombinant protein and antibody to the tag are available. The following procedure describes detection of an unlabeled protein probe with specific antibody. Many variations of immunoblotting exist; additional information and procedures may be found in UNIT 10.8. Additional Materials (also see Basic Protocol) Recombinant protein or unlabeled in vitro translated–protein for probe 5% (w/v) non-fat instant dry milk in 1× TBST (see recipe for TBST) Primary antibody specific for protein probe TBST (see recipe) Alkaline phosphatase (AP)–conjugated secondary antibody against Ig of species from which specific antibody was obtained Alkaline phosphatase buffer (see recipe) Developing solution (see recipe) 100 mM EDTA, pH 8.0 (APPENDIX 2) Prepare blot and probe 1. Prepare and block the blot (see Basic Protocol, steps 1 to 9). 2. Prepare the probe protein by diluting in vitro–translated or recombinant protein in 3 ml of 1× probe dilution buffer. The amount of recombinant protein must be empirically determined for each protein. Various researchers have used from 0.5 to 20 ìg recombinant protein/ml of probe dilution buffer.

Expose blot to probe 3. Bind the probe to blot and wash (see Basic Protocol, steps 12 and 14 to 15). Do not dry the membrane after washing. 4. Incubate blot in 200 ml of 5% non-fat milk in 1× TBST for 1 hr, room temperature, with gentle rotation on an orbital shaker. Expose to antibodies 5. Dilute the primary antibody in 5% milk in 1× TBST. Incubate blot in 5 to 10 ml diluted antibody at room temperature with gentle agitation to ensure blot is evenly covered with the antibody solution. Incubations are usually carried out in heat-sealed plastic bags or hybridization bottles to minimize the volume necessary to completely cover the blot. A volume of 5 to 10 ml of diluted antibody is sufficient to cover most blots. Appropriate antibody concentrations vary for each antibody and must be determined empirically. Identification of Protein Interactions by Far Western Analysis

6. Wash for 10 min in ≥200 ml 1× TBST by agitating on an orbital shaker. Repeat an additional two times.



7. Dilute the AP-conjugated secondary antibody in 5 to 10 ml of 5% milk in 1× TBST and incubate blot for 1 hr as in step 4. Suppliers generally provide an estimate of appropriate dilution for the secondary antibody.

8. Wash blot six times for 5 min each in ≥200 ml TBST, with agitation. Detect antibodies 9. Briefly, rinse blot in 50 ml alkaline phosphatase buffer. 10. Incubate blot in 20 ml developing solution for 1 to 15 min and rinse blot with 100 ml water. 11. Wash blot for 5 min with 100 ml of 100 mM EDTA, pH 8.0, to stop the development reaction. Rinse with 100 ml water, dry, and photograph.

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An example of a far western blot of proteins is shown in Figure 20.6.1. Lane 1 shows a Coomassie blue stain of a protein sample enriched for histone proteins separated on a 22% SDS-PAGE gel. Lane 2 shows a far western autoradiogram of a parallel lane probed with the yeast Tup1 protein according to this protocol. Lane 3 shows a parallel lane probed with an unlabeled Tup1 protein and detected with antibody specific to Tup1 as in Alternate Protocol 2. Both protocols yield the same result—Tup1p interacts with H3 and H4 but not with H2A or H2B. Lanes 4 and 5 show immunoblots of parallel lanes using anti-H3 and anti-H4 antibodies to identify histories H3 and H4 unequivocally.

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H3 H2B H2A H3* H4

Figure 20.6.1 Far western of blotted SDS-PAGE gel. Lane 1, Coomassie blue–stained gel showing locations of histone bands. Lane 2, far western of a parallel lane using radiolabeled in vitro–translated probe. Lane 3, far western using unlabeled probe detected with probe-specific antibody. Lane 4, western blot using anti-histone H3 specific antibody. Lane 5, western blot using anti-histone H4 specific antibody. Analysis of Protein Interactions


Supplement 55


USING PEPTIDES TO IDENTIFY SPECIFIC INTERACTING SEQUENCES IN A FAR WESTERN BLOT Synthetic peptides can also be used in far western analyses. The use of peptides enables the identification of specific interacting sequences. Specific post-translational modifications can be examined for their effect on protein-protein interactions. Peptides as small as 11 amino acids have been used successfully as far western targets. Peptide far westerns differ from other far westerns only in the preparation of the blots. Peptide dilutions are prepared, then dot or slot blotted onto the support membrane. Blocking and probing of peptide blots are identical to procedures used for traditional far westerns. Duplicate blots are stained to verify that comparable quantities of different peptides have been loaded. Because Ponceau S staining is temporary, staining of duplicate blots with India ink is used to provide a permanent record for peptide blots. In the authors’ experience, only peptides that have been synthesized on MAP resins have worked well for peptide blots. MAP resins consist of branched lysine chains whose chemically active groups have been blocked. Although peptides prepared in other ways do give reproducible results, the peptide concentrations required are several orders of magnitude higher than those required for MAP peptides, making these blots very costly to perform. The reason why increased peptide is needed is unclear, but perhaps the MAP resin “presents” the peptide in such a way that it is more accessible for interaction. Additional Materials (also see Basic Protocol) Peptides 0.4% Tween 20/PBS (see recipe) India ink solution (see recipe) Slot or dot blot apparatus (e.g., Bio-Rad Bio-Dot SF or Schleicher & Schuell Minifold II) 1. Make dilutions of peptides between 5 ng and 5 µg in a final volume of 100 to 200 µl of distilled water. 2. Prepare slot or dot blotter and support membrane (PVDF or nitrocellulose) as described by the manufacturer. Load the peptide dilutions into wells. Prepare duplicate blots, one for far western and one for India ink staining. After all samples are loaded, apply vacuum to draw the peptide samples through the manifold device and onto the support membrane. 3. Block, bind, wash, and autoradiograph one blot for far western (see Basic Protocol, steps 7 to 16). 4. Incubate the second blot in 100 ml of 0.4% Tween 20/PBS for 5 min at room temperature with gentle agitation. Repeat incubation. 5. Stain blot by incubating 15 min to overnight with 100 ml India ink solution at room temperature. 6. Wash the filter for 2 hr in 4 changes of 1× PBS. Dry and store the membrane. This stain is permanent.


Figure 20.6.2 shows an example of results from this alternate protocol, using peptides as a substrate for a far western. The right-hand panel is a blot stained with India ink verifying that comparable quantities of peptide were loaded on the blot. The left-hand panel is a far western of a duplicate blot demonstrating the effect of acetylation of lysine residues of histone peptides on Tup1p/histone interaction.



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5 µg 1 µg 0.2 µg 0.04 µg 0.008 µg far western blot

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Figure 20.6.2 Far western blot of peptides (A) dot blotted onto PVDF membrane and (B) stained with India ink. Figure reproduced with permission of Cold Spring Harbor Laboratory Press.

REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Alkaline phosphatase buffer 100 ml 1 M Tris⋅Cl, pH 9.5 (APPENDIX 2) 20 ml 5 M NaCl (APPENDIX 2) 5 ml 1 M MgCl2 (APPENDIX 2) Add H2O to 1 liter Store up to 1 year at room temperature 5-Bromo-4-chloro-3-indolyl phosphate (BCIP) stock solution Dissolve 0.5 g of BCIP in 10 ml of 100% dimethylformamide. Store at 4°C or in small aliquots at −20°C. Discard when solution turns color. Developing solution Add 66 µl of NBT stock (see recipe) to 10 ml of alkaline phosphatase buffer (see recipe). Mix well. Add 33 µl of BCIP stock solution (see recipe) and mix again. Prepare fresh. India ink solution Add 100 µl India ink (Pelikan or Higgans) to 100 ml 0.4% Tween 20/PBS (see recipe). Prepare fresh. Nitroblue tetrazolium chloride (NBT) stock solution Dissolve 0.5 g of NBT in 10 ml of 70% dimethylformamide. Store at 4°C or in small aliquots up to 6 months at −20°C. Phosphate-buffered saline (PBS), 10× 80 g NaCl 2.2 g KCl 9.9 g Na2HPO4 2.0 g K2HPO4 Add H2O to 1 liter

Analysis of Protein Interactions continued



Adjust pH to 7.4 Store indefinitely at room temperature Prior to use, dilute to 1× by mixing 1 part 10× PBS with 9 parts water Leftover 1× PBS should be stored at 4°C to discourage bacterial growth.

Ponceau S staining solution, 10× 2 g Ponceau S 30 g trichloroacetic acid 30 g sulfosalicylic acid Add H2O to 100 ml Store indefinitely at room temperature Just prior to use dilute to 1× by mixing 1 part 10× Ponceau S with 9 parts water Probe dilution buffer, 10× 3.0 g bovine serum albumin (BSA) 10 ml normal goat serum 10 ml 10× PBS (see recipe) H2O to 100 ml Store indefinitely at −20°C Just prior to use, dilute to 1× by mixing 1 part 10× stock with 9 parts 1× PBS Probe purification buffer 400 µl 1 M HEPES, pH 7.4, 400 µl 1 M dithiothreitol (DTT) 9.2 ml H2O Prepare fresh TBST, 10× 90 g NaCl 100 ml 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2) 10 g Tween 20 Add H2O to 1 liter For 1× TBST dilute 1 part 10× TBST with 9 parts water prior to use Store indefinitely at room temperature Tween 20/PBS, 0.4% (w/v) Dissolve 0.4 g Tween 20 in 100 ml 1× PBS (see recipe). Store up to 1 week at room temperature. COMMENTARY Background Information


The far western blot (also called a west western and a ligand blot) has been widely used to examine the interactions of many diverse proteins. For example, it has been used to examine the interactions between the subunits of eukaryotic initiation factors (Kimball et al., 1998), to look at interactions between basic helix-loop-helix DNA-binding proteins (Chaudhary et al., 1997), and to examine the interactions of keratin intermediate filaments with desmosomal proteins (Kouklis et al., 1994). Far westerns have been particularly useful in examining interactions of histones with regulatory proteins. Far westerns have been

used to look at interactions of WD repeat proteins with histones (Edmondson et al., 1996; Palaparti et al., 1997), the interaction of Epstein-Barr virus nuclear antigen 2 with histone H1 (Grasser et al., 1993), and the interaction of histones with the Xenopus oocyte protein N1 (Kleinschmidt and Seiter, 1988). In addition, far westerns have been used to study receptorligand interactions and to screen libraries for interacting proteins (Grulich-Henn et al., 1998; Hsiao and Chang, 1999). Sometimes, the nature of the proteins being examined is such that standard methods of studying protein-protein interactions are not possible. For example, some proteins are diffi-



cult to solubilize or to extract from cells except under conditions that disrupt protein-protein interactions and therefore, are difficult to assay by immunoprecipitation. Other proteins cannot be expressed in bacteria or yeast due to toxicity problems, thus making the production of recombinant proteins or the use of two-hybrid assays impossible. Far westerns are particularly useful in such cases. Since far westerns are performed totally in vitro, they circumvent these types of problems. Another advantage of the far western blot is its flexibility. Proteins prepared in a variety of ways can be used for the assay. Cell extracts, recombinant proteins, and peptides can all be used as both probe and target proteins. For example, Palaparti et al. (1997) used cell extracts to probe a semipurified histone sample and detected bound proteins of interest with antibodies. Hsiao and Chang (1999) used phage-expressed proteins immobilized on filter lifts for a far western library screen. Unpurified E. coli extracts containing recombinant protein have been successfully used as probes (Fischer et al., 1997). In addition, many different detection techniques, either radioactive or nonradioactive, can be used. Kimball and co-workers used recombinant protein probes that were labeled radioactively with kinases and recombinant proteins labeled with biotin and subsequently detected with a strepavidin-biotin detection scheme (Grulich-Henn et al., 1998; Kimball et al., 1998). Thus, techniques and reagents already in hand can frequently be adapted for use with this assay. Finally, the far western can be modified to define the protein domains and amino acid residues that are important in protein-protein interactions. Mutagenized clones can be used to produce variants of protein probes. A single SDS-PAGE gel can be run with identical lanes and cut into strips, and a different in vitro–translated probe can be used for each strip. In this way, multiple variants of a protein can be tested simultaneously for their ability to interact with a target protein. Non-SDS polyacrylamide gels can also be used to separate proteins for far westerns. For example, acid urea gels, which separate on the basis of both size and charge, have been successfully employed. Finally, peptides corresponding to specific interacting sequences can be synthesized with specific post-translational modifications to test their effects on proteinprotein interactions.

Critical Parameters Blocking nonspecific binding sites on the membranes is critical to achieving good results with far westerns. Too little blocking results in high background, while extended time in blocking solutions results in weakened or lost signal. The reason for the diminished signal is unclear, but protein renaturation apparently takes place during the blocking step, so an optimal renaturation may require limited blocking. The best time may well be different for each protein and require empirical optimization. In addition, different lots of BSA appear to result in diminished signal. Therefore, it is important to purchase high-quality BSA from a reputable manufacturer. An important consideration is the inclusion of appropriate controls to rule out nonspecific interactions that might result in false positives. Suitable controls should be furnished for both the target proteins and the protein probe. When using a complex mixture of proteins, such as cell extracts, as target, “negative” control proteins are already present. However, when using a mixture of only a few proteins, it is important to provide a protein that does not interact to serve as a negative control for nonspecific binding. The ideal negative control should be similar to the protein of interest in charge and size. Appropriate controls should be subjected to SDS-PAGE and blotted in parallel with the samples of interest. Another important control is the use of an unprogrammed translation lysate as a probe. Translation of an unrelated protein as a control probe is also often helpful.

Troubleshooting Precise conditions for far westerns vary from procedure to procedure and probably reflect the nature of the individual proteins being examined. Optimal conditions for each protein may need to be determined empirically. If background staining is too high, there are several possible remedies. The probe may be diluted or the sample concentration lowered. Other “blocking” reagents may be tested.The blocking reagent, nonfat dry milk, ranging in concentration from 1% to 5%, has been used successfully. Other detergents such as NP-40 and Triton X-100 are commonly employed for this procedure and may help to decrease background. Also, increasing the length of the blocking step may aid in background reduction. Most procedures that call for extended blocking times suggest incubation at 4°C.



Supplement 55

If no specific staining is observed, confirm the quality of the radiolabeled probe by SDSPAGE and autoradiography. Make sure the reticulocyte lysate has not been repeatedly frozen and thawed. If using a recombinant protein/antibody scheme, confirm the affinity of that interaction by immunoblotting. Sometimes a decrease in blocking time or use of a different blocking reagent will increase signal. Some proteins may interact more readily following a denatuation/renaturation cycle. In this case, the membrane is incubated for 1 hr in PBS-buffered 7 M guanidine or 8 M urea and renatured overnight. Renature in blocking buffer I (see Basic Protocol) without agitation at 4°C with several changes of buffer. Finally, the length of the binding reaction can be increased.

Anticipated Results Sensitivity of the far western blot is dependent on the affinity of the protein-protein interactions being investigated and on the quality of the probe. Thus, extra attention given to the preparation of a high-quality probe is almost invariably worthwhile. When using a radiolabeled probe, a positive interaction can typically be visualized after overnight exposure to X-ray film.

Time Considerations The basic protocol can be performed in 2 days. An SDS-PAGE gel can be set up and run in 4 to 6 hr. The rest of the procedure can be performed in 8 to 10 hr. It is often convenient to set up the SDS-PAGE gels on one day and run them slowly overnight. The rest of the basic protocol can then be performed the next day. Semidry transfer requires ∼2 hr, the blocking steps take ∼4 hr, binding of the probe takes ∼2 hr, washing, drying, and setting up the autoradiography cassette takes ∼1 hr. Detection of interactions using antibodies to detect the probe protein requires an additional day. Peptide blots can be performed in one day. It is possible to store the blots after the blocking steps for up to 2 weeks at 4°C. The blots should be stored in airtight containers or wrappings so they do not dry out. In addition, in vitro–translated probes may be produced ahead of time and stored unpurified at −20°C, although the efficiency may be reduced for some probes. Identification of Protein Interactions by Far Western Analysis

Literature Cited Chaudhary, J., Cupp, A.S., and Skinner, M.K. 1997. Role of basic-helix-loop-helix transcription factors in Sertoli cell differentiation: Identification

of an E-box response element in the transferrin promoter. Endocrinology 138:667-675. Edmondson, D.G., Smith, M.M., and Roth, S.Y. 1996. Repression domain of the yeast global repressor Tup1 interacts directly with histones H3 and H4. Genes & Dev. 10:1247-1259. Fischer, N., Kremmer, E., Lautscham, G., MuellerLantzsch, N., and Grasser, F. A. 1997. EpsteinBarr virus nuclear antigen 1 forms a complex with the nuclear transporter karyopherin alpha2. J. Biol. Chem. 272:3999-4005. Grasser, F. A., Sauder, C., Haiss, P., Hille, A., Konig, S., Gottel, S., Kremmer, E., Leinenbach, H. P., Zeppezauer, M., and Mueller-Lantzsch, N. 1993. Immunological detection of proteins associated with the Epstein-Barr virus nuclear antigen 2A. Virology 195:550-560. Grulich-Henn, J., Spiess, S., Heinrich, U., Schonberg, D., and Bettendorf, M. 1998. Ligand blot analysis of insulin-like growth factor-binding proteins using biotinylated insulin-like growth factor-I. Horm. Res. 49:1-7. Hsiao, P. W., and Chang, C. 1999. Isolation and characterization of ARA160 as the first androgen receptor N-terminal-associated coactivator in human prostate cells. J. Biol. Chem. 274:2237322379. Kimball, S. R., Heinzinger, N. K., Horetsky, R. L., and Jefferson, L. S. 1998. Identification of interprotein interactions between the subunits of eukaryotic initiation factors eIF2 and eIF2B. J. Biol. Chem. 273:3039-3044. Kleinschmidt, J. A., and Seiter, A. 1988. Identification of domains involved in nuclear uptake and histone binding of protein N1 of Xenopus laevis. EMBO J. 7:1605-1614. Kouklis, P. D., Hutton, E., and Fuchs, E. 1994. Making a connection: Direct binding between keratin intermediate filaments and desmosomal proteins. J. Cell Biol. 127:1049-1060. Luna, E.J. 1996. Biotinylation of proteins in solution and on cell surfaces. In Current Protocols in Protein Science. (J.E. Coligan, B.M. Dunn, H.L. Ploegh, D.W. Speicher, and P.T. Wingfield, eds.) pp. 3.6.1-3.6.15. John Wiley & Sons, New York. Palaparti, A., Baratz, A., and Stifani, S. 1997. The Groucho/transducin-like enhancer of split transcriptional repressors interact with the genetically defined amino-terminal silencing domain of histone H3. J. Biol. Chem. 272:26604-26610. Schumacher, T.N.M. and Tsomides, T.I. 1995. In vitro radiolabeling of peptides and proteins. In Current Protocols in Protein Science (J.E. Coligan, B.M. Dunn, H.L. Ploegh, D.W. Speicher, and P.T. Wingfield. eds.) pp. 3.3.1-3.3.19. John Wiley & Sons, New York.

Contributed by Diane G. Edmondson and Sharon Y. Roth M.D. Anderson Cancer Center Houston, Texas



Two-Hybrid Dual Bait System

UNIT 20.7

In characterizing a given protein’s function, it is frequently desirable to identify other proteins with which it interacts. The yeast two-hybrid system, or interaction trap, is one of the most versatile methods available with which to identify/establish protein-protein interactions. This system relies on the modular nature of many transcription factors, wherein different domains (e.g., DNA binding, transcription activating) function independently. The two-hybrid system takes advantage of this functional independence of transcription factor domains by expressing the DNA binding domain and transcription activation domain as components of separate fusion proteins, which can be coalesced via noncovalent interactions to reconstitute a functional transcription factor. For example, in the two-hybrid system, a protein of interest (the bait) may be expressed as a fusion protein with a DNA binding domain. In parallel, the interacting protein (the prey) is expressed as a fusion protein with a transcription activation domain. Co-expression of the bait and prey fusion proteins in the appropriate yeast strain reconstitutes transcription activity through noncovalent bait-prey interactions. The yeast two-hybrid system was conceived to determine the affinity of single pair protein-protein interactions (Fields and Song, 1989), but rapidly evolved into an efficient means to identify interacting protein binding pairs in both directed and nondirected screens. The latter feature enables the use of the two-hybrid system to screen a bait library against a prey library to identify interacting protein binding pairs. The two-hybrid system, therefore, lends itself to genomic and proteomic level screens. While the two-hybrid system is well suited to high-throughput screening methodology, it is an ideal system for such broad-based screening projects. More recently, the basic system has evolved to encompass a variety of adaptations, which enhance the versatility of the system by including modifying/accessory proteins and compatibility with peptide libraries and RNA-protein screens (reviewed in Serebriiskii et al., 2001). The dual bait system (Serebriiskii et al., 1999), which is one such adaptation of the classic two-hybrid system, is the focus of this unit. The dual bait system facilitates the simultaneous comparison of two distinct baits with one prey. Briefly, in the dual bait system one protein of interest is expressed as a fusion to the DNA binding protein LexA (bait 1), while a second protein of interest is expressed as a fusion to the DNA binding protein cI (bait 2). Strains of yeast engineered for screening of these dual baits possess four separate reporter genes: GusA and LYS2, which are transcriptionally responsive to a cI operator (cIop); and LacZ and LEU2, which are transcriptionally responsive to a LexA operator (lexop). A plasmid expressing an activation domain–fused protein (prey), which can be either a defined protein interactor or a cDNA library, is also expressed to allow dual hybrid-mediated transcriptional activation. Selective interaction of the prey with one of the two baits is scored by observing transcriptional activation of either the LexA operator–responsive reporters or the cI operator–driven reporter genes—e.g., if the prey preferentially interacts with bait 1 (as shown in Fig. 20.7.1), then LacZ and LEU2 will be more activated than LYS2 and GusA. This selective activation can be detected by comparing yeast growth on plates lacking leucine or lysine, and by comparing quantitative results of LacZ and GusA, respectively. By facilitating the simultaneous assay of one prey with multiple baits, the dual bait system overcomes two major problems inherent to the two-hybrid approach. First, since many proteins are members of large protein families which share considerable sequence similarity, the degree to which two-hybrid systems can differentiate between specific partners (e.g., individual members of a protein family) and less specific or nonspecific Contributed by Elena Kotova, Ilya Serebriiskii, and Thomas Coleman Current Protocols in Molecular Biology (2002) 20.7.1-20.7.37 Copyright © 2002 by John Wiley & Sons, Inc.



AD

Prey Bait1

1

integrated

growth w/o leucine

AD

Prey Bait1 LexA

3

LEU2

LexA LexA-op

2

LacZ

LexA-op

blue on XGal

3

plasmid-borne 4

AD

Prey Prey

AD

AD

Prey

Bait2 1

gusA

cI cI-op

white on XGluc

3

plasmid-borne

Bait2

cI cI-op

5

LYS2

no growth w/o lysine

3

integrated 6

Figure 20.7.1 Outline of the dual bait system and the key control points. An activation domain (AD)-fused prey interacts with Bait1 (LexA-fused) to drive transcription of lexAop-responsive reporters (LEU2 and LacZ), but does not interact with Bait2 (cI-fused) and thus does not turn on transcription of cIop-responsive reporters (LYS2 and GusA). As shown in this figure, the cI-Bait represents a negative control for prey binding; the system can also be configured so prey interacts with only cI-bait or both baits. To maximize the utility of the system, one can adjust experimental parameters affecting expression of baits and reporter readouts. Points of flexibility in the dual bait-hybrid system are indicated in the figure by circled numbers and include: (1) varying expression level of baits, (2) varying cellular localization of baits, (3) varying sensitivity of reporters, (4) diversifying plasmid markers to broaden compatibility with different yeast two-hybrid systems and to facilitate isolation of library plasmid in E. coli, (5) enriching polylinkers and choice of reading frames to facilitate cloning of baits, and (6) development of robust yeast strains suitable both for bait testing and interaction mating.


partners (e.g., all of the members of the protein family) for a given bait can be an issue. This notion is of paramount importance since multiple members of a protein family are frequently co-expressed in a single cell type and assigning a single physiological interactor with a particular function can be challenging. Second, the majority of two-hybrid library screens yield one or more nonspecific interactors (see http://www.fccc.edu/ research/labs/golemis/InteractionTrapInWork.html). These false positives are proteins with broad interaction capabilities that result in nonspecific protein binding. Sorting out the physiologically meaningful interactors from the false positives can be time consuming. The dual bait system, therefore, provides a number of significant advantages, including the ability to readily distinguish interactions specific for individual members of related protein families and specific from nonspecific protein-protein interactions.



construct LexAbait plasmid

7–9 days

obtain or construct cDNA library

construct cI-bait plasmid

transform cDNA library into mating partner yeast strain under noninducing conditions

transform yeast; characterize transcriptional activity of baits testing Lys and Leu reporters

freeze and store transformants

characterize expression of bait proteins by immunoblot

determine plating efficiency of pretransformed library

combine two baits in one strain and complete characterization

6 – 8 days

set up mating

induce library expression and select for interacting proteins

7–10 days

characterize clones by replica plating

characterize inserts by PCR and restriction mapping, choose independent clones

sequence the inserts, choose prospective isolates

test for specificity using in vivo recombination and mating

analyze the interactors by other means

Figure 20.7.2 Flow chart of the two-hybrid screen done by interaction mating. The third stage allows some flexibility, reflected in the availability of different protocols (see Basic Protocol 4 and Alternate Protocol).



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The following protocols can be used to investigate questions related to specific protein interactions that are difficult to clarify by other means. This method is a derivation of the classical interaction trap/two-hybrid system and readers are referred to Chapters 1 to 3, 10, and 13 for a more detailed description of the basic methodology related to the execution of the two-hybrid technique (e.g., preparation of yeast medium, immunoblotting). It is recommended that an investigator who is attempting to perform a dual bait screen as a first effort in the field of yeast interaction trap screens consult a basic protocol manual describing the interaction trap system (UNIT 20.1). The outline of the protocol is presented in Figure 20.7.2. In the first protocol (see Basic Protocol 1), baits are transformed into the yeast reporter strains and characterized for protein expression and transcriptional activation. In parallel, a library is introduced into a yeast strain of the opposite mating type, to enable mating (see Basic Protocol 2). In the next method (see Basic Protocol 3), the bait-expressing yeast strain is mated to the library-bearing strain. Expression of library-encoded proteins is induced by growing the diploids in galactose-containing medium, and yeast containing interacting pairs of proteins are identified on the selection plates. Primary candidates are analyzed for their ability to activate the correct set of reporters selected for further study. In the final protocol (see Basic Protocol 4), selected primary candidates are further analyzed by PCR to detect redundant clones, and then the library inserts are isolated and characterized to confirm the specific interaction with the bait protein(s). Clones which satisfy the user’s criteria are ready for further analysis. Three additional protocols are provided as support material. The first (see Support Protocol 1) outlines the Xgal or Xgluc overlay technique, which provides a convenient and sensitive assay to assess β-galactosidase or β-glucuronidase activity of yeast. The second (see Support Protocol 2) describes the detection of bait protein from growth of yeast through immunoblot analysis. Establishing both the expression level and proteolytic status for each novel bait construct is critical for a meaningful interpretation of bait-prey interactions. In the last (see Support Protocol 3) a method to assess the quality of digested library plasmid is described. This protocol is recommended in order to assure minimal background in the testing specificity of interaction using homologous recombination. Together with the basic protocols, these support protocols provide the necessary steps for performing and interpreting the two-hybrid dual bait system. BASIC PROTOCOL 1

PREPARATION OF BAITS AND LIBRARY: CHARACTERIZING BAIT PROTEINS To utilize the dual bait system, constructs encoding fusion proteins of independent DNA-binding domains (either LexA or cI) and the two baits must be prepared. A protein to be used as a primary bait should be fused to LexA, since more options exist to optimize LexA bait performance.


Materials DNA encoding the proteins of interest LexA-fusion plasmids and controls (Table 20.7.1): pMW103 (Figs. 20.7.3 and 20.7.4), pEG202-hsRPB7 (control), pSH17-4, and pEG202-Ras (control) cI-fusion plasmids and controls (Table 20.7.2): pGBS10 (Figs. 20.7.4 and 20.7.5) , pGBS10-Krit (control), and pGBS10-Krev (control) Yeast strains (Table 20.7.3): SKY191 and SKY48 YPD plates and liquid media (UNIT 13.1) with and without 200 µg/ml geneticin (G418; APPENDIX 1K) pLacGus (Fig. 20.7.6)



Table 20.7.1

LexA-Fusion Plasmids and Controls

Plasmid name

Yeast selection

E. coli selection

LexA Fusion plasmids pMW101 HIS3

CmR

pMW103

HIS3

KmR

pEG202

HIS3

ApR

pJK202

HIS3

ApR

pNLexA

HIS3

ApR

pEG202I

HIS3

ApR

pGilda

HIS3

ApR

pDD

HIS3

KmR

pHybLex/Zeo

ZeoR

ZeoR

pCGLex/p2GLex

ZeoR

ZeoR

Control LexA-fused baits pEG202-Ras HIS3

ApR

pEG202-hsRPB7

HIS3

ApR

pEG202-Krit1 pRFHM1 (control)

HIS3 HIS3

ApR ApR

pSH17-4 (control)

HIS3

ApR

Comment/description

Contact information

Basic plasmids to clone bait as fusion with LexA. Expression is driven by the ADH1 promoter. Basic plasmids to clone bait as fusion with LexA. Expression is driven by the ADH1 promoter. Basic plasmids to clone bait as fusion with LexA. Expression is driven by the ADH1 promoter. pEG202 derivative, incorporating nuclear localization sequences between LexA and polylinker (enhanced ability to translocate bait to nucleus) Polylinker is upstream of LexA, which allows fusion of LexA to C terminus of bait, leaving amino-terminal residues of bait unblocked pEG202 derivative (see above), which can be integrated into yeast HIS3 gene after digestion with KpnI. Ensures lower levels of bait expression. GAL1 promoter and CEN-ARS backbone facilitate a tightly controlled, galactose-inducible bait expression; should be used if continuous presence of the bait is toxic to yeast GAL1 promoter and CEN-ARS backbone facilitate a tightly controlled, galactose-inducible bait expression; should be used if continuous presence of the bait is toxic to yeast Bait cloning vector, compatible with IT and all other two-hybrid systems. Minimal ADH1 promoter expresses LexA followed by extended polylinker. Gal-inducible bait vector, compatible with IT and all other two-hybrid systems. GAL1 promoter expresses LexA followed by extended polylinker. Both high and low copy number versions available.

RBa

A negative control for activation and positive control for interaction with Raf1 and RalGDS A weak positive control for activation

I. Serebriiskiig

A moderate positive control for activation The homeodomain of bicoid cloned into pEG202 backbone (see above); the resulting nonactivating fusion is recommended as a negative control for activation and interaction assays, and as a positive control for repression assay. GAL4 activation domain cloned into pEG202 backbone (see above) is recommended as a positive control for transcriptional activation.

RBa Origeneb, MoBiTecc Origeneb Origeneb RBa Origeneb

R. Hopkinsd

Invitrogene J. Huangf

I. Serebriiskiig I. Serebriiskiig Origeneb

Origeneb

aContact R. Brent at [email protected]. bSee the Origene Web-site at http://www.origene.com. cSee the MoBiTec Web-site at http://www.mobitec-germany.com. dContact R. Hopkins at [email protected]. eSee the Invitrogen Web-site at http://www.invitrogen.com. fFor more information, refer to Huang and Schreiber (1997) or contact S. Schreiber at [email protected]. gContact Ilya Serebriiskii at [email protected].


Supplement 60

R pB R

NcoI

c ba

/N eo

95

0

00

00

00

A

85

00

SpeI

MluI

75 00

pMW103 10101 nt

EcoRI SmaI BamHI SalI NcoI NotI XhoI SalI

7

0

000

30 0

BsaI

A DH t er m

2500

SmaI

0

8000

200

NruI

le x

Km

o ne

15

AatII

90

kb

HpaI

PacI BsrGI A DH BsaI 1p rom ot er 50 0 10 00

65

00

00

35

XbaI ri

00

o

µ

450

60

0

50 0 0

55 0

00

0 KpnI

40

S3 HI

BsiWI

SpeI

2

HpaI SacI

KpnI

AvrII

Figure 20.7.3 Plasmid map of the 10101-nt basic dual bait vector pMW103. This plasmid, a derivative of pEG202, uses the strong constitutive ADH1 promoter to express bait proteins as fusions to the DNA binding protein LexA. The plasmid contains the HIS3 selectable marker and the 2 µ origin of replication to allow selection and propagation in yeast, the kanamycin resistance gene and the pBR origin of replication to allow selection and propagation in E. coli. A detailed map of the LexA-fusion polylinker is given in Figure 20.7.4. More maps and sequences are available on the Web at http://www.fccc.edu/ research/labs/golemis/InteractionTrapInWork.html.

90- or 100-mm complete minimal (CM) medium dropout plates (UNIT 13.1) with and without 350 µg/ml G418 lacking the following nutrients and supplemented with 2% (w/v) glucose (glu) or 2% (w/v) galactose (gal) and 1% raffinose (raff): Glu minus uracil and histidine (Glu/CM −Ura −His) Glu minus lysine (Glu/CM −Lys) Gal and raff minus lysine (Gal-Raff/CM −Lys) Gal and raff minus uracil, histidine, and leucine (Gal-Raff/CM −Ura −His −Leu) Gal and raff minus uracil and histidine (Gal-Raff/CM −Ura −His) Gal and raff minus uracil, histidine, and lysine (Gal-Raff/CM −Ura −His −Lys) Glu minus tryptophan (Glu/CM −Trp; also in 24 × 24–cm plates) H2O, sterile


96-well microtiter plate Insert grid from a rack of 200-µl micropipet tips Tape 200-µl micropipet tips, sterile Metal frogger (e.g., Dankar Scientific) or plastic replicator (Bel-Blotter; Bel-Art Products or Fisher)



A pMW103 (pEG202) SalI SalI . NotI * EcoRI BamHI NcoI XhoI . GAA TTC CCG GGG ATC CGT CGA CCA TGG CGG CCG CTC GAG TCG AC

pNLexA NotI BamHI . EcoRI XhoI . g aat tcg cgg ccg cct cga ggg atc caa ttc ATG AAA N S R P P R G I Q F M K

B cI-fusion plasmids (e.g., pGBS 10) A = AAT frame BglII ApaI NotI SalI EcoRI SacI PvuII KpnI XhoI__ G AAT TCA AGC TTG AGC TCA GAT CTC AGC TGG GCC CGG TAC CGC GGC CGC TCG AGT CGA Cct gca N S S L S S D L S W A R Y R G R S S R P A

B = GAA frame EcoRI

BglII ApaI NotI SalI . SacI PvuII KpnI XhoI__. G AAT Ttg GAA TTC GAG CTC AGA TCT CAG CTG GGC CCG GTA CCG CGG CCG CTC GAG TCG ACC TGC E F E L R S Q L G P V P R P L E S T C N L

C pJG4-5 (aka pB42AD, displayTarget) ATG GGT GCT CCT CCA AAA AAG AAG ... M G A P P K K K ...

EcoRI XhoI . CCC GAA TTC GGC CGA CTC GAG AAG CTT ... P E F G R L E K L ...

pYesTrp2 KpnI BamHI . HinDIII SacI . ATG GGT AAG CCT ... AAG CTT GGT ACC GAG CTC GGA TCC ACT AGT AAC GGC N G M G K P K L G T E L G S T S EcoRI NotI SphI . BstXI BstXI XhoI . CGC CAG TGT GCT GGA ATT CTG CAG ATA TCC ATC ACA CTG GCG GCC GCT CGA GGC ATG C R Q C A G I L Q I S I T L. A A A R G M H

Figure 20.7.4 Polylinkers of the two-hybrid basic vectors. (A) The LexA-fusion vectors polylinker for (top) pMW103 (also see Fig. 20.7.3) and (bottom) pNLexA. (B) The cI-fusion vectors polylinker. All cI-fusion plasmids with antibiotic resistance markers (Zeo or G418) share this polylinker (e.g., pGBS10; Fig. 20.7.5). Note both the A (top) and B (bottom) reading frames are shown. These are the AAT and GAA reading frames, respectively. (C) The AD-fusion (library) vectors polylinker for (top) pJG4-5 (also see Fig. 20.7.8), which is also known as pB42AD and displayTarget, and pYesTrp2. Only restriction sites that are available for insertion of coding sequences are shown; those shown in bold type are unique. The asterisk (*) in panel A denotes that NcoI is unique in pEG202 but not pMW103.



Supplement 60

Table 20.7.2

cI-Fusion Plasmids and Controls

Plasmid name (reading frames)

Yeast selection

E. coli selection

cI-Fusion plasmids pGKS3 (A, B) pGKS4 (A, B) pHybcI/HK (B) pGKS8 (A, B)

HIS3 HIS3 HIS3 ZeoR

ApR KmR KmR ZeoR

pGKS6 (A, B, C) pGKS7 (A, B)

ZeoR ZeoR

ZeoR ZeoR

G418R G418R

KmR KmR

pGMS11 (A)

ZeoR

ZeoR

pGMS12 (B)

G418R

KmR

Control cI-fused baits pGKS3-Krev

HIS3

ApR

pHybcI/HK-Krev

HIS3

KmR

pGBS9-Krev

G418R

KmR

pGBS10-Krev

G418R

KmR

pGKS6-Krev

ZeoR

ZeoR

pGKS6-Krit

ZeoR

ZeoR

pGKS7-Krit

ZeoR

ZeoR

pGKS3-Krit

HIS3

ApR

pGBS9-Krit

G418R

KmR

pGBS10-Krit

G418R

KmR

pGMS12-Krit

G418R

KmR

pGBS9 (A, B) pGBS10 (A, B)

Comment/descriptiona

ADH1 promoter expresses cI followed by polylinker ADH1 promoter expresses cI followed by polylinker ADH1 promoter expresses cI followed by polylinkerb Dual purpose vector: ADH1 promoter expresses cI followed by polylinker, while cI-responsive gusA reporter cassette is integrated into the same backbone. ADH1 promoter expresses cI followed by polylinker Modified ADH1 promoter ensures higher level of expression of cI ADH1 promoter expresses cI followed by polylinker Modified ADH1 promoter ensures higher level of expression of cI GAL promoter expresses cI followed by polylinker, for use with baits whose continuous presence is toxic to yeast GAL promoter expresses cI followed by polylinker, for use with baits whose continuous presence is toxic to yeast Expresses cI-Krev fusion protein. Use as negative control for activation assay and positive control for interaction with Krit1. Expresses cI-Krev fusion protein. Use as negative control for activation assay and positive control for interaction with Krit1b. Expresses cI-Krev fusion protein. Use as negative control for activation assay, and positive control for interaction with Krit1. Expresses cI-Krev fusion protein. Use as negative control for activation assay, and positive control for interaction with Krit1. Expresses cI-Krev fusion protein. Use as negative control for activation assay, and positive control for interaction with Krit1. Expresses cI-Krit fusion protein. Use as positive control for activation assay. Expresses cI-Krit fusion protein. Use as positive control for activation assay. Expresses cI-Krit fusion protein. Use as positive control for activation assay. Expresses cI-Krit fusion protein. Use as positive control for activation assay. Expresses cI-Krit fusion protein. Use as positive control for activation assay. Expresses cI-Krit fusion protein. Use as positive control for activation assay.

aContact Ilya Serebriiskii at [email protected]. bAvailable from Invitrogen (http://www.invitrogen.com).




SphI

MluI

i or

C

E1 ol

mo

te

r

30 0

0

EcoRI 60

0

C

45

90

3900

R

36

0

00

33

0

270 0

30 0

0

PstI

SalI SphI

PstI

2µ

BspHI BstEII

MunI

240

0 om

00

F

pr

XhoI

21

TE

MluI

NotI

18

00

18 / G4 Km

150 0

5310 nt

NruI NcoI

SacII A DH t er m

pGBS10

Bgl II KpnI

12 0 0

42 0 0

0

HindIII

SacI ApaI

ne

cy t

HindIII

ge

00

te

pr o

cI

EcoRV

0

0 48

rm

BsrGI

A DH1

ApaLI

BsaI

o ri

ApaLI

Figure 20.7.5 Plasmid map of the 5310-nt basic dual bait vector pGBS10. This plasmid uses the strong constitutive ADH1 promoter to express bait proteins as fusions to the DNA binding protein cI. The plasmid contains the 2 µ origin of replication to allow propagation in yeast and the ColE1 origin of replication to allow propagation in E. coli. The same Tn903-encoded gene confers kanamycin resistance in E. coli and geneticin (G418) resistance in yeast. More maps and sequences are available on the Web at http://www.fccc.edu/research/labs/golemis/InteractionTrapInWork.html.

Table 20.7.3

LEU2/LYS2 Selection Strains

Name

Genotype

No. operators

Comment/description

SKY48

MATα, trp1, his3, ura3, lexAop-LEU2, cIop-Lys2

6 lexA, 3 cI

SKY191

MATα, trp1, his3, ura3, lexAop-LEU2, cIop-Lys2

2 lexA, 3 cI

SKY473

MATa, his3, leu2, trp1, ura3, lexAop-LEU2, cIop-LYS2

4 lexA, 3 cI

Provides a more stringent selection for interaction partners of cI-fused baits, and more sensitive LexA-responsive LEU2 reporter than the one in SKY191 Provides a more stringent LexA-responsive LEU2 reporter, and more sensitive cI-responsive LYS2 reporter than the one in SKY48. Mating partner for SKY strains; can be also used as a reporter strain itself. Sensitivity of LEU2 reporter is intermediate between sensitivity of LEU2 in SKY48 and SKY191. Sensitivity of LYS2 reporter is the same as sensitivity of LYS2 in SKY191.

Contact information I. Serebriiskiia

I. Serebriiskiia

I. Serebriiskiia

aE-mail at [email protected].


Supplement 60

HindIII 3 URA

BsaI ri

sA

700

00

14

0 91 0 84 0 0

cytC term

t er 2 8 0 0

KpnI EcoRI

77

350

00

ps

mo

pLacGus (pDR8) 11099 nt

cI o

pro

Kan R

XbaI

00

98

Co lE I

o

gu

0

0

70

00

t er

5600

SalI

mo

EcoRI HindIII BamHI

00

EcoRI

op s

49

i

pro

xA

42

or

00

2µ

Le

lac

Z

AatII

Figure 20.7.6 Plasmid map of the 11099-nt basic dual bait vector pLacGus, which is also known as pDR8. pLacGus is a dual reporter plasmid. Eight LexA operators upstream of the lacZ reporter make it the most sensitive of the lexA-responsive lacZ reporters. The cI-responsive gusA reporter (three cI operators) has a comparable sensitivity range. These two cassettes are separated by cytC terminator. The plasmid contains the URA3 selectable marker and the 2 µ origin of replication to allow selection and propagation in yeast, and the kanamycin resistance gene and the pUC origin of replication to allow selection and propagation in E. coli. More maps and sequences are available on the Web at http://www.fccc.edu/research/labs/golemis/InteractionTrapInWork.html.

Additional reagents and equipment for subcloning DNA fragments (UNIT 3.16) or alternative cloning strategies (e.g., in vivo recombination; Ma et al., 1987), lithium acetate transformation of yeast (UNIT 13.7), gal overlay assay (see Support Protocol 1), and immunoblotting and immunodetection (see Support Protocol 2) 1. Using standard techniques for subcloning DNA fragments (UNIT 3.16) or alternative cloning strategies (e.g., in vivo recombination), insert the DNA encoding one of the two proteins of interest into the polylinker of the LexA-fusion plasmid pMW103 (Figs. 20.7.3 and 20.7.4A) to make an in-frame protein fusion to LexA. Call this plasmid pMW103-Bait1. A number of modified versions of the plasmid exist which contain additional sites, altered reading frame, alternate antibiotic resistance markers, or include an in-frame nuclear-localization motif between the LexA protein and the polylinker (see Background Information in the Commentary and Table 20.7.1).


2. Similarly, clone the DNA encoding the second protein of interest into the polylinker of the cI-fusion plasmid pGBS10 (Figs. 20.7.4B and 20.7.5) to enable synthesis of an in-frame protein fusion to cI. Call this plasmid pGBS10-Bait2.



3. Lithium acetate transform yeast strain SKY191 (UNIT plasmids:

13.7)

using the following

pGBS10-Bait2 (step 2) pGBS10-Krit (positive control for activation) pGBS10-Krev (negative control for activation). Plate each transformation mixture on YPD plates containing 200 µg/ml geneticin (YPD G418), and incubate two days at 30°C to select for yeast colonies containing transformed plasmid. Incubating yeast in YPD liquid medium 6 hr to overnight at 30°C prior to plating significantly increases plating efficiency.

4. Transform SKY48 (UNIT 13.7) using the following combinations of LexA-fusion and lexAop-LacZ (pLacGus) plasmids: pMW103-Bait1 (step 1) + pLacGus pEG202-hsRPB7 + pLacGus (weak positive control for activation) pSH17-4 + pLacGus (strong positive control for activation) pEG202-Ras + pLacGus (negative control for activation). Transfer the transformations to 90- to 100-mm Glu/CM −Ura −His plates without G418 to select for yeast that contain both plasmids, and incubate 2 to 3 days at 30°C. If bait transformants either grow noticeably slower or in reduced number (compared to the controls), the bait protein is likely toxic to the yeast (see Critical Parameters and Troubleshooting, Table 20.7.11 and Table 20.7.12 for recommended modifications). Increased heterogeneity of yeast colonies compared to controls (e.g., a mix of large and small colonies) is also suggestive of bait toxicity. If only a small number of colonies are obtained even in controls, or colonies are not apparent within three to four days, the transformation efficiency is low. (While the transformation efficiency at this particular step is not paramount, it will be critical for the library transformation). In this case, all solutions, media, and conditions must be doublechecked or prepared fresh, and the transformation repeated. An efficient transformation should yield ∼1 × 104 transformants per microgram DNA (when two plasmids are being simultaneously transformed), and up to 1 × 105 for transformation with a single plasmid.

5. Replica plate to assess activation of cI-and LexA-fused baits: a. Add 50 to 75 µl sterile water to each well of one-half (6 × 8 wells) of a 96-well microtiter plate. Place an insert grid from a rack of 200-µl micropipet tips over the top of the microtiter plate and attach with tape. The grid should be elevated ∼1 cm from the plate and the holes in the insert grid should be aligned with the wells of the microtiter plate, as this will keep the tips in the upright position in the plate and allow their simultaneous removal. A convenient pipet-tip brand which accomplishes this is the Rainin RT series.

b. Pick six 1- to 2-mm-diameter yeast colonies from each of the transformation plates (steps 3 and 4) using a different sterile plastic 200-µl micropipet tip for each colony. Leave the tips supported in a near-vertical position by the insert grid until all the colonies have been picked. c. Swirl the plate gently to ensure the yeast are mixed into suspension. Remove the sealing tape and lift the insert grid along with all of the tips. d. Use a metal frogger or plastic replicator to plate yeast suspensions (each spoke leaves an ∼3-µl drop) on the following plates, marked for orientation:



Supplement 60

YPD containing 200 µg/ml G418 (master plate for cI-fused baits) Glu/CM −Lys Gal-Raff/CM −Lys Glu/CM −Ura −His (master plate for LexA-fused baits) Gal-Raff/CM −Ura −His −Leu Gal-Raff/CM −Ura −His (for Xgal overlay assay). e. Incubate the plates up to 4 days at 30°C, and save the YPD G418 and Glu/CM −Ura −His master plates at 4°C. Yeast containing cI-fused bait and controls should grow on YPD G418 plates, but should not grow on any −Ura −His plates. On Glu/CM −Lys and Gal-Raff/CM −Lys plates, yeast containing the positive control (pGBS10-Krit) should exhibit growth within 4 days, while yeast containing the negative control (pGBS-Krev) should not grow. Yeast containing LexA-fused bait and controls should not grow on either YPD G418 or CM −Lys plates, and should grow on Glu/CM −Ura −His plates. On Gal-Raff/CM −Ura −His −Leu plates, the strong positive control (pSH17-4 + pLacGus) should show detectable growth in 1 to 2 days, the weak positive control (pEG202-hsRPB7 + pLacGus) should be growing within 4 days, and the negative control (pEG202-Ras + pLacGus) should not grow. If the yeast containing the bait under test (pMW103-Bait1 + pLacGus) shows no growth in this period, it should be suitable for library screening. If it activates LEU2 reporter similar to the transformants from the pEG202-hsRPB7 reaction, it may be suitable, but is likely to have a high background in library screening (which may be slightly reduced by the use of a less sensitive strain). If it is similar to the pSH17-4 + pLacGus transformants, it must be modified before screening.

6. Approximately 24 to 30 hr after the plating, overlay the Gal-Raff/CM −Ura −His with Xgal agarose as described (see Support Protocol 1). Assess the ability of LexA-fused bait to activate transcription of LEU2 and LacZ reporters. In assessing the control transformants (step 4), strongly activating baits will be detectable as dark blue colonies in 20 to 60 min, while negative controls should be faint blue or white; an optimal bait would be either as white as the negative control or only a faint blue color. At this step, the ability of LexA-fused bait to activate transcription is tested on both LEU2 and LacZ reporters, while potential cI self-activation is only tested for LYS2 reporters. Auxotrophic reporters are, however, the most important for the library screening, because they allow the direct selection for interaction phenotype. In addition, there is normally a good correlation between activation of the two reporters, so it is very unlikely that the bait which is not activating LYS2 will significantly activate GusA. Therefore, if no activation is detected on −Lys plates, one should proceed further; if bait causes growth on −Lys plates, it should be modified. The ability of the cI-fused bait to activate GusA reporter will be tested in step 9 below.

7. Select from the master plate at least two primary transformants for each novel bait construct and perform immunoblot analysis as described (see Support Protocol 2). Establishing protein expression levels by immunoblot analysis is critical for several reasons. Proteins expressed at low levels, and apparently inactive in transcriptional activation assays, can be up-regulated to much higher levels under the auxotrophic selection and unexpectedly demonstrate a high background of transcriptional activation. Moreover, where proteins are proteolytically clipped, screens might inadvertently be performed with LexA fused only to the amino-terminal end of the larger intended bait. Adding positive controls for protein expression such as pGBS10-Krev (step 3) and pEG202Ras (step 4) is helpful.


8. Note which cI-bait colonies on the master plate express bait appropriately, and use two of these colonies to introduce LexA-fused bait and reporter plasmids. Use standard transformation procedure (UNIT 13.7), except grow the yeast in YPD liquid medium containing 200 µg/ml G418.



This transformation should combine in a single strain both of the baits and a double reporter plasmid—i.e., pMW103-Bait1 + pLacGus + pGBS10-Bait2 (clones 1 and 2).

In parallel, transform SKY191 expressing pGBS10-Krit and pGBS10-Krev to obtain pEG202-Ras + pLacGus + pGBS10-Krev and pEG202-hsRPB7 + pLacGus + pGBS10-Krit. 9. Plate each transformation mixture on Glu/CM −Ura −His G418 dropout plates, and maintain at 30°C for 2 to 4 days to select for yeast colonies containing desired plasmid combinations. 10. Replica-plate (step 5) twelve colonies of each transformant on the following set of plates: Glu/CM −Ura −His G418 (new master plate) Gal-Raff/CM −Ura −His G418 (Xgal overlay assay) Gal-Raff/CM −Ura −His G418 (Xgluc overlay assay) Gal-Raff/CM −Ura −His −Leu Gal-Raff/CM −Ura −His −Lys Incubate the plates up to 4 days at 30°C, and save the Glu/CM −Ura −His G418 master plate at 4°C. 11. Overlay with Xgal and Xgluc agarose as described (see Support Protocol 1) and analyze the results. Self-activation abilities are expected to be the same as in the characterization of separate baits.

12. Reconfirm expression of the baits by immunoblot (see Support Protocol 2). An investigator who is proficient in yeast transformations may wish to cotransform all baits and reporter plasmids into a single strain simultaneously. If the resultant strain behaves favorably, this single transformation can save time. It should be cautioned, however, that simultaneous transformation with three plasmids can be problematic and the authors recommend the step-wise strain construction and verification.

PREPARATION OF BAITS AND LIBRARY: TRANSFORMING AND CHARACTERIZING THE LIBRARY

BASIC PROTOCOL 2

A list of some of the libraries currently available for use with the Dual Bait system can be found at http://www.fccc.edu/research/labs/golemis/InteractionTrapInWork.html. Currently, the most convenient source of libraries suitable for the interaction trap is commercial, and can be viewed at the companies’ web-sites. The protocol outlined below describes the steps used to perform a screen that should saturate a cDNA library derived from a genome of mammalian complexity. Fewer plates will be required for screens with libraries derived from organisms with less complex genomes; therefore, the protocol should be scaled back accordingly. Currently, mating the library-pretransformed strain with the desired bait strain is recommended as the most convenient strategy. The main advantage of this approach is that if the investigator wishes to use the same library to screen multiple baits, only a single large-scale transformation is required, followed by relatively easy mating steps. This approach is also useful when analyzing a toxic bait, as yeast-expressing toxic proteins can be difficult to transform with high efficiency. Finally, direct transformation in the bait strain requires media not only selective for the library plasmid, but also maintaining selective pressure to keep both baits and reporter. Large-scale transformation plating on G-418 medium will make screening much more expensive.



Supplement 60

As outlined in Figure 20.7.7, it is recommended to perform the large-scale library transformation (plated on multiple 24 × 24–cm plates) in parallel with several small-scale control transformations (each plated on a single 90-mm plate). These small-scale controls are critical in calculating the frequency of false positives and will be useful as positive and negative controls during interactor characterization. Materials Yeast strains (Table 20.7.3), fresh: SKY473 YPD liquid media (UNIT 13.1) without G418 (APPENDIX 1K) H2O, sterile TE buffer (APPENDIX 2)/0.1 M lithium acetate Library DNA in pJG4-5 (Fig. 20.7.8) or pYesTrp (Table 20.7.4) Carrier DNA, freshly denatured Negative control plasmids (Table 20.7.4; optional): pJG4-5 or pYesTrp2, pJG4-5-Raf1, pJG4-5-Krit, and pYesTrp2-RalGDS 40% (w/v) PEG 4000/0.1 M lithium acetate/TE buffer, pH 7.5 Dimethyl sulfoxide (DMSO) 90-mm and 24 × 24–cm Glu/CM −Trp plates (UNIT 13.1) TE buffer (APPENDIX 2), sterile (optional) Glycerol solution: 65% (w/v) sterile glycerol/0.1 M MgSO4/25 mM Tris⋅Cl, pH 8.0 (APPENDIX 2) Orbital shaker, 30°C 50-ml conical tubes, sterile 42°C heat block 3- to 4-mm glass balls, sterile (Thomas Scientific or Fisher) Additional reagents and equipment for lithium acetate transformation of yeast (UNIT 13.7) Prepare competent yeast 1. Inoculate a colony of SKY473 in ∼20 ml liquid YPD medium and grow overnight on an orbital shaker at 30°C. It is important to use fresh yeast (i.e., thawed from −70°C and streaked to a single colony less than ∼7 days previously) and maintain sterile conditions throughout all subsequent procedures.

2. Dilute the overnight culture into ∼300 ml YPD liquid medium such that the diluted culture has an OD600 of ∼0.15. Incubate at 30°C on an orbital shaker until the culture has reached an OD600 of ∼0.50 to 0.7. 3. Transfer the culture to six sterile 50-ml conical tubes, and centrifuge 5 min at 1000 to 1500 × g, room temperature. Gently resuspend the pellets in ∼5 ml sterile water each, and combine all slurries into one of the conical tubes. Add sterile water to 50 ml and mix. Perform transformations 4. Centrifuge cells again 5 min at 1000 to 1500 × g, room temperature. Decant water and resuspend yeast in 1.5 ml TE buffer/0.1 M lithium acetate (UNIT 13.7). 5. Mix 30 µg library DNA in pJG4-5 or pYesTrp with 1.5 mg freshly denatured carrier DNA in a microcentrifuge tube and add this mixture to the yeast. Mix gently and aliquot ∼60 µl DNA/yeast suspension into each of 30 microcentrifuge tubes. Two-Hybrid Dual Bait System

6. Optional: Use aliquots of competent yeast from step 4 to transform control plasmids—i.e., empty library plasmid (pJG4-5 or pYesTrp2), pJG4-5-Raf1, pJG4-5-



SKY473

prepare competent yeast

large-scale transformation

small-scale transformations pJG4-5Raf1

pYesTrp2RalGDS

pJG4-5Krit1

pJG4-5 or pYesTrp2

one 90-mm Glu –Trp plate each

library in pJG4-5 or pYesTrp2

thirty large 24 x 24–cm Glu –Trp plates

collect cells

collect cells aliquot

SKY191 pEG202-Ras + pLacGus + pGBS10-Krev1

SKY191 pMW103-Bait1 + pLacGus + pGBS10-Bait2

large-scale matings

small-scale matings collect cells titer on Glu/CM –Ura –His –Trp streak on selection G418 Glu/CM –Ura –His –Trp G418 selection one plate ten plates at 1×106 cells/plate one plate at 1×106 cells/plate ten plates at 1×107 cells/plate pos. controls at 1×107 cells/plate

neg. control test clones

Figure 20.7.7 Detailed library screening flow chart. See Basic Protocols 2 and 3 for details.



Supplement 60

KpnI SacI

00

400

0

EcoRI XhoI

fusion cassette

52 0

0

56

0

pU C

60

0

160 0

48 0 0

pJG4-5

20 0

4

0

0 40

A m p ic llin R

BsaI

kb

e on

GA promoL 1 ter

H at or A D m in r Te 00 12

ba c

Afl III

BclI

SpeI

0

0

O

ri

0

µ

28

40

00

3200

36 0

0

0 TR

BamHI SalI NotI

24

AatII

SphI

2

P1

fusion cassette: NLS

ATG

B42 domain

EcoRI

HA Tag

XhoI

Figure 20.7.8 Plasmid map of the basic dual bait vector pJG4-5. This plasmid expresses cDNAs or other coding sequences inserted into the unique EcoRI and XhoI sites as translational fusion to a cassette consisting of the SV40 nuclear localization sequence, the acid blob B42, and the hemagglutinin (HA) epitope tag. Expression of sequences is under the control of the GAL1 galactose-inducible promoter. The plasmid contains the TRP1 selectable marker and the 2 µ origin of replication to allow selection and propagation in yeast, and the ampicillin resistance gene and pUC origin of replication to allow selection and propagation in E. coli. The fusion cassette is shown at the bottom. A detailed map of the AD-fusion polylinker is given in Figure 20.7.7. More maps and sequences are available on the Web at http://www.fccc.edu/research/labs/golemis/Interaction TrapInWork.html.

Krit1, and pYesTrp2-RalGDS—into SKY473. Transfer to a 100-mm Glu/CM − Trp plate and collect the transformed cells as for the library. The control strain with an empty library plasmid can be safely reamplified in liquid medium. When using a new bait strain, it is recommended to set up a parallel mating with negative and positive controls as outlined in Figure 20.7.7. The negative control strain is the same strain used for the library (e.g., SKY473), but containing the library vector with no cDNA insert. Positive control strains are also very helpful and will be later used in mating with a positive control bait strain (e.g., pEG202-Ras + pLacGus + pGBS10-Krev). Two-Hybrid Dual Bait System



Table 20.7.4

Activation Domain Fusion Plasmids and Controls

Plasmid name

Yeast selection

E. coli selection Comment/description

pJG4-5

TRP1

ApR

pJLo

TRP1

ApR

pJG4-5I

TRP1

ApR

pYesTrp

TRP1

ApR

pNB42 series

TRP1

ApR

pMW102

TRP1

KmR

pMW104

TRP1

CmR

pCGB42/p2GB42

G418R

KmR

Control activation domain–fusions pJG4-5-Raf TRP1 pYesTrp-RalGDS TRP1

ApR ApR

pJG4-5-Krit

ApR

TRP1

Library construction plasmid; GAL1 promoter provides efficient expression of a gene fused to a cassette consisting of nuclear localization sequence, transcriptional activation domain, and HA epitope tag A derivative of pJG4-5 that has a lower copy number (CEN/ARS ori) A derivative of pJG4-5 that can be integrated into yeast TRP1 gene after digestion with Bsu36I; designed to study interactions that occur physiologically at low protein concentrations (in combination with pEE202I) GAL1 promoter expresses nuclear localization domain, transcriptional activation domain, V5 epitope tag, multiple cloning sites; contains f1 ori and T7 promoter/flanking site. Used to express cDNA libraries Allow fusion to the N terminus of an AD, leaving N-terminal residues of Prey unblocked; various multiple cloning sites. No libraries yet available Same as pJG4-5, but with altered antibiotic resistance markers; no libraries yet available Same as pJG4-5, but with altered antibiotic resistance markers; no libraries yet available The same Tn903-encoded gene confers kanamycin resistance in E. coli and geneticin (G418) resistance in yeast; both high- and low copy number versions available. Multiple cloning site.

Contact information Origenea, MoBiTecb

R. Hopkinsc RBd

Invitrogene

M. Brownf

RBd RBd J. Huangg

A positive control for interaction with Ras I. Serebriiskiih A positive control for interaction with Ras and I. Serebriiskiih Krev A positive control for interaction with Krev I. Serebriiskiih

aSee the Origene homepage at http://www.origene.com. bSee the MoBiTec Website at http://www.mobitec-germany.com. cE-mail at [email protected]. dE-mail at [email protected]. eSee the Invitrogen Website at http://www.invitrogen.com. fSee Brown and MacGillivray (1997). gFor more information, refer to Huang and Schreiber (1997). hE-mail at [email protected].



Supplement 60

7. To each tube (steps 5 and 6), add 300 µl sterile 40% (w/v) PEG 4000/0.1 M lithium acetate/TE buffer pH, 7.5. Gently invert the tubes several times (do not vortex) and incubate 30 to 60 min at 30°C. 8. To each tube, add 40 µl DMSO and mix by inversion. Place tubes in a 42°C heat block for 10 min. 9. Evenly spread the contents of each tube onto a 24 × 24–cm Glu/CM −Trp plate, using sterile 3- to 4-mm glass balls. Invert plates and incubate at 30°C until colonies appear (usually 2 to 4 days). It is acceptable and efficient to keep the glass balls on the lids while incubating the plates; they will be needed to harvest the library transformants (see below). One to two dozen glass balls is sufficient.

10. Select a couple of representative transformation plates and draw a 23 × 23–mm square (1% of plate bottom surface) over an average density spot. Count colonies in each grid section and recalculate for the whole transformation. An efficient transformation done by this protocol should yield ∼20,000 to 40,000 colonies per large plate, and represent ∼1 × 105 transformants per microgram library DNA. Doing transformations in small aliquots helps reduce the likelihood of contamination, and for reasons that are not clear, provides significantly better transformation efficiency than scaled-up versions. Note, it is not recommended to use excess transforming library DNA per aliquot competent yeast cells, as each competent cell may take up multiple library plasmids, complicating subsequent analysis. Under the conditions described here, 10 days old. Otherwise, these yeast must be replated or the whole transformation process repeated. In any case, single colonies from the fresh bait strains have to be characterized by immunoblotting to confirm the expression of bait proteins. A major strength of this protocol is that it will identify redundant clones prior to plasmid isolation and bacterial transformation, which in some cases greatly reduces the amount of work required. However, accurate records should be maintained as to how many of each class of cDNA are obtained; and if any ambiguity is present as to whether a particular cDNA is part of a set or unique, investigators should err on the side of caution. Materials Glu/CM −Ura −His −Trp G418 master plate (see Basic Protocol 3) β-glucuronidase solution: dilute crude β-glucuronidase type HP-2 from H. pomatia (Sigma) 1:50 in 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2)/10 mM EDTA; prepare fresh Library plasmid specific primers—e.g., for JG4-5: Forward primer (FP1): 5′-CTG AGT GGA GAT GCC TCC Reverse primer (FP2): 5′ CTG GCA AGG TAG ACA AGC CG HaeIII (UNIT 3.1) SKY191 containing pMW103-Bait1 + pLacGus + pGBS10-Bait2 (see Basic Protocol 1, steps 8 to 12) SKY 191 containing pEG202-Ras + pLacGus + pGBS10-Krev (see Basic Protocol 1, steps 8 to 12; optional) Raf1 and Krit1 PCR fragment (see Support Protocol; optional) Glu/CM −Ura −His −Trp G418 dropout plates (UNIT 13.1) Metal frogger (e.g., Dankar Scientific) or plastic replicator (i.e., Bel-Blotter; Bel-Art Products or Fisher) 96-well microtiter plate Horizontal shaker, 37°C Tape 150- to 212-µm glass beads (e.g., Sigma G-1145) Vortex with flat-top surface Additional reagents and equipment for PCR (UNIT 15.1), restriction endonuclease digestion (UNIT 3.1), agarose gel electrophoresis (UNIT 2.5A), purification of DNA fragments (UNIT 2.6), lithium acetate transformation of yeast (UNIT 3.17), sequencing (Chapter 7), rapid miniprep isolation of yeast DNA (UNIT 3.6), and miniprep isolation of bacterial DNA (UNIT 1.6).



Supplement 63

Rapid screen for interaction trap positives 1. Starting from a Glu/CM −Ura −His −Trp G418 master plate, use a metal frogger or plastic replicator to resuspend yeast in 25 µl β-glucuronidase solution in a 96-well microtiter plate (also see Basic Protocol 3, step 14). Seal the wells using tape, and incubate 1.5 to 3.5 hr on a horizontal shaker at 37°C. Alternatively, pipet tips can be used to transfer yeast in the wells (similar to replica plating; see Basic Protocol 1, step 5). This option is especially useful when the initial set of candidates was spread over a few master plates, and only a fraction of them has to be tested further. Do not take more than approximately the volume of one middle-sized yeast colony; otherwise, quality of isolated DNA may suffer. The master plate does not need to be absolutely fresh: plates that have been stored for a week at 4°C have been successfully used.

2. Remove the tape, add ∼25 µl (i.e., a small scoop) of 150- to 212-µm glass balls to each well, and seal again. Attach the microtiter plate to a vortex with a flat top surface (e.g., using rubber bands) and mix vigorously for 5 min. 3. Add 100 µl sterile water to each well. Take 1 to 2 µl as a template for each PCR reaction. Reseal the plate with tape and keep the remainder frozen at −70°C. In many cases, PCR products can be obtained directly from the yeast colonies even without enzymatic treatment (e.g., by introducing a 10-min 94°C step at the beginning of the PCR program). However, the crude yeast lysate obtained in this protocol (step 2) can also be used as a source of plasmid DNA (use 1 to 2 ìl for electroporation into E. coli), making it worth preparing this simple lysate.

4. Prepare master mix (25 to 30 µl/reaction) as described in UNIT 15.1 using library-specific primers. For example, use primer FP1 and FP2 for the JG4-5-based library or commercially available primers (Invitrogen) for pYesTrp2-based libraries.

5. Perform a PCR amplification (UNIT 15.1) in 25 to 30 µl using the following program: Initial step: 35 cycles:

2 min 45 sec 45 sec 5 sec

94°C 94°C 56°C 72°C

(denaturation) (denaturation) (annealing) (extension).

Modified versions of this protocol with extended elongation times were also found to work. The variant given above has been shown to amplify 1.8-kb fragments well. To interpret the results of the PCR, it is helpful to have the following ∼1 ng control templates: empty library plasmid, yeast from the positive control colonies treated in parallel with experimental clones, and the same amounts of library plasmid and positive control yeast, mixed together. Interpretation of possible PCR outcomes is summarized in Table 20.7.7. If the library being screened is based on JG4-5 plasmid, only clones containing Raf1 and Krit plasmids would produce products. For a pYesTrp-based library, use a RalGDS clone as positive control.

6. Load 2/3 (~20 µl) of the PCR reaction onto a 0.7% agarose gel (UNIT 2.5A). Identify fragments which appear to be of the same size. Based on insert sizes, group obtained interactors in families (i.e., identify potential multiple isolates of the same cDNAs). Store gel in the refrigerator until ready to isolate fragments.


7. While the gel with the uncut PCR products is running, perform a restriction digest of the remainder (~10 µl) of the PCR product with 15 U HaeIII in a total volume of 20 µl. Based on classification of undigested PCR products (step 6), rearrange the loading



Table 20.7.7

Analysis of PCR Reaction

Template Empty Control Empty library + library yeast control yeast plasmid

Clone of interest

−

−

−

−

+ + +

− − ±

+ − ±

− − ±

Possible cause

Solution

Bad master mix, wrong settings, or a faulty amplifier were used Not enough template Lysed yeast inhibited PCR Uneven template load/ digestion

Double check reaction and repeat Add more template/improved Add less template Adjust template, PCR from obtained bands again

order of HaeIII-digested samples in such a way that the digests of the PCR products thought to be identical are run side-by-side. Load the HaeIII digestion products on a 1.0% agarose gel (UNIT 2.5A) and electrophoresis to optimize resolution of DNA products in the 200 to 1000 bp range. HaeIII is recommended only because it tolerates the presence of PCR buffer well. If the PCR product is purified, any four-base cutter can be used. This technique will generally yield distinctive and unambiguous groups of inserts, confirming whether multiple isolates of a small number of cDNAs have been obtained. As mentioned above, sometimes a single yeast will contain two or more different library plasmids. If this happens it is typically revealed by PCR. Thus, after bacterial transformation, and increased number of clones should be checked to avoid the loss of the “real” interactor.

8. Purify undigested fragments from the agarose gel using standard techniques (UNIT 2.6). In cases where a very large number of isolates have been obtained from a small number of cDNAs, the investigator may choose to sequence the PCR product directly. PCR products obtained from this step will also be used for reconfirmation of an interaction. To get enough PCR products, reamplify if necessary. Only the forward primer, FP1, works reliably in sequencing PCR fragments, while the reverse primer will only work in sequencing from purified plasmid (Chapter 7). In general, the TA-rich nature of the ADH terminator sequences downstream of the polylinker in the pJG4-5 vector makes it difficult to design high quality primers in this region.

Perform specificity testing 9. Lithium acetate transform (UNIT 3.17) combinations of digested library plasmid and selected PCR products (optionally including Raf1 and Krit1; see Support Protocol 3) into the naïve bait strain SKY191 (pMW103-Bait1 + pLacGus + pGBS10-Bait2). Optionally, transform the same combinations in parallel into the control bait strain SKY191 (pEG202-Ras + pLacGus + pGBS1-Krev). 10. Plate each transformation mix on Glu/CM −Ura −His −Trp G418 dropout plates and incubate at 30°C until colonies grow (2 to 3 days). 11. Prepare a master plate for each library plasmid being tested. Each plate should contain at least ten colonies of the transformed PCR-insert/digested plasmid into each of the bait strains. Analysis of Protein Interactions


Supplement 60

12. Test for the activity of colorimetric and auxotrophic reporters as described (see Basic Protocol 3, steps 14 and 15). At least ten to twelve colonies from each transformation should be tested. True positives should show a LEU+ LacZ+ phenotype with pMW103-Bait1 + pLacGus + pGBS10-Bait2, but not with pEG202-Ras + pLacGus + pGBS10-Krev. Clones transformed with Raf control PCR will provide both positive and negative controls: pMW103-Bait1 + pLacGus + pGBS10-Bait2 should be negative while pEG202-Ras + pLacGus + pGBS10Krev should be positive when assayed for β-galactosidase and growth on −Leu plates, but negative for β-glucuronidase activity and growth on −Lys plates. Vice versa, clones of pEG202-Ras + pLacGus + pGBS10-Krev transformed with Krit1 control PCR products should be blue on Xgluc, but not Xgal, and grow on −Lys, but not on −Leu dropout plates. Again, pMW103-Bait1 + pLacGus + pGBS10-Bait2 should be completely negative. The fraction of the correct clones in positive controls is indicative of the efficiency of recombination in this particular experiment. Normally, it should be between 85% and 95%.

13. Proceed with sequencing (Chapter 7) and biological characterization. 14. If recovery of yeast plasmids for archival purposes is desired, isolate yeast plasmids using tradition phenol-chloroform technique (UNIT 3.6) or a commercially available kit. Transform selected positives into E. coli, using 1 to 2 µl β-glucuronidase-treated frozen yeast (see Basic Protocol 3). Use of electroporation is highly recommended. An alternative protocol can be used in case PCR technology is not readily available, or in case of failure to obtain a specific PCR product using the library vector primers. It is based on plasmid isolation, its amplification in E. coli, and then transformation back into yeast (see Alternate Protocol). A database of common false positives has been compiled and made available on the World Wide Web, at http://www.fccc.edu/research/labs/golemis/interactiontrapinwork.html. Particularly for cDNAs only isolated a single time, or which do not appear to make biological sense in the context of the starting bait, it may be helpful to consult the database to make sure the clone has not been reported by multiple additional groups. ALTERNATE PROTOCOL

SCREENING FOR INTERACTION TRAP POSITIVES BY YEAST PLASMID RECOVERY This protocol can be used instead of the rapid PCR screen (see Basic Protocol 4). See outline in Figure 20.7.9 for additional details. For materials see Basic Protocol 4. Also see UNIT 1.6 for additional reagents and equipment for preparation of plasmid DNA. 1. Isolate plasmid DNA from selected yeast clones (see Basic Protocol 4, steps 1 and 2) and transform into E. coli. A transformation protocol described in UNIT 13.11 can be used. Alternatively, a number of kits for yeast minipreps are commercially available (e.g., from Clontech and others). Some companies (e.g., Qbiogene; http://www.qbiogene.com/services/two-hybrid.html) will isolate plasmid from the yeast cells, transform, and amplify the plasmid in E. coli to produce a sequencing template.

2. Select at least two bacterial clones for each yeast clone, and prepare a small quantity of plasmid DNA (UNIT 1.6) from each bacterial clone. If using a specialized ApR LexA-fusion bait plasmid in combination with ApR library plasmid, it will be necessary to select specifically for transformants containing a library plasmid by the ability of the yeast TRP1 gene to complement the E. coli trpC mutation in a KC8 strain (see UNIT 20.1). Two-Hybrid Dual Bait System



3. Analyze the isolated plasmids by restriction digest (UNIT 3.1), to identify redundant clones and to confirm that both isolates from the same yeast clone are identical. 4. Follow the instructions given above (see Basic Protocol 4, steps 9 to 14) as described, except transform with purified library plasmids, instead of a mixture of PCR product and digested library vector. COLORIMETRIC ASSAY OF â-GALACTOSIDASE AND â-GLUCURONIDASE ACTIVITY BY AGAROSE OVERLAY

SUPPORT PROTOCOL 1

There are a variety of the standard assays used to assess β-galactosidase and β-glucuronidase activity in yeast, including colony overlay, colony lift (filter) assays, and growth on plates containing Xgal or Xgluc. Although popular, assays in which yeast are grown in the presence of substrate yield sometimes erratic results. One of the reasons for this is that expression of some combinations of bait and prey may result in greater permeability of yeast cells, reflected by the increased degree of uptake of Xgal from the surrounding media (yeast seed density may also play a role). In addition, Xgal plates are buffered to approach pH 7.0 (optimal for β-galactosidase activity), while yeast grow optimally on media of pH 5.5. Thus, yeast grow suboptimally under these conditions. Overlay and colony lift (filter) assays avoid these issues since yeast are first grown under optimal conditions and then exposed to substrates following permeabilization of yeast. Of these techniques, the overlay technique provides the most consistent results in the hands of the authors, in part because of the avoidance of issues such as the sometimes uneven transfer of colonies to filters. Xgal and Xgluc overlay plates are quick and extremely sensitive, and normally will give a strong signal within an hour. Materials Plates containing spotted yeast colonies to be assayed Chloroform Xgal agarose: prepare 1% low-melting agarose in 100 mM KHPO4, pH 7.0, and heat to boil; cool to ∼60°C and add Xgal to 0.25 mg/ml; prepare fresh Xgluc agarose: prepare as described for Xgal agarose, except replace Xgal with Xgluc 1. Gently overlay each plate containing spotted yeast colonies to be assayed with a minimal amount of chloroform (CHCl3), pipetting slowly in from the side so as not to smear colonies. Leave colonies completely submerged for 5 min, but do not cover the dish with the lid. CAUTION: CHCl3 is quite toxic and should neither be inhaled nor come into contact with skin. Wear gloves and work in a chemical hood. Try to minimize the amount of CHCl3 used, just enough to cover the colonies. Try to avoid contact with the walls of the plate, as the plastic will melt.

2. Optional: Briefly rinse the plates with another ~5 ml chloroform, drain, and allow to dry uncovered for another 5 min at 37°C or for 10 min in a hood. 3. Overlay the plate with ~10 ml Xgal or Xgluc agarose, making sure that all yeast spots are completely covered. Plates will be chilled after CHCl3 evaporation, so it will be difficult to spread 50-fold over background, indicating a suitable dynamic range of signal for most applications. The pLacGus plasmid is an example of a plasmid which combines both colorimetric reporters on a single backbone. It utilizes eight lexA operators to direct LacZ and three cI operators to direct GusA, and provides a well balanced readout for many general applications. Other plasmids, combining LacZ and GusA reporters of various sensitivity, can be easily constructed.


Auxotrophic reporter strains Ideally, an auxotrophic reporter yeast strain will be suitable for genomic level applications (e.g., compatible with interaction mating) and possess modulated sensitivity (e.g., variable

operator number). To date, three strains with an expanded range of response for lexAop-LEU2 are available. These strains demonstrate high (SKY48), moderate (SKY473), or low (SKY191) sensitivity to a LexA-fused activator, based on the number of lexA operators present. Although all three strains contain the same number of cI operators, SKY473 and SKY191 are fortuitously more sensitive to activation by a cI-fusion. Of these strains, SKY473 has the best balance of sensitivity to both LexA- and cI-fusions. It has the mating type opposite to that of other strains, and thus can be used in combination with them to set up interaction mating. The dual bait system has been extensively tested in a variety of applications (see Table 20.7.10), and proved to be robust and versatile (Serebriiskii et al., 2002). It was first validated in analysis of the selective interactions of Raf, Ral-GDS, and Krit1 (AD-fused components) with related Ras family GTPases—i.e., Ras (cI-fused Bait 2) and Krev (LexA-fused Bait 1). Raf interacts preferentially with Ras, Krit1 with Krev, and RalGDS with both. The dual bait system successfully identifies correct interactions among these proteins, and these reagents are now used as a set of controls in the described protocol. The authors have also documented the ability of the system to sort highaffinity interactors for either LexA- or cI-fused baits from a pool of low-affinity interactors, and to identify mutants in an activation domainfused prey that selectively alter affinity for discrete LexA- versus cI-fused partners (Serebriiskii et al., 1999; Reeder et al., 2001). The new reagents described here have been used effectively to study interactions between known sets of proteins (E. Benevolenskaya and W.Kaelin, pers. comm.), and have been utilized in a number of library screening applications. In these library screens, the two baits used to screen either a bait of interest and a nonspecific control (E. Kotova, pers. comm.), or a wild-type and a mutated form of the same bait (Serebriiskii et al., 2002). Both approaches yielded specific interactive partners that were either known to be biological or have been validated (by GST pulldown or co-immunoprecipitation). Baits used in these studies range from cytoplasmic serine/threonine regulatory kinases, morphoregulatory GTPases (e.g., Cdc42, Rac), or cell cycle regulators (e.g., Cdc6, wild-type/mutant RB protein). Screens have been performed by direct library transformation, mating, or both. Significantly, substan-



Table 20.7.11 Possible Modifications to Enhance LexA-Fused Bait Performance in Specific Applications

Problema Responseb

Not transported to Strongly Weakly the nucleus or activating activating low expression level

Continuous expression of LexA-fusion is toxic to yeast

Bait protein requires unblocked amino-terminal end for function

Bait protein expressed at high levels, unstable or interacts promiscuously

Truncate/modify baitc

+

+

−

+

−

+?

Use more stringent strain/reporter combinationd

−

+

−

−

−

−

Fuse to nuclear localization sequence pJK202

−

−

+

−

−

−

Put LexA-fused protein under GAL1-inducible promoter pGildae

−

−

−

+

−

+?

Fuse LexA to the carboxy terminus of the bait pNLexAf Integrate bait, reduce concentration pEG202Ig

−

+?

−

+?

+

+?

−

+?

−

+?

−

+

aPlus sign (+) indicates the response usually helps; plus sign followed by question mark (+?) indicates the response may help, and a minus sign (−)

indicates the response will not help. bAll of the alternative LexA bait expression vectors remain on an AmpR selection for bacteria. If using them as is, the investigator may need to use KC8 bacteria to isolate the library plasmid after a library screen. cIt may be necessary to subdivide bait into two or three overlapping constructs, each of which must be tested independently. dUse of very stringent interaction strains may eliminate detection of biologically relevant interactions. eCan no longer use GAL-dependence of reporter phenotype to indicate cDNA-dependent interaction. fGenerally, LexA poorly tolerates attachment of the N-terminal fusion domain. Only ∼60% of constructs are expressed correctly. gReduced bait protein concentration may lead to reduced assay sensitivity.

tially overlapping sets of interactors have been obtained both through transformation and interaction mating (B. Spodik and T. Coleman, pers. comm.), supporting the comparable nature of these approaches. Other work has indicated that these reagents will be useful in pharmaceutical screening applications, as the presence of two baits provides a significant shield against artifactual results based on potential global dysregulation of transcription (Serebriiskii et al., 2002). As with other new technologies, and particularly gene discovery screens, it will be several years until the identified interactors and general use of the system will be completely established. However, at present, the assembled reagents appear to offer an efficient, manipulable set of tools to analyze specific protein-protein interactions.

Critical Parameters and Troubleshooting To make an ideal bait, a protein of interest should satisfy the following four criteria: 1. It should have little or no intrinsic ability to activate transcription. 2. It should be expressed at reasonably high levels. 3. It should be nontoxic for yeast. 4. It must be able to enter the yeast nucleus and bind DNA. The first three properties are analyzed in the first protocol (see Basic Protocol 1), so that problems can be identified and potentially corrected before screening. Suggested modifications are summarized in Table 20.7.11 and Table 20.7.12. Currently, there are more options to enhance the performance of LexAfused baits, than the cI-fused. Therefore, a pro-



Supplement 60

Table 20.7.12

Possible Modifications to Enhance cI-fused bait Performance in Specific Applications

Problema Response

Strongly Weakly Continuous expression of activating activating cI-fusion is toxic to yeast

Bait protein expressed at high levels or interacts promiscuously

Truncate or modify baitb

+

+?

+?

−

Use more stringent strain/reporter combinationc

−

+

−

−

Put cI-fused protein under GAL1inducible promoter pGMS12d Use low-level expression vector pGBS9e

−

−

+

+?

−

−

+?

+

aPlus sign (+) indicates the response usually helps, plus sign followed by question mark (+?) indicates the response may help, and a minus sign (−)

indicates the response will not help. bIt may be necessary to subdivide bait into two or three overlapping constructs, each of which must be tested independently. cUse of very stringent interaction strains may eliminate detection of biologically relevant interactions. dCan no longer use GAL-dependence of reporter phenotype to indicate cDNA-dependent interaction. eReduced bait protein concentration may lead to reduced assay sensitivity.


tein to be used as a primary bait should be fused to LexA. The inability of the bait to go to the nucleus cannot be detected as reliably as its activation properties. Although the repression assay (described in UNIT 20.1) was developed to assess this property, the current trend is to proceed with the screening even if the results of this assay are negative. In addition, this assay is not available for cI-fused baits, and therefore it is omitted in this protocol. However, a researcher should be aware of this possibility, and avoid (or truncate) proteins that have extensive transmembrane domains. In screening the library, it is very important to obtain it for the tissue source in which the bait protein(s) are known to be biologically relevant. A second prerequisite is to screen a sufficient number of clones to fully represent the library. This latter goal is achieved in two steps. First, it requires an efficient transformation of the library in S. cerevisiae. In contrast to E. coli, the maximum efficiency of transformation for yeast is up to 1× 105 cfu/µg DNA, so it is extremely important to optimize the transformation procedure prior to attempting library transformation. At least 1 × 106 clones should be obtained for a cDNA library derived from a genome of mammalian complexity. A second step requires an efficient mating of the pretransformed library strain with the bait strain. The number of diploids to obtain depends on the number of clones in the library and the number of primary yeast transformants. The objective is to screen a number of transfor-

mants at least equivalent to the size of the library; if the size of the library was larger than the number of transformants, then all of the yeast transformants should be screened. Under the optimal conditions, ∼10% of the library strain cells will mate with the bait cells, forming diploids. Thus, a complete screen of 1 × 107 library transformants requires a single mating with about 1 × 108 cfu pretransformed library strain (and twice as much bait strain). Colony formation on the selection plates is one of the most variable parts of the screening. For strong interactors, colonies will grow up in 2 days on −Leu plates or 3 days on −Lys plates. However, new colonies will continue to appear for up to one week if the plates are left in the incubator. Growth speed depends on many factors, including the expression levels of the library-encoded protein and the strength of interaction, so not necessarily the most rapidly growing colonies are more biologically relevant to the bait protein. Dual bait provides a “built-in” control for the specificity of interaction. In addition, second confirmation of positives described in this protocol will test the isolated preys against two more unrelated baits. However, it is generally informative to retest positive clones on more than one bait protein. If possible, library-derived clones should be tested against a set of proteins ranging from almost identical to the original bait to clearly unrelated proteins. If the bait used in the screen showed weak transcriptional activity, it is expected to have greater difficulties with false positive background. In



this case it is advisable to choose a nonspecific control bait that can weakly activate transcription on its own in order to match the background level of transcription activation.

Anticipated Results Depending on the properties of the bait protein(s) and the type of selection applied, screening a library of ∼1 × 106 primary transformants may yield from zero to hundreds of specific interactors.

Time Considerations In general, it is good practice to move quickly through the steps of the described protocol. While plasmids will be retained in yeast for a long time, expressed protein levels will gradually drop, making the outcome of the whole screening less predictable. Under the favorable conditions, it will take about 3 weeks to complete the screening, as is outlined in Figure 20.7.2.

Literature Cited Brown, M.A. and MacGillivray, R.T. 1997. Vectors for expressing proteins at the amino-terminus of an activation domain for use in the yeast two-hybrid system. Anal. Biochem. 247:451-452 Estojak, J., Brent, R., and Golemis, E.A. 1995. Correlation of two-hybrid affinity data with in vitro measurements. Mol. Cell. Biol. 15:58205829. Fields, S. and Song, O. 1989. A novel genetic system to detect protein-protein interaction. Nature 340:245-246. Fujita, A., Tonouchi, A., Hiroko, T., Inose, F., Nagashima, T., Satoh, R., and Tanaka, S. 1999. Hsl7p, a negative regulator of Ste20p protein kinase in the Saccharomyces cerevisiae filamentous growth-signaling pathway. Proc. Natl. Acad. Sci. U.S.A. 96:8522-8527. Gyuris, J., Golemis, E.A., Chertkov, H., and Brent, R. 1993. Cdi1, a human G1 and S phase protein phosphatase that associates with Cdk2. Cell 75:791-803. Huang, J. and Schreiber, S.L. 1997. A yeast genetic system for selecting small molecule inhibitors of protein-protein interactions in nanodroplets. Proc. Natl. Acad. Sci. U.S.A. 94:13396-13401. Inouye, C., Dhillon, N., Durfee, T., Zambryski, P.C., and Thorner, J. 1997. Mutational analysis of STE5 in the yeast Saccharomyces cerevisiae: Application of a differential interaction trap assay for examining protein-protein interactions. Genetics 147:479-492.

Jiang, R. and Carlson, M. 1996. Glucose regulates protein interactions within the yeast SNF1 protein kinase complex. Genes Dev. 10:3105-3115. Ma, H., Kunes, S., Schatz, P.J., and Botstein, D. 1987. Plasmid construction by homologous recombination in yeast. Gene 58:201-216. Petermann, R., Mossier, B.M., Aryee, D.N., and Kovar, H. 1998. A recombination-based method to rapidly assess specificity of two-hybrid clones in yeast. Nucl. Acids Res. 26: 2252-2253. Reeder, M.K., Serebriiskii, I.G., Golemis, E.A., and Chernoff, J. 2001. Analysis of small GTPase signaling pathways using Pak1 mutants that selectively couple to Cdc42. J. Biol. Chem. 276:40606-40613. Serebriiskii, I., Khazak, V., and Golemis, E.A. 1999. A two-hybrid dual bait system to discriminate specificity of protein interactions. J. Biol. Chem. 274:17080-17087. Serebriiskii, I., Estojak, J., Berman, M., and Golemis, E.A. 2000. Approaches to detecting two-hybrid false positives. Biotechniques 28:328-336. Serebriiskii, I.G., Khazak, V., and Golemis, E.A. 2001. Redefinition of the yeast two-hybrid system in dialogue with changing priorities in biological research. BioTechniques 30:634-655. Serebriiskii, I.G., Mitina, O., Pugacheva, E., Benevolenskaya, E., Kotova, E., Toby, G.G., Khazak, V., Kaelin, W.G., Chernoff, J. and Golemis, E.A. 2002. Detection of peptides, proteins, and drugs that selectively interact with protein targets. Genome Res. In press. Xu, C.W., Mendelsohn, A.R. and Brent, R. 1997. Cells that register logical relationships among proteins. Proc. Natl. Acad. Sci. U.S.A. 94:1247312478.

Internet Resource http://www.fccc.edu/research/labs/golemis/Interac tionTrapInWork.html Analysis of the two-hybrid usage; database of false positives detected in two-hybrid experiments; database of available libraries, strains and specialized vectors (maps, sequences) for Interaction Trap/Dual Bait system; protocols related to the twohybrid screening and β-galactosidase assays in yeast

Contributed by Elena Kotova, Ilya Serebriiskii, and Thomas Coleman Fox Chase Cancer Center Philadelphia, Pennsylvania



Supplement 60

Interaction Trap/Two-Hybrid System to Identify Loss-of-Interaction Mutant Proteins

UNIT 20.8

To understand how a specific protein functions, it can be useful to identify mutants that no longer interact with a known partner. This is especially useful for proteins that interact with multiple partners, because it allows dissection of the importance of specific proteinprotein interactions. By taking advantage of the ease of the interaction trap/two-hybrid system (UNIT 20.1; Gyuris et al., 1993) and of PCR mutagenesis (UNIT 8.3; Cadwell and Joyce, 1992), it becomes possible to rapidly make and identify proteins that no longer bind to a specific partner protein. These mutants usually result from a single missense mutation, which can easily be identified. IDENTIFYING LOSS-OF-INTERACTION MUTANT PROTEINS In this protocol, two-hybrid analysis in S. cerevisiae is used to confirm interaction of a bait, a cDNA encoding a protein of interest fused to a DNA binding domain, with a prey, a cDNA encoding an interacting protein fused to a transcription activation domain (see UNIT 20.1 for more details). The open reading frame (ORF) from the cDNA encoding the prey protein is PCR-amplified with primers designed to remove its termination codon to prepare it for fusion with green fluorescent protein (GFP; Fig. 20.8.1, top left). The ORF is mutagenized by Mn-doped PCR—also known as error-prone (EP)-PCR—to introduce an average of one mutation per DNA molecule (Fig. 20.8.1, middle left), and then PCR-ligated at its 3′ end to DNA encoding GFP (Fig. 20.8.1, top right and middle). GFP expression is used as a reporter to ensure that the mutagenesis has not introduced a stop codon into the ORF. The resulting construct is then introduced back into a two-hybrid prey plasmid (pJG4-5) by recombination in S. cerevisiae (Fig. 20.8.1, bottom) or by standard cloning techniques (see Alternate Protocol). This mutagenized prey minilibrary is then screened for preys that do not interact with their former partner protein expressed as bait. Colonies of cells which fluoresce green and that carry missense mutant proteins that do not interact with a specific bait protein in plasmid pEG202, as shown by two-hybrid analysis, are chosen for further study. DNA derived from these colonies is sequenced and corresponding single-point mutations are introduced into the original prey plasmid by directed PCR mutagenesis. The noninteracting phenotype is then confirmed by two-hybrid analysis. Materials Genes for proteins of interest Two-hybrid interaction trap plasmids (Table 20.1.1): Prey plasmid pJG4-5 Bait plasmid pEG202 LacZ reporter plasmid pSH18-34 Two-hybrid interaction trap yeast strain EGY48 (UNIT 20.1) CM dropout plates (UNIT 13.1) substituted with 2% glucose (Glc), 2% galactose(Gal), and/or 1% raffinose (Raff) as indicated: Glc/CM −Ura Glc/CM −Ura −His Glc/CM −Ura −His, −Trp Glc/CM/Xgal −Ura −His −Trp Gal/Raff/CM/20 µg/ml Xgal in Z buffer −Ura, −His, −Trp: prepare as described for CM plates (UNIT 13.1) using a 1 mg/ml Xgal stock (20 µg/ml final) in Z buffer (UNIT 13.6) Contributed by Andrew R. Mendelsohn Current Protocols in Molecular Biology (2004) 20.8.1-20.8.10 Copyright © 2004 by John Wiley & Sons, Inc.

BASIC PROTOCOL



1

5 ORF

GFP

*

2 3

6

PCR 3 ORF

PCR 4

PCR ORF′

Mn2+

GFP 6

PCR

42

ORF′

GFR

PGal1 HA NLS B42

Figure 20.8.1 Interaction trap/two-hybrid system loss-of-interaction mutant proteins. Circled numbers indicate primers (see Basic Protocol). The asterisk (*) on primer 2 indicates that the stop codon has been deleted from the ORF.

2.5 mM (each) dNTP stock solution (UNIT 15.1) 2.5 U/µl high-fidelity PCR-compatible DNA polymerase (e.g., PfuTurbo, Kod) and 10× buffer 1 U/µl Taq DNA polymerase (UNIT 3.5) and 10× Mg2+-free reaction buffer 2.5 mM MgCl2 50 mM MnCl2 Plasmid carrying EGFP (e.g., pEGFP; BD Biosciences) CM −Trp liquid medium (UNIT 13.1) TrpC E. coli strain KC8 (hsdR, leuB600, trpC9830, pyrF::Tn5, hisB463, lac∆X74, strA, galU, galK) (UNIT 20.1; constructed by K. Struhl and available from R. Brent), competent M9 minimal plates (UNIT 1.1) containing 100 µg/ml amp, 40 µg/ml Leu, 40 µg/ml His, and 40 µg/ml Ura Commercial miniprep kit (e.g., Qiagen; optional) 30°C incubator Long-wavelength UV lamp, or fluorescence microscope equipped with fluorescein or GFP filter set Additional reagents and equipment for subcloning into plasmid vectors (UNIT 3.16), lithium acetate transformation (UNIT 13.7), PCR (UNIT 15.1), agarose gel electrophoresis (UNIT 2.5A), restriction digestion (UNIT 3.1), replica plating using velvet filters (UNITS 1.3 & 13.2), interaction trap (UNIT 20.1), purification of DNA by glass beads (UNIT 3.11), DNA minipreps (UNIT 2.1; optional), bacterial transformation (UNIT 1.8), sequencing DNA by dideoxy chain termination (UNIT 7.4), and site-directed PCR mutagenesis (UNIT 8.5)

Identification of Loss-of-Interaction Mutant Proteins



Confirm interaction of bait protein with prey protein by two-hybrid analysis Once interaction has been confirmed, the experiment can be started with step 7. 1. Insert the gene (UNIT 3.16) for the first protein of interest into the prey plasmid pJG4-5. This protein will later be mutagenized (see step 10) so that it no longer interacts with its protein partner.

2. Insert the gene (UNIT 3.16) for the protein partner into bait plasmid pEG202. 3. Transform EGY48 with ∼1 µg lacZ reporter plasmid pSH18-34 by the lithium acetate yeast protocol (UNIT 13.7) and select for yeast that grow on CM −Ura dropout medium. 4. Transform 1 µg of both plasmids simultaneously into interaction-trap strain EGY48 carrying pSH18-34. Select for yeast carrying all three plasmids by growing on Glc/CM −Ura −His −Trp triple dropout plates. Selection for pSH18-34 is achieved by omitting uracil (−Ura), pEG202-derived bait plasmid by omitting histidine (−His), and pJG4-5-derived prey plasmid by omitting tryptophan (−Trp).

5. As a negative control, transform EGY48/pSH18-34 (step 3) with prey vector pJG4-5 and bait vector pEG202 and select colonies on Glc/CM −Ura −His −Trp triple dropout plates. 6. Pick colonies and replica plate onto the following fresh plates: Glc/CM −Ura −His −Trp Glc/CM/Xgal −Ura −His −Trp Gal/Raff/CM/Xgal −Ura −His −Trp. Colonies carrying interacting bait and prey should appear blue on Xgal/Gal/Raff plates and white on Xgal/Glc plates. Compare with the pJG4-5/pEG202 negative control in which colonies should appear white on all plates. If the interaction is very strong or if the bait self-activates transcription to some extent (see Critical Parameters and Troubleshooting), the colonies may appear somewhat blue on the control Xgal/Glc plates. If the amount of blue on Xgal/Glc plates is substantial and similar to that seen on Xgal/Raff plates, it may not be possible to use this particular bait in this protocol. However, there are several ways to reduce interaction. For example, one way to reduce background is to use a less sensitive lacZ reporter plasmid, such as pJK103 (see Table 20.1.1). For a detailed discussion on how to reduce background from the bait self-activating the lacZ reporter, see UNIT 20.1, Commentary.

Remove the termination codon from the prey ORF 7. Synthesize the following PCR primers (Fig. 20.8.1) to remove the 3′ translation termination codon from the prey by directed PCR mutagenesis: a. Primer 1: Design to contain 5′ sequence homologous to the B42 region of pJG4-5 (for later homologous recombination) fused to ∼21 bp of the prey ORF beginning at the initiation codon (ATG). An example sequence is 5′-GATTATGCCTCTCCCGAATTC-ATG-ORF-3′, where the ORF is 21 bp directly downstream of the ORFs initiation codon (ATG). It is also possible to design an alternate primer 1 for later conventional cloning by including an appropriate restriction site such as EcoRI instead of sequence homologous to the B42 region of pJG4-5 (e.g., CTCTCTCTCTCTCTGAATTC-ORF-3′) where the ORF is 21 bp directly downstream of the ORFs ATG. Analysis of Protein Interactions


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b. Primer 2: Design to contain 5′ sequence homologous to the reverse complement of the sequence homologous to the 5′ end of EGFP fused to the reverse complement of the 3′ end of the ORF just upstream of, but not including, the termination codon. An example sequence is 5′-CTCCTCGCCCTTGCTCACCAT-ORF-3′, where ORF is the reverse complement of the sequence 21 bp upstream of the termination codon.

8. Set up the termination-codon-removal PCR reaction: 1 ng prey plasmid carrying ORF 0.2 µM primer 1 0.2 µM primer 2 2 µl 10× (2.5 mM) dNTPs 2.5 µl 10× polymerase buffer 1 µl 2.5 units/µl high-fidelity PCR-compatible DNA polymerase (e.g., PfuTurbo; add last). Use distilled water to bring the final reaction volume to 25 µl. 9. Perform PCR (UNIT 15.1) using the following cycling conditions: Initial step: 20 cycles:

Final step:

30 sec 30 sec 60 sec 60 sec 10 min

94°C 94°C 56°C 72°C 72°C.

10. Check for appropriate-sized PCR fragment on a 1% agarose gel with TBE as the buffer (UNIT 2.5A). Mutagenize the prey ORF 11. Synthesize the following primers for mutagenesis of the prey ORF, with termination codon removed, by Mn-doped PCR (Fig. 20.8.1): a. Primer 3: Design to contain sequences homologous to the 5′ part of primer 1 (step 7; i.e., homologous to the B42 region of pJG4-5), but without any sequence homologous to the prey ORF. This primer is designed to extend the homology with pJG4-5 to 40 bp for later homologous recombination (see below)—e.g., 5′-CTACCCTTATGATGTGCCAGATTATGCCTCTCCCGAATTC-3′. If the minilibrary (see below) is to be made by conventional cloning, primer 3 can be designed to be homologous only to the 5′ region of alternate primer 1 (step 7), such that it does not contain any sequence homologous to the ORF (e.g., 5′CTCTCTCTCTCTCTGAATTC-3′).

b. Primer 4: Design to be identical to the 3′ section of primer 2—i.e., homologous to the reverse complement of the 5′ end of the EGFP gene (step 7; e.g., 5′CTCCTCGCCCTTGCTCACCAT-3′). If the region of interaction is known and defined by a contiguous region on the prey protein, then it may be useful to restrict the mutagenesis to this region by appropriate primer design. Mn-doped PCR mutagenesis has been described by Lin-Goerke et al. (1997), and Cadwell and Joyce (1992). Identification of Loss-of-Interaction Mutant Proteins



12. Set up the Mn-doped PCR reaction as follows: 16 µl 2.5 mM (each) dNTPs 2 µl primer 3 (5 µM final concentration) 2 µl primer 4 (5 µM final concentration) 5 µl 10× Mg2+-free Taq reaction buffer 5 µl 2.5 mM MgCl2 2.5 µl 50 mM MnCl2 1 µl 10 ng of 1.3-kb DNA fragment (step 10) 1 µl 1 U/µl Taq DNA polymerase 15.5 µl H2O. Mutagenesis occurs by increasing the misincorporation rate of Taq DNA polymerase; it is often referred to as error-prone PCR (see UNIT 8.3 for a more in depth discussion of this methodology).

13. Perform PCR (UNIT 15.1) using the following cycling conditions: Initial step: 36 cycles:

Final step:

2 min 30 sec 1 min 1 min 10 min

94°C 94°C 55°C 72°C 72°C.

Mn-doped PCR mutagenesis needs to be optimized in order to obtain an average of one mutation per DNA molecule. Optimization can be performed by varying the concentration of Mn2+ and Mg2+ in the protocol. It should be noted that even mutagenesis that results in as many as ten mutations per DNA molecule can be successfully used in this protocol, as the key mutation can be identified by sequencing followed by individually introducing mutations into the prey ORF and checking whether the resulting prey does not interact with the bait. While it can be useful to vary the both the Mn2+ and Mg2+ concentrations, the authors have had some success varying just the Mg2+ concentration from 1 mM to 7 mM. It is suggested that several reactions be performed in parallel using various Mg2+ concentrations. Efficiency of mutagenesis can be later determined by sequencing resulting clones from a minilibrary (see below).

Fuse prey ORF with GFP 14. Synthesize the following primers (Fig. 20.8.1) for amplifying the full-length EGFP gene: a. Primer 5: Design to contain a sequence homologous to the 5′ end of the EGFP gene. An example sequence is 5′-ATGGTGAGCAAGGGCGAGGAG-3′

b. Primer 6: Design a sequence where the 5′ end is homologous to the reverse complement of the polylinker region of pJG4-5 fused to the reverse complement of the 3′ end of the EGFP gene. The 5′ end is prepared in this fashion in order to facilitate homologous recombination (see below). An example sequence is 5′-TTGACCAAACCTCTGGCGAAGAAGTCCAAAGCTTCTCGAGTTACTTGTACAGCTCGTCCATG-3′. An alternate primer 6 for conventional cloning (see Alternate Protocol) would consist of sequence creating a compatible site for cloning—e.g., XhoI fused to the reverse complem en t of th e 3′ e n d o f th e E G F P gen e (5′-CTCTCTCTCTCGAGTTACTTGTACAGCTCGTCCATG-3′).



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15. Set up the reaction to amplify EGFP: 1 ng plasmid carrying EGFP (e.g., pEGFP) 1 µl primer 5 (1 µM final) 1 µl primer 6 (1 µM final) 2 µl 2.5 mM dNTPs 2.5 µl 10× high-fidelity PCR-compatible DNA polymerase buffer 1 µl 2.5 U/µl high-fidelity PCR-compatible DNA polymerase (e.g., PfuTurbo; add last). H2O to 25 µl. 16. Perform PCR (UNIT 15.1) using the following cycling conditions: Initial step: 19 cycles:

Final step:


94°C 94°C 56°C 72°C 72°C.

17. Set up PCR reaction to fuse mutated prey ORF with EGFP: 1 ng plasmid carrying EGFP (e.g., pEGFP) 1 ng mutated prey ORF DNA (step 13) 1 µM primer 3 1 µM primer 6 2 µl 10× (2.5 mM) dNTPs 2.5 µl 10× PfuTurbo buffer 1 µl PfuTurbo (2.5 units/µl; add last) H2O to 50 µl. This reaction can be scaled up if necessary. Primers 2 and 4 have previously been used to introduce sequence into the prey ORF such that the mutated ORF and GFP now overlap. This allows fusion of the two pieces of DNA by PCR amplification using primers that are homologous to 5′ end of the mutated ORF DNA (primer 3) and the 3′ end of the EGFP gene (primer 6; see Fig. 20.8.1, middle).

18. Perform PCR (UNIT 15.1) using the following cycling conditions: Initial step: 19 cycles:

Final step:


94°C 94°C 56°C 72°C 72°C.

At least 1 ìg DNA is needed for the minilibrary construction if using the yeast recombination method, or 100 ng DNA if using conventional cloning (see Alternate Protocol). The number of cycles or volume of the reaction should be adjusted accordingly. The yield can be checked by samples removed for gel electrophoresis or by real-time PCR.

Create prey minilibrary by recombination in yeast See Alternate Protocol if constructing minilibrary by conventional means. 19. Make competent EGY48 carrying pSH18-34 and appropriate bait in pEG202 using the lithium acetate method (UNIT 13.7). 20. Plate and select on Glc/CM −Ura −His. Identification of Loss-of-Interaction Mutant Proteins



21. Add 1 µg mutated ORF-EGFP DNA (step 18) and 1 µg pJG4-5 linearized with EcoRI (UNIT 3.1) to 200 µl competent yeast (step 19). 22. Incubate 30 min at 30°C. 23. Heat shock 2 min at 37°C. 24. Plate on Glc/CM −Ura −His −Trp plates, as appropriate. 25. Determine the total number of independent transformants by counting colonies. At least 1000 independent transformants will be needed for a successful screen. A lesser amount may be satisfactory if the library contains multiple mutations per DNA molecule.

Screen library for loss-of-interaction prey mutants 26. Replica plate the yeast transformed with the prey minilibrary onto Gal/Raff/CM/Xgal −Ura −His −Trp and Glc/CM/Xgal −Ura −His −Trp using velvet filters (UNIT 1.3 & 13.2), or by patching colonies onto the Xgal-containing plates (UNIT 20.1). 27. Incubate 24 to 48 hr at 30°C. 28. Screen the library for the following two features at the same time: a. Search for mutant preys that no longer interact with the original bait by looking for white colonies on Gal/Raff/CM/Xgal −Ura −His −Trp plates. b. Hunt for clones with uninterrupted ORFs by looking for colonies that remain green when viewed under a long-wavelength UV lamp or, more desirably, a fluorescence microscope equipped with a fluorescein or GFP filter set. Green fluorescent or white colonies represent candidate loss-of-interaction mutants and should be picked for further analysis.

Identify resulting candidate mutations by sequencing 29. Grow candidate colony overnight in 5 ml CM −Trp medium. 30. Microcentrifuge yeast 20 sec to pellet. 31. Use the glass bead method (UNIT 13.11) to isolate DNA from candidates. 32. Transform E. coli trp− strain KC8 (hsdR, leuB600, trpC9830, pyrF::Tn5, hisB463, lac∆X74, strA, galU, galK) with DNA (UNIT 1.8) and plate on M9 minimal plates containing 100 µg/ml amp, 40 µg/ml Leu, 40 µg/ml His, and 40 µg/ml Ura. This works because there is enough expression of the TRP1 gene in E. coli to complement the trpC mutation in KC8.

33. Perform DNA minipreps (UNITS miniprep).

1.7 or 2.1B)

or use a commercial kit (e.g., Qiagen

34. Sequence DNA by dideoxy chain termination method (UNIT 7.4). Confirm candidate mutations by directed PCR mutagenesis and two-hybrid analysis 35. Analyze the results of sequencing the candidate DNAs: a. For single mutation: If a candidate clone has only one missense mutation, that mutation must have resulted in the loss-of-interaction phenotype (i.e., no further analysis is necessary). b. For multiple mutations: If a candidate clone has multiple mutations, introduce these mutations into the prey individually to determine which one actually causes the loss-of-interaction phenotype (see below).



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36. Use site-directed PCR mutagenesis (UNIT 8.5) to introduce single specific mutations into the original prey ORF. 37. Individually transform preys into EGY48 carrying pSH18-34 and the original bait cloned into pEG202 (steps 19 and 20). 38. Replica-plate colonies and screen candidates for loss-of-interaction prey mutants as described above (steps 21 to 32). ALTERNATE PROTOCOL

CREATE PREY MINILIBRARY BY CONVENTIONAL CLONING The following steps describe preparing minilibraries by conventional cloning. These steps are used in place of the minilibrary construction procedure described above (see Basic Protocol, steps 19 to 25). Additional Materials (also see Basic Protocol) T4 DNA ligase (UNIT 3.14) LB medium and plates containing amp (UNITS 1.1 & 1.4) Additional reagents and equipment for gel purification of DNA fragments (UNIT 2.6), multiple endonuclease digestion (UNIT 3.2), DNA ligation (UNITS 3.14 & 3.16) 19. Gel purify both cut fragments (UNIT 2.6). Digest mutated ORF-EGFP DNA (see Basic Protocol, step 18) and prey plasmid pJG4-5 with EcoRI and XhoI (see UNITS 3.1 & 3.2). This presumes that there are no internal EcoRI and XhoI sites in the prey ORF. If these sites are present in the ORF, then alternates must be chosen that are compatible with the unique sites in the polylinker of pJG4-5.

20. Gel purify both cut fragments (UNIT 2.6). Ligate 100 ng cut DNA using T4 DNA ligase (UNIT 3.14 & 3.16). 21. Transform E. coli with library DNA (UNIT 1.8). Plate serial 1:10 dilutions of E. coli transformed with the library DNA onto LB plates containing amp to determine the number of independent transformants. 22. Amplify library by diluting an aliquot 1:10 into 100 ml LB medium containing amp and growing overnight at 37°C. 23. Make high-quality prey DNA from culture using a maxiprep method (UNIT 1.7) or commercial kit. 24. Transform competent EGY48 carrying pSH18-34 and appropriate bait in pEG202 (see Basic Protocol, step 19) with prey DNA by lithium acetate method (UNIT 13.7). Plate on Glc/CM −Ura −His −Trp plates, as appropriate. 25. Determine the total number of independent transformants by counting colonies. At least 1000 independent transformants will be needed for a successful screen. A lesser amount may be satisfactory if the library contains multiple mutations per DNA molecule.



The methodology described in this unit was used to isolate mutants of caspase 2 that no longer interacted with cyclin D3 (Mendelsohn et al., 2002). The basic techniques involved, the two-hybrid system (UNIT 20.1), and PCR mu-

tagenesis (UNITS 8.3 & 8.5) are established techniques and are easy to perform. The basic advantage of this methodology is the addition of EGFP as a reporter to identify clones that have not been truncated by termination codons introduced by PCR mutagenesis. EGFP also



serves to eliminate mutants that destabilize the prey protein in yeast.

Critical Parameters and Troubleshooting There are two critical parameters that must be considered to use this methodology successfully. The first is that the interacting proteins must be well-behaved in the interaction trap/two-hybrid system. If the bait is self-activating, or if the interaction is so strong as to activate the lacZ reporter on the glucose plates that should greatly repress the synthesis of the prey, then steps must be taken to reduce the sensitivity of the assay. UNIT 20.1 discusses this in more depth. The bottom line is that some proteins are not amenable to being used as baits in the two-hybrid system. Therefore, perhaps the simplest way to resolve the problem of a self-activating bait is to switch the bait and prey proteins. It is quite possible that when the prey protein is used as a bait, it will not self-activate transcription. The second critical parameter is the condition of the in vitro mutagenesis. Given a specific rate of mutagenesis, the length of the ORF to be mutated is an important consideration. The same rate of mutagenesis that would cause an average of one mutation in a 100-bp ORF will cause an average of ten mutations in a 1000-bp ORF. Critically, there can be great variation from published ideal conditions for error-prone (EP)-PCR. It is strongly recommended that the magnesium and/or manganese concentration(s) be titrated to obtain the desired rate of mutagenesis on the specific template chosen. Another factor to consider is that Mn2+ ions may associate with the DNA and interfere with subsequent DNA manipulations. After the completion of the reaction, it may be useful to add a large excess of Mg2+ ions to compete out the Mn2+ ions, or EDTA to bind up the Mn2+ ions, or to subsequently purify DNA by glasspowder isolation from an agarose gel. One factor to consider is that the screen described here for intact ORFs could be made into a selection by substituting an auxotrophic yeast marker such as Lys2 for EGFP in the PCR fusion step and then transforming a Lys2 auxotrophic mutant derived from EGY48. In this case, selection could be made on CM dropout −Ura −His −Trp −Lys plates. The absence of lysine would select for the intact ORF. Protein interaction itself is still probably best measured by screening for white colonies on plates containing Xgal, galactose, and raffinose.

Another factor to think about is that it may prove valuable to test the mutated prey against a battery of known interacting proteins as baits in the two-hybrid system. This may be especially useful if the mutated proteins are to be characterized functionally in their native biological context. Any change of phenotype would likely be the result of the loss of a specific protein-protein interaction. The proteins that no longer interact would thus represent potential key players in causing the observed phenotype. In this regard, it is important to note that it may not be the loss of the specific protein interaction selected in this procedure that has true functional significance, but rather the simultaneous loss of interaction of the mutated protein with another interacting protein that would be critical. Furthermore, an interesting extension of this technology would be to perform a series of experiments to prove the relevance of a particular protein-protein interaction by (1) isolating a loss-of-function version of the prey protein, (2) associating this mutant with a change of phenotype, (3) isolating a mutant version of the partner protein that does interact with the initial mutant, and then (4) showing that the presence of both mutant proteins restores the original phenotype. The methodology described here could easily be modified to this task by the following modification. After isolating the loss-of-function prey mutant, insert it into the pEG202 bait vector, then insert the original bait into the pJG4-5 prey vector and follow the procedure described, but screen for blue colonies that fluoresce green. In other words, look for full-length mutant proteins that interact with the original mutant protein. After identifying such proteins, if simultaneous expression of both mutant proteins in the original biological system restores the original phenotype, then the specific protein-protein interaction between the nonmutated versions of these proteins must be critical for the function associated with the mutant phenotype.

Anticipated Results Between one and several hundred loss-offunction mutants should be obtained from successful application of this methodology.

Time Considerations Altogether, this methodology should require ∼2 weeks to obtain and characterize lossof-function mutants, if the cDNAs for the interacting protein pair are available and behave well in the two-hybrid system.



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Literature Cited Cadwell, R.C. and Joyce, G.F. 1992. Randomization of genes by PCR mutagenesis. PCR Methods Appl. 2:28-33. Gyuris, J., Golemis, E., Chertkov, H., and Brent, R. 1993. Cdi1, a human G1 and S phase protein phosphatase that associates with Cdk2. Cell 75:791-803. Lin-Goerke, J.L., Robbins, D.J., and Burczak, J.D. 1997. PCR-based random mutagenesis using manganese and reduced dNTP concentration. Biotechniques 23:409-412.

Mendelsohn, A.R., Hamer, J.D., Wang, Z.B., and Brent, R. 2002. Cyclin D3 activates caspase 2, connecting cell proliferation with cell death. Proc. Natl. Acad. Sci. U.S.A. 99:6871-6876.

Contributed by Andrew R. Mendelsohn The Molecular Sciences Institute Berkeley, California




CHAPTER 21 Chromatin Assembly and Analysis INTRODUCTION

M

any nuclear processes require that DNA be recognized by sequence-specific DNA binding proteins. The packaging of DNA into chromatin inhibits DNA binding by most proteins, and also influences the efficiency of events such as transcription, replication, recombination, and DNA repair. These considerations have led to an increased interest in characterizing chromatin structure in general and the precise structural changes that can occur over specific chromosomal domains. The basic unit of chromatin is the nucleosome, composed of DNA and histones (small basic proteins that are highly conserved in all eukaryotes). Each nucleosome consists of two copies each of histones H2A, H2B, H3, and H4, arranged into an octamer that organizes 146 bp of DNA. The DNA is wrapped 1.65 times around the histones, and so is severely bent compared to DNA in solution. Each histone has an N-terminal “tail” that extends outside the DNA, and these tails are the sites of numerous covalent modifications that contribute significantly to the regulation of chromatin structure. The most extensively studied of these modifications is acetylation, which occurs on lysine residues. Chromatin structure can be changed by covalently modifying the nucleosome tails or by noncovalently “remodeling” nucleosome structure. Remodeling is an ATP-dependent process that can alter nucleosome mobility and increase the access of DNA binding proteins and nucleases to the nucleosome. Both remodeling and covalent modification of nucleosomes can lead to local changes in chromatin structure that have been associated with activation or repression of transcription. These structural changes can be studied by characterizing the specific chromatin structure over a defined region of the chromosome in vivo, and by recapitulating chromatin structure on DNA fragments in vitro. This chapter presents techniques for analyzing chromatin structure. UNIT 21.1 describes the use of micrococcal nuclease (MNase), an important reagent for chromatin research, in characterizing nucleosome structure across any specific gene in intact cells. Because MNase cleaves preferentially in the linker DNA that lies between individual nucleosomes, MNase cleavage can be used to map the boundaries of nucleosomes and determine their spacing. To perform a detailed analysis of chromatin structure, it is frequently necessary to assess whether the nucleosomes in a chromosomal domain are covalently modified, and to determine which other DNA binding proteins are bound nearby. The most commonly studied histone modification is the acetylation of lysine residues. Many cellular enzymes are capable of catalyzing this modification, and several of these have been implicated in transcriptional activation. To separate the differentially acetylated histone species, it is necessary to use a specialized gel system (UNIT 21.2). This technique can be used to quantify the amount of each acetylated species in a mixture of histones, either a crude histone fraction prepared from cells or a mixture of histones modified in vitro. To determine the location of covalently modified histones, or to localize any other DNA binding protein, immunoprecipitations of cross-linked chromatin can be performed (UNIT 21.3). This widely used technique, called Chromatin Immunoprecipitation (or ChIP), can determine the relative concentrations of any protein epitope across a region of chromatin, provided a

Contributed by Robert E. Kingston Current Protocols in Molecular Biology (2002) 21.0.1-21.0.2 Copyright © 2002 by John Wiley & Sons, Inc.

Chromatin Assembly and Analysis


good antisera is available. This technique has been used to analyze relative levels of covalently modified histones across broad regions of chromatin using antisera that recognize specific histone modifications, to localize sequence-specific DNA binding factors, and to analyze components of the general transcription and replication machinery. Analysis of the precise position and structure of nucleosomes both in vitro and in vivo can be accomplished using either enzymatic cleavage reagents, such as DNase, or chemical cleavage reagents (UNIT 21.4). These agents produce specific patterns of cleavage that reflect the location of DNA on the surface of the histone octamer, and thus give more precise information than MNase cleavage. While these procedures are most frequently used to analyze nucleosomes formed in vitro (see UNITS 21.5-21.7), they have also been combined with PCR-based technologies to analyze detailed nucleosome structure in intact cells (as in UNIT 21.1). To characterize the functional roles of chromatin structure in nuclear processes, it is necessary to use in vitro systems and templates that have been assembled into appropriate chromatin structure. In order to form chromatin in vitro, it is first necessary to have a source for core histones (UNIT 21.5). Either isolated histones or nucleosomal DNA can be used to transfer histones onto a specific template, although isolated histones can be used in a wider variety of protocols. UNIT 21.5 also provides methods for isolating intact polynucleosomes from cells, which have a wide variety of uses in vitro. Templates can be assembled into nucleosomes in vitro using either salt dialysis (UNIT 21.6) or an enzymatic chromatin assembly system (UNIT 21.7). Salt dialysis offers the advantage that no specialized proteins are needed, and the final nucleosome preparation is not contaminated by additional factors that might influence further mechanistic studies. It has the significant disadvantage that appropriate nucleosomal spacing on natural templates or on arrays of nucleosomes will not be achieved unless very specialized templates (such as 5S arrays, see UNIT 21.6) are used. Enzymatic assembly of nucleosomal arrays (UNIT 21.7) will achieve physiological spacing of nucleosomes on virtually any template, and thus is considerably more flexible than salt dialysis. Care must be taken, however, to either purify the template away from the assembly factors following assembly, or to carefully consider the potential role of the assembly factors in any process that is under study. Robert E. Kingston

Introduction



Micrococcal Nuclease Analysis of Chromatin Structure

UNIT 21.1

Micrococcal nuclease (MNase) assays of chromatin are relatively simple procedures for obtaining information about the locations of nucleosomes along DNA strands. When nuclei or permeabilized cells are exposed to MNase in the presence of a divalent cation, the enzyme makes double-stranded cuts between nucleosome particles (see UNIT 3.12). Treatment of chromatin substrates with very high concentrations of MNase gives rise mostly to mononucleosome-length DNA, whereas low concentrations of the enzyme will produce one double-stranded cut every 10 to 50 nucleosomes (depending on the exact concentration). MNase can also make single-stranded DNA cuts on the histone octamer, and thus experiments to map the positions of nucleosomes are usually performed with native double-stranded DNA. In this unit, Basic Protocols 1 and 2 describe how to permeabilize cells or prepare nuclei from tissues, respectively, and then perform MNase digestion. Basic Protocol 3 describes how to perform MNase digests of purified genomic DNA, which will serve as a control for the inherent MNase cleavage pattern over a sequence of interest. Support Protocol 1 describes the purification of DNA from chromatin digests in a manner suitable for subsequent analysis. Support Protocol 2 describes different ways of analyzing the purified DNA at the Southern blot level of resolution, and Support Protocol 3 describes a modification of ligation-mediated PCR (LM-PCR) to map genomic, double-stranded MNase cleavages at the nucleotide level of resolution. STRATEGIC PLANNING In an experiment to probe chromatin structure, the chromatin is made accessible to MNase, the chromatin is digested with the enzyme for a brief period of time, the reaction is stopped by the addition of detergent and proteinase, and DNA within the chromatin is isolated free from protein. Prior to initiating such an analysis, it is important to consider the following three parameters, which will dictate a specific experimental strategy. Chromatin Preparation Method The quality of an MNase analysis is critically dependent upon the integrity of the chromatin substrate. Isolated nuclei are most commonly used, but there is clear evidence that chromatin components are perturbed during the nuclear isolation procedure. Permeabilized cells retain chromatin components that can be lost during nuclear isolation, and therefore permeabilization is preferable to nuclear isolation (Pfeifer and Riggs, 1991). The method of preparing a chromatin substrate is generally dictated by the source of cells: tissues must usually be disrupted mechanically, and therefore nuclear isolation is required, whereas cell suspensions or cells on tissue culture plates are uniformly accessible and therefore are suitable for cell permeabilization. The latter protocol is also far faster and simpler. The two procedures are described in Basic Protocols 1 and 2. Type of Information Desired The extent of MNase digestion of the sample determines the kind of inferences that can be made about the chromatin structure of the genomic region of interest. An extensive MNase digestion, which yields predominantly mononucleosomal-sized DNA fragments, can indicate the frequency with which a DNA sequence is nucleosomal among a population of templates in the reaction. A partial MNase digestion, which yields lengthy polynucleosomal-sized DNA fragments, can indicate the positions of nucleosomes with Contributed by Ken Zaret Current Protocols in Molecular Biology (1999) 21.1.1-21.1.17 Copyright © 1999 by John Wiley & Sons, Inc.



respect to a defined point in the DNA, such as a restriction site. Each of these approaches employs a different Southern blotting or LM-PCR strategy to map MNase cleavages, described in Support Protocols 2 and 3, respectively. It is essential to plan a cleavage mapping strategy in order to determine the controls needed for the MNase digestion reactions. Controls Several controls are critical for the strategic planning of any MNase study. First, it is essential to analyze an MNase digestion of protein-free DNA, as described in Basic Protocol 3, because MNase cleaves DNA in a highly nonrandom manner. The range of MNase digestion of free DNA should be comparable to the range of digestion of the target sequence seen in chromatin MNase digests. Only MNase cleavages that are induced or enhanced in chromatin, compared to free DNA, can be presumed to provide conclusive information about the presence of nucleosomes and their positions (see Fig. 21.1.1 in Support Protocol 2). Second, with isolated nuclei or permeabilized cells, it is necessary to include a sample that was incubated in the MNase digestion buffer for the same amount of time and at the same temperature as the MNase-treated samples, but without added enzyme. Cleavages that occur over the genomic region of interest in this control will be due to endogenous nucleases. The Commentary following the protocols describes how to minimize the effects of these enzymes (see Troubleshooting). In summary, a well-controlled MNase assay must include the following, preferably all on the same blot or LM-PCR gel, and in the following order: (1) MNase digestion of free DNA at several enzyme concentrations, including a no-enzyme control; (2) MNase digestion of chromatin at several enzyme concentrations, including a no-enzyme control; (3) MNase digestion of a chromatin sample from a different cell type or physiological condition, to determine the specificity of nucleosomal arrangements observed in the first chromatin sample; and (4) an internal marker lane (see Support Protocol 2). BASIC PROTOCOL 1

MICROCOCCAL NUCLEASE DIGESTION OF CHROMATIN IN PERMEABILIZED CELLS In this protocol, chromatin substrates are prepared by permeabilizing cultured cells using the detergent lysolecithin (either in situ for cells adherent on plates, or in suspension for suspension cultures). The chromatin is then digested with micrococcal nuclease, and the DNA purified so that it is suitable for Southern blot (see Support Protocol 2) or LM-PCR analysis (see Support Protocol 3). Before proceeding with the full-scale experiment, optimal conditions should be determined by performing a preliminary set of trials, both to determine the appropriate lysolecithin incubation conditions and to select a workable range of MNase that yields the desired level of digestion. Materials Cells cultured on 100-mm petri dishes or cells in suspension Permeabilization solution 1, room temperature and 37°C (see recipe) 1 mg/ml lysolecithin (Sigma) in permeabilization solution 1 (mix fresh before use) Permeabilization solution 2 (see recipe), with and without MNase (see step 4 below; mix fresh before use) Permeabilization stop solution (see recipe) Lysis dilution buffer: 150 mM NaCl/5 mM EDTA


Phase-contrast microscope 12-ml conical polypropylene tubes with caps (e.g., Sarstedt) NOTE: All solutions, pipets, and pipet tips should be at room temperature so that the permeabilization reaction proceeds rapidly.



Carry out cell permeabilization 1a. Adherent cells (per 100-mm plate): Working at room temperature (e.g., in a tissue culture hood), aspirate the medium from the cells and add 5 ml room temperature permeabilization solution 1. Do not remove cells from the plates. Use a separate plate of cells for each MNase digestion condition, remembering to include a zero-enzyme condition (see Strategic Planning). Try to harvest cells at 70% to 90% confluence.

1b. Suspension cultures: Pellet by centrifugation under conditions appropriate for cell type and resuspend cell pellet in 5 ml permeabilization solution 1. The remainder of the procedure is the same for suspension cells as for adherent cells, except that centrifugation is required to change solutions.

2. Aspirate medium from each plate, then treat with lysolecithin (diluted from 1 mg/ml stock in 37°C permeabilization solution 1 to 2.8 ml total volume) at room temperature as follows: either use 0.025% lysolecithin and incubate 2 min, or use 0.05% lysolecithin and incubate 1 min. Lysolecithin has been the preferred reagent for permeabilizing cells (Zhang and Gralla, 1990; Pfeifer and Riggs, 1991), although other reagents have not been explored in depth for this purpose. Different cell lines or the same cell line cultured under different conditions may require different concentrations of lysolecithin, or different times or temperatures (30° or 37°C instead of room temperature) for the detergent incubation (see Critical Parameters).

3. Aspirate solution from plate, then add 5 ml room temperature permeabilization solution 1 (without lysolecithin). Examine the cells by phase-contrast microscopy to be sure that cell lysis has not occurred. If it has, repeat the assay with a lower concentration of lysolecithin. Carry out MNase digestion of permeabilized cells 4. Aspirate solution from plate, then add 2.8 ml of room temperature permeabilization solution 2 containing either 0, 7.5, 15, or 30 U MNase for a partial digestion assay, or 0, 50, 100, or 200 U MNase for an extensive digestion assay. Incubate 5 min at room temperature. Dilute the MNase into aliquots of permeabilization solution 2 just prior to use.

5. Aspirate solution from plate, then add 2.8 ml permeabilization stop solution. Swirl around plate to complete cell lysis. If the benchtop is not level, occasionally tilt the plate during the MNase digestion.

6. Add 2.8 ml lysis dilution buffer. Swirl around plate to complete mixing, then transfer mixture to a 12-ml conical polypropylene tube. Cap tube and incubate overnight at 37°C. 7. Proceed to purification and characterization of the DNA (see Support Protocol 1).



Supplement 45

BASIC PROTOCOL 2

MICROCOCCAL NUCLEASE DIGESTION OF CHROMATIN IN ISOLATED NUCLEI In this protocol, chromatin substrates are prepared by isolating cell nuclei, the chromatin within the nuclei is digested with micrococcal nuclease, and the DNA is purified so that it is suitable for Southern blot (UNIT 2.9A) or LM-PCR analysis (UNIT 15.5). Different nuclear isolation conditions are described for liver, kidney, and spleen, each of which exhibit different characteristics during homogenization and thus should serve as examples for a wide variety of tissue types. Materials Animal tissues (e.g., liver, kidney, and/or spleen) from sacrificed animal(s) or biopsy Nuclear buffers A, B, and C (see recipes) Calcium- and magnesium-free (CMF) PBS (e.g., APPENDIX 2) 1 M NaOH 2× TNESK solution (see recipe) 0.1 M CaCl2 Micrococcal nuclease (MNase) stock solution (see recipe) Razor blades or scalpels 100-mm petri dishes on ice 10- and/or 15-ml tissue homogenizers with Teflon-coated pestles Tissue grinder motor and chuck Phase-contrast microscope Cheesecloth 15- and 30-ml glass centrifuge tubes (e.g., Corex) Refrigerated high-speed centrifuge with fixed-angle rotor Ultracentrifuge with SW50.1 rotor and 1⁄2 × 2–in. Ultraclear tubes Ultraviolet light spectrophotometer NOTE: All solutions, test tubes, pipets, and pipet tips should be ice cold. Carry out nuclear isolation 1. Immediately after sacrificing animal or obtaining biopsy, rinse isolated tissues in 25 ml nuclear buffer A in a 100-ml glass beaker on ice. For mice, combine tissues from four mice >1 month old. Do not freeze tissue or nuclei.

2. Transfer tissues to 100-mm plastic petri dishes on ice containing the following amounts of nuclear buffer A: liver, 8 ml buffer A; kidney, 5 ml; spleen, 4 ml. Mince tissues into chunks several millimeters in size using razor blades or scalpels in a scissors-like motion. 3. Decant each minced preparation into a separate homogenizer. For the aforementioned amounts of tissue, use a 15-ml apparatus for liver or kidney and a 10-ml apparatus for spleen. Let spleen fragments settle on ice ∼10 min before decanting, allowing red blood cells to remain in the supernatant. For liver and kidney fragments, decant supernatants immediately. 4. Add 5 ml fresh nuclear buffer A to all samples. Mix, decant supernatants again, and resuspend the tissue fragments into 5 ml fresh nuclear buffer A. Micrococcal Nuclease Analysis of Chromatin Structure

IMPORTANT NOTE: From this point on, perform all manipulations in the cold room.



5. Homogenize tissues with five to ten strokes of a motor-driven tissue grinder, followed by an additional five sharp strokes by hand. Dilute a small portion of the homogenate into CMF PBS and view under a phase-contrast microscope. Nuclei appear as grey, evenly round blobs. If intact tissue or cells remain, perform an additional five strokes or so with the homogenizer and check once again for quantitative cell breakage. Some tissues, such as spleen, require considerably less effort to disrupt than liver. Do not proceed until the cells are quantitatively broken.

6. Filter each homogenate through eight layers of folded cheesecloth that has been prewet with nuclear buffer A into a 30-ml Corex tube on ice. Wearing gloves, twist the cheesecloth to elute most of the liquid. 7. Layer homogenates onto cushions of 1:1 nuclear buffer A/nuclear buffer B in 15-ml Corex tubes, using the following volumes of the buffer A + B mixture: liver, 1.4 ml; kidney, 1 ml; spleen, 0.6 ml. 8. Centrifuge 15 min at 12,000 × g, 4°C, in a fixed-angle rotor (e.g., 10,000 rpm in 55-34 rotor). The liver and kidney pellet should be large, with a bright red center surrounded by brown. The spleen pellet should be small and brown.

9. Discard supernatants and resuspend pellets into the following volumes of nuclear buffer B: liver and kidney, 11 ml; spleen, 3.6 ml. First add ∼2 ml to each pellet and gently resuspend into a thick slurry, then add remaining nuclear buffer B. 10. Layer 3.6 ml of each suspension over 1.2-ml cushions of nuclear buffer B in 1⁄2 × 2–in. Ultraclear SW50.1 centrifuge tubes (liver and kidney in three tubes; spleen in one). 11. Spin 90 min at 120,000 × g (e.g., 37,000 rpm in SW50.1 rotor) 4°C. 12. Decant by inverting tube and blotting rim on a Kimwipe tissue. Gently resuspend each pellet into the following volumes of nuclear buffer C: liver and kidney, 0.2 ml; spleen, 0.25 ml. Combine suspensions from a single tissue type into a single tube and keep on ice. 13. Dilute 5 µl of each nuclear suspension into 2 ml of 1 M NaOH and measure the OD260 by spectrophotometry. Dilute the nuclear suspensions with nuclear buffer C to obtain the following diluted OD260 values: liver, OD260 = 0.25; kidney and spleen, OD260 = 0.085. Carry out MNase digestion of isolated nuclei 14. Remove one-fifth of each nuclear suspension and mix with an equal volume of 2× TNESK. Mix very well. Incubate overnight at 37°C. This sample serves as a “quench” control for the extent of nuclear digestion prior to warming up the nuclei.

15. Divide remaining volume into the following number of reaction tubes on ice: liver and kidney, up to six; spleen, about three. Be sure to agitate samples frequently to prevent clumping. Each reaction tube will be used for a different concentration of MNase.

16. Add 0.1 M CaCl2 to the first tube to a final concentration of 3 mM, incubate tube 3 min at 37°C, then add equal volume of 2× TNESK and incubate overnight at 37°C. This samples serves as an essential control to assess the level of endogenous nuclease. Swish the tube around in the 37°C water bath to facilitate warming.



Supplement 45

17. Add 0.1 M CaCl2 to the next tube to a final concentration of 3 mM and incubate tube 1.5 min at 37°C to warm nuclei. 18. Add the desired amount of MNase to each tube. For partial MNase digestions, try the following ranges: liver and kidney, 0.1 U MNase per ml reaction, 0.2 µ/ml, 0.5 µ/ml, 1 µ/ml, and 2 µ/ml; spleen, 0.5 µ/ml and 1 µ/ml. For extensive MNase digestions, try a range of 10 to 100 U/ml. Mix, and incubate tube 1.5 min at 37°C. 19. Terminate reactions by adding an equal volume of 2× TNESK and shaking the tube sharply a few time to mix, then incubate overnight at 37°C. 20. Repeat steps 17 to 19 in succession for each MNase concentration to be used. Do not add the CaCl2 to all tubes at the same time. Do each CaCl2 addition and MNase digestion separately.

21. Proceed to purification and characterization of DNA (see Support Protocol 1). BASIC PROTOCOL 3

MICROCOCCAL NUCLEASE DIGESTION OF PURIFIED GENOMIC DNA The enzyme MNase has a nonrandom sequence preference for cleaving DNA. It is therefore essential to compare the MNase cleavage pattern of chromatin with that for free genomic DNA in order to determine how the inherently nonrandom pattern is influenced by nuclear proteins such as the histones. MNase-digested free DNA, digested to the same relative extent as DNA from a chromatin digest, should be analyzed side-by-side in the cleavage mapping studies described in Support Protocols 2 and 3 below. Materials 0.5 to 1 mg purified genomic DNA (see Support Protocol 1) Nuclear buffer C (see recipe) 0.5 M CaCl2 Micrococcal nuclease (MNase) 0.25 M EGTA Chloroform Heating block set to 68°C Bucket of ice Additional reagents and equipment for agarose minigel electrophoresis (UNIT 2.5A) 1. Dilute 100-µg aliquots of genomic DNA into a final volume of 300 µl nuclear buffer C at room temperature. Set up four or five such aliquots. IMPORTANT NOTE: For each aliquot, carry out reaction through step 4 before starting the next. The reaction is carried out at room temperature, rather than 37°C, to provide better control over the digestion conditions.

2. Add CaCl2 to 3 mM final concentration. 3. Add MNase to 0.5, 1, 2, or 3 U/ml. Mix sharply to distribute the enzyme evenly (the solution may be viscous). Incubate 3 min. 4. Aspirate solution with a micropipettor and transfer to a tube containing 50 µl of 0.25 M EGTA, all prewarmed at 68°C. Keep tube in heat block for 10 min. 5. Transfer tube to ice bucket. Micrococcal Nuclease Analysis of Chromatin Structure

6. Electrophorese 200-ng samples on a 1.2% agarose minigel to check the extent of digestion. It is useful to run these MNase digestion products alongside chromatin DNA samples (see Support Protocol 1), so that the ranges of digestion are similar.



7. Extract the samples of interest once with 1 vol chloroform. 8. Proceed to precipitation, quantitation, and assessment of the DNA (see Support Protocol 1, step 7 onward). PURIFICATION AND CHARACTERIZATION OF DNA FROM CHROMATIN DIGESTIONS

SUPPORT PROTOCOL 1

This protocol has been optimized to maximize the recovery of small amounts of DNA from chromatin digests, and so that the resulting DNA is suitable for direct analysis and should not require further concentration. The protocol begins with a cell or nuclear lysate in an SDS solution that has been incubated with proteinase K for several hours or overnight (this can be used for any small-scale DNA preparation; e.g., see Basic Protocols 1 and 2). Materials MNase-digested cell or nuclear lysate (see Basic Protocols 1, 2, and 3) TE buffer, pH 7.9 (APPENDIX 2) Neutralized phenol (see recipe) Chloroform Ether (for permeabilized cell preparations) 5 mg/ml RNase A (for permeabilized cell preparations) 3 M sodium acetate 95% and 70% ethanol, room temperature 10- or 12-ml polypropylene tubes with tight caps (e.g., 12-ml Sarstedt tubes) Shaker or rocking device 6-in. Pasteur pipets 30-ml Corex tubes, silanized (APPENDIX 3B; for permeabilized cell preparations) 6000- to 8000-MWCO dialysis tubing (for permeabilized cell preparations) 1- to 5-µl glass capillary pipets Glass capillary pipettor Plastic wrap (e.g., Saran Wrap) Ultraviolet light spectrophotometer Additional reagents and equipment for dialysis (APPENDIX 3C) and agarose minigel electrophoresis (UNIT 2.5A) Carry out organic extractions of MNase-digested DNA 1. Dilute the cell or nuclear lysate with 1 vol TE buffer, pH 7.9. If the resulting volume is ≤0.5 ml, perform the following steps in a 1.5-ml microcentrifuge tube; if the volume is ≥0.5 ml, transfer to 12-ml polypropylene tube(s). Diluting the material will make the solution less viscous and reduce loss of DNA during the subsequent manipulations. A marked decrease in viscosity of the MNase-treated samples should be evident at this step, and indicates successful cleavage.

2. In a fume hood, add 1 vol neutralized phenol. Invert tube sharply several times, then place on a gentle shaker or rocking device for 10 to 20 min. 3. Centrifuge the samples 30 sec in a microcentrifuge for those in 1.5-ml tubes, or 5 min at 500 × g (2000 rpm in 55-34 rotor) for those in 12-ml tubes. Using a 6-in. Pasteur pipet, remove the upper aqueous layer and transfer to a fresh tube. There is little worry about shearing the DNA here, as it should be significantly digested by the MNase and the “0 MNase” samples will be digested with restriction enzymes during analysis.



Supplement 45

4. Add 1 vol chloroform. Invert tube sharply several times, then gently shake for 10 to 20 min. Spin and obtain upper aqueous phase as in step 3. Leave the interface behind. 5. Repeat chloroform extractions for a total of two to four extractions, until the interface between the organic and aqueous phases is clear. For samples that were treated with MNase, two or three chloroform extractions should be sufficient; for samples that did not receive MNase, the aqueous phase should be very viscous and may not clear completely even after four extractions. For MNase digestions of isolated nuclei and purified DNA, proceed directly to step 7 of this protocol. For MNase digestions of permeabilized cells, proceed to step 6 below.

6. For permeabilized cell preparations: Add 1 vol ether, then invert tube sharply several times, burp the tube, and gently shake or rock for 10 to 20 min. Heat uncapped for 10 to 15 min in a 65°C water bath, shaking periodically to help evaporate the ether, then dialyze overnight in 6000- to 8000-MWCO dialysis tubing against two changes of 100 vol TE buffer, pH 7.9, at 4°C. Transfer the dialysate, which will have increased in volume by ≥50%, to a 30-ml silanized Corex tube and add RNase to 25 µg/ml. Incubate 1 hr at room temperature. The dialysis procedure alleviates the problem of lysolecithin (which can inhibit restriction enzymes) precipitating with the DNA in later steps, and the RNase treatment digests the cytoplasmic RNA present in the permeabilized cell lysates.

7. Add 1⁄10 vol of 3 M sodium acetate. Mix well by inversion. Precipitate, spool, and resuspend DNA precipitate IMPORTANT NOTE: For each sample, perform steps 8 to 12 in succession without stopping. 8. Add 2.5 vol of 95% ethanol. Invert gently 10 to 30 times, until the Schlieren patterns in the liquid are completely gone, and assess the solution to determine how to proceed. a. If the DNA has not been extensively digested with MNase, a stringy white precipitate should be visible; continue with steps 9 to 13 (and omit steps 14 to 17). b. If the DNA has been extensively digested with MNase, a precipitate may not be visible; proceed directly to step 14 (beginning with overnight incubation at −20°C). c. With a moderate amount of MNase digestion, there may be a small stringy precipitate; spool this out by following steps 9 to 13, and proceed directly to step 14 (beginning with overnight incubation at −20°C) to recapture DNA from the remaining solution. The DNA from MNase-digested permeabilized cells will probably be too dilute to form the stringy precipitate, in which case proceed directly to step 14.

9. Use a 1- to 5-µl glass microcapillary tube to spool out the DNA precipitate from a single tube. Use a microcapillary pipettor to expel liquid that rises into the capillary while touching the DNA to the inside of the tube to remove excess liquid. Do this step over a sheet of plastic wrap spread on the laboratory bench. If the spooled DNA falls off the microcapillary, simply use the microcapillary to pick it up from the plastic wrap and carry on.

10. Immediately dip the DNA in a 1.5-ml microcentrifuge tube containing 70% ethanol. Swish around for a few seconds, then again blow out liquid from the tube while removing excess liquid from the DNA. Micrococcal Nuclease Analysis of Chromatin Structure

Work quickly; do not allow the DNA to dry out or it will be impossible to resuspend.

11. Blot the DNA on the inside of an empty 1.5-ml tube. Air dry for ∼5 sec.



12. Place the DNA in a 1.5-ml tube containing 75 to 300 µl TE buffer (depending on the amount of DNA spooled) for isolated nuclei or free DNA experiments, or 400 µl TE for permeabilized cells. Discard the capillary tube. Remember to go on to the next sample, starting at step 8. Do not add ethanol to multiple samples at the same time; if the DNA is kept as a stringy precipitate for too long it will be very difficult to resuspend.

13. Let DNA dissolve overnight at room temperature. Vortex lightly a few times, then store at 4°C. Do not freeze. The DNA is stable for years at 4°C.

Precipitate dilute DNA remaining in solution 14. If the DNA does not precipitate at step 8, or if only a small precipitate can be spooled out, incubate the remaining mixture at −20°C overnight. 15. If using 12-ml polypropylene or 30-ml Corex tubes, centrifuge 10 min at 10,000 × g in a fixed-angle rotor (e.g., 9000 rpm in 55-34 rotor); if using 1.5-ml tubes, spin 5 min in a microcentrifuge. Decant supernatant. Add 0.5 ml of 70% ethanol and gently invert the tube once. Repeat centrifuge spin for 1 min. 16. Decant supernatant. Vacuum dry pellet for 5 min. For experiments with isolated nuclei or free DNA, resuspend in 75 to 300 µl TE buffer (depending on the size of the pellet and anticipated amount of DNA); for experiments with permeabilized cells, resuspend the DNA in 400 µl TE. If some DNA from the sample was spooled out and resuspended in TE, use this solution to resuspend the DNA pellet, so the portions are pooled and kept concentrated. Allow solution to dissolve overnight at room temperature and then store at 4°C, as in step 13. 17. For DNA prepared from permeabilized cells: Repeat the precipitation starting at step 7 and resuspend the final precipitate in 75 to 150 µl TE buffer, so that the DNA is sufficiently concentrated for subsequent analysis. Quantitate and assess DNA 18. Dilute 4 µl DNA solution in 400 µl (total) TE buffer and read the OD260. Calculate DNA concentration and total yield, assuming that an OD260 reading of 1 equals 50 µg/ml of DNA. If DNA is very viscous, take up 4 ìl of water in pipet tip and mark the meniscus with a felt tip pen; expel and then use marked tip to take up 4 ìl of the DNA solution.

19. Apply 0.5-µg samples of each DNA to a 1.2% agarose minigel and assess the level of endogenous nuclease and MNase cleavage of the chromatin. A ladder of bands consistent with the nucleosomal repeat length should be evident.

NUCLEASE CLEAVAGE MAPPING STRATEGIES Very different approaches are taken to map extensive versus partial MNase digestions, and it is important to know what patterns to look for in the final data, as described below. The methodology for running the DNA samples on agarose gels, blotting to filters, and hybridizing to radiolabeled probes involves well-established technology described in UNITS 2.5A, 2.9A & 2.10, respectively. Mapping MNase cleavages at the nucleotide level of resolution requires special modifications of existing protocols and is described in Support Protocol 3. Cleavage Mapping of an Extensive MNase Digest DNA from a chromatin sample that was extensively digested with MNase is typically assessed by performing gel electrophoresis (e.g., 1.2% to 1.5% agarose or 5% polyacry-

SUPPORT PROTOCOL 2



Supplement 45

lamide) to resolve mononucleosomal fragments, blotting to a nylon filter, and then hybridizing to an oligonucleotide or short DNA probe corresponding to the DNA region of interest. Visualization of a mononucleosome-sized hybridization band is evidence that some fraction of the templates in the chromatin population was nucleosomal during the MNase digestion. If an oligonucleotide probe from a different part of the genome is hybridized to another portion of the original chromatin digest, then the signals from the two probes can provide information about the relative frequencies with which the two DNA sequences were nucleosomal in the original population of chromatin templates. The specific activities of the oligonucleotide probes must be comparable or accounted for. Very highly labeled probes are required to observe mononucleosome-sized bands from single-copy higher-eukaryotic gene sequences. Cleavage Mapping of a Partial MNase Digest by Indirect End Labeling DNA that is isolated from a chromatin sample that was only partially digested with MNase is typically assessed by an indirect end-labeling experiment (Wu, 1980; see Fig. 21.1.1). The indirect end-labeling approach is designed to map, at the Southern blot level of

A

free DNA [MNase] 0

chromatin 1 chromatin 2

0

0 "parental" EcoRI fragment

electrophoresis

–

MNase-generated sub-bands

+

B

MNase cleavages

EcoRI

EcoRI

HindIII

HindIII

probe DNA DNA extent of MNase-generated sub-bands in chromatin sample 2

DNA extent of internal control sub-bands from HindIII partial digest in lane C


Figure 21.1.1 (A) Hypothetical Southern blot of indirect end-label analysis of MNase-treated chromatin. Expected variation in band intensities are shown with increasing MNase digestion. Lack of sub-bands in free DNA is due to lack of intrinsic MNase hypersensitivity, and lack of sub-bands in chromatin sample 1 is due to lack of nucleosome phasing in the region probed. (B) Mapping strategy and interpretation of bands depicting a positioned nucleosome array.



resolution, the positions of frequent cleavage by MNase at DNA sites in a population of chromatin templates. If multiple MNase cleavage sites occur 160 to 200 bp apart, then nucleosomes may be positioned over the sequence of interest. The variation in the nucleosome repeat length depends upon the organism and other parameters, such as the presence of linker histone in the local chromatin domain. It is also possible to map double-stranded MNase cleavages at the nucleotide level of resolution, using a modification of the LM-PCR technique (see Support Protocol 3). For indirect end-labeling, the MNase-treated, purified DNA is digested with one or more restriction enzyme(s) that cut(s) at least 1 kb away on both sides of the region of interest (Fig. 21.1.1; see example for EcoRI). The genomic DNA is then electrophoresed, blotted to nylon or nitrocellulose membrane, and then hybridized to a purified DNA fragment probe that is 0.5 to 1 kb in length and that abuts one restriction-cut end of the genomic DNA target sequence (Fig. 21.1.1; EcoRI-HindIII fragment). Autoradiographic or phosphorimager exposures should reveal the “parent” restriction fragment from control chromatin samples that were not treated with MNase, and the presence of sub-bands in MNase-treated samples. Discrete sub-bands will appear if MNase cleaves at the same position(s) on most of the substrates in the population (Fig. 21.1.1; chromatin sample 2). The positions of cleavages can be determined by the distance in kilobases from the restriction enzyme site abutted by the probe fragment (Fig. 21.1.1, Mapping strategy). If a ladder of sub-bands is evident with the aforementioned spacing, it suggests the presence of a positioned nucleosome or a nucleosome array (Fig. 21.1.1, ovals in diagram in Mapping Strategy). Inclusion of an internal marker control allows for a definitive mapping of the position of MNase cleavage in an indirect end-label assay. An aliquot of uncut genomic DNA (designated the internal marker control), equal to or less than the amount of the chromatin samples to be loaded onto the gel for Southern blotting, is digested to completion with the restriction enzyme(s) used for the chromatin samples. The internal marker control sample is then digested partially with restriction enzyme(s) that cleave in the vicinity of the region of interest: i.e., where nucleosome phasing is being investigated (Fig. 21.1.1; HindIII partial digest). The marker control sample is then loaded onto the same gel as the chromatin samples. It should yield hybridization sub-bands due to the partial restriction enzyme cleavages. These sub-bands provide the ideal standard for the mobility of sub-bands in the MNase digested samples, and allow accurate mapping of the MNase cleavage sites with respect to the positions of the restriction sites from the partial digest (see Liu et al., 1988; McPherson et al., 1993). Regarding technical details: 12 µg of genomic DNA per lane are sufficient for the Southern blots; large (e.g., 25-cm) 1.2% agarose gels run overnight at 40 to 50 V give the best resolution; and DNA probes should be labeled with 32P to a specific activity of at least 0.5 × 109 cpm per microgram of DNA, preferably in the range of 109 cpm/µg.



Supplement 45

SUPPORT PROTOCOL 3

USING A MODIFIED LM-PCR PROCEDURE TO MAP DOUBLE-STRANDED MNase CLEAVAGES AT THE NUCLEOTIDE LEVEL OF RESOLUTION The ligation-mediated PCR (LM-PCR; UNIT 15.5) procedure is used for genomic footprinting and therefore can be used to map MNase cleavages at the nucleotide level of resolution. However, conventional LM-PCR detects single-stranded genomic cleavages, whereas MNase makes single-stranded DNA cleavages on nucleosomes. In order to selectively map these double-stranded intranucleosomal MNase cleavages, modifications to the LM-PCR procedure have been developed (McPherson et al., 1993), as described here. See Figure 21.1.2 for a comparison of nucleosome and conventional LM-PCR footprinting. The essential change that is made in the LM-PCR protocol is to eliminate the initial primer extension step of LM-PCR and directly ligate the asymmetric linker to the MNase-cleaved double-stranded DNA. But because MNase cleavage leaves a 5′ hydroxyl, it is necessary to first phosphorylate the cleaved double-stranded substrate so that the long linker strand can be ligated to it (see Fig. 15.5.3). Note that a DMS cleavage pattern with genomic DNA, generated by conventional LM-PCR and run on the same gel as the final MNase LM-PCR products, is necessary to determine the positions of MNase cleavages along the sequence. Materials DNA from MNase digest of chromatin (e.g., see Basic Protocols 1 and 2; see Support Protocol 1 for purification procedure) DNA from MNase digest of purified genomic DNA (e.g., see Basic Protocol 3) 10 U/µl T4 polynucleotide kinase (e.g., New England Biolabs) and 10× buffer (supplied with enzyme)

NUCLEOSOME FOOTPRINTING

CONVENTIONAL FOOTPRINTING

MNase-treated DNA

DNase- or DMS-treated DNA

(skip steps 1-5 of UNIT 15.5)

denature; anneal primer 1 (UNIT 15.5, steps 1-3)

5' phosphorylate with kinase (UNIT 21.1, Support Protocol 3)

perform first-strand synthesis (UNIT 15.5, step 3)

ligate double-stranded linker to blunt ends (UNIT 15.5, steps 4-9)

PCR amplify with linker-oligo and primer 2 (UNIT 15.5, steps 10-16)

primer extend with labeled primer 3 (UNIT 15.5, steps 17-24)

analyze on sequencing gel


Figure 21.1.2 Nucleosome (MNase; left) versus conventional footprinting (right) by ligation-mediated PCR. Step numbers in UNIT 15.5 refer to the basic protocol.



10 mM ATP 0.5 M EDTA, pH 8 (APPENDIX 2) 3 M sodium acetate 95% and 70% ethanol, room temperature 1. Set up phosphorylation reactions as follows in 1.5-ml microcentrifuge tubes—one for each MNase enzyme digestion point, as well as for no-enzyme and MNasetreated, purified DNA controls: 5 µl 10× kinase buffer 0.5 µl 10 mM ATP H2O to give 50 µl final volume 5 µg MNase-cleaved, purified genomic DNA. 2. Add 1 µl of 10 U/µl T4 polynucleotide kinase (10 U) and incubate 1 hr at 37°C. 3. Add 1 µl of 0.5 M EDTA and incubate 20 min at 68°C. 4. Add 5 µl of 3 M sodium acetate, mix, then add 125 µl of 95% ethanol, mix, and incubate 30 min at −20°C to precipitate DNA. 5. Spin tube 5 min in a microcentrifuge, decant supernatant, add 125 µl of 70% ethanol, and gently invert once. Spin 2 min, decant supernatant, and dry briefly under vacuum. 6. Suspend the DNA in the tube in water at 0.1 to 0.2 µg/µl and store at −20°C until use. This DNA can be used directly for LM-PCR; proceed directly to the linker ligation step (step 4 in the basic protocol of UNIT 15.5). Use several more cycles of the PCR reaction (e.g., 22 cycles instead of 20) than would be used for conventional LM-PCR.


MNase stock solution Dissolve 50 activity units MNase (Worthington Biochemicals) per microliter in 5 mM Tris⋅Cl (pH 7.5; APPENDIX 2)/0.01 mM CaCl2. Divide into aliquots in 0.5-ml tubes and store at −20°C. One activity unit of MNase = 85 OD A260 units.

Neutralized phenol Combine 20 ml phenol with 20 ml of 100 mM Tris⋅Cl (pH 7.5) in a 50-ml tube. Mix by inversion. After the phases separate, the lower, organic phase will contain neutralized phenol. Nuclear buffer A 15 mM HEPES, pH 7.5 60 mM KCl 15 mM NaCl 2 mM EDTA 0.5 mM EGTA 0.34 M sucrose 0.15 mM 2-mercaptoethanol (add just before use) 0.15 mM spermine (add just before use) 0.5 mM spermidine (add just before use) The above was designed for homogenizing liver tissue (Hewish and Burgoyne, 1973; Kornberg et al., 1989). For other tissues, add 0.5% nonfat dry milk (e.g., Carnation).



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Nuclear buffer B 15 mM HEPES, pH 7.5 60 mM KCl 15 mM NaCl 0.1 mM EDTA 0.1 mM EGTA 2.1 M sucrose 0.15 mM 2-mercaptoethanol (add just before use) 0.15 mM spermine (add just before use) 0.5 mM spermidine (add just before use) Nuclear buffer C 15 mM HEPES, pH 7.5 60 mM KCl 15 mM NaCl 10 mM NaHSO3 0.34 M sucrose 0.15 mM 2-mercaptoethanol (add just before use) 0.15 mM spermine (add just before use) 0.5 mM spermidine (add just before use) Permeabilization solution 1 150 mM sucrose 80 mM KCl 35 mM HEPES, pH 7.4 5 mM K2HPO4 5 mM MgCl2 0.5 mM CaCl2 Permeabilization solution 2 150 mM sucrose 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 50 mM NaCl 2 mM CaCl2 Permeabilization stop solution 20 mM Tris⋅Cl, pH 8 (APPENDIX 2) 20 mM NaCl2 20 mM EDTA 1% SDS 0.6 mg/ml proteinase K (add just before use) TNESK, 2× 20 mM Tris⋅Cl, pH 7.4 (APPENDIX 2) 0.2 M NaCl 2 mM EDTA 2% SDS 0.2 mg/ml proteinase K (add just before use)




COMMENTARY Background Information It is increasingly appreciated that chromatin structure is integral to the mechanisms of transcriptional regulation, DNA replication, and DNA repair (van Holde, 1989; Felsenfeld, 1992). The basic repeating unit of chromatin is the nucleosome core particle, which consists of an octamer of two each of the four core histone proteins, along with ∼146 bp of DNA. The addition of linker histone to the core particle creates the nucleosome and increases the amount of associated DNA by ∼20 bp. Nucleosomes and nucleosome cores occur in extensive arrays that span the eukaryotic genome and elicit higher levels of chromatin compaction. Since the histone octamers may span DNA sequences that are important for transcription factors, polymerases, and DNA modification enzymes, understanding precisely where the nucleosomes occur can provide insight into the role of chromatin structure during physiological conditions of interest. With the modifications to standard DNA footprinting procedures described in Support Protocol 3, it is now possible to map doublestranded genomic MNase cleavages at the nucleotide level of resolution. This is useful for comparing the apparent positions of nucleosome boundaries to the positions of regulatory-protein binding sites and other genetic landmarks (see McPherson et al., 1993). This method can also be used to compare nucleosome boundaries in vivo with those obtained from in vitro chromatin reconstitution reactions (Shim et al., 1998).

Critical Parameters and Troubleshooting For the cell permeabilization in Basic Protocol 1, which is essentially the procedure described by Pfeifer and Riggs (1991), it is best to start with trials to determine the optimal lysolecithin concentration and incubation temperature and select a workable range of MNase that yields the desired level of digestion. The cells must be checked at various stages of the experiment to be sure that they have not lysed. If cell lysis has occurred, repeat the experiment with a lower concentration of lysolecithin or at a lower temperature. Lysolecithin seems to be more potent when fresh than after it has been stored for several months. Conversely, if agarose gel analysis of the bulk chromatin samples reveals a weak ladder of MNase-generated bands and a considerable amount of undigested

genomic DNA, it is likely that an insufficient fraction of the cells were permeabilized, and a higher lysolecithin concentration, longer incubation, or higher incubation temperature may be needed. Endogenous nucleases can be a major problem when isolating nuclei from certain tissues and cell lines using Basic Protocol 2. A simple way to reduce their effect is to work quickly during the nuclear isolation and chromatin digestion procedures and keep the chromatin on ice as much as possible. It is also helpful to minimize the time taken to warm the sample for the MNase digestion and to use a sufficient concentration of enzyme that the desired amount of digestion is complete within a few minutes. The nuclear isolation buffer contains EDTA to chelate divalent cations that activate endogenous nucleases; spermine and spermidine are included to provide counter-ions that stabilize the chromatin. To make the nuclear isolation in Basic Protocol 2 readily adaptable for formaldehyde cross-linking experiments, the original protocols of Hewish and Burgoyne (1973) and Kornberg et al. (1989) have been modified here with a Hepes buffer system instead of Tris, because formaldehyde reacts with primary amines in the latter. Spermine and spermidine must be avoided for the same reason, so for formaldehyde-based cross-linking, the cell lysis and nuclear isolations buffers should contain 2 mM MgCl2 to stabilize the chromatin and lack both EDTA and molecules with primary amines. For partial MNase digestion studies with indirect end-label probes, as in Support Protocol 2, sub-bands are best seen when ∼25% to 75% of the parent restriction fragment is cleaved (i.e., depleted) by MNase. It will be necessary to empirically determine the appropriate concentration of MNase by varying enzyme levels in two-fold steps over the suggested range. Excessive MNase digestion can lead to the artifactual appearance of a positioned nucleosome array because short oligonucleosome fragments bounded by MNase cleavages on both sides can hybridize to the indirect end-label probe. Note that the activity of MNase is highly dependent upon the concentration of enzyme; a two-fold difference in concentration can yield a marked difference in the average size of double-stranded DNA cleavage product. Another important parameter is the activity state of the genetic region of interest. Silent



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chromatin is often highly compacted and may require a ≥10-fold enzyme concentration to cleave a specific genomic sequence to the same extent as when the region is “open” or expressed. Thus, it may require a large difference in bulk chromatin MNase digestion in Basic Protocols 1 and 2 for two samples to exhibit the same extent of cleavage of a specific target gene. For LM-PCR mapping of nucleosome boundaries, it is critical to first define the general region to be analyzed by mapping MNase cleavages at the Southern blot level of resolution. Chromatin digest samples that just begin to exhibit a laddering band pattern on a Southern blot (see Fig. 21.1.1A, first lane or two of chromatin sample 2) serve as the best substrates for the modified LM-PCR protocol; i.e., underdigested DNA rather than over-digested. Once a set of genomic DNA samples is obtained where the extents of bulk chromatin cleavage by MNase are known to give interesting, chromatin-specific results, it is convenient to use these samples as standards for subsequent experiments. That is, 0.5-µg aliquots of the MNase digested chromatin samples from the known experiment can be run on a 1.2% agarose minigel alongside DNA aliquots from a new chromatin experiment from cells under similar physiological conditions. The gel can be stained with ethidium bromide and samples in the new chromatin experiment which exhibit the same extent of bulk genomic digestion, or laddering, can often be assumed to be worthy of further analysis by the aforementioned hybridization approaches.

Anticipated Results


For an extensive MNase digest, it is critical to compare the intensity of mononucleosomesized bands hybridizing to DNA probes from different parts of the genome. For two probes of comparable specific activity, if the hybridizing bands are of similar intensity, it indicates that the same fraction of target sequences in the population were nucleosomal. Alternatively, if the hybridizing band is markedly more intense for one probe than for another, then more gene copies in the population hybridizing to the first probe were nucleosomal than in the population hybridizing to the second probe. It is important to note that nucleosomes do not have to be phased, or positioned, over a DNA sequence in chromatin in order to give clear hybridization signals indicating that the sequence is nucleosomal. Nucleosomes that are randomly positioned over the region spanned

by an oligonucleotide will, on average, yield a nucleosomal-sized band upon hybridization to extensively digested chromatin. Thus, if indirect end-labeling analysis indicates a lack of nucleosome phasing over a region, it is inappropriate to conclude that the region is free of nucleosomes. The extensive MNase digestion assay can address this issue. For a partial MNase digest and indirect endlabeling analysis, it is critical that each subband generated by MNase treatment has one end resulting from restriction enzyme cleavage. If the chromatin sample generates a ladder of hybridizing sub-bands that differ in size by ∼180 to 200 bp, it suggests the presence of a phased nucleosome array. In this case, the size of a sub-band indicates the distance between the restriction enzyme cleavage site and the position of double-stranded MNase cleavage. The hybridizing ladder of bands should not be smaller than the indirect end-labeled probe; if it is, the band sizes are probably not generated by restriction enzyme cleavage on one side and thus fragment endpoints cannot be predicted. Ideally, internal marker controls are chosen so that the marker bands occur in the region of MNase-generated sub-bands, so that the positions of MNase cleavage can be determined with accuracy. A high level of accuracy in mapping MNase cleavage sites in chromatin, by this method, is useful for planning a primer probe strategy for LM-PCR analysis. The clusters of MNase cleavages that define apparent linker regions between nucleosomes, by indirect end-labeling, should be evident in the LM-PCR analysis. That is, what appears as a single sub-band on a Southern blot should appear as a cluster of bands on an LM-PCR sequencing gel at about the positions that are predicted by the internal control marker on the Southern blot. Additional bands will probably be seen in the LM-PCR analysis, but if they do not occur in a cluster they may be insufficient to generate a sub-band on the Southern blot. It is worth noting that further evidence, such as may be provided by protein-DNA cross-linking experiments, should be obtained before concluding that an apparent “nucleosomal” ladder of MNase-generated bands is indeed caused by histone octamers on the DNA.

Time Considerations Cell permeabilization (Basic Protocol 1) is much easier and quicker to perform than nuclear isolation (Basic Protocol 2). The entire cell permeabilization and MNase digestion procedure should take 90%) should be in the size range of 100 to 1000 bp. As an alternative to sonication, DNA fragment size can be reduced by treatment of the cross-linked chromatin with micrococcal nuclease (UNIT 3.12). Micrococcal nuclease preferentially cleaves DNA located in the linker regions between nucleosomes. By varying the concentration of micrococcal nuclease, it is possible to generate samples in which average DNA size varies. The minimal useful size is about 150 bp, which corresponds to a mononucleosome. However, cleavage to mononucleosome-sized fragments may also result in a preferential loss of certain genomic regions due to the sequencespecificity of micrococcal nuclease.

Immunoprecipitation The success of this procedure relies on the use of an antibody that will specifically and tightly bind its target protein in the buffer and wash conditions used. In addition, antibody should be present in excess with respect to its target protein so that differences in the amounts of the protein-DNA complexes of interest will be accurately measured. Perform preliminary experiments to confirm avid immunoprecipitation and determine an approximate amount of antibody to use. Chromatin extracts should be prepared without prior cross-linking of the cells and subjected to immunoprecipitation with varying concentrations of antibody (20 µg/ml antibody may be a good starting point). The amount of the protein of interest in the extracts before and after immunoprecipitation should be analyzed by immunoblotting (UNIT 10.8) to determine the lowest antibody concentration that depletes >90% of the protein of interest from the extract. This antibody concentration is a good starting point for the full protocol and may later be modified to maximize the signalto-noise ratio (see Anticipated Results). With cross-linked chromatin, immunodepletion of the target protein is less efficient (∼50%), presumably due to masking or modification of the epitopes, and a significant amount of the protein remains refractory to immunoprecipitation even with higher antibody concentrations. Thus, the ideal antibody concentration is ultimately determined empirically to maximize the yield of specific coprecipitated DNA while minimizing precipitation of nonspecific DNA. Both monoclonal and polyclonal antibodies have been used in this procedure. The monoclonal antibodies 12CA5 (anti-HA), 17D09 (anti-HA), and 9E10 (anti-myc) have been used successfully in different laboratories. In general, triple-HA epitope tags work well (Hecht et al., 1996; Aparicio et al., 1997; Tanaka et al., 1997), and larger multi-myc epitope tags have also been successful (e.g., myc-9, myc-18; Tanaka et al., 1997). Protein G–Sepharose, Protein A–Sepharose, and anti-mouse immunoglobulin-coupled magnetic beads have all been used to precipitate the immune complexes, although it should be noted that certain classes of mouse and rat immunoglobulins are not strongly bound by protein A (Harlow and Lane, 1988). For optimal results, it is critical to minimize the background level of material that inevitably comes down during the immunoprecipitation. The procedures described here work well with a diverse set of antibodies, but it might be



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B

α-Orclp-HA

α-Orc2p/ α-Orc3p

precipitates W orT orc1 c W2 T or c or 1 c2

W T A– B1 – B2 – B3 –

no t no ag w X- l it h in X- k lin

k

A

ARS1 ARS305 URA3 1 2 3 4 5 6

7 8 9 10 11

121314 1516 17 37o 23o

Input DNA

ARS1 ARS305 URA3

Figure 21.3.2 Anticipated results from chromatin immunoprecipitation analysis of origin recognition complex (ORC) with replication origin and nonorigin DNA sequences.

Determining the Association of Proteins with Specific Genomic Sequences

necessary to modify the binding and elution conditions in specific cases. Peptide elution is clearly preferred over heat elution, as it is more specific and results in lower experimental backgrounds and hence higher-fold inductions. However, peptide elution is only possible for experiments using antibodies against peptides (typically for analyzing epitope-tagged proteins, but analysis of native proteins should also be possible). In performing peptide elution, it is important to add enough peptide such that the protein-DNA complexes are efficiently eluted from the beads. Another consideration is that the epitope of interest in the chromatin-bound protein might be inaccessible to the antibody due to associated proteins or DNA structures. In such a case, one might obtain a false-negative result. Whereas the majority of a given protein may be efficiently immunoprecipitated from the cross-linked cells, the fraction that is actually cross-linked might be undetectable. The use of polyclonal antibodies (which often recognize multiple determinants within a protein) or epitope-tagged proteins (the epitope is unlikely to have a specific interaction with other proteins or DNA sequences, particularly if the epitope does not affect the biological function as determined by genetic complementation) minimizes, but does not eliminate this concern. Because of this caveat, negative results should be interpreted cautiously and alternative meth-

ods (e.g., in vitro DNA binding or association of the protein with bulk chromatin) should be tried. This concern is particularly relevant when a protein of interest does not appear to interact with any genomic sequence. However, if a protein selectively associates with some genomic sequences, this concern is significantly reduced—i.e., it is unlikely that epitope masking will occur at some loci, but not others. PCR strategy The choice of primers depends on the experimental goals. If binding to a specific site is being tested, a primer pair that flanks the site and at least one control primer pair recognizing a DNA sequence not expected to bind the protein of interest are the minimal requirements (see Fig. 21.3.2). When choosing primers, it is important to remember that resolution between adjacent sequences is limited by the shear size of the DNA. For an average DNA size of 500 bp, a significant fraction of the DNA molecules will be in the 500 to 1000 bp range, and hence DNA sequences 1000 bp distal from the actual protein binding site may be coprecipitated. Therefore, primer pairs used as controls should amplify a region of DNA that is far enough away from the expected binding site (e.g., >1 kbp) that coprecipitation of adjacent DNA is not detected. A good strategy is to design multiple sets of primers at increasing distances from a suspected binding site. Such a strategy



has also been used to probe the “spreading” and “movement” of proteins on chromatin (Hecht et al., 1996; Aparicio et al., 1997; StrahlBolsinger et al., 1997). Success in obtaining high-quality quantitative data is critically dependent on good primer design! In general, primers should be 20 to 30 bases long with a Tm of 55° to 60°C. Most primers require no purification or special treatment prior to PCR. Amplification products should be 75 to 300 bp. Longer PCR products should be avoided, because the amplification efficiency is substantially lower, and DNA fragments that do not bind to both primers will not be amplified (this can be a significant problem since the size of DNA fragments in the samples averages ∼500 bp and ranges between 100 to 1000 bp). A final primer concentration of 1 µM works well for most primers, but in some instances, improved product specificity may be obtained by lowering the final primer concentration 5 to 10 fold. The design of good primers is greatly facilitated by commercially available software packages such as Oligo 6.6 or Primer Express 1.5. These packages allow for extensive customization of many different parameters, including Tm, oligonucleotide length, GC content, and more. While the success of each individual primer pair in the specific amplification of its target sequence is dependent on many variables, special care must be taken to minimize primer-dimers and hairpins. Finally, it is a good idea to check primers for hybridization to other genomic sequences through the use of a webbased program such as BLAST. Newly obtained primer pairs must be tested for amplification specificity and performance under the conditions that will be used in quantitative PCR. Primer pairs that are suitable for reactions performed by the Basic Protocol might not be suitable for real-time PCR reactions using SYBR Green, because SYBR Green can inhibit Taq polymerase. It is particularly informative to analyze input DNA amplification by the primers in question on high-percentage agarose or polyacrylamide gels after completion of the PCR. The presence of multiple product bands indicates poor specificity, and will invariably lead to unreliable results. For the Basic Protocol, the best test for quality of a given primer pair is to carry out a standard curve using different dilutions of DNA. For a high-quality primer pair, the amount of PCR product should be directly proportional to the amount of DNA, with an error of less than ±20%. The number of PCR

cycles is determined empirically. Usually, 25 to 28 cycles is appropriate. More than 28 cycles can result in detection of nonspecifically precipitated sequences and/or lead to variable results due to inactivation of Taq polymerase. Multiple primer pairs can be used in combination if the PCR products are separable by gel electrophoresis (as many as five have been used), but some combinations interfere with efficient amplification of one or more products. It is essential to test primer pairs singly and in combination, with titrations of template DNA, to determine if this is a problem. The advantage of using multiple primer pairs is that individual reactions can generate data for multiple genomic regions in an internally controlled manner. In addition, the Basic Protocol can be used to simultaneously analyze two alleles of a given locus in an internally controlled manner, provided the individual alleles result in different-sized PCR products. When quantitative PCR will be performed in real time using SYBR Green (see Alternate Protocol 2), high-quality primer pairs should result in ∼1.9-fold amplification/cycle. Such amplification efficiency can be determined from quantitative analysis of raw fluorescence data for each cycle. Amplification efficiencies 1.

If the higher complexes form a significant fraction of assembly, one can perform histone titration as above. If these complexes are not a significant fraction of the total assembly, they can be separated away from standard mononucleosomes on a glycerol gradient (Basic Protocol 2).

Less than 30% of the DNA is incorporated into nucleosomes.

The histone:DNA ratio is significantly less than 1.

Perform a titration with increasing amounts of histones.

Nucleosomes look “smeary” when The gel temperature is high causing run on gel. dissociation of nucleosomes. EcoRI digestion of array gives smeary bands on agarose gel.

Assembly of Nucleosomal Templates by Salt Dialysis

Maintain gel at room temperature for agarose gel and run acrylamide gel in cold room.

The running and gel buffer composition Use 45 mM Tris-borate, 1 mM EDTA is different from recipe. buffer (see Support Protocol 7).

30 nm in diameter. Unfortunately, details of this compaction, such as the conformation of the linker DNA within the 30-nm fiber or the packing of nucleosome cores within the fiber, have not been elucidated. These fibers are organized into even higher-order structures to form the metaphase chromosome. Efforts to recapitulate in vivo the processes which occur within the eukaryotic nucleus require the reconstruction of proper chromatin substrates. Thus, methods for the reconstitution of chromatin complexes have become widely used (Camerini-Otero, 1976). Moreover, reconstitution methods allow the construction of model chromatin complexes containing defined DNA sequences. Salt dialysis reconstitution has been useful in this respect. It is known that at 2 M salt, the histone octamer forms its canonical structure even when not bound to DNA. As the salt concentration falls below 2 M, the histone octamer dissociates into an H3/H4 and two H2A/H2B dimers. The tetramer also starts to bind to DNA at this salt concentration. At ~0.8 M salt, a single H2A/H2B dimer binds to each H3/H4 tetramer and by 0.6 M

nucleosome assembly is complete. Samples of reconstituted chromatin complexes can be prepared that exhibit less structural heterogeneity than preparations of natural chromatin complexes. This homogeneity has allowed detailed investigations of nucleosomal DNA structure and the salt-induced folding properties of nucleosomal arrays (Hayes et al., 1990, Schwarz and Hansen, 1994). Reconstituted nucleosomes exhibit physical and structural properties identical to native complexes isolated from eukaryotic nuclei. Hydroxyl radical cleavage analysis and micrococcal nuclease digestions have shown that the DNA structure within reconstituted complexes is identical to that found in bulk preparations containing random DNA sequences (Hayes et al., 1990). Cross-linking experiments have demonstrated that the histone protein–DNA contacts within reconstituted nucleosomes are identical to that of native complexes (Lee and Hayes, 1997). Reconstituted oligonucleosomal complexes exhibit the same hydrodynamic shape and folding behavior as native complexes (Schwarz and Hansen, 1994). Further, the



stoichiometry of polypeptides within reconstituted complexes is identical to that found in native chromatin (Camerini-Otero et al., 1976). A key element in many of these studies is the use of a well-defined nucleosome positioning sequence. Sequence-dependent variations in the B-DNA structure within these fragments cause the fragments to adopt a defined rotational and translational orientation with respect to the surface of the nucleosome core. Since the core histone–DNA contacts occur at specific locations on the surface of the octamer, these two positional variables are related. However, a single preferred rotational orientation may accommodate several translational frames related by approximately 10n base pairs. Therefore, reconstitutions with stably curved DNAs will exhibit a strong rotational preference but may yield a population with a heterogeneous distribution of translational frames. DNA fragments that contain 5S RNA gene sequences have been found to strongly direct translational positioning (Rhodes, 1985). A 5S DNA fragment from Xenopus exhibits has been found to exhibit exceptional nucleosome positioning properties. When this fragment is reconstituted with purified core histones, approximately 80% of the population of nucleosomes within the sample adopt a unique translational position (Hayes et al., 1990). The molecular basis of this translational positioning effect is not yet understood.

Critical Parameters and Troubleshooting Critical variables and ways to overcome experimental difficulties are presented in Table 21.6.1.

Anticipated Results The results obtained from the analysis should reveal the quality of the nucleosomal template (Support Protocol 3). If the purity of the histones and DNA is high and consideration is given to the relative stoichiometry, the assembly should generate suitable template for further experiments. The efficiency of assembly is obtained by quantifying the percentage of naked DNA assembled into nucleosomes as described in Support Protocols 5, 6, and 7. This should be between 30% and 80%.

Time Considerations Theoretically, step salt dialysis can be completed in 10 hr. Step dialysis has the advantage of being quick and convenient to set up. However, the drawbacks are that it does not always

generate properly assembled nucleosomes when the nucleosome concentration is high (∼1 mg) and when nucleosomal arrays are generated. An alternative method is to use the gradient salt dialysis method (see Basic Protocol 2). The assembly through gradient dialysis takes ~50 hr. It is a relatively long procedure but once it is set up, there are no further manipulations until the reconstitution is finished. The purification of histones from native chromatin takes roughly two full days after the necessary equipment is set up. The isolation of recombinant histones from bacteria requires 3 to 4 half days starting from the inoculation to the final chromatography steps. The analysis of the nucleosomes takes about 3 to 4 hr.

Literature Cited Camerini-Otero, R.D., Sollner-Webb, B., and Felsenfeld, G. 1973. The organization of histones and DNA in chromatin: evidence for an argininerich histone kernel. Cell 3:333-347. Carruthers, L.M., Tse, C., Walker, K.P., and Hansen, J. 1999. Assembly of defined nucleosomal and chromatin arrays from pure components. Methods Enzymol. 304:19-35. Hayes, J.J. and Lee, K.M. 1997. In vitro reconstitution and analysis of mononucleosomes containing defined DNAs and proteins. Methods 12:2-9. Hayes, J.J., Tullius, T.D., and Wolffe, A.P. 1990. The structure of DNA in a nucleosome. Proc. Natl. Acad. Sci. U.S.A. 87:7405-7409. Lee, K.M. and Hayes, J.J. 1997. The N-terminal tail of histone H2A binds to two distinct sites within the nucleosome core. Proc. Natl. Acad. Sci. U.S.A. 94:8959-8964. Lee, K.M., Chafin, D.R., and Hayes, J.J. 1999. Targeted cross-linking and DNA cleavage within model chromatin complexes. Methods Enzymol. 304:231-251. Luger, K., Rechsteiner, T.J., Flaus, A.J., Waye, M.M., and Richmond, T.J. 1997a. Characterization of nucleosome core particles containing histone protein made in bacteria. J. Mol. Biol. 272:301-311. Luger, K., Mader, A.W., Richmond, R.K., Sargent, D.F., and Richmond, T.J. 1997b. Crystal structure of the nucleosome core particle at 2.8 Å resolution. Nature 389:251-260. Luger, K., Rechsteiner, T.J., and Richmond, T.J. 1999. Preparation of nucleosome core particle from recombinant histones. Methods Enzymol. 304:3-19. Neely K.E., Hassan, A.H., Wallberg, A.E.,Steger, D.J., Cairns, B.R., Wright, A.P. and Workman, J.L. 1999. Activation domain-mediated targeting of the SWI/SNF complex to promoters stimulates transcription from nucleosome arrays. Mol. Cell. 4:649-655.



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Rhodes, D. 1985. Structural analysis of a triple complex between the histone octamer, a Xenopus gene for 5S RNA and transcription factor IIIA. EMBO J. 4:3473-3482. Schwarz, P.M. and Hansen, J.C. 1994. Formation and stability of higher order chromatin structures.contributions of the histone octamer. J. Biol. Chem. 269:16284-16289. van Holde, K.E. 1989. Chromatin. Springer-Verlag, New York

Contributed by Kyu-Min Lee and Geeta Narlikar Massachussets General Hospital Boston, Massachusetts

Assembly of Nucleosomal Templates by Salt Dialysis



Chromatin Assembly Using Drosophila Systems

UNIT 21.7

To study chromatin structure and activity in vitro, it is necessary to use appropriate methods for the preparation of chromatin substrates. The goal of chromatin assembly procedures is to prepare extended nucleosome arrays from cloned DNA templates (usually plasmid DNA) and purified chromatin proteins (core and linker histones, DNA-binding proteins). The assembled chromatin should be highly defined in its protein content and resemble bulk chromatin isolated from living cell nuclei in terms of periodicity and nucleosome positioning. In this unit we describe two systems that assemble minichromosome templates in an ATP-dependent fashion from circular plasmid DNA and purified core histones. These systems can be used to assemble minichromosomes from linear DNA (plasmid and lambda) and can also incorporate proteins other than core histones (linker histone H1, HMG17, and DNA-binding transcription factors). The products of these chromatin assembly reactions have been used directly (or after purification) in assays to study transcription, DNA replication, recombination, and repair. The first of the two systems employs a Drosophila embryonic extract (S-190) as a source of the assembly factors (see Basic Protocols 1 to 3). Well established and easy to troubleshoot, this protocol should be considered the first stop for researchers with limited experience in chromatin analysis. The second system utilizes purified recombinant Drosophila chromatin assembly factors (see Basic Protocols 4 to 6 and Alternate Protocol 1). It provides an additional advantage of strictly defined protein content of assembled chromatin. Determining the optimal ratio of core histone to DNA is important for either system; this is presented in Alternate Protocol 2. Chromatin assembly on relaxed circular DNA requires the presence of a topoisomerase activity in the reaction. An example of expression and purification of the core catalytic domain of Drosophila topoisomerase I is presented in the Support Protocol. PREPARATION OF THE DROSOPHILA S-190 CHROMATIN ASSEMBLY EXTRACT

BASIC PROTOCOL 1

This protocol describes the preparation of the S-190 Drosophila cell extract. When supplemented with core histones and ATP, the S-190 extract can mediate the in vitro assembly of regularly spaced arrays of nucleosomes on a DNA template. Materials 0- to 6-hr Drosophila embryos Bleach wash: 50% (v/v) household bleach in distilled H2O Embryo wash: 0.7% (w/v) NaCl/0.4% (v/v) Triton X-100, room temperature and 4°C Saline wash: 0.7% (w/v) NaCl, 4°C Buffer R (see recipe), 4°C 1 M MgCl2 (APPENDIX 2) Liquid N2 Fine nylon mesh (e.g., Sefar America 03-80/37) 800-ml glass beaker Glass rod Vacuum aspirator 40 ml Wheaton Dounce homogenizer with “A” and “B” pestles 17 × 100–mm polypropylene tubes (e.g., Falcon 2059) Sorvall Superspeed centrifuge with SS-34 rotor (or equivalent) Contributed by Dmitry V. Fyodorov and Mark E. Levenstein Current Protocols in Molecular Biology (2002) 21.7.1-21.7.27 Copyright © 2002 by John Wiley & Sons, Inc.



50-ml conical tubes 10-ml plastic syringes and 18.5-G needles Beckman ultracentrifuge with SW 41 and SW 55 rotors Thin-walled 14 × 89–mm (for SW 41 rotor) and 13 × 51–mm (for SW 55 rotor) Beckman ultracentrifuge tubes Prepare the embryos 1. Secure a fine nylon mesh in a support and place ∼100 g of 0- to 6-hr Drosophila embryos on the mesh. Wash thoroughly with cold tap water to eliminate yeast and contaminants. Do not freeze the embryos before extraction. 100 g of the starting material should yield ∼50 ml of S-190 extract. S-190 extracts prepared from embryos older than 6 hr are significantly less active. It is important to select an appropriate mesh that is fine enough to retain the embryos but large enough to allow the buffers and yeast to flow through. A coarser mesh placed above the fine mesh can be used to separate adult fly bodies and parts as well as pieces of food agar from the embryos.

2. Soak the embryos 90 sec in 3 liters bleach wash at room temperature. Quickly rinse with 1 liter embryo wash at room temperature. Rinse extensively with distilled water. Transfer embryos to an 800-ml glass beaker on ice. Bleach wash will remove chorions from the embryos. After this point, all subsequent steps should be carried out on ice or in a cold room, unless otherwise noted.

3. Add 500 ml of embryo wash at 4°C and suspend the embryos by stirring with a glass rod. Allow the embryos to settle to the bottom of the beaker for ∼2 min. Aspirate the embryo wash using a vacuum aspirator, avoiding the settled embryos at the bottom of the beaker. Be sure to aspirate the chorions and other floating non-embryo material at the top of the beaker.

4. Repeat step 3 with an additional 500 ml of 4°C embryo wash. 5. Repeat step 3 twice, each time with 500 ml of 4°C saline wash. Allow the embryos to settle ∼5 min in saline wash. It is important to avoid aspirating the now floating embryos in the beaker.

6. Repeat step 3 twice, each time with 500 ml of 4°C buffer R. The embryos will require up to 10 min to settle in buffer R. Again, it is important to avoid aspirating floating embryos as much as possible. After the second aspiration, the final volume should be no more than 80 ml. While it may require some loss of the starting material, it is more desirable to have a concentrated extract than a larger volume of the extract.

Prepare the extract 7. Transfer the material to a 40 ml Wheaton Dounce homogenizer on ice. Homogenize with 15 strokes of the “B” pestle and additional 40 strokes of the “A” pestle. Repeat with any remaining material. Be certain to homogenize with full strokes. It is important to achieve complete homogenization of the embryos. This can be verified by placing a drop of the homogenate on a glass slide and examining for the presence of intact embryos.

8. Transfer the homogenate to 17 × 100–mm polypropylene tubes on ice. Spin 5 min at 7500 × g (8000 rpm in a Sorvall SS-34 rotor), 4°C. Extract the aqueous supernatant from the middle of the tubes and pool in 50-ml conical tube(s). Chromatin Assembly Using Drosophila Systems

To avoid the pellet at the bottom and the white lipid layer at the top of the tube, use a 10-ml syringe and an 18.5-G needle inserted through the tube just above the pellet.



9. Using 1 M MgCl2, adjust the concentration of magnesium to 7 mM. Note that buffer R contains 1.5 mM MgCl2. This step significantly reduces the amount of RNA in the final extract.

10. Transfer the supernatant to thin-walled 14 × 89–mm ultracentrifuge tubes. Ultracentrifuge for 2.25 hr at 200,000 × g (40,000 rpm in an ultracentrifuge with Beckman SW 41 rotor), 4°C. For smaller volumes, use 13 × 51–mm tubes and spin at 192,000 × g (45,000 rpm in a Beckman SW 55 rotor), 4°C.

11. Remove the white upper lipid layer from each tube with a metal spatula and discard. Collect the golden-brown liquid layer and pool in a 50-ml conical tube(s) on ice. Freeze in liquid nitrogen. It is important to avoid the beige, upper layer of the pellet, which will inhibit the assembly activity. At this point, the extract can be stored at −80°C indefinitely until the final spin. However, freezing of the extract is essential even if you proceed directly to step 12. Handle the 50-ml polypropylene tubes gently in liquid N2 to avoid cracking.

12. Thaw the extract in a room temperature water bath. Spin 2.25 hr at 192,000 × g (45,000 rpm in an ultracentrifuge with Beckman SW 55 rotor), 4°C. Note that the SW 55 rotor consistently yields a more active extract than the SW 41 for this spin.

13. Remove the white upper lipid layer (if any) and pool the extract in a 50-ml conical tube. Aliquot 1-ml fractions into microcentrifuge tubes and freeze in liquid nitrogen. Store at −80°C. The S-190 extract will remain active for up to 3 years at −80°C.

PURIFICATION OF CORE HISTONES FROM THE DROSOPHILA EMBRYOS Nuclei are isolated from the Drosophila embryos. Chromatin is partially digested with micrococcal nuclease to generate ∼2000-bp fragments. The chromatin fragments are isolated, bound to hydroxylapatite resin, and core histones are separated from the DNA by salt elution. Materials 0- to 12-hr Drosophila embryos Bleach wash: 50% (v/v) household bleach in distilled H2O Embryo wash: 0.7% (w/v) NaCl/0.04% (v/v) Triton X-100 Buffer B (see recipe) Buffer A (see recipe) 1 M NaOH 0.1 M CaCl2 200 U/ml micrococcal nuclease stock solution (see recipe) 10 mM and 500 mM EDTA 10% (w/v) SDS 5 M NaCl (APPENDIX 2) 24:1 chloroform/isoamyl alcohol Linear 5% to 30% sucrose gradients (see recipe) T50E4 buffer: 50 mM Tris⋅Cl, pH 7.9 (APPENDIX 2)/4 mM EDTA Hydroxylapatite resin (e.g., BioGel HT gel; Bio-Rad) HA chromatography buffer (see recipe) with 0 M, 0.35 M, and 2.5 M NaCl Core histone storage buffer (see recipe) BCA assay kit (Pierce)

BASIC PROTOCOL 2



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Fine nylon mesh (e.g., Sefar America 03-80/37) Weigh boats Yamato LH-21 homogenizer (alternatively, Potter-Elvehjem homogenizer) 500-ml centrifuge bottles for Sorvall GSA rotor (or equivalent) Mira cloth (Calbiochem-Novabiochem) Sorvall Superspeed centrifuge with GSA and SS-34 rotors or equivalents Beckman ultracentrifuge with SW-28 rotor and appropriate tubes 12,000 to 15,000 and 3,500 MWCO dialysis tubing FPLC apparatus Additional reagents and equipment for extraction of DNA (UNIT 2.1A), agarose gel electrophoresis (UNIT 2.5A), SDS-PAGE (UNIT 10.2), Coomassie blue staining and destaining of proteins in gels (UNIT 10.6), and dialysis (APPENDIX 3C) Prepare the embryos 1. Secure a fine nylon mesh in a support and place ∼100 g of 0- to 12-hr Drosophila embryos on the mesh. Wash thoroughly with cold tap water to eliminate yeast and contaminants. The embryos can be frozen in liquid nitrogen and stored up to 1 year at −80°C before processing. 100 g of the starting material should yield between 5 to 10 mg of core histones. See step 1 of Basic Protocol 1 for information on the nylon mesh.

2. Soak the embryos 90 sec in 3 liters bleach wash at room temperature. Quickly rinse with 1 liter of embryo wash at room temperature. Rinse extensively with distilled water. Blot embryos dry through the mesh with paper towels. Transfer to a weigh boat and weigh. Bleach wash will remove chorions from the embryos. After this point, all subsequent steps should be carried out on ice or in a cold room, unless otherwise noted.

Isolate the nuclei 3. Transfer the embryos to a glass beaker. Resuspend in 3 ml buffer B per 1 g of embryos. Pour the suspension through a Yamato LH-21 homogenizer at 1000 rpm and filter the effluent into GSA bottles through Mira cloth. If a Yamato homogenizer is not available, the embryos can be disrupted with six to eight strokes in a motorized Potter-Elvehjem homogenizer (serrated Teflon pestle in a glass vessel), followed by several strokes in a Wheaton Dounce homogenizer with the “B” pestle.

4. Wash the original beaker with 1 ml buffer B per 1 g of embryos and pass through the homogenizer and Mira cloth as in step 3. Repeat with 1 ml buffer B per 1 g of embryos. Centrifuge the filtrate 20 min at 10,000 × g (8000 rpm in a GSA rotor), 4°C. The final volume of the suspension before spinning should be 5 ml buffer B per 1 g of embryos.

5. Discard the supernatant. Resuspend the loose pellet of nuclei in 200 ml buffer A. Centrifuge 10 min at 10,000 × g (8000 rpm in a GSA rotor), 4°C.


Avoid the yellow yolk protein pellet when resuspending the nuclei. A firmer, “yellow” yolk pellet should form at the center of the looser nuclei pellet; it is this pellet that should be avoided. A Pasteur pipet can be used to trace the outline of the yolk pellet, after which gentle resuspension of the nuclear pellet can be carried out with a battery-powered pipetting device (e.g., Pipet-Aid) and buffer A.



6. Discard the supernatant. Resuspend the nuclei in 100 ml buffer A. Centrifuge 10 min at 10,000 × g (8000 rpm in a GSA rotor), 4°C. Again, avoid the yolk protein, if any remains.

7. Discard the supernatant. Resuspend the nuclear pellet in 30 ml buffer A. Digest a sample of the chromatin 8. Dilute 10 µl of suspended nuclei with 990 µl of 1 M NaOH. Measure the optical density at 260 nm (OD260). Dilute the suspension of nuclei to 100 OD260 units/ml with buffer A. 9. Perform a test micrococcal nuclease digestion as follows: a. In a 1.5 ml microcentrifuge tube, prewarm 1 ml of the nuclear suspension to 37°C. b. Add 10 µl of 0.1 M CaCl2 and 2 µl of 200 U/ml micrococcal nuclease stock solution. c. Incubate at 37°C. At 1, 2, 4, 6, 8, 10, 15, and 20 min, remove a 100-µl aliquot and stop enzyme activity in the aliquot with 2.5 µl 500 mM EDTA. 10. To each aliquot add 30 µl water and 20 µl 10% SDS. Mix thoroughly and add 40 µl 5 M NaCl. Mix thoroughly. Extract each aliquot with 200 µl of 24:1 chloroform/isoamyl alcohol. UNIT 2.1A

describes general techniques for extraction of DNA.

11. Mix 4 µl of the aqueous phase from each aliquot with agarose gel loading buffer. Run each of the aliquots (representing time points from step 9) in a separate lane of a 1% agarose gel. Stain with ethidium bromide and photograph (see UNIT 2.5A for reagents and techniques used in this step). Use a 1-kb ladder as a reference and select the time point displaying a prevalence of ∼2000-bp fragments.

Perform bulk digest of the chromatin 12. Warm the nuclear suspension to 37°C and add 1/100 vol of 0.1 M CaCl2. Add 1/500 vol of 200 U/ml micrococcal nuclease and mix by swirling. Incubate at 37°C with occasional swirling for the time determined in steps 8 to 11. Stop digestion by adding of 1/50 vol of 500 mM EDTA and place at 4°C. 13. Centrifuge 10 min at 12,000 × g (10,000 rpm in an SS-34 rotor), 4°C. Discard the supernatant. Resuspend the pellet in 10 mM EDTA for a total volume of 9 ml. Add 1 ml of 5 M NaCl and swirl 5 min. Upon the addition of NaCl, extensive lysis should be observed and the solution should become more viscous.

14. Centrifuge 5 min at 12,000 × g (10,000 rpm in an SS-34 rotor), 4°C. Save the supernatant. Dilute 10 µl into 990 µl of 1 M NaOH and measure the OD260. 15. Load ≤500 OD260 units of supernatant onto linear gradients (18 to 20 ml each) of 5% to 30% sucrose in sucrose gradient buffer in SW 28 tubes. Ultracentrifuge 16 hr at 90,000 × g (26,000 rpm in an SW 28 rotor), 4°C. 16. Cut the gradients into 2-ml fractions and run 8 µl per fraction on a 15% SDS-PAGE gel (UNIT 10.2). Coomassie stain and destain (UNIT 10.6). Pool the peak core histone containing fractions and dialyze in 12,000 to 15,000 MWCO tubing twice, each time for 2 hr against 2 liters T50E4 buffer (APPENDIX 3C) at 4°C.



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Like fractions from each gradient tube can be pooled in 50-ml conical tubes. If any debris is suspended in sucrose-containing fractions, it can be spun down after dialysis. The fractionated digested chromatin can be stored for several months at 4°C without loss of core histone integrity. When selecting fractions for further purification, be careful to avoid histone H1-containing fractions.

Purify core histone octamers by hydroxylapatite chromatography 17. Dilute 10 µl dialyzed sample with 990 µl of 1 M NaOH and measure the OD260. Calculate the required volume of hydroxylapatite resin as 1 ml column volume per 1.5 mg of DNA. 18. Pack an appropriately sized hydroxylapatite column on an FPLC and equilibrate in 3 column volumes HA chromatography buffer without NaCl. 19. Apply the dialyzed sample to the column and wash with an additional 3 column volumes of HA chromatography buffer without NaCl. Wash with an additional 3 column volumes of HA chromatography buffer containing 0.35 M NaCl. The 0.35 M NaCl wash elutes DNA-binding proteins, other than histones, off the resinbound chromatin.

20. Elute the core histones with 2 column volumes of HA chromatography buffer containing 2.5 M NaCl. Alternatively, the core histones can be eluted with a salt gradient from 0.35 to 2.5 M NaCl in HA chromatography buffer. This separates any residual histone H1 that was collected from the sucrose gradient. A variation of this procedure can also be used to fractionate histone dimers and tetramers. Additional information on hydroxylapatite chromatography of histones is found in UNIT 21.5.

21. Analyze 2 µl of peak fractions on a 15% SDS-PAGE gel (UNIT 10.2). Coomassie stain and destain (UNIT 10.6). Pool the peak core histone–containing fractions and dialyze in 3,500 MWCO tubing twice, each time for 2 hr against 2 liters core histone storage buffer at 4°C. 22. After dialysis, determine the histone concentration by BCA assay. Aliquot into microcentrifuge tubes and freeze in liquid nitrogen. Store at −80°C. The core histones can be stored for several years at −80°C. BASIC PROTOCOL 3


CHROMATIN ASSEMBLY WITH THE S-190 EXTRACT The extract (from Basic Protocol 1) and core histones (from Basic Protocol 2) are combined and preincubated at room temperature to facilitate binding of histones and chaperone(s). A DNA template, ATP, and ATP regeneration system are added, and chromatin is assembled at 27°C. Chromatin is analyzed by partial micrococcal nuclease digestion. Materials Buffer R (see recipe) S-190 extract (see Basic Protocol 1) Core histones (see Basic Protocol 2) ATP mix (see recipe) 0.1 M MgCl2 DNA template (2.5 to 25 kbp circular or linearized plasmid, or λ DNA) 0.1 M CaCl2 200 U/ml micrococcal nuclease stock solution (see recipe)



0.5 M EDTA 10 mg/ml RNase A Glycogen stop buffer (see recipe) 2.5 mg/ml proteinase K 50:49:1 (v/v/v) phenol/chloroform/isoamyl alcohol, equilibrated with 10 mM Tris⋅Cl, pH 8.0 (see UNIT 2.1A for equilibration technique) 2.5 M ammonium acetate 70% and 100% ethanol 1.2% agarose gel (UNIT 2.5A) TBE buffer (APPENDIX 2) 123 bp DNA ladder (Life Technologies) 27° and 37°C water baths Additional reagents and equipment for extraction of DNA (UNIT 2.1A) and agarose gel electrophoresis (UNIT 2.5A) Preincubate core histones with the extract 1. Combine 40 µl buffer R, 30 µl S-190 extract, and 400 ng of core histones. Incubate 30 min at room temperature. A single reaction produces enough chromatin for one micrococcal nuclease digestion analysis (two lanes on an agarose gel). The reaction can be scaled up. Typically, the S-190 extract comprises 25% to 40% of the reaction volume and the core histones are in a 0.8:1 mass ratio with the DNA template (5 ng/ìl). The final Mg2+ concentration is 7 mM.

Add ATP and the DNA template 2. Add 10 µl ATP mix, 0.1 M MgCl2 to 7 mM final concentration, and 500 ng DNA template. Incubate 5 hr at 27°C. If necessary, make up to a final volume of 100 µl with buffer R. Note that both buffer R and the S-190 extract contain MgCl2.

Perform partial micrococcal nuclease digestions 3. Add 3 µl of 0.1 M CaCl2 to the assembly reaction. Divide the reaction in two equal parts (“a” and “b”). Dilute the micrococcal nuclease (200 U/ml) 1:50 and 1:150 with buffer R. Prepare the nuclease dilutions within a few minutes before use.

4. In a controlled manner (at certain time intervals, e.g., 15 sec), add 5 µl of the 1:150 dilution to the “a” tube and 5 µl of the 1:50 dilution to the “b” tube. Digest 10 min at room temperature. 5. Terminate the reactions with 7 µl of 0.5 M EDTA. Add 1 µl of 10 mg/ml RNase A solution and incubate for 10 min at room temperature. This step eliminates RNA present in the S-190 extract.

Deproteinate the chromatin samples 6. To each aliquot add 95 µl glycogen stop buffer and 5 µl of 2.5 mg/ml proteinase K solution. Incubate 30 min at 37°C. 7. Extract with 200 µl of 50:49:1 phenol/chloroform/isoamyl alcohol. Precipitate the DNA from 150 µl of the aqueous phase with 15 µl of 2.5 M ammonium acetate and 450 µl of 100% ethanol. Microcentrifuge for 15 min at maximum speed, room temperature. Wash the pellets with 200 µl of 70% ethanol. Microcentrifuge 5 min at maximum speed, room temperature. Dry by exposing to air for 5 min. UNIT 2.1A

describes general techniques for extraction and precipitation of DNA.



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S-190 Histones ATP DNA

+ + + +

1

2

3

—

+

+ + +

—

+ +

+ +

4

5

6

7

—

+ + +

+

—

8

9

10

Figure 21.7.1 Chromatin assembly by the S-190 extract. Lanes 1, 2—complete system, as described in Basic Protocol 3; lanes 3, 4—no S-190 extract; lanes 5, 6—no core histones (note residual assembly activity with the endogenous histones of the extract); lanes 7, 8—no ATP; lanes 9, 10—no DNA template.

Analyze micrococcal nuclease ladders 8. Resuspend the pellets in 6 µl gel loading buffer and run on a 1.2% agarose gel in 1× TBE buffer. Use 123 bp DNA ladder as a marker. Stain with ethidium bromide (see UNIT 2.5A for the reagents and techniques used in this step).


Run the gel at ∼5 to 10 V/cm. Running too quickly will cause uneven heating of the gel and tilting of the DNA bands, which will interfere with detection of the bands at the top of the ladder. Running the gel too slowly will cause excessive diffusion of the bands at the bottom of the ladder. The electrophoresis should be terminated when the bromphenol blue dye front reaches the bottom quarter of the gel. Bromphenol blue migrates at ∼250 bp DNA size, while the mononucleosome band has a size of ∼170 bp. For the best results, stain the gel 15 to 20 min in 2 gel volumes of 0.75 ìg/ml ethidium bromide in distilled water. Do not overstain. Destain the gel for 1 to 3 hr in distilled water with several changes. A ladder of at least 7 to 8 nucleosomal bands, counting from the bottom up, should be apparent. The repeat length should be 165 base pairs or more. An example of the type of results obtained is shown in Figure 21.7.1.



EXPRESSION AND PURIFICATION OF THE RECOMBINANT DROSOPHILA ACF

BASIC PROTOCOL 4

Drosophila ACF (ATP-utilizing chromatin assembly and remodeling factor) is prepared by coexpression of carboxyl-terminally FLAG-tagged Acf1 subunit with untagged ISWI subunit in baculovirus. The complex is then purified in one step by FLAG immunoaffinity chromatography and eluted with a buffer containing the FLAG peptide. This procedure typically results in a stoichiometric complex of p185 isoform of Acf1 and ISWI. In a slight alteration of this protocol, it is possible to express and purify the individual FLAG-tagged ISWI subunit. Materials High titer Acf1-FLAG and ISWI baculovirus stocks (Orbigen; also see UNITS 16.9 & 16.10) Late log phase Sf9 cells cultured in suspension (>2 × 106 cells/ml; also see UNITS 16.9 & 16.10) Phosphate-buffered saline (PBS; APPENDIX 2), ice-cold Lysis buffer F (see recipe) 1:1 (v/v) slurry of FLAG-M2 resin (Sigma-Aldrich), equilibrated in lysis buffer F Dilution buffer F (see recipe) Wash buffer F (see recipe) Elution buffer F (see recipe) Liquid nitrogen Bovine serum albumin (BSA) standard, 2 mg/ml (Pierce, Cat. No. 23209) Clinical centrifuge with swinging-bucket rotor, 4°C 250-ml conical centrifuge bottles appropriate for clinical centrifuge, or 50-ml conical tubes 14- and 50-ml disposable conical centrifuge tubes 15-ml Wheaton dounce homogenizer, “A” pestle Sorvall Superspeed centrifuge with SS-34 rotor (or equivalent) 15-ml capped polypropylene tubes Siliconized 1.5-ml polypropylene tubes (e.g., ISC BioExpress, Cat. # C-3302-1) Additional reagents and equipment for baculovirus culture (UNITS 16.9 & 16.10), SDS-PAGE (UNIT 10.2), and staining of gels (UNIT 10.6) Infect and harvest Sf9 cells 1. Amplify baculovirus stocks several days before the infection (UNIT 16.10). Viruses are supplied as cell culture supernatants and are to be stored at 4°C. Titers of the original stocks may vary (1–5 × 108 pfu per ml; see Orbigen’s protocols; http://www.orbigen.com). To amplify the viruses, Sf9 cells are plated on 150-mm culture plates at 2 × 107 cells/plate in a total of 25 ml of appropriate serum-containing medium, and infected at multiplicity of infection (MOI) of 0.1 to 0.5 (10 to 20 ìl viral supernatant per plate). To pass a virus stock for storage, allow the infection to proceed for 60 hr and collect the medium supernatant aseptically. For high-titer stock (to be further used for expression infection), the infection is allowed to progress until cell lysis is apparent (∼72 hr for His-NAP-1 virus, ∼84 hr for ISWI and Acf1-FLAG). This procedure produces viral stocks with titers close to or above 109 pfu per ml. The medium is aspirated off the plates aseptically and stored in sterile 50-ml tubes in the dark at 4°C for up to 12 months without significant loss of titer.

2. Grow Sf9 cells (see UNIT 16.10) in 150 or 500 ml spinner flasks for 2 to 3 days after seeding at 0.5 × 106 cells/ml. Plate 5 to 25 plates of Sf9 cells at 2.5 to 3 × 107 cells/plate in a total of 25 ml of appropriate insect medium per plate. Allow the cells to settle



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for 20 min in the tissue culture hood and infect with recombinant Acf1-FLAG and ISWI baculoviruses at an MOI of 5 to 10 each. It is not necessary to completely replace the old medium. Typically, equal volumes (0.3 to 0.7 ml) of both high titer virus stocks are used. Occasionally, the ratio of Acf1-FLAG and ISWI viruses has to be adjusted empirically to ensure a proper stoichiometry.

3. At 44 to 46 hr subsequent to infection, aspirate the medium and wash the cells off the plates with 10 ml ice-cold PBS per plate. Centrifuge 5 min at 2000 rpm in a clinical centrifuge in 250-ml conical bottles or 50-ml tubes at 4°C. Do not exceed the 2-day infection time, since it may result in lower yields and accumulation of complexes with partially degraded Acf1 and ISWI. From this point on, work in a cold room or on ice. Cell pellets can be frozen in liquid nitrogen and stored at −80°C for several weeks before further processing.

4. Resuspend the cell pellet in 8 ml of lysis buffer F and disrupt with a Wheaton Dounce homogenizer (“A” pestle; 3 series of 10 strokes over a 30 min period, on ice). Cell nuclei are lysed by 500 mM salt. Both nuclei and cytosol are extracted. Cell lysis can be monitored microscopically, if desired. However, the appearance of the cell debris pellet after centrifugation, with distinct yolk and nuclear phases, may serve as an equally good indication of lysis completeness.

Prepare and analyze ACF protein 5. Pellet insoluble material by centrifuging in 14-ml disposable conical tubes 10 min at 14,500 × g (11,000 rpm in an SS-34 rotor), 4°C. Combine the supernatant with 250 µl of FLAG-M2 resin (as a 1:1 slurry equilibrated in lysis buffer F) and 7 ml dilution buffer F. Mix the slurry on a rocking platform for 3 to 4 hr at 4°C in a 15-ml capped polypropylene tube. The salt concentration is decreased to allow efficient binding of the FLAG-tag to the M2 antibody.

6. Wash the resin four times, each time with 12 ml wash buffer F by successive cycles of centrifugation for 3 min at 2000 rpm in the clinical centrifuge at 4°C, followed by aspiration and resuspension. Mix by inverting; avoid vigorous shaking.

7. Elute protein as follows: a. b. c. d. e. f.

Add 100 µl elution buffer F to resin pellet from step 6 and resuspend. Transfer resin to a 1.5-ml siliconized microcentrifuge tube. Incubate on ice 10 min. Microcentrifuge 30 sec at maximum speed. Transfer the supernatant to another tube (to be pooled with subsequent elutions). Continuing in the same siliconized microcentrifuge tube, repeat steps 7a, 7c, 7d, and 7e, three additional times, pooling all of the eluates together. Insulin is thought to stabilize the recombinant protein at low concentrations.

8. Freeze the protein in liquid nitrogen in small aliquots (20 to 50 µl) and store at −80°C. The recombinant ACF is stable for several years and can withstand multiple (5 to 10) freeze-thaw cycles. Prepare and freeze several aliquots of wash buffer F containing 0.4 mg/ml insulin. This solution will be used further as ACF dilution buffer. Chromatin Assembly Using Drosophila Systems



9. Optional: Estimate protein concentration by SDS-PAGE (UNIT 10.2) along with a range of concentrations of the BSA standard, followed by staining with Coomassie Brilliant Blue R-250 (UNIT 10.6). Typical yields of ACF are ∼5 to 10 ìg per 150-mm plate.

EXPRESSION AND PURIFICATION OF THE RECOMBINANT DROSOPHILA NAP-1

BASIC PROTOCOL 5

Sf9 cells are infected with dNAP-1 expressing baculovirus. The chaperone protein is purified by affinity chromatography through a 6-Histidine tag, followed by anion exchange chromatography on Source 15Q resin (Pharmacia). Materials Late-log-phase Sf9 cells cultured in suspension (>2 × 106 cells/ml; also see UNITS 16.9 & 16.10) High-titer His-NAP-1 baculovirus stock (Orbigen; also see UNITS 16.9 & 16.10) Phosphate-buffered saline (PBS; APPENDIX 2), ice-cold Lysis buffer H (see recipe) Wash buffer H (see recipe) Elution buffer H: wash buffer H (see recipe) containing 480 mM imidazole HEGD buffer containing 0.1 M NaCl (see recipe) NAP-1 purification buffer (see recipe) containing 0.0, 0.1, and 1.0 M NaCl Ni-NTA agarose resin (Qiagen) Bovine serum albumin (BSA) standard, 2 mg/ml (Pierce, Cat. No. 23209) Source 15Q resin (Amersham Pharmacia Biotech) 20% (v/v) ethanol 8% and 15% SDS-PAGE gels (UNIT 10.2) Liquid nitrogen Clinical centrifuge with swinging-bucket rotor, 4°C 250-ml conical centrifuge bottles appropriate for clinical centrifuge 40-ml Wheaton Dounce homogenizer with “A” pestle Sorvall Superspeed centrifuge with SS34 rotor (or equivalent) 15- and 50-ml conical tubes End-over-end rotator 12,000 to 15,000 MWCO dialysis tubing HR-5 or HR-10 FPLC column (Amersham Pharmacia Biotech) FPLC apparatus Siliconized 1.5 ml polypropylene tubes (e.g., ISC BioExpress, Cat. # C-3302-1) Additional reagents and equipment for baculovirus culture and infection (UNITS 16.9-16.11), dialysis (APPENDIX 3C), and SDS-PAGE (UNIT 10.2) Infect and harvest Sf9 cells 1. Grow Sf9 cells in 500 or 1000 ml spinner flasks to a density >2.0 × 106 cells/ml in culture medium (UNIT 16.10). Dilute with the medium to 1.0 × 106 cells/ml. Infect with 25 ml amplified His-NAP-1 virus per 1 liter cell culture. 2. At 72 hr subsequent to infection, collect the cells by spinning 5 min at 2000 rpm in 250 ml conical tubes in a clinical centrifuge at 4°C. Resuspend each pellet in cold PBS (1/10 of the original culture volume). Repeat the centrifugation. Exercise caution not to discard the cell pellet, which becomes quite loose after washing with PBS. From this point on, all steps should take place on ice or in a cold room, unless otherwise noted. All buffers should be at 4°C unless otherwise noted. Cell pellets can be frozen in liquid nitrogen and stored at −80°C for several weeks before further processing.



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3. Resuspend the cell pellet in lysis buffer H (1/40 original culture volume). Homogenize in a Dounce homogenizer using 40 strokes of the “A” pestle over 30 min. Centrifuge 10 min at 14,500 × g (11,000 rpm in an SS-34 rotor), 4°C. Pool all supernatants in a 50-ml conical tube. Purify NAP-1 by nickel affinity chromatography 4. Equilibrate 1 ml Ni-NTA agarose resin in lysis buffer H per 500 ml original cell culture volume. Add the cell extract and incubate for 3 to 4 hr on an end-over-end rotator. Pellet the resin by centrifuging 5 min at 2000 rpm in a clinical centrifuge, 4°C. Wash the resin twice with 100 ml lysis buffer H, then twice with 100 ml wash buffer H, resuspending the resin by inverting the tube after each addition of buffer, and pelleting the resin by centrifuging 3 min at 2000 rpm in the clinical centrifuge after each wash. 5. To elute protein, resuspend the resin in 2 ml elution buffer H by gentle vortexing. Incubate on ice for 5 min. Centrifuge 3 min at 2000 rpm in clinical centrifuge, then remove the supernatant to a fresh tube on ice. Repeat this elution cycle three more times, pooling the eluates. 6. Dialyze the eluted NAP-1 twice in 12,000 to 15,000 MWCO tubing, each time for 2 hr against 4 liters HEGD buffer containing 0.1 M NaCl. General techniques for dialysis are described in APPENDIX 3C. This dialysis step removes phosphate from the buffer.

7. Dialyze for an additional 2 hr against 4 liters NAP-1 purification buffer containing 0.1 M NaCl. Significant protein precipitation at this step is reduced by complete dialysis in step 6. The dialyzed protein can be frozen in liquid nitrogen and stored at −80°C before further processing.

8. Remove the precipitate by spinning in a 15-ml conical tube at 14,500 × g (11,000 rpm in an SS-34 rotor), 4°C. Analyze the dialyzed NAP-1 by SDS-PAGE (UNIT 10.2) with BSA standard to estimate the amount of protein. A 14-kDa contaminating Sf9 protein band will be apparent if analyzed on a 15% (or higher) gel. Although dNAP-1 is >95% pure after the affinity chromatography step, the contaminants appear to moderately inhibit chromatin assembly.

Purify NAP-1 to apparent homogeneity by anion-exchange chromatography 9. Using an FPLC, pack Source 15Q resin in an HR-5 or HR-10 column according to the manufacturer’s instructions. Use 1 ml of packed resin per 5 mg of NAP-1 from step 8. Equilibrate the Source 15Q column in 10 column volumes of NAP-1 purification buffer containing 0.1 M NaCl. 10. Load the NAP-1 onto the Source 15Q column. Wash the sample with 10 column volumes of NAP-1 purification buffer containing 0.2 M NaCl. Elute the protein with a 20-column-volume gradient of NAP-1 purification buffer from 0.2 M to 0.5 M NaCl. NAP-1 should elute in two distinct peaks. The early (lower-salt) peak is inhibitory towards assembly while the later (higher salt) peak is active. The 14-kDa band (see step 8) elutes with the early (inhibitory) peak. Collect fractions of 0.25 to 0.5 column volumes.


11. Run 2 µl of each fraction on 15% SDS-PAGE gel (UNIT 10.2) to identify pure NAP-1-containing fractions. Combine peak fractions and dialyze twice (APPENDIX 3C), each time for 2 hr against 2 liters NAP-1 purification buffer containing 0.1 M NaCl. A 15% gel will allow visualization of the 14-kDa contaminating band.



12. Analyze the dialyzed NAP-1 on an 8% SDS-PAGE gel (UNIT 10.2) along with BSA standard to determine the concentration. Divide the material into 100- to 200-µl aliquots in 1.5-ml siliconized tubes and freeze in liquid nitrogen. Store at −80°C. The expected yield of the active NAP-1 fraction is 1 to 3 mg per 1 liter of the cell culture volume.

EXPRESSION AND PURIFICATION OF THE RECOMBINANT DROSOPHILA NAP-1 (NTA SUPERFLOW RESIN)


If culture volumes are large, it can be helpful to perform the affinity chromatography purification with NTA Superflow resin (Qiagen) on an FPLC, rather than using batch purification on Ni-NTA agarose. Additional Materials (also see Basic Protocol 5) NTA Superflow resin (Qiagen) Superflow chromatography buffer (see recipe) containing 20 mM and 500 mM imidazole C-10/10 or C-10/20 FPLC column (Amersham Pharmacia Biotech) 1. Using an FPLC, pack an NTA Superflow column in 20% ethanol using 1 ml NTA Superflow resin per 500 ml of the original culture volume. Attach column to FPLC apparatus. 2. Equilibrate the column with 5 column volumes of Superflow chromatography buffer containing 20 mM imidazole. Load the supernatant from Basic Protocol 5, step 3 onto the column. If you plan to reuse the column, it may be helpful to briefly spin the extract in an ultracentrifuge 30 min at 80,000 × g (25,000 rpm in an SW-28 rotor), 4°C, in order to reduce the particulate matter being injected on the FPLC column. The injection on the column should be very slow (overnight at 0.05 to 0.1 ml/min). If the flowthrough contains significant amounts of NAP-1, it can be used for an additional round of purification.

3. Wash with 5 column volumes of Superflow chromatography buffer containing 20 mM imidazole. Elute the protein with a 10 column volume gradient from 20 mM to 500 mM imidazole in Superflow chromatography buffer. 4. Run 5 µl per fraction on a 15% SDS-PAGE gel (UNIT 10.2) and pool the protein peak. Continue as in Basic Protocol 5, step 6. A 15% gel will allow visualization of the 14-kDa contaminating band.

CHROMATIN ASSEMBLY WITH PURIFIED RECOMBINANT DROSOPHILA FACTORS

BASIC PROTOCOL 6

Core histones are preincubated on ice with NAP-1 to form histone-chaperone complexes. The chromatin assembly reaction is initiated at 27°C by addition of ACF, ATP, and plasmid DNA (circular supercoiled or relaxed, or linear). The extent of chromatin assembly can be monitored by analysis of circular DNA supercoiling, while the “quality” of chromatin is assayed by micrococcal nuclease digest assay. Materials HEG buffer (see recipe) 300 mM KCl (store in aliquots of 0.1 to 1 ml at −20°C) PvOH/PEG solution (see recipe) 2 mg/ml BSA solution (store in aliquots of 0.1 to 1 ml at −20°C)



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0.5 to 4.0 mg/ml recombinant NAP-1 (see Basic Protocol 5) 0.3 to 2.0 mg/ml purified Drosophila core histones, 0.3 to 2.0 mg/ml (see Basic Protocol 2) 0.5 M ATP (store in aliquots of 0.1 to 1 ml at −20°C) 0.5 M creatine phosphate (see recipe) 5 mg/ml creatine kinase (see recipe) 100 mM MgCl2 (store in aliquots of 0.1 to 1 ml at −20°C) 0.3 to 2.0 mg/ml plasmid DNA, double CsCl-purified (UNIT 1.7), in TE buffer (APPENDIX 2) 10× topoisomerase I buffer (see recipe) Recombinant topoisomerase I working solution (see Support Protocol) 0.002 to 0.2 mg/ml recombinant ACF (see Basic Protocol 4) ACF dilution buffer: wash buffer F (see recipe) containing 0.4 mg/ml recombinant human insulin (Roche) 200 U/ml micrococcal nuclease stock solution (see recipe) Buffer R (see recipe) 10 mM CaCl2 (store in aliquots of 0.1 to 1 ml at −20°C) 0.5 M EDTA 10 mg/ml RNase A Glycogen stop buffer (see recipe) 2.5 mg/ml proteinase K 50:49:1 (v/v/v) phenol/chloroform/isoamyl alcohol, equilibrated with 10 mM Tris⋅Cl, pH 8.0 (see UNIT 2.1A for equilibration technique) 2.5 M ammonium acetate 100% ethanol Siliconized 1.5 ml polypropylene tubes (e.g., ISC BioExpress, Cat. # C-3302-1) 27° and 30°C water baths Additional reagents and equipment for extraction of DNA (UNIT 2.1A) and agarose gel electrophoresis (UNIT 2.5A) Prepare master solutions for chromatin assembly 1. Thaw all buffers and proteins. It is recommended that all buffers be equilibrated to room temperature (for consistent pipetting). Proteins must be quick-thawed (in a room temperature water bath, mixed by tapping or extremely gentle vortexing, and transferred on ice) and quick-frozen (in liquid nitrogen) after use. ACF, NAP-1, core histones, and micrococcal nuclease (but not topoisomerase I or ISWI expressed and purified as an individual polypeptide) can withstand multiple freeze-thaw cycles (up to 10). Small working aliquots of topoisomerase I (which contains 50% glycerol) can be stored for up to 3 months at −20°C without freezing.

2. Prepare the master mix of NAP-1 and core histones (NH) by combining the following in a siliconized 1.5-ml microcentrifuge tube: 172 µl of HEG 70 µl of 300 mM KCl 84 µl of PvOH/PEG solution 4.2 µl of 2 mg/ml BSA 6.4 µl of 2 mg/ml NAP-1 3.03 µl of 0.7 mg/ml core histones. Vortex gently for 2 to 3 sec. Chromatin Assembly Using Drosophila Systems

Use siliconized 1.5-ml tubes throughout the protocol. The provided recipe is calculated for six standard reactions, and should be used to perform five or fewer reactions (to allow for imprecise pipetting). A single reaction is calculated to contain 0.353333 ìg DNA, 0.353333



ìg core histones, 2.12 ìg NAP-1, 10 units ACF (0.22 pmol), 1.4 ìg BSA in 70 ìl of a buffer, containing ∼50 mM KCl, 3 mM ATP, and ∼5 mM MgCl2. The final volume of the reaction is made up to 70 ìl with HEG. The NAP-1 to core histone mass ratio is 5:1 and should be sufficient to eliminate unbound histones in the reaction. PvOH/PEG and BSA are not obligatory—simply use or do not use them consistently in all assembly experiments. Preparing the master mix for several assembly reactions minimizes pipetting small volumes of core histones. Use a P2 Pipetman (Gilson) to pipet the core histones.

3. Pipet a 56.6-µl aliquot of the NH mix prepared in step 2 (at room temperature) into each of five siliconized 1.5-ml microcentrifuge tubes. Incubate on ice for ≥20 min to allow histone-NAP-1 binding. 4. Prepare the master mix of ATP and Mg2+ (AM) by combining 3 µl of 0.5 M ATP with 30 µl of 0.5 M creatine phosphate, 16.5 µl distilled water, and 25 µl 100 mM MgCl2. Add 0.5 µl 5 mg/ml creatine kinase immediately before use. The ATP regeneration system (creatine phosphate and creatine kinase) is optional (it is only required for assembly times >2 hr and high ACF concentrations). Keep the mix at room temperature at all times to avoid creatine phosphate precipitation. Discard the thawed aliquots of creatine kinase.

5. Prepare the DNA template by combining: 7.64 µl of plasmid DNA (at 0.42 mg/ml) 2 µl 10× topoisomerase I buffer 2.36 µl recombinant topoisomerase I working solution 8 µl distilled H2O. Relax the DNA for 10 min at 30°C; keep at room temperature until ready to use. Chromatin inhibits DNA relaxation by topoisomerase I. Thus, topoisomerase I in the reaction should be in 5- to 10-fold excess over the amount that is necessary to completely relax the supercoiled plasmid after a 10-min incubation at 30°C. The purified system will efficiently assemble chromatin on linear supercoiled DNA in the absence of the topoisomerase. In the latter case, the template must contain more than 95% supercoiled DNA (achieved by two rounds of CsCl banding).

6. Prepare ACF dilution(s) in ACF dilution buffer (2 to 10 units/µl). Keep on ice. 1 unit of ACF equals 22 fmol of protein.

Assemble chromatin and analyze by micrococcal nuclease assay 7. Start 5 assembly reactions as follows: a. To 56.6 µl NH (step 3) add 1 µl ACF (2 to 10 units) b. Transfer from ice and equilibrate to room temperature. c. Add 10.5 µl AM master mix (step 4) and 2 µl DNA template (relaxed with excess topoisomerase I or supercoiled; step 5). Immediately vortex, gently, for 2 to 3 sec. d. Allow the assembly to proceed at 27°C for 1.5 to 2.5 hr. Mix the reactions by gentle vortexing after adding each component. Microcentrifuge the tubes briefly after vortexing to collect the solution in the bottom of the tube. It is especially important to mix the reactions immediately upon the addition of the DNA template.

8. Immediately before use, prepare two dilutions of micrococcal nuclease in buffer R, 1:500 and 1:1500. Note the difference in dilution ratios from step 3 of Basic Protocol 3. Micrococcal nuclease digests purified chromatin more efficiently in the absence of inhibitory components within the crude system.



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9. Add 17.5 µl 10 mM CaCl2 to each reaction. Divide each reaction in two equal parts (“a” and “b”). In a controlled manner (at certain time intervals, e.g., 15 sec), add 5 µl of the 1:1500 dilution prepared in step 8 to each “a” tube and 5 µl of the 1:500 dilution to each “b” tube. Allow the digestion to progress for 10 min at room temperature for every tube. The assembly reaction can also be monitored by the DNA supercoiling assay. Stop one quarter of the 70 ìl assembly reaction (∼0.177 µg DNA in 17.5 ìl) by addition of 3 ìl 0.5 M EDTA. Deproteinate and precipitate the DNA as in step 11 below. Run, along with supercoiled and relaxed DNA samples, 1 kbp DNA ladder on a 0.8% agarose, 1× TBE gel until the xylene cyanol dye front reaches the bottom third of the gel. Stain and destain with ethidium bromide.

10. Prepare stop solution (ST) by mixing 55 µl of 0.5 M EDTA and 11 µl of 10 mg/ml RNaseA. Stop micrococcal digestions by adding 6 µl ST to each tube and then vortexing. Allow samples to stand 5 min at room temperature to digest contaminating RNA. From this point on, the reactions do not have to be timed precisely.

11. Prepare proteinase solution (PR) by mixing 1.1 ml glycogen stop buffer and 55 µl of 2.5 mg/ml proteinase K solution. Add 105 µl PR to each tube, then vortex. Digest the histones and soluble proteins at 37°C for ≥30 min. Extract with 200 µl of 50:49:1 phenol/chloroform/isoamyl alcohol. Precipitate the DNA with 25 µl of 2.5 M ammonium acetate and 475 µl of 100% ethanol. Do not wash the pellets with 70% ethanol. UNIT 2.1A describes general techniques for extraction and precipitation of DNA.

12. Perform agarose gel electrophoresis as described (see Basic Protocol 3, step 8, and UNIT 2.5A). ALTERNATE PROTOCOL 2

TITRATION OF THE RATIO OF CORE HISTONES TO DNA IN THE RECOMBINANT CHROMATIN ASSEMBLY REACTION For every new plasmid DNA and core histone preparation, the optimal core histone to DNA ratio should be determined experimentally. For materials, see Basic Protocol 6 1. Proceed as in Basic Protocol 6, steps 1 to 6. As an approximation for actual concentrations, use the DNA concentration as calculated from A260 and the histone concentration as determined by the BCA assay.

2. Start 5 assembly reactions as follows: a. To 56.6 µl NH (see Basic Protocol 6, step 3) add 1 µl ACF (2 to 10 units) b. Transfer from ice and equilibrate to room temperature. c. Add 10.5 µl AM master mix (see Basic Protocol 6, step 4) and 2.5, 2.22, 2.0, 1.82, or 1.67 µl DNA template (relaxed with excess topoisomerase I or supercoiled; see Basic Protocol 6, step 5). Immediately after adding DNA, vortex each tube gently for 2 to 3 sec. d. Allow the assembly to proceed at 27°C for 1.5 to 2.5 hr. Chromatin Assembly Using Drosophila Systems

The assumed histone to DNA mass ratios in these reactions are 0.8:1, 0.9:1. 1:1. 1.1:1, and 1.2:1, respectively. Use a P2 Pipetman (Gilson) to pipet the DNA solution.

3. Proceed as in steps 8 to 12 of Basic Protocol 6.



1

2

3

4

5

6

7

:1 1.2

:1 1.1

:1 1.0

:1 0.9

:1 0.8

Core histone to DNA mass ratio

8

9 10

Figure 21.7.2 Chromatin assembly in the recombinant system. The reactions were performed as described in Alternate Protocol 2. The core histone to DNA ratios were initially estimated as: lanes 1, 2—0.8:1; lanes 3, 4—0.9:1; lanes 5, 6—1.0:1; lanes 7, 8—1.1:1; lanes 9, 10—1.2:1.

4. From the gel, determine the highest histone-to-DNA mass ratio that still results in an unsmeared ladder (lanes 5, 6 in Fig. 21.7.2, 1.0:1 estimated ratio). The next lowest ratio (i.e., 0.9:1 as in lanes 3, 4 of Fig. 21.7.2) is postulated to be the optimal ratio for further chromatin assembly experiments. The effective concentration of the core histones is recalculated so that the new calculated optimal ratio is set to 1.0:1. For instance, for the experiment in Figure 21.7.2, the original (estimated) histone concentration is multiplied 1.111 times. Thus the histone to DNA ratio in lanes 3, 4 calculated from the new (effective) histone concentration is 1.0:1.

EXPRESSION AND PURIFICATION OF THE CORE CATALYTIC DOMAIN OF THE DROSOPHILA TOPOISOMERASE I

SUPPORT PROTOCOL

The smallest active N-terminal truncation (Shaiu and Hsieh, 1998) of the Drosophila topoisomerase I was cloned into pET-28a expression vector (Novagen) from a cDNA that was kindly provided by Tao-shih Hsieh at Duke University. The coding sequence was subcloned into NcoI/XhoI restriction sites in frame with the C-terminal 6-His tag. The construct is referred to as pET-NDH6; the protein is referred to as ND423. Chromatin Assembly and Analysis


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Materials Competent BL21(DE3) bacteria (Novagen) PET-NDH6 plasmid (available on request from Dmitry Fyodorov; [email protected]) LB plates and medium (UNIT 1.1) containing 50 µg/ml kanamycin (add from 10 mg/ml kanamycin stock) 100 mM IPTG Liquid nitrogen Lysis buffer T (see recipe) Ni-NTA resin (Qiagen) Elution buffer T: lysis buffer T containing 0.5 M imidazole Dialysis buffer T (see recipe) Storage buffer T (see recipe) 8% SDS-PAGE gel (UNIT 10.2) Bovine serum albumin (BSA) standard, 2 mg/ml (Pierce, Cat. No. 23209) 0.8% agarose gel (UNIT 2.5A) 10× topoisomerase I buffer (see recipe) Sorvall Superspeed centrifuge with GSA and SS-34 rotors (or equivalents) Microtip sonicator (e.g., Branson Sonifier 450; VWR Scientific) 10-ml polypropylene chromatography column 0.5 to 3.0 ml Slide-A-Lyzer, 10,000 MWCO (Pierce) Additional reagents and equipment for transformation of bacteria (UNIT 1.8), growth of bacteria in solid (UNIT 1.3) and liquid (UNIT 1.2) media, SDS-PAGE (UNIT 10.2), and agarose gel electrophoresis (UNIT 2.5A) Express ND423 in E. coli 1. Transform BL21(DE3) cells with pET-NDH6. Plate on LB plates containing 50 µg/ml kanamycin; incubate overnight at 37°C. Transformation is performed immediately prior to the expression.

2. Pick one average size colony into 0.5 liter LB medium (containing 50 µg/ml kanamycin) and grow while shaking at 37°C for 6 to 8 hr. At a bacterial density equivalent to A600 ∼0.5, induce by adding 100 mM IPTG to a final concentration of 0.42 mM. 3. Incubate while shaking at 30°C for 5 hr. Harvest the cells by centrifugation for 10 min at 8500 × g (7000 rpm in a GSA rotor), 4°C. Note the temperature change from 37° to 30°C in this incubation.

4. Freeze the cell pellets in liquid nitrogen. The pellets can be stored at −80°C overnight, if desired.

Extract soluble bacterial proteins and isolate ND423 by affinity chromatography 5. Thaw and resuspend the cell pellets in 10 to 20 ml lysis buffer T (20 to 40 ml per 1 liter of original culture). Freeze again in liquid nitrogen and thaw in a water bath at room temperature. 6. Sonicate on ice with a microtip with 4 burst of 30 sec at a setting of “6.5.” Centrifuge the lysate in 40 ml polycarbonate tubes for 10 min at 19,000 × g (16,000 rpm in an SS-34 rotor), 4°C. Chromatin Assembly Using Drosophila Systems

7. Equilibrate 2 ml Ni-NTA resin with the lysis buffer T. Mix and incubate the cell lysate with the resin for 3 hr in the cold room on a rocking platform.



8. Load the resin onto a disposable 10-ml polypropylene column by gravity flow (in a cold room). Wash 3 times with 10 ml of cold lysis buffer T. Elute three times, each time with 1 ml elution buffer, discarding the void volume (the first 350 µl). 9. Dialyze in a 10,000 MWCO Slide-A-Lyzer against 2 liters dialysis buffer T for 2 hr at 4°C. Dialyze again against 1 liter storage buffer T for 2 hr at 4°C, then freeze in 100-µl aliquots in liquid nitrogen. Store at −80°C. Determine the concentration by visualizing on an 8% SDS-PAGE gel (UNIT 10.2) along with the BSA mass standard. The typical yield is 1.5 to 2.0 mg protein in 1.0 to 1.2 ml.

Prepare the working stock of topoisomerase I and assay the enzymatic activity 10. Prepare the working solution by 100-fold dilution in storage buffer T containing 0.2 mg/ml human recombinant insulin (add from 50 mg/ml stock). Upon dilution, store in 100-µl aliquots at −80°C. Keep one aliquot at −20°C for daily use. 11. Assay the activity of the protein by relaxing 0.5 µg supercoiled plasmid DNA with different dilutions of the enzyme. Run on a 0.8% agarose gel; stain with ethidium bromide and destain (UNIT 2.5A). 1 ìl of the working stock should completely relax ∼10 ìg of 3 kbp plasmid in a 100-ìl reaction in 1× topoisomerase buffer after 10 min incubation at 30°C.


ATP mix 300 mM creatine phosphate 30 mM ATP 10 µg/ml creatine kinase Store up to 2 years at −80°C Buffer A 15 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 15 mM NaCl 60 mM KCl 0.34 M sucrose 0.1% (v/v) 2-mercaptoethanol Store solution with above components up to 24 hr at 4°C 0.5 mM spermidine (add immediately prior to use) 0.15 mM spermine (add immediately prior to use) 0.25 mM PMSF (add immediately prior to use) Buffer B To buffer A (see recipe above), add: 2 mM EDTA 0.5 mM EGTA Store up to 24 hr at 4°C Buffer R 10 mM potassium HEPES, pH 7.6 10 mM KCl 1.5 mM MgCl2 0.5 mM EGTA continued



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10% (v/v) glycerol Store solution with above components up to 24 hr at 4°C 10 mM β-glycerophosphate (add immediately prior to use) 1 mM DTT (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) Aliquots of 0.1 to 1 ml can be stored at −20°C for up to 2 years.

Core histone storage buffer 10 mM potassium HEPES, pH 7.6 1 mM EDTA 10 mM KCl 10% (v/v) glycerol Store solution with above components up to 24 hr at 4°C 1 mM DTT (add immediately prior to use) Creatine kinase solution 5 mg/ml creatine kinase (Sigma-Aldrich) 10 mM potassium phosphate, pH 7.0 (APPENDIX 2) 50 mM NaCl 50% (v/v) glycerol Store in 5- to 10-µl aliquots up to 2 years at −80°C Creatine phosphate, 0.5 M 106 mg/ml creatine phosphate (phosphocreatine) 20 mM potassium HEPES, pH 7.6 Adjust pH to 7.0 Store in aliquots of 0.1 to 1 ml up to 2 years at −20°C Dialysis buffer T HEG buffer (see recipe below) containing: 50 mM NaCl 0.01% NP-40 Store solution with above components up to 24 hr at 4°C 0.5 mM DTT (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 0.5 mM benzamidine (add immediately prior to use) 5 mM Na2S2O5 (add immediately prior to use) Dilution buffer F 20 mM Tris⋅Cl, pH 7.9 (APPENDIX 2) 10% (v/v) glycerol 0.02% (v/v) NP-40 Store up to 24 hr at 4°C Elution buffer F Wash buffer F (see recipe below) containing: 0.4 mg/ml FLAG peptide (Sigma-Aldrich) 0.4 mg/ml recombinant human insulin (Roche) Use immediately Add FLAG peptide from 10 mg/ml stock in STE buffer and add insulin from 50 mg/ml stock in TE buffer (see APPENDIX 2 for buffers). Chromatin Assembly Using Drosophila Systems

Store stock solution of FLAG peptide in 20-ìl aliquots up to 2 years at −80°C. Store insulin stock solution up to 1 year at 4°C.



Glycogen stop buffer 20 mM EDTA 0.2 M NaCl 1% (w/v) SDS 0.25 mg/ml glycogen Store up to 2 years at room temperature HA chromatography buffer 40 mM sodium phosphate, pH 6.8 (APPENDIX 2) 0, 0.35, or 2.5 M NaCl Store solution with above components up to 24 hr at 4°C 1 mM DTT (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) HEG buffer 25 mM potassium HEPES, pH 7.6 0.1 mM EDTA 10% (v/v) glycerol Store in aliquots of 0.1 to 1 ml up to 2 years at −20°C HEGD buffer with 0.1 M NaCl 25 mM potassium HEPES, pH 7.6 1 mM EDTA 10% (v/v) glycerol 0.1 M NaCl 0.01% (v/v) NP-40 Store solution with above components up to 24 hr at 4°C 1 mM DTT (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 10 mM β-glycerophosphate (add immediately prior to use) Lysis buffer F 20 mM Tris⋅Cl, pH 7.9 (APPENDIX 2) 500 mM NaCl 20% (v/v) glycerol 4 mM MgCl2 0.4 mM EDTA Store solution with above components up to 24 hr at 4°C 2 mM DTT (add immediately prior to use) 20 mM β-glycerophosphate (add immediately prior to use) 0.4 mM PMSF (add immediately prior to use) 1 mM benzamidine hydrochloride (add immediately prior to use) 4 µg/ml leupeptin (add immediately prior to use) 2 µg/ml aprotinin (add immediately prior to use) Lysis buffer H 50 mM sodium phosphate, pH 7.6 (APPENDIX 2) 0.5 M NaCl 15% (v/v) glycerol 20 mM imidazole 0.01% (v/) NP-40 Store solution with above components up to 24 hr at 4°C 10 mM β-glycerophosphate (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 0.5 mM benzamidine (add immediately prior to use)



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Lysis buffer T 50 mM sodium phosphate, pH 7.0 (APPENDIX 2) 0.5 M NaCl 15% (v/v) glycerol 15 mM imidazole 0.1% (v/v) NP-40 Store solution with above components up to 24 hr at 4°C 0.2 mM PMSF (add immediately prior to use) 0.5 mM benzamidine (add immediately prior to use) 10 mM Na2S2O5 (add immediately prior to use) Micrococcal nuclease stock solution, 200 U/ml 1.56 mg/ml (200 U/ml) micrococcal nuclease (Sigma-Aldrich) 5 mM sodium phosphate, pH 7.0 (APPENDIX 2) 2.5 µM CaCl2 Store in aliquots of 0.1 to 1 ml up to 1 year at −20°C NAP-1 purification buffer Buffer R (see recipe above) containing: 0.0, 0.1, or 1.0 M NaCl (add from 5 M NaCl stock or as solid NaCl) 0.01% NP-40 (add from 10% v/v stock) Store up to 24 hr at 2°C PvOH/PEG solution HEG buffer (see recipe above) containing: 5% polyvinyl alcohol (mol. wt. 10,000, Sigma-Aldrich P-8136) 5% polyethylene glycol (mol. wt. 8,000, Sigma-Aldrich P-2139) Store in aliquots of 0.1 to 1 ml up to 2 years at −20°C Storage buffer T 10 mM potassium HEPES, pH 7.6 0.1 mM EDTA 50 mM NaCl 0.01% (v/v) NP-40 50% (v/v) glycerol Store solution with above components up to 24 hr at 4°C 10 mM β-mercaptoethanol (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 1 mM benzamidine (add immediately prior to use) 1 µg/ml leupeptin (add immediately prior to use) Sucrose gradients, 5% to 30% 10 mM potassium HEPES, pH 7.6 1 mM EDTA 0.5 M NaCl 5% or 30% (w/v) sucrose 0.02% (w/v) NaN3 Store solution with above components up to 1 year at 4°C 0.2 mM PMSF (add immediately prior to use) Use a gradient maker to prepare 5% to 30% sucrose gradients in SW 28 ultracentrifuge tubes Chromatin Assembly Using Drosophila Systems



Superflow chromatography buffer 50 mM sodium phosphate, pH 7.6 (APPENDIX 2) 500 mM NaCl 15% (v/v) glycerol 0.01% (v/v) NP-40 Store solution with above components up to 24 hr at 4°C 10 mM β-glycerophosphate (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 0.5 mM benzamidine (add immediately prior to use) 2 µg/ml leupeptin (add immediately prior to use) 2 µg/ml aprotinin (add immediately prior to use) Topoisomerase I buffer, 10× 0.5 M Tris⋅Cl, pH 7.5 (APPENDIX 2) 100 mM MgCl2 1 mM EDTA 0.5 mg/ml BSA 5 mM DTT Store in aliquots of 0.1 to 1 ml up to 2 years at −20°C Wash buffer F 20 mM Tris⋅Cl, pH 7.9 (APPENDIX 2) 150 mM NaCl 15% (v/v) glycerol 2 mM MgCl2 0.2 mM EDTA 0.01% (v/v) NP-40 Store solution with above components up to 24 hr at 4°C 1 mM DTT (add immediately prior to use) 10 mM β-glycerophosphate (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 0.5 mM benzamidine-HCl (add immediately prior to use) 2 µg/ml leupeptin (add immediately prior to use) 1 µg/ml aprotinin (add immediately prior to use) Wash buffer H 50 mM sodium phosphate, pH 7.6 (APPENDIX 2) 100 mM NaCl 20 mM imidazole 15% glycerol (v/v) 0.01% NP-40 (v/v) Store solution with above components up to 24 hr at 4°C 10 mM β-glycerophosphate (add immediately prior to use) 0.2 mM PMSF (add immediately prior to use) 0.5 mM benzamidine (add immediately prior to use) COMMENTARY Background Information Several procedures to assemble nucleosome arrays on circular or linear DNA in vitro have been developed (reviewed in Ito et al., 1997a). Some of these procedures employ a histone transfer process with a histone transfer vehicle (histone-binding proteins, polyglutamic acid,

RNA, or high salt concentration; UNIT 21.6). They yield well-defined templates, yet the nucleosomes are irregularly distributed and lack the periodicity of bulk native chromatin. The second type of these procedures utilizes crude cell extracts and yields extended periodic arrays of nucleosomes. This reaction was



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achieved with cytosolic extracts from Xenopus oocytes (Glikin et al., 1984), HeLa cells (Banerjee and Cantor, 1990), or Drosophila embryos (Becker and Wu, 1992; Bulger and Kadonaga, 1994). The third type of chromatin assembly procedure is the assembly during viral DNA replication (Stillman, 1986). In this reaction, crude cell extracts (from human 293 cells) are complemented by SV40 T antigen helicase and histone chaperones CAF-1 and RCAF (Tyler et al., 1999) to achieve chromatin assembly of the newly replicated SV40 DNA. The early realization that chromatin assembly in crude cell extracts is an ATP-dependent process has triggered the quest to purify individual protein factors that could mediate this reaction. Fractionation of Drosophila extracts resulted in purification of two essential assembly components, a histone chaperone NAP-1 (Bulger et al., 1995) and a SWI/SNF-like chromatin remodeling factor, ACF (Ito et al., 1997b). This work was supported by the NIH grant GM58272 to James T. Kadonaga at University of California, San Diego. ACF consists of two subunits, a SWI2/SNF2-related ATPase ISWI (p140) and a subunit termed Acf1 (p170 and p185). The chromatin assembly reaction can be reconstituted in a purified recombinant system that contains plasmid DNA, purified native or recombinant histones, recombinant NAP-1, recombinant ACF, and ATP (Ito et al., 1999; Levenstein and Kadonaga, 2002). This system combines the advantages of histone transfer procedures (well-defined reaction products) with the advantages of ATP-dependent crude extract assembly procedures (nucleosome array regularity similar to that of the native bulk chromatin). The pair of cofactors (an ATP-dependent remodeling factor and a histone chaperone) is likely to represent the minimal required reaction system that assembles physiologically relevant chromatin. The histone chaperone NAP-1 is necessary to prevent rapid nonspecific histone-DNA aggregation and to present histones to ACF. Consistent with its role as a molecular sink that sequesters positively charged histones, NAP-1 has to be present at 1:1 or higher stoichiometry with respect to core histone polypeptides. Although NAP-1 has a higher affinity to H2A/H2B dimers, it also binds to H3/H4 tetramers and can facilitate the assembly of nucleosomes from dimers and tetramers. On the other hand, ACF performs multiple functions: core histone deposition on the DNA, establishing proper histone-DNA contacts in the nucleosome, and spacing nu-

cleosome arrays. All these tasks are carried out catalytically (substoichiometric relative to the core histones) and at the expenditure of ATP hydrolysis by ISWI. Although histone chaperones (and specifically NAP-1) have been reported to mediate nucleosome assembly with supercoiled plasmid DNA, it is important to note that they do not deposit histones efficiently on relaxed DNA templates. Furthermore, electron microscopy of “chromatin” assembled by NAP-1 alone reveals that it is composed of large histone-DNA particles that are not canonical nucleosomes (Nakagawa et al., 2001). Finally, the replication-dependent chromatin assembly of the SV40 DNA by CAF-1 and RCAF histone chaperones may also be mediated by an ATPdependent chromatin remodeling complex that is present in the 293 cell extract.

Critical Parameters and Troubleshooting S-190 extract A concentrated extract, stemming from a high ratio of embryos to buffer R, tends to be a highly active extract. Aspirating buffers too cautiously to prevent the loss of embryo material may result in a dilute extract. This can lead to low quality of assembly or require the S-190 to assume a significant portion of the reaction volume. Homogenizing embryos in a Dounce homogenizer generates sizeable resistance on down strokes as well as considerable back-pressure on up strokes. Be careful not to lose grip of the pestle during homogenization, or else the vessel may break and the embryos may be lost in the ice bucket. Be certain to use full strokes in order to homogenize all embryos. Between the first and second spins in the ultracentrifuge, the S-190 extract should be frozen in liquid nitrogen and then thawed in a room-temperature water bath. This step presumably precipitates unwanted materials and improves the activity of the assembly extract. Core histone purification When isolating the nuclei, the yellow yolk protein is undesirable. It will form a tight pellet at the bottom of the tube. A Pasteur pipet can be used to carefully separate the loose nuclear pellet from the yolk before resuspending. The addition of 5 M NaCl to the nuclear suspension (step 13) may not result in obvious lysis. Continue to the next step under the assumption that the lysis was successful. When selecting fractions from the sucrose gradient, avoid those



containing histone H1. H1 will be a significant band at ∼40 kDa. Dialyzing the sucrose gradient fractions will result in significant swelling within the dialysis tubing. Take this into consideration when cutting the tubing size and consider using double clips to avoid mishaps. Hydroxylapatite resin crushes easily and needs to be run on the FPLC at low pressure. If the core histone volume is large, this is a good opportunity to inject the sample overnight at a very slow flow rate (0.1 ml/min or less). As an alternative to crystalline hydroxylapatite, ceramic hydroxylapatite resin is now available from Bio-Rad Laboratories (CHT Type I Ceramic Hydroxyapatite). This resin performs well at higher pressures and yields otherwise equivalent results. S-190 chromatin assembly Three key parameters tend to influence assembly with the S-190 extract: (1) the proportion of the extract in the reaction volume, (2) the Mg2+ concentration, and (3) the ratio of core histones to DNA. Variations among extracts result in a range of adequate S-190 proportions, typically falling between 25% to 40% of the reaction volume. The most desirable percentage should be determined for each extract. The optimal Mg2+ concentration lies between 3.5 and 7 mM. This should also be determined for each extract. The ratio of histones to DNA is affected by the volume of the S-190 used. The S-190 extract contains a residual pool of core histones (in the realm of 20% to 30% required for assembly). Typically, a 0.8:1 mass ratio yields high quality chromatin (as assayed by partial micrococcal nuclease digestion), but this ratio can be examined from 0.6:1 to 1.2:1. If low-quality agarose is used for gel electrophoresis, ladders can appear “smeared.” SeaKem LE agarose (BioWhittaker) is suggested. Complete destaining can significantly improve the appearance of nucleosomal ladders, especially those at the top of the ladder. Purification of recombinant ACF and NAP-1 Lysing of the Sf9 cells must be thorough and complete. Low yield is likely to be a direct result of incomplete lysis. Sonication can be used in place of homogenization on the Dounce homogenizer to disrupt Sf9 cells (2 to 3 bursts of 30 sec with a microtip, on ice). The resulting extract will be cruder, and thus the affinity resins should be washed more thoroughly after the protein binding step.

Typically, purified ACF will contain equimolar amounts of p185 (Acf1-FLAG) and p140 (ISWI). However, ACF complexes with substoichiometric ISWI will also be suitable for most chromatin assembly applications. Calculate ACF molar concentration based on the concentration of the ISWI subunit. After affinity chromatography, the dNAP-1 protein is often more than 95% pure but performs poorly in assembly reactions. At this stage, it exists as a mixture of pure active NAP-1 multimers and an inhibitory NAP-1 fraction, which is contaminated with a 14-kDa Sf9 protein. It is therefore critical to perform the anion-exchange chromatography to separate active NAP-1 from this inhibitory species. Recombinant chromatin assembly The purified system performance is relatively harder to fine-tune than the S-190 based system. While the success in the latter is determined primarily by the quality of the extract, the purified system requires more extensive optimization of reaction parameters to achieve the best performance. The single most critical parameter in the reaction is the ratio of core histones and template DNA. Variations as low as 5% to 10% can inhibit the reaction or result in lower-quality chromatin as evidenced by the micrococcal digest analysis. Standard methods to determine DNA and protein concentrations (spectrophotometric, colorimetric) are almost never accurate enough to establish the correct histone-to-DNA ratio. Thus, it has to be determined experimentally for every pair of DNA and histone preparations. Alternate Protocol 2 provides an idea of such a titration experiment. The range and the step of this titration can be expanded, if necessary. In general, low histone content results in short ladders (2 or 3 apparent nucleosome bands at the bottom) and large nucleosome repeat (180 bp and longer). Excessive histone concentration will completely inhibit the reaction and result in histone-DNA aggregates that will not be efficiently digested by the micrococcal nuclease (see Fig. 21.7.2). The assembly reaction is much less sensitive to the concentration of NAP-1. It should be noted, however, that it is safer to add an excess of the chaperone. If the mass ratio of NAP-1 to core histones drops below 3.8 to 4, the extra free histones that are present in the solution can form histone-DNA aggregates, which cannot serve as a substrate for ACF. The assembly reaction is tolerant to over an order of magnitude variation in ACF concentration. ACF will



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assemble chromatin within a range of concentrations from 0.05 to 1 ACF protomers per 1 kbp DNA. It is recommended that the reaction be first optimized for a short plasmid template (3 kbp or less) before attempting to assemble chromatin on larger templates. Chromatin may be purified after the assembly by gel filtration or sucrose gradient centrifugation. However, unpurified chromatin templates perform equally well in transcription assays. Several components of the assembly reaction are semi-optional. They include PvOH/PEG, BSA, ATP regeneration system, and topoisomerase I (if the DNA template is predominantly supercoiled). The reactions can be formulated and optimized accordingly (the changes will alter other reaction parameters, such as the optimal histone to DNA and NAP-1 to histone ratios). The suggested reaction conditions and micrococcal dilutions will produce nucleosome ladders with an apparent repeat length of ∼165 bp. The nucleosomes can be packed more loosely or more tightly (down to 145 bp repeat length) by varying KCl concentration and the amount of histones in the assembly reaction. Tighter packed nucleosomes will require higher micrococcal concentration to be comparably digested. The purified system appears insensitive to histone modification states: it assembles chromatin equally well with purified native or unmodified bacterially expressed Drosophila core histones (Levenstein and Kadonaga, 2002). Moreover, the system does not exhibit specificity towards Drosophila core histones and can assemble chromatin with histones purified from other species.

Anticipated Results


Both ATP-dependent chromatin assembly systems described here can provide chromatin substrates for use in studies of chromatin structure, transcription, and DNA metabolism. Micrococcal nuclease analysis of the reaction products should reveal highly periodic nucleosome arrays. A DNA-binding transcription factor, when added to the assembly reaction, can position nucleosomes with respect to its binding sites. The number of negative supercoils in the assembled DNA (for circular plasmid DNA templates) can be estimated by oneor two-dimensional supercoiling analyses and may be used as a measure of the nucleosome content of the assembled minichromosomes. The assembled chromatin should migrate differently and can be separated from the unassembled DNA on sucrose gradients.

Time Considerations Allow 2 days each for core histone purification and preparation of the S-190 extract. S-190 mediated chromatin assembly is extremely reproducible and does not require extensive troubleshooting. Typically, chromatin is assembled, then digested with micrococcal nuclease (6 to 7 hr) and analyzed by electrophoresis on the next day (3 to 4 hr). Allow 3 to 5 days for baculovirus amplification and 2 to 3 days for infection of Sf9 cells before harvesting. The recombinant ACF prep takes 1 day, and the NAP-1 prep takes 2 days. Recombinant chromatin assembly may require several pilot experiments to optimize concentrations of the components. Many researchers have reported that the quality of chromatin assembly in the recombinant system improves significantly after several trials.

Acknowledgments The studies of the Drosophila chromatin assembly factors were supported by the NIH grant GM58272 to James T. Kadonaga. We thank Dr. Kadonaga for critical reading of the manuscript.

Literature Cited Banerjee, S. and Cantor, C.R. 1990. Nucleosome assembly of simian virus 40 DNA in a mammalian cell extract. Mol. Cell. Biol. 10:2863-2873. Becker, P.B. and Wu, C. 1992. Cell-free system for assembly of transcriptionally repressed chromatin from Drosophila embryos. Mol. Cell. Biol. 12:2241-2249. Bulger, M. and Kadonaga, J.T. 1994. Biochemical reconstitution of chromatin with physiological nucleosome spacing. Methods Mol. Genet. 5:241-262. Bulger, M., Ito, T., Kamakaka, R.T., and Kadonaga, J.T. 1995. Assembly of regularly-spaced nucleosome arrays by dCAF-1 and a 56 kDa histone-binding protein. Proc. Natl. Acad. Sci. U.S.A. 92:11726-11730. Glikin, G.C., Ruberti, I., and Worcel, A. 1984. Chromatin assembly in Xenopus oocytes: In vitro studies. Cell 37:33-41. Ito, T., Tyler, J.K., and Kadonaga, J.T. 1997a. Chromatin assembly factors: A dual function in nucleosome formation and mobilization? Genes Cells 2:593-600. Ito, T., Bulger, M., Pazin, M.J., Kobayashi, R., and Kadonaga, J.T. 1997b. ACF, an ISWI-containing and ATP-utilizing chromatin assembly and remodeling factor. Cell 90:145-155. Ito, T., Levenstein, M.E., Fyodorov, D.V., Kutach, A.K., Kobayashi, R., and Kadonaga, J.T. 1999. ACF consists of two subunits, Acf1 and ISWI, that function cooperatively in the ATP-depend-



ent catalysis of chromatin assembly. Genes & Dev. 13:1529-1539.

Stillman, B. 1986. Chromatin assembly during SV40 DNA replication in vitro. Cell 45:555-565.

Levenstein, M.E. and Kadonaga, J.T. 2002. Biochemical analysis of chromatin containing recombinant Drosophila core histones. Submitted.

Tyler, J.K., Adams, C.R., Chen, S.-R., Kobayashi, R., Kamakaka, R.T., and Kadonaga, J.T. 1999. The RCAF complex mediates chromatin assembly during DNA replication and repair. Nature 402:555-560.

Nakagawa, T., Bulger, M., Muramatsu, M., and Ito, T. 2001. Multistep chromatin assembly on supercoiled plasmid DNA by nucleosome assembly protein-1 and ATP-utilizing chromatin assembly an d remo delin g factor. J. Biol. C hem. 276:27384-27391. Shaiu, W.L. and Hsieh, T.S. 1998. Targeting to transcriptionally active loci by the hydrophilic N-terminal domain of Drosophila DNA Topoisomerase I. Mol. Cell. Biol. 18:4358-4367.

Contributed by Dmitry V. Fyodorov and Mark E. Levenstein University of California, San Diego La Jolla, California



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Chapter 22 Nucleic Acid Arrays INTRODUCTION

N

ucleic acid arrays present a near-textbook case for a technology in which an increase in throughput results in a qualitative change in the usefulness of the technique. Arrays of nucleic acids on nitrocellulose have been around for many years (consider dot blots and slot blots, used for gene expression monitoring since the late 1970’s). Nucleic acid arrays have followed a now-typical “genomic” pattern, in which, above some point where the representation of genes on the array begins to comprise a substantial fraction of the genes of a genome or of a set of important genes, the experimental value gained from the use of that array becomes much larger.

For insight into current applications of microarray technology in molecular biology, the reader is referred to UNIT 22.1. In this unit, Joseph DeRisi describes how and why methods that use these arrays have gone from costly—and requiring great technical sophistication—to mainstream. The first procedural unit in this chapter details the preparation of RNA for expression monitoring (UNIT 22.2). Both methods and ideology (i.e., “heuristics”) for use of DNA arrays, micorarrays, and chips are developing rapidly, and the editors will be particularly vigilant in adding new methods as those prove useful. Units currently in development will describe how researchers can make their own nucleic acid arrays, how they can use these and commercially available arrays to perform gene expression monitoring (GEM) and comparative genome hybridization (CGH), and how to collect and interpret the data. Roger Brent

Nucleic Acid Arrays Contributed by Roger Brent Current Protocols in Molecular Biology (2000) 22.0.1 Copyright © 2000 by John Wiley & Sons, Inc.


Overview of Nucleic Acid Arrays Nucleic acid array technology refers to the fabrication and use of arrays containing thousands of nucleic acid samples bound to solid substrates, such as glass microscope slides or silicon wafers. Because the physical area occupied by each sample is usually 50 to 200 µm in diameter, nucleic acid samples representing entire genomes, ranging in size from 3,000 to 32,000 genes, may be efficiently packaged onto a single regular microscope slide in an area easily covered by a coverslip (Fig. 22.1.1). Such “genomes on a chip” then serve as a target to which fluorescently labeled nucleic acid probes can be applied. Nucleic acid arrays, or microarrays, allow all genes of a given genome to be simultaneously monitored with respect to some experimental condition of interest. This fact has fundamentally changed the manner in which the study of genomics and gene expression can be pursued. The majority of applications discussed in this overview relate to DNA microarrays fabricated by the mechanical deposition of nucleic acid samples onto glass. Typically, these samples are in the form of PCR products, ranging in size from 100 bps to 9 kb. However, the term “DNA microarray” may apply to several different forms of the technology, each differing in the type of nucleic acid applied and the method of application. For example, Affymetrix sells DNA arrays produced by photolithographic synthesis of individual short oligonucleotides directly on the substrate. (Fodor et al., 1991).

WHAT ARE MICROARRAYS GOOD FOR? Gene Expression Analysis Undoubtedly the most common use for DNA microarrays is for monitoring gene expression levels. The broad appeal of this approach stems from the fact that it can be applied to virtually any organism, tissue, or cell line from which RNA may be isolated. In a typical experiment, total RNA or mRNA is collected from two or more individuals, cultures, or conditions. The amount of RNA needed for a microarray experiment depends on many factors, such as genome complexity and message content. Most experiments use anywhere from 100 ng to as much as 20 µg of RNA. The next step is the separate conversion of the RNA samples into cDNA by reverse transcription. This is

Contributed by Joseph DeRisi Current Protocols in Molecular Biology (2000) 22.1.1-22.1.7 Copyright © 2000 by John Wiley & Sons, Inc.

usually accomplished by priming with randomized oligonucleotides or, in the case of organisms that produce polyadenylated messages, an oligo-dT primer. The basic principle behind these manipulations is to convert the RNA from each sample into a form that can be readily distinguished from another RNA sample. This is usually accomplished by labeling the cDNA samples with different fluorescent dyes, either during the reverse transcription process through direct incorporation by reverse transcriptase, or afterwards by chemical conjugation. The resulting pools of cDNA are mixed together, and when the pool is hybridized to a microarray, the ratio between the intensities observed for two of the fluorophores at any given location in the array is a direct measure of the relative abundance of the corresponding cDNA transcript (see Fig. 22.1.2). By using a single reference sample as the control for a series of experimental samples collected over time, one can compare relative levels of transcript abundance among samples (see Fig. 22.1.3). This in turn allows one to identify gene expression trends. For those who wish to study global regulation of gene expression, the most significant data lies in these trends and patterns. Many instances where this methodology has been successfully put into practice can be found in the literature (DeRisi et al., 1997, Alizadeh et al., 1998, Cho et al., 1998; Chu et al., 1998, Eisen et al., 1998, Spellman et al., 1998; Amundson et al., 1999; Iyer et al., 1999; Perou et al., 1999). It is certain that the use of microarrays to analyze gene expression will continue to increase. Aside from looking at developmental time courses, mutations, and other genetic modifications, many other novel expression experiments will undoubtedly be developed. One recent variation includes the use of expression analysis to reverse-engineer the changes that occurred during a 200-generation yeast evolution experiment (Ferea et al., 1999). In addition to well-established model systems, gene expression analysis involving microarrays may be used as a method to attack problems that were formerly too cumbersome to approach using standard molecular biology techniques. This is especially true for potentially dangerous and difficult-to-culture organisms such as Mycobacterium tuberculosis and

UNIT 22.1

Nucleic Acid Arrays


Plasmodium falciparum (Hayward et al., 1999, Wilson et al., 1999). The primary reason for this is that only relatively small amounts of RNA are required to determine expression profiles for entire genomes. In other words, the data return on the up-front labor investment is much more substantial than with older techniques for studying gene expression, thus enlarging the potential scope and depth of experiments. For example, consider the case of Plasmodium: genetic crosses are extremely difficult and transformation efficiencies are abysmal. These facts make many of the tools researchers have relied upon for cloning new genes nearly useless. Therefore, determining which genes are induced by various drugs becomes a large un-

Overview of Nucleic Acid Arrays

dertaking, especially considering that the Plasmodium genome has yet to be sequenced completely. However, it is a relatively simple matter to collect RNA samples, so DNA microarrays (utilizing known or random genomic fragments) provide an easy means to answer these questions.

WHAT ELSE ARE NUCLEIC ACID MICROARRAYS GOOD FOR? Despite all the recent emphasis on gene expression analysis, the number of different uses for DNA microarrays is currently limited by our imagination. The most fundamental feature of microarray analysis is that meaningful data can be derived from any set of biological

Figure 22.1.1 Yeast “genome on a chip.”



experiments that result in a differential recovery of nucleic acid material. Aside from gene expression, these uses fall into several basic categories: genetic mapping and genotyping, assessment of genome structure and copy number, polysome analysis, and assays of DNAprotein interaction. Given that there are dozens of variations in experimental design and approach within each of these categories, the following is meant only to pique the imagination and alert readers to the diverse opportunities afforded by microarray technology.

Genotyping and Genetic Mapping Efficient genotyping of hundreds or thousands of markers for the purpose of mapping multigenic traits may be carried out using DNA microarrays. In one approach, genotyping by array analysis is accomplished by directly detecting hybridization differences caused by single-nucleotide polymorphisms (SNPs). Arrays utilizing short oligo features are well suited for the former method due to their low annealing

temperatures (Hacia et al., 1996). Another approach, genomic mismatch scanning (GMS), uses mismatch repair enzymes to recognize and selectively degrade hybrid DNA fragments between two individuals that possess SNPs (Nelson et al., 1993). The hybrid DNA is created by denaturing and reannealing fragments from two related individuals. The resulting perfectly matched DNA fragments can then be fluorescently labeled and applied to a microarray, revealing which segments are identical by descent (Cheung et al., 1998; McAllister et al., 1998). Regardless of the method used, genotyping by array analysis has the potential to increase the throughput by several orders of magnitude when compared to traditional gel electrophoresis–based methods.

Comparative Genomic Hybridization Differences in gene copy number have been associated with various tumorigenic phenotypes. These polymorphisms, which consist of deletions and/or amplifications, can easily be

grow yeast under two differing conditions

isolate mRNA from each sample

reverse transcribe mRNA in the presence of fluorescent nucleotides

mix fluorescent cDNA and hybridize to the microarray

rinse away unbound material, read with a laser microscope, and quantitate

Figure 22.1.2 Scheme for a typical gene expression experiment.

Nucleic Acid Arrays


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assayed using DNA microarrays, which provide a viable alternative to traditional comparative genomic hybridization (CGH) techniques. The primary limitation of traditional cytogenetic CGH techniques lies in the fact that the resolution to which an amplification or deletion may be mapped is ∼20 Mb (Kallioniemi et al., 1992). With array-based CGH methods, the resolution depends only on how many DNA elements are present on the microarray. Therefore, it is entirely feasible that all human genes may be represented as discrete elements on future microarrays, taking the resolution of the map to its logical limit. Indeed, with the conclusion of the human genome sequencing project, it should be feasible to detect the deletion or amplification of virtually any and all genes simultaneously for a given human cell population. Several examples already exist in the literature, and advances in the use of the technology are sure to follow (Solinas-Toldo et al., 1997; Trent et al., 1997; Pinkel et al., 1998; Behr et al., 1999; Pollack et al., 1999).

Polysome Analysis Although transcriptional regulation is the primary focus of most microarray experiments, changes in translation can also be readily assayed. This may be accomplished by fluorescently labeling the nucleic acid portion of polysomes, which are typically isolated as lowvelocity nucleoprotein fractions separated on sucrose gradients. Higher-molecular-weight material, representing messages bound by several ribosomes, may be differentially labeled

hr:

1

1

2

with respect to lower-molecular-weight material, representing those messages with few or no bound ribosomes. The resulting hybridization can then be used to assay simultaneously the degree to which each individual mRNA is associated with translation machinery. Tracking these associations provides a means to measure the degree to which the expression of any particular message is likely to be controlled at this level (K. Kuhn and P. Sarnow, pers. comm.). In one interesting variation on this approach, membrane-bound polysomes may also be collected and analyzed. Because membrane-bound polysomes contain messages undergoing cotranslational secretion, it is then possible to quickly identify these gene products with respect to any given experimental condition, whether it be time, cell type, or environment (Diehn et al., 1999).

DNA-Protein Interaction The separation of nucleoprotein components for array analysis is not limited to mRNA bound to polysomes. Indeed, any method that creates a stable or covalent cross-link between the nucleic acid and the protein component may be used. Subsequent separation by immunoprecipitation (UNIT 20.5), filter binding, or other chromatographic procedures may then produce a nucleic acid fraction that can be directly labeled and applied to a microarray representation of the genome. Such genome-wide studies are underway for epitope-tagged DNA binding proteins and for proteins that cross-link enzymatically to DNA as part of their normal

3

4

5

mRNA

fluorescent cDNA

array


Figure 22.1.3 Monitoring levels of transcript abundance. Time zero is the reference probe for each comparison.



function. Obvious applications of this approach include the study of DNA binding proteins (Chapter 12), endonucleases, and chromatinassociated proteins (UNIT 21.3). These methods often produce vanishingly small amounts of nucleic acid source material, often in the subnanogram range. For this reason, there is likely to be much development in the area of linear DNA and RNA amplification. Indeed, amplification technology coupled to microarray analysis will allow the study of single-cell expression patterns (Eberwine, 1996).

www-genome.stanford.edu Stanford Genomic Resources. One-stop shopping for genome-wide expression analysis. This site contains links to several yeast expression databases and research article companions in areas including cell-cycle analysis, sporulation, the diauxic shift, yeast evolution, and yeast clustering. Links to several human expression databases are also available here, including databases relevant to serum starvation, breast cancer expression patterns, and microarray-based CGH.

WHAT ABOUT DATA ANALYSIS?

rana.stanford.edu/software Stanford Genome Analysis Group software download area. This site provides access to several free microarray-analysis tools that run on your PC, written by Mike Eisen. They include ScanAlyze, an advanced image-analysis package, and clustering/data-visualization applications.

In addition to the many ways microarray experiments may be carried out, there are equally many ways to analyze the resulting data. Especially for gene expression, it is likely that genome-wide expression analysis will become a field of its own and that an ever-increasing number of innovative approaches will be developed to address the problem of reverse engineering the mechanisms guiding the observed expression patterns. Several methods for visualization and analysis already exist, and many of the necessary tools are readily available to be downloaded from the Internet. The method known as clustering, whereby similar expression patterns are grouped together, has been particularly successful. (Eisen et al., 1998; Bassett et al., 1999; Brown and Botstein., 1999; Iyer et al., 1999). Its success relies upon the fact that genes which participate in a common process, pathway, or function often share common regulatory mechanisms and this, in turn, results in similar expression profiles. Therefore, the functions of previously uncharacterized genes may be revealed by what other genes cluster with it over a broad range of experiments.

WHERE CAN I GET MORE INFORMATION? The field of microarray technology is evolving rapidly, and therefore the uses and methods of this field will be continuing to change as researchers innovate and disseminate new techniques and ideas. One way to stay connected to this rapid progression is to frequent the various noncommercial web sites listed below. The sites listed contain directions for the fabrication of microarray robots, up-to-date protocol revisions, public access to genome wide expression data, free microarray software tools, forums, and database information.

cmgm.stanford.edu/pbrown/mguide The MGuide. Home of a do-it-yourself guide to building your own microarraying robot, maintained by Patrick Brown’s lab at Stanford University. A step-by-step guide, technical drawings, and parts lists, as well as free software, are available for downloading here. www.nhgri.nih.gov/DIR/LCG/15K/HTML National Human Genome Research Institutes (NHGRI) Microarray Project. This site features database and data handling information, as well as protocols and research projects. web.wi.mit.edu/young/expression Genome-Wide Expression Homepage maintained by Rick Young of the Whitehead Institute. Access to various yeast transcription experiments, protocols, and experiment designs are available. industry.ebi.ac.uk/∼alan/MicroArray Large-Scale Gene Expression and Microarray Links and Resources. A “personal not-forprofit web site” containing literally hundreds of links to microarray articles, databases, and companies, maintained by Alan Robinson, a researcher at the EMBL–European Bioinformatics Institute (EBI).

LITERATURE CITED Alizadeh, A., Eisen, M., Botstein, D., Brown, P.O., and Staudt, L.M. 1998. Probing lymphocyte biology by genomic-scale gene expression analysis. J. Clin. Immunol. 18:373-379.

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Amundson, S.A., Bittner, M., Chen, Y., Trent, J., Meltzer, P., and Fornace, A.J., Jr. 1999. Fluorescent cDNA microarray hybridization reveals complexity and heterogeneity of cellular genotoxic stress responses. Oncogene 18:3666-3672. Bassett, D.E. Jr., Eisen, M.B., and Boguski, M.S. 1999. Gene expression informatics—it’s all in your mine. Nature Genet. 21:51-55. Behr, M.A., Wilson, M.A. Gill, W.P., Salamon, H., Schoolnik, G.K., Rane, S., and Small, P.M. 1999. Comparative genomics of BCG vaccines by whole-genome DNA microarray. Science 284:1520-1523. Brown, P.O. and Botstein, D. 1999. Exploring the new world of the genome with DNA microarrays. Nature Genet. 21:33-37. Cheung, V.G., Gregg, J.P., Gogolin-Ewens, K.J., Bandong, J., Stanley, C.A., Baker, L., Higgins, M.J., Nowak, N.J., Shows, T.B., Ewens, W.J., Nelson, S.F., and Spielman, R.S. 1998. Linkagedisequilibrium mapping without genotyping. Nature Genet. 18:225-230. Cho, R.J., Campbell, M.J., Winzeler, E.A., Steinmetz, L., Conway, A., Wodicka, L., Wolfsberg, T.G., Gabrielian, A.E., Landsman, D., Lockhart, D.J., and Davis, R.W. 1998. A genome-wide transcriptional analysis of the mitotic cell cycle. Mol. Cell 2:65-73. Chu, S., DeRisi, J., Eisen, M., Mulholland, J., Botstein, D., Brown, P.O., and Herskowitz, I. 1998. The transcriptional program of sporulation in budding yeast. Science 282:699-705. DeRisi, J.L., Iyer, V.R., and Brown, P.O. 1997. Exploring the metabolic and genetic control of gene expression on a genomic scale. Science 278:680686. Diehn, M., Eisen, M., Brown, P., and Botstein, D. 1999. Large-scale identification of membraneassociated gene products using DNA microarrays, submitted. Eberwine, J. 1996. Amplification of mRNA populations using aRNA generated from immobilized oligo(dT)-T7 primed cDNA. Biotechniques 20:584-591. Eisen, M.B., Spellman, P.T., Brown, P.O., and Botstein, D. 1998. Cluster analysis and display of genome-wide expression patterns. Proc. Natl. Acad. Sci. U.S.A. 95:14863-14868. Ferea, T.L., Botstein, D., Brown, P.O., and Rosenzweig, R.F. 1999. Systematic changes in gene expression patterns following adaptive evolution in yeast. Proc. Natl. Acad. Sci. U.S.A. 96:97219726. Fodor, S.P., Read, J.L., Pirrung, M.C., Stryer, L., Lu, A.T., and Solas, D. 1991. Light-directed, spatially addressable parallel chemical synthesis. Science 251:767-773.


Hacia, J.G., Brody, L.C., Chee, M.S., Fodor, S.P., and Collins, F.S. 1996. Detection of heterozygous mutations in BRCA1 using high density oligonucleotide arrays and two-colour fluorescence analysis. Nature Genet. 14:441-447.

Hayward, R., DeRisi, J., Alfadhli, S., Kaslow, D., Brown, P., and Rathod, P. 1999. Shot-gun DNA microarrays and stage specific gene expression in Plasmodium falciparum, submitted. Iyer, V.R., Eisen, M.B., Ross, D.T., Schuler, G.T., Moore, J.C., Lee, F., Trent, J.M., Staudt, L.M., Hudson, J., Jr., Boguski, M.S., Lashkari, D., Shalon, D., Botstein, D., and Brown, P.O. 1999. The transcriptional program in the response of human fibroblasts to serum. Science 283:83-87. Kallioniemi, A., Kallioniemi, O.P., Sudar, D., Rutovitz, D., Gray, J.W., Waldman, F., and Pinkel, D. 1992. Comparative genomic hybridization for molecular cytogenetic analysis of solid tumors. Science 258:818-821. McAllister, L., Penland, L., and Brown, P.O. 1998. Enrichment for loci identical-by-descent between pairs of mouse or human genomes by genomic mismatch scanning. Genomics 47:7-11. Nelson, S.F., McCusker, J.H., Sander, M.A., Kee, Y., Modrich, P., and Brown, P.O. 1993. Genomic mismatch scanning: A new approach to genetic linkage mapping. Nature Genet. 4:11-18. Perou, C.M., Jeffrey, S.S., van de Rijn, M., Rees, C.A., Eisen, M.B., Ross, D.T., Pergamenschikov, A., Williams, C.F., Zhu, S.X., Lee, J.C., Lashkari, D., Shalon, D., Brown, P.O., and Botstein, D. 1999. Distinctive gene expression patterns in human mammary epithelial cells and breast cancers. Proc. Natl. Acad. Sci. U.S.A. 96:9212-9217. Pinkel, D., Segraves, R., Sudar, D., Clark, S., Poole, I., Kowbel, D., Collins, C., Kuo, W.L., Chen, C., Zhai, Y., Dairkee, S.H., Ljung, B.M., Gray, J.W., and Albertson, D.G. 1998. High resolution analysis of DNA copy number variation using comparative genomic hybridization to microarrays. Nature Genet. 20:207-211. Pollack, J.R., Perou, C.M., Alizadeh, A.A., Eisen, M.B., Pergamenschikov, A., Williams, C.F., Jeffrey, S.S., Botstein, D., and Brown, P.O. 1999. Genome-wide analysis of DNA copy-number changes using cDNA microarrays. Nature Genet. 23:41-46. Solinas-Toldo, S., Lampel, S., Stilgenbauer, S., Nickolenko, J., Benner, A., Döhner, H., Cremer, T., and Lichter, P. 1997. Matrix-based comparative genomic hybridization: Biochips to screen for genomic imbalances. Genes Chrom. Cancer 20:399-407. Spellman, P.T., Sherlock, G., Zhang, M.Q., Iyer, V.R., Anders, K., Eisen, M.B., Brown, P.O., Botstein, D., and Futcher, B. 1998. Comprehensive identification of cell cycle–regulated genes of the yeast Saccharomyces cerevisiae by microarray hybridization. Mol. Biol. Cell 9:3273-3297. Trent, J.M., Bittner, M., Zhang, J., Wiltshire, R., Ray, M., Su, Y., Gracia, E., Meltzer, P., De Risi, J., Penland, L., and Brown, P. 1997. Use of microgenomic technology for analysis of alterations in DNA copy number and gene expression in malignant melanoma. Clin. Exp. Immunol. 107(Suppl 1):33-40.



Wilson, M., DeRisi, J., Kirstensen, H., Imboden, P., Rane, S., Brown, P., and Schoolnik, G. 1999. Exploring drug-induced alterations in gene expression in Mycobacterium tuberculosis by microarray hybridization. Proc. Natl. Acad. Sci. U.S.A. (in press).

Contributed by Joseph DeRisi University of California, San Francisco San Francisco, California

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Preparation of mRNA for Expression Monitoring

UNIT 22.2

The ability to construct comprehensive gene expression profiles comprising hundreds to thousands of genes whose RNA levels are monitored simultaneously represents an exciting new capability in molecular biology. This is accomplished by hybridizing mRNA, which has been quantitatively amplified and labeled with biotin, to DNA chips that display thousands of oligonucleotides complementary to the mRNAs of interest. An overview of the entire process is shown schematically in Figure 22.2.1. The Strategic Planning section outlines considerations in starting with poly(A)+ versus total RNA, and also discusses oligonucleotide selection for chip design. The Basic Protocol outlines RNA amplification and labeling, which entails cDNA synthesis followed by in vitro transcription (IVT), and hybridization of the resulting biotinylated antisense RNA to the chip. An Alternate Protocol is included for purification of cDNA and in vitro transcription products on carboxy-coated magnetic beads, making the amplification reaction amenable to automation. Support Protocols 1 and 2 provide methods for quantifying the cDNA product and for producing in vitro transcripts of control genes. CAUTION: Diethylpyrocarbonate (DEPC) is a suspected carcinogen and should be handled with care. STRATEGIC PLANNING As this is an RNA-based procedure, care should be taken at every step to avoid contamination of reagents and materials with RNases (UNIT 4.1). New plasticware can be used without decontamination processes if kept free of dust and handled exclusively with gloved hands. Filtered pipet tips should be used routinely to avoid contamination with the micropipettor. Any reagent that needs to be made in the laboratory, or that is not supplied RNase-free by the manufacturer, should be treated with DEPC (UNIT 4.1). The amplification procedure is absolutely dependent on clean, intact starting RNA. For cultured cells, the authors use a protocol of lysis in guanidine followed by resin-based purification (such as the Qiagen RNeasy kit). The authors’ results using one-step protocols (in which guanidine and phenol are combined) have been variable, presumably because inhibitors of the amplification reaction enzymes can be carried along. Tissues are immediately snap-frozen in liquid nitrogen, then ground to a powder in a dry ice–embedded mortar and pestle. RNA is then extracted in guanidine reagent in a Polytron mixer, followed by extraction with phenol (e.g., Promega RNAgents kit, Ambion Totally RNA kit). The mRNA amplification can be performed starting with either poly(A)+ or total RNA. Although poly(A)+ RNA gives higher sensitivity because it largely eliminates mispriming on rRNA during the cDNA reaction, the ability to use total RNA makes it possible to analyze biological systems in which cells or tissues are limited. A key modification in using total RNA is elevation of the first-strand cDNA reaction temperature from 37° to 50°C. Although this will decrease cDNA yield somewhat, it dramatically decreases mispriming on rRNA. The protocol outlined here will preferentially amplify the 3′ end of long mRNAs, owing to its dependence on priming with oligo(dT). It is therefore best to use either custom or commercially available chips that display probes toward the 3′ end of the mRNA. In general, the authors restrict probe selection to the last 600 bases of coding sequence unless the 3′-untranslated region (3′-UT) is >800 bases, in which case some untranslated sequence is also included. Contributed by Michael C. Byrne, Maryann Z. Whitley, and Maximillian T. Follettie Current Protocols in Molecular Biology (2000) 22.2.1-22.2.13 Copyright © 2000 by John Wiley & Sons, Inc.

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tissue

AAAA

GIT poly(A)+ RNA

oligo(dT)

T7T24 annealing

or cells TTTT T7 AAAA cDNA synthesis

biotinylated antisense RNA B B T7 T7

ds cDNA

B B

B B

B

SPRI purification T7 T7

T7 RNA polymerase bio-UTP bio-CTP

SPRI purification

fragment (Mg2+, heat)

50- to 100-base RNA fragments hybridize to chip

wash

stain

scan

(PE-streptavidin)

analyze pattern

Figure 22.2.1 Chip analysis overview. Abbreviations: GIT, guanidine isothiocyanate, PE, phycoerythrin; SPRI, solid-phase reversible immobilization.

BASIC PROTOCOL

AMPLIFICATION OF mRNA FOR EXPRESSION MONITORING AND HYBRIDIZATION TO OLIGONUCLEOTIDE ARRAY CHIPS The Basic Protocol details the quantitative amplification and biotin labeling of mRNA to produce antisense RNA for the purpose of gene expression monitoring. Starting RNA is first converted to double-stranded cDNA using a primer containing a T7 RNA polymerase site, so that amplified and labeled RNA can be produced directly in an in vitro transcription reaction. The resulting RNA is hybridized to a DNA chip.


Materials SuperScript cDNA kit (Life Technologies), including: 5× First Strand Buffer 200 U/µl SuperScript II reverse transcriptase 5× Second Strand Buffer 10 mM dNTPs 10 U/µl E. coli ligase 2 U/µl E. coli RNase H 10 U/µl E. coli DNA polymerase 5 U/µl T4 DNA polymerase T7T24 primer: 5′-GGCCAGTGAATTGTAATACGACTCACTATAGGGAGGCGG TTTTTTTTTTTTTTTTTTTTTTTT-3′ (HPLC purification is recommended) RNase inhibitor (Life Technologies or Ambion) RNase-free H2O (see UNIT 4.1 for DEPC treatment of solutions; prepare from glass-distilled H2O) Sample RNA: poly(A)+ or total RNA Sense control transcript pool (see Support Protocol 1)



25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol (molecular biology grade; UNIT 2.1A) 7.5 M ammonium acetate (APPENDIX 2) Absolute ethanol 70% (v/v) ethanol in RNase-free H2O, prechilled to −20°C 10× transcription buffer (Ambion) 10× rNTP mix (see recipe) 100 mM dithiothreitol (DTT) 10 mM Bio-11-CTP and Bio-11-UTP (Enzo Diagnostics) 2500 U/µl T7 RNA polymerase (Epicentre) RNeasy mini columns with RLT and RPE buffers and collection tubes (Qiagen) 5× fragmentation buffer (see recipe) 20× SSPE (Bio-Whittaker) 0.5% (v/v) Triton X-100 (molecular biology grade; Sigma) in RNase-free H2O 10 mg/ml herring sperm DNA (Promega) 500 pM Bio948 (see recipe) 20× antisense control transcript pool (see Support Protocol 1) 6× SSPET: 6× SSPE containing 0.005% (v/v) Triton X-100 Thermocycler (e.g., Perkin-Elmer 9600 PCR machine with heated lid) 0.1- to 10-µl filtered micropipet tips (Continental) Lyophilizer Small, thin-walled PCR tubes GeneChip (Affymetrix) 1- to 200-µl filtered gel-loading micropipet tips (Fisher) Rotisserie-type rotator (Appropriate Technical Resources) 50°C oven Additional reagents and equipment for quantitation of cDNA (see Support Protocol 2), for quantitation of DNA by spectrophotometry (APPENDIX 3D), and for washing, staining, and scanning the GeneChip (see manufacturer’s instructions) NOTE: Many buffers and enzymes are supplied with the SuperScript II cDNA kit. Kit enzymes that are limiting may be ordered separately from Life Technologies. NOTE: All temperature-controlled reactions are performed in an appropriate thermocycler. Perform first-strand synthesis 1. Set up a linked program on a thermocycler as follows: 10 min 65 min 150 min indefinitely

70°C 37°C (or 50°C for total RNA) 15.8°C 4°C (hold).

2. Prepare 10 µl first-strand reagent cocktail for each sample RNA by combining the following reagents, using filtered pipet tips: 4 µl 5× First Strand Buffer 200 pmol T7T24 primer 1 µl RNase inhibitor 1 µl 200 U/µl SuperScript II reverse transcriptase RNase-free H2O to 10 µl.

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3. Combine each sample RNA with sense control transcript pool. Use 5 µl sense control transcript pool per 1 µg sample poly(A)+ RNA or 1 µl control pool per 10 to 20 µg total RNA. Lyophilize to reduce the volume per tube to 8000 × g, room temperature. Qiagen RNeasy columns work well for removal of unincorporated nucleotides. It is very important to remove the nucleotides in order to accurately quantitate the IVT RNA. This protocol follows the manufacturer’s protocol with the following changes: no 2-mercaptoethanol is used in the RLT buffer, and final elution is done twice using 50 ìl RNase-free H2O each time.

23. Transfer column to a new collection tube. Add 500 µl RPE buffer. The RPE working solution is prepared as described in the kit directions; the supplied stock is diluted with 4 vol absolute ethanol.

24. Centrifuge 15 sec at >8000 × g, room temperature. 25. Discard flowthrough and replace column on the same collection tube. Add 500 µl RPE buffer and microcentrifuge 2 min at maximum speed. 26. Transfer column to a new collection tube. Add 50 µl RNase-free H2O and centrifuge 1 min at >8000 × g. Repeat elution step, pooling the second eluate with the first. 27. Quantitate RNA yield spectrophotometrically at 260 nm (APPENDIX 3D). Fragment RNA 28. Bring volume of 10 µg IVT RNA to 24 µl with RNase-free H2O. 29. Add 6 µl of 5× fragmentation buffer. Mix carefully and incubate 35 min at 95°C in the thermocycler. Allow to cool to room temperature. Fragmented RNA can be stored up to 1 year at −80°C. Thaw at 37°C for 5 min before preparing hybridization solution.

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Hybridize RNA to chip 30. For each reaction, prepare 170 µl hybridization master mix as follows: 51 µl 20× SSPE 1.7 µl 0.5% Triton X-100 1.7 µl 10 mg/ml herring sperm DNA 20 µl 500 pM Bio948 10 µl 20× antisense control transcript pool 85.6 µl RNase-free H2O. 31. Add 170 µl hybridization master mix to 30 µl of each fragmented IVT RNA. 32. Heat to 99°C for 10 min in the thermocycler, then move to 37°C for ≥5 min. 33. Microcentrifuge 5 min at maximum speed. 34. Insert a filtered micropipet tip into the upper septum of a GeneChip to provide a vent. Fill the chip from the bottom septum with the 200 µl hybridization solution using a filtered gel-loading micropipet tip. Remove the vent and cover both septa with transparent tape. Use one GeneChip for each IVT RNA.

35. Incubate in a 40°C oven overnight (16 to 18 hr) on a rotisserie-type rotator running at ∼60 rpm. 36. Transfer the chip to a 50°C oven and continue rotating the chip for exactly one hour. 37. Remove chip from the oven and insert a filtered micropipet tip into the upper septum to vent the chip. 38. Using a micropipettor with plunger fully depressed, insert a 200-µl gel-loading pipet tip into the lower septum. Holding the chip vertically, slowly draw out all of the hybridization solution and store it in a microcentrifuge tube at −20°C. 39. Fill the chip with 6× SSPET. 40. Wash and stain the chip with phycoerythrin according to the manufacturer’s protocols. Scan the chip as soon as possible. If the chip cannot be washed right away, seal the two septa with transparent tape and store at 4°C for up to a few hours. Once the chips have been stained with phycoerythrin they should be kept wrapped in foil to avoid photobleaching. Although it is recommended that they be scanned as soon as possible, if necessary, they can be stored at 4°C, wrapped in foil, for several hours before scanning. SUPPORT PROTOCOL 1

IN VITRO TRANSCRIPTION OF CONTROL GENES AND PREPARATION OF TRANSCRIPT POOLS It is essential to include labeled antisense control RNAs of known concentration in each hybridization reaction to normalize chip-to-chip variation and to allow construction of a standard curve for converting hybridization intensity to mRNA frequency. It is also very helpful to add unlabeled sense transcripts to monitor performance of the amplification protocol. This protocol describes the preparation of these two control pools.




Additional Materials (also see Basic Protocol) Plasmids (Table 22.2.1; ATCC #87482 to 87490) 2500 U/µl T3 RNA polymerase (Enzo Diagnostics) 25 mM 4rNTP mix: 25 mM each rGTP, rCTP, rATP, and UTP (Ultrapure; Pharmacia Biotech) in RNase-free H2O Additional reagents and equipment for purifying IVT products (see Alternate Protocol) Prepare sense and antisense transcripts 1. Prepare linearized plasmid templates according to Table 22.2.1. See UNIT 3.1 and manufacturer’s instructions for additional information on reaction conditions.

2. Purify plasmid DNA by phenol/chloroform extraction and ethanol precipitation (see Basic Protocol, steps 12 to 16). Resuspend at ∼0.1 mg/ml in RNase-free H2O and quantitate DNA (see Support Protocol 2). 3. For labeled antisense transcripts, prepare four tubes containing a master mix that includes all components except plasmid DNA: 6 µl 10× transcription buffer 6 µl 10× rNTP mix 3 µl 100 mM DTT 2.4 µl 10 mM Bio-11-UTP 2.4 µl 10 mM Bio-11-CTP 2 µl RNase inhibitor 2 µl 2500 U/µl T7 RNA polymerase RNase-free H2O to 60 µl. The 60-ìl volume must include the 100 ng DNA that will be added in step 5. The master mix can be prepared in advance and held on ice for up to 3 hr, but should be warmed to room temperature before adding to plasmid DNA to avoid precipitates.

Table 22.2.1

Preparation of Plasmid Template Controls

Namea

ATCC #

pGIBS-LYSb pGIBS-PHEb pGIBS-THRb pGIBS-TRPb pGIKS-BioB pGIKS-BioC pGIKS-BioD pGIKS-CRE

87482 87483 87484 87485 87487 87488 87489 87490

Sense RNA Antisense RNA Transcript size (kb) Linearize Polymerize Linearize Polymerize with with with with 1.0 1.3 2.0 2.5 1.1 0.8 0.7 1.0

NotI NotI NotI NotI

T3 T3 T3 T3 XhoI XhoI XhoI XhoI

T7 T7 T7 T7

aAbbreviations: BioB, BioC, and BioD are cloned fragments from the E. coli bioB, bioC, and BioD genes,

respectively. LYS, PHE, THR, and TRP are fragments from the Bacillus subtilis lysA, pheA, thrBC, and trpEDCF genes, respectively. CRE is a fragment from the Cre recombinase derived from E. coli bacteriophage P1. PGIBS and pGIKS are derived from the Bluescript KS II vector (Stratagene). bpGIBS-LYS, -PHE, -THR, -TRP, and DAP contain a 40-nucleotide synthetic poly(A) tract at the 3′ end of the respective genomic fragments derived from B. subtilis. Sense IVT transcripts derived from NotI-linearized plasmid templates will contain the artificial poly(A) tail. Plasmids linearized with BamHI prior to T3 IVT will generate sense transcripts without the synthetic poly(A) tract.

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4. For unlabeled sense transcripts, prepare four tubes of master mix as in step 3, but substitute 25 mM 4rNTP mix for the 10× rNTP mix, eliminate the biotinylated nucleotides, and substitute T3 for T7 RNA polymerase. 5. Add 100 ng of each linearized plasmid DNA to a separate tube and incubate at 37°C for 8 hr to overnight. The transcripts can be stored for up to 48 hr at −80°C, if desired, before purification.

6. Purify IVT products as described (see Alternate Protocol). 7. Quantitate carefully by absorbance at 260 nm (APPENDIX 3D). 8. Dilute unlabeled sense transcripts to 200 nM stock solutions in RNase-free H2O for long-term storage (up to 1 year) at −80°C. 9. Fragment biotin-labeled antisense transcripts for standard chip controls (BioB, BioC, BioD, CRE) as described (see Basic Protocol, steps 18 and 19). 10. Dilute fragmented antisense transcripts to 20 nM stock solutions in 6× SSPET with 0.1 mg/ml herring sperm DNA for long-term storage (up to 1 year) at −80°C. Prepare control transcript pools 11. Prepare 20× antisense control transcript pool by combining the following transcripts in 6× SSPET with 0.1 mg/ml herring sperm DNA. Store up to 6 months at −80°C. 30 pM fragmented BioB transcript 100 pM fragmented BioC transcript 500 pM fragmented BioD transcript 2 nM fragmented CRE transcript. 12. Prepare sense control transcript pool by combining the following transcripts in RNase-free H2O. Store up to 6 months at −80°C in aliquots of 50 to 100 µl. 10 pM LYS transcript 30 pM PHE transcript 90 pM THR transcript 180 pM TRP transcript. ALTERNATE PROTOCOL

SOLID-PHASE REVERSIBLE IMMOBILIZATION (SPRI) PURIFICATION OF cDNA AND IVT PRODUCTS Because of the labor-intensive nature of the mRNA amplification protocol, the procedure has been adapted for automation. Although the assembly of the reactions is easily converted to liquid handling robotics, purification of final products should be free of organic solvents and the requirement for centrifuges. A magnetic bead–based purification protocol of cDNA and IVT products has been developed to meet this requirement. This protocol was adapted from a protocol for the automated preparation of DNA sequencing templates (DeAngelis et al., 1995). It is equivalent to the purification methods described in the Basic Protocol (steps 12 to 17 or steps 20 to 27).


Materials Carboxy-coated magnetic beads (PerSeptive BioSystems for cDNA purification; Bangs Laboratories for IVT purification) 0.5 M EDTA (APPENDIX 2) Sample to be purified: cDNA (see Basic Protocol, step 11) or IVT RNA (see Basic Protocol, step 19, or see Support Protocol 1, step 6) 2.5 M NaCl/20% (w/v) PEG 8000 (molecular biology grade; RNase free)



70% (v/v) ethanol in RNase-free H2O 10 mM Tris acetate, pH 7.8 (RNase free) Magnetic stand (CPG) Additional reagents and equipment for determining concentration of cDNA (see Support Protocol 2) or RNA (APPENDIX 3D) Purification of cDNA 1a. Aliquot 10 µl PerSeptive carboxy-coated magnetic beads per 150-µl cDNA reaction, dispensing the total volume of beads into a single microcentrifuge tube. 2a. Place the tube on a magnetic stand and allow the beads to separate to the side of the tube. Carefully remove supernatant with a micropipet. 3a. Add 0.5 M EDTA equal to the starting volume and resuspend the beads by gentle vortexing or agitation. Replace tube on magnetic stand, wait for beads to separate, and remove supernatant. Repeat this washing procedure two more times. 4a. Resuspend beads in 0.5 M EDTA equal to the starting volume. 5a. To each tube of cDNA, add 150 µl of 2.5 M NaCl/20% PEG 8000 and 10 µl beads and mix by gentle vortexing or agitation. Incubate 10 min at room temperature. 6a. Place tubes on magnetic stand and allow beads to separate to the side of the tube (∼2 min for the original separation, faster for the washes). 7a. Draw off the supernatant, then wash the beads twice with 150 µl of 70% ethanol. Remove as much of the final ethanol wash as possible and allow to air dry for 2 min. 8a. Elute RNA by adding 25 µl of 10 mM Tris acetate, pH 7.8, and incubating 5 min at room temperature. 9a. Place tube on magnetic stand and save supernatant. 10a. Determine cDNA concentration by PicoGreen fluorescence (see Support Protocol 2). Purification of IVT RNA 1b. Aliquot 20 µl Bangs Laboratories carboxy-coated magnetic beads per 60-µl IVT reaction, dispensing the total volume of beads into a single microcentrifuge tube. 2b. Place the tube on a magnetic stand and allow the beads to separate to the side of the tube. Carefully remove supernatant with a micropipet. 3b. Add 0.5 M EDTA equal to the starting volume and resuspend the beads by gentle vortexing or agitation. Replace tube on magnetic stand, wait for beads to separate, and remove supernatant. Repeat this washing procedure two more times. 4b. Resuspend beads in 1.25 M NaCl/10% PEG equal to the starting volume. 5b. To each tube of IVT RNA, add 60 µl of 2.5 M NaCl/20% PEG 8000 and 20 µl beads and mix by gentle vortexing or agitation. Incubate 10 min at room temperature. 6b. Place tubes on magnetic stand and allow beads to separate to the side of the tube (∼2 min for the original separation, faster for the washes). 7b. Draw off the supernatant, then wash the beads twice with 150 µl of 70% ethanol. Remove as much of the final ethanol wash as possible and allow to air dry for 3 min. Nucleic Acid Arrays


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8b. Elute RNA by adding 25 µl of 10 mM Tris acetate, pH 7.8, and incubating 5 min at room temperature. 9b. Place tube on magnetic stand and save supernatant. 10b. Determine RNA concentration by absorbance at 260 nm (APPENDIX 3D). SUPPORT PROTOCOL 2

QUANTITATION OF cDNA If IVT yield is low, it can be helpful to determine the amount of cDNA that is being produced. For optimal IVT yield, do not exceed 100 ng cDNA in the standard reaction. Quantitation of cDNA can be performed with Molecular Probes PicoGreen reagent. Materials PicoGreen dsDNA Quantitation Kit (Molecular Probes), including 100 ng/µl standard DNA stock solution 20× TE buffer PicoGreen reagent cDNA to be quantitated (see Basic Protocol and Alternate Protocol) Black-walled 96-well plate (Corning) Fluorimager (Molecular Dynamics, model FSI) 1. Prepare 1 ml of 2 µg/ml diluted standard DNA by diluting 20 µl of 100 ng/µl standard DNA stock solution in 980 µl of 1× TE buffer. 2. Dilute PicoGreen reagent 1:200 (v/v) with 1× TE buffer. Prepare enough diluted reagent so that 100 µl can be placed in each well. 3. Pipet 100, 50, 20, 10, 5, 2, and 0 µl dilute standard DNA into seven wells in a black-walled 96-well plate. Bring each to a total of 100 µl with 1× TE buffer. The amounts of DNA in the standard wells are 200, 100, 40, 20, 10, 4, and 0 ng, respectively.

4. Pipet 2 µl cDNA to be quantitated into additional wells, as needed. 5. Add 100 µl diluted PicoGreen reagent to each well. 6. Read fluorescence of each well using a fluorimager. 7. Generate a volume report on each well according to manufacturer’s instructions. Use volume number versus ng per well to generate a standard curve. 8. Calculate the concentration of cDNA in test wells. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.


Bio948, 500 pM 500 pM biotinylated control oligonucleotide Bio948 (5′-GTCAAGATGCTACCGTTCAG-3′) 6× SSPE (Bio-Whittaker) 0.1 mg/ml herring sperm DNA (Promega) 0.005% (v/v) Triton X-100 (molecular biology grade; Sigma) Prepare in RNase-free H2O Store up to 1 year at −20°C



Fragmentation buffer, 5× Dissolve 6.06 g Tris base (Sigma; molecular biology grade) in 175 ml RNase-free H2O. Adjust pH to 8.1 with glacial acetic acid. Add 12.3 g potassium acetate (from an unopened or dedicated bottle) and 8.04 g magnesium acetate (Sigma; molecular biology grade) and adjust volume to 250 ml (final pH ∼8.4). Filter sterilize with a 0.2-µm filter. Store up to 6 months at −20°C. rNTP mix, 10× 30 mM rGTP 15 mM rATP 12 mM rCTP 12 mM UTP Prepare using RNase-free H2O and Ultrapure reagents from Pharmacia Biotech. Store up to 3 months at −20°C in small aliquots (e.g., 50 to 100 µl) to prevent multiple freeze/thaw cycles. COMMENTARY Background Information The availability of thousands of gene sequences through high-throughput sequencing efforts, including the Human Genome Project, created an immediate need for highly parallel methods for assessing RNA levels of many genes simultaneously. One approach that is being practiced by an ever-expanding number of laboratories is the hybridization of labeled cellular RNA or cDNA to microarrays displaying DNA probes complementary to the genes of interest. Both cDNA (Schena et al., 1995) and oligonucleotide (Lockhart et al., 1996) probe arrays are used, and each type has its relative merits and shortcomings. The strength of cDNA arrays lies in the ability to display probes on the array prior to obtaining any sequence information, allowing the possibility of novel gene discovery. The disadvantages are that (1) quantitation is possible in a relative sense (i.e., fold-induction) but not an absolute sense (i.e., number of mRNA molecules per million), and (2) it is difficult to discern expression of different members of multigene families due to cross-hybridization. Construction of oligonucleotide arrays, on the other hand, requires that DNA sequence information be available for every gene to be monitored, precluding use of the arrays for novel gene discovery. This limitation is partially mitigated, however, by the recent availability of chips monitoring tens of thousands of expressed sequence tags (ESTs) with no known function. Advantages of oligonucleotide arrays include (1) the possibility of absolute quantitation, by virtue of averaging across multiple probes for every gene being monitored, and (2) the ability to inde-

pendently follow the expression of closely related gene family members through judicious probe selection.

Critical Parameters The most critical parameter affecting the success of the protocol is the quality of the starting RNA. It is absolutely essential to start with RNA that is intact and free from inhibitors of the enzymes in the amplification procedure. For sample comparisons, e.g., control versus treated sample, it is necessary that the RNA samples be of similar quality and, preferably, that samples are hybridized to arrays within the same lot to minimize chip-to-chip variation.

Troubleshooting The problems that can arise during expression monitoring procedures are detailed in Table 22.2.2.

Anticipated Results The yield of amplified in vitro transcription (IVT) product reflects the quality and quantity of the starting RNA and, in general, is an indicator of the success of the subsequent chip hybridization. A good amplification reaction from 1 µg poly(A)+ or 10 µg total RNA can yield 50 to 60 µg IVT product. Successful hybridizations can be obtained with lesser yields, but yields of ≤10 µg, when the quantity of starting RNA was not limiting, are generally predictive of nonoptimal IVT product.

Time Considerations The entire protocol is generally carried out over a three-day period, starting with purified

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Table 22.2.2

Troubleshooting Guide for Expression Monitoring

Problem

Possible cause

Little or no cDNA yield

Beginning RNA quality was poor

Little or no IVT yield

Blank or low signal on GeneChip

Action

Visualize size range of starting RNA on a gel. There should be a smear extending well above 5 kb, and not much 7 mg/ml) anchored primer (2 µg/µl) DEPC-treated H2O

For Cy5 labeling 150 to 200 µg 1 µl up to 17 µl

For Cy3 labeling 50 to 80 µg 1 µl up to 17 µl.

19b. If using an oligo d(T)12-18 primer: Anneal the primer to the RNA in the following 17-µl reaction (use a 0.2-ml thin-wall PCR tube so that incubations can be carried out on a thermal cycler): Component total RNA (>7 mg/ml) dT(12-18) primer (1 µg/µl) DEPC-treated H2O

For Cy5 labeling 150 to 200 µg 1 µl up to 17 µl

For Cy3 labeling 50 to 80 µg 1 µl up to 17 µl.

The incorporation rate for Cy5-dUTP is less than that of Cy3-dUTP, so more RNA is labeled to achieve a more equivalent signal from each species.

20. Heat to 65°C for 10 min and cool on ice for 2 min. 21. Prepare a master mix with the following components (total volume, 23 µl). 8 µl 5× first strand buffer 4 µl 10× low-T dNTP mix 4 µl 1 mM Cy5 or Cy3 dUTP 4 µl 0.1 M DTT 1 µl 30 U/µl RNasin 2 µl 200 U/µl Superscript II. Profiling Human Gene Expression with cDNA Microarrays

Superscript polymerase is very sensitive to denaturation at air/liquid interfaces, so be very careful to suppress foaming in all handling of this reaction.



22. Add 23 µl of reaction mixture containing either Cy5-dUTP or Cy3-dUTP nucleotides to each sample, mix well by pipetting, and centrifuge briefly to concentrate in the bottom of the tube. 23. Incubate 30 min at 42°C, then add 2 µl more Superscript II. Make sure the enzyme is well mixed in the reaction volume and incubate for 30 to 60 min at 42°C. 24. Add 5 µl of 0.5 M EDTA, pH 8.0, to stop the reaction. Be sure to stop the reaction with EDTA before adding NaOH, since nucleic acids precipitate in alkaline magnesium solutions.

25. Add 10 µl of 1 N NaOH, then incubate for 60 min at 65°C to hydrolyze residual RNA. Cool to room temperature. The purity of the sodium hydroxide solution used in this step is crucial. Slight contamination or long storage in a glass vessel can produce a solution that will degrade the Cy5 dye molecule, turning the solution yellow. Some researchers achieve better results by reducing the time of hydrolysis to 30 min.

26. Neutralize by adding 25 µl of 1 M Tris⋅Cl, pH 7.5. 27. Desalt the labeled cDNA by adding 400 µl of TE buffer, pH 7.5, and 20 µg of human C0t-1 DNA in a Microcon YM-100 cartridge. Pipet to mix, spin for 10 min at 500 × g, room temperature. 28. Wash again by adding 200 µl TE buffer, pH 7.5, and concentrating in a Microcon YM-100 cartridge to about 20 to 30 µl (∼8 to 10 min at 500 × g). Alternatively, a smaller-pore Microcon YM-30 can be used to speed the concentration step. In this case, centrifuge the first wash for ∼4.5 min at 16,000 × g and the second (200-ìl wash) for ∼2.5 min at 16,000 × g.

A B

600 bp

Figure 22.3.1 Fluorescence scan of a 2 Cy5 labeled cDNAs electrophoresed on a 2% agarose gel.

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29. Recover by inverting the concentrator over a clean collection tube and spinning for 3 min at 500 × g, room temperature. In some cases, the Cy5-labeled cDNA will form a gelatinous blue precipitate that is recovered in the concentrated volume. The presence of this material signals the presence of contaminants. The more extreme the contamination, the greater the fraction of cDNA that will be captured in this gel. Even if heat solubilized, this material tends to produce uniform nonspecific binding to the DNA targets. When concentrating by centrifugal filtration, the time required to achieve the desired final volume is variable. Overly long spins can remove nearly all the water from the solution being filtered. When fluor-tagged nucleic acids are concentrated onto the filter in this fashion, they are very hard to remove, so it is necessary to approach the desired volume by conservative approximations of the required spin times. If control of volumes proves difficult, the final concentration can be achieved by evaporating liquid in the Speedvac evaporator. Vacuum evaporation, if not carried to dryness, does not degrade the performance of the labeled cDNA.

30. Take a 2- to 3-µl aliquot of the Cy5-labeled cDNA for analysis, leaving 18 to 28 µl for hybridization. 31. Run this probe on a 2% agarose gel (e.g. UNIT 2.5A) in TAE buffer. For maximal sensitivity when running samples on a gel for fluor analysis, use loading buffer with minimal dye and do not add ethidium bromide to the gel or running buffer.

32. Scan the gel on a Molecular Dynamics Storm fluorescence scanner (setting: red fluorescence, 200-µm resolution, 1000 V on PMT). Successful labeling produces a dense smear of probe from 400 bp to >1000 bp, with little pile-up of low-molecular-weight transcripts (as in Fig. 22.3.1, Lane A). Weak labeling and significant levels of low-molecular-weight material indicates a poor labeling (as in Fig. 22.3.1, Lane B). A fraction of the observed low-molecular-weight material is unincorporated fluor nucleotide. BASIC PROTOCOL 3

Profiling Human Gene Expression with cDNA Microarrays

HYBRIDIZATION AND DATA EXTRACTION This protocol describes the conditions for hybridizing the fluor-tagged cDNA representations of the mRNA pools of samples (see Basic Protocol 2) to the EST PCR products immobilized on the glass microarrays (see Basic Protocol 1). The format for these hybridizations is simultaneous hybridization of both labeled species to a single microarray. The amount of immobilized cDNA on the slide is in excess of the amount of labeled sample that can hybridize to it, so that the amounts of labeled cDNA that do hybridize to any given immobilized cDNA are proportional to their original relative abundance in the cellular message pool. By measuring the ratio of fluorescent intensities at each immobilized cDNA spot, one obtains a measure of relative levels of messages in one sample pool versus the other. If a number of samples are measured against a common reference, the relative amounts of the messages across all of the samples can be compared for similarities and differences. Materials Glass microarrays (see Basic Protocol 1) Cy3- and Cy5-labeled cDNAs (see Basic Protocol 2) DEPC-treated water (UNIT 4.1) 8 mg/ml poly(dA)40-60 (Amersham Pharmacia Biotech) 4 mg/ml yeast tRNA (see recipe) 10 mg/ml human C0t-1 DNA (see recipe) 20× SSC (APPENDIX 2)



50× Denhardt’s solution (APPENDIX 2) 10% SDS 0.5× SSC/0.01% SDS wash buffer (see recipe) 0.06× SSC wash buffer (see recipe) 0.2-ml thin-wall PCR tubes Thermal cycler 24-mm × 50-mm glass cover slips Microarray hybridization chamber 65°C water bath Microarray scanner Image analysis software NOTE: Use RNase-free water (e.g., DEPC-treated water, UNIT 4.1) to make up all solutions, unless indicated otherwise. Hybridize fluorescent cDNA to slide 1. Determine the volume of hybridization solution required. The rule of thumb is to use 0.033 ìl for each mm2 of slide surface area covered by the coverslip used to cover the array. An array covered by a 24-mm by 50-mm coverslip will require 40 ìl of hybridization solution. The volume of the hybridization solution is critical. When too little solution is used, it is difficult to seat the coverslip without introducing air bubbles over some portion of the arrayed ESTs, and the coverslip will not sit at a uniform distance from the slide. If the coverslip is bowed toward the slide in the center, there will be less labeled cDNA in that area and hybridization will be nonuniform. When too much volume is applied, the coverslip will move easily during handling, leading to misplacement relative to the arrayed ESTs, and nonhybridization in some areas of the array.

2. For a 40-µl hybridization, pool the Cy3- and Cy5-labeled cDNAs into a single 0.2-ml thin-wall PCR tube and adjust the volume to 30 µl by either adding DEPC-treated water or removing water in a Speedvac evaporator. If using a vacuum device to remove water, do not use high heat or heat lamps to accelerate evaporation because this could degrade the fluorescent dyes.

3. For a 40-µl hybridization combine the following components: Cy5 + Cy3 probe 8 mg/ml poly(dA) 4 mg/ml yeast tRNA 10 mg/ml C0t-1 DNA 20× SSC 50× Denhardt’s solution Total volume

high sample blocking 30 µl 1 µl 1 µl 1 µl 6 µl 1 µl (optional) 40 µl

high array blocking 28 µl 2 µl 2 µl 0 µl 6 µl 2 µl 40 µl

Arrays and samples can vary somewhat, making it necessary to vary the composition of the hybridization cocktail. In cases where there is residual hybridization to control repeat DNA samples on the array, more C0t-1 DNA can be used, as in the high sample blocking formulation. When there is diffuse background or a general haze on all of the array elements, more of the nonspecific blocker components can be used, as in the high array blocking formulation. The authors generally try the high sample blocking formulation first.

4. Mix the components well by pipetting, heat at 98°C for 2 min in a thermal cycler, cool quickly to 25°C, and add 0.6 µl of 10% SDS. Nucleic Acid Arrays


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5. Centrifuge for 5 min at 16,000 × g, 20° to 25°C. The fluor-labeled cDNAs have a tendency to form small, very fluorescent, aggregates that result in bright, punctate background on the array slide. Hard centrifugation will pellet these aggregates, allowing one to avoid introducing them to the array.

6. Apply the labeled cDNA to a 24-mm × 50-mm glass coverslip and then touch with the inverted microarray. Applying the hybridization mix to the array and coverslipping it is an operation that requires some dexterity to get the positioning of the coverslip and the exclusion of air bubbles just right. It is helpful to practice this operation with buffer and plain slides before attempting actual samples. The hybridization solution is added to the coverslip first, since some aggregates of fluor remain in the solution and will bind to the first surface they touch.

7. Place the slide in a microarray hybridization chamber, add 5 µl of 3× SSC in the reservoir, if the chamber provides one, or else at the scribed end of the slide, and seal the chamber. Submerge the chamber in a 65°C water bath and allow the slide to hybridize for 16 to 20 hr. There are a wide variety of commercial hybridization chambers. It is worthwhile to prepare a mock hybridization with a blank slide, load it in the chamber, and incubate it to test for leaks or drying of the hybridization fluid, either of which will cause severe fluorescent noise on the array.

Wash off unbound fluorescent cDNA 8. Remove the hybridization chamber from the water bath, cool, and carefully dry off. Unseal the chamber and remove the slide. As there may be negative pressure in the chamber after cooling, it is necessary to remove water from around the seals so that it is not pulled into the chamber and onto the slide when the seals are loosened.

9. Place the slide, with the coverslip still affixed, into a Coplin jar filled with 0.5× SSC/0.01% SDS wash buffer. Allow the coverslip to fall from the slide and then remove the coverslip from the jar with forceps. Allow the slide to wash for 2 to 5 min. 10. Transfer the slide to a fresh Coplin jar filled with 0.06× SSC. Allow the slide to wash for 2 to 5 min. The sequence of washes may need to be adjusted to allow for more aggressive noise removal, depending on the source of the sample RNA. Useful variations are to add a first wash that is 0.5× SSC/0.1% SDS or to repeat the normal first wash twice.

11. Transfer the slide to a slide rack and centrifuge 3 min at low speed—167 × g (700 to 1000 rpm) in a clinical centrifuge equipped with a horizontal rotor for microtiter plates. If the slide is simply air dried, it frequently acquires a fluorescent haze. Centrifuging off the liquids results in a lower fluorescent background. As the rate of drying can be quite rapid, it is suggested that the slide be placed in the centrifuge immediately upon removal from the Coplin jar.


Acquire a fluorescent image of the slide and extract the signal data The particulars of adjusting a microarray imaging device will vary considerably depending upon the type used and the manufacturer. Similarly, a number of methods for identifying the signal in the image, adjusting for the level of background, and normalizing the values between the two channels can be used. This section is intended only to provide some guidelines to the general considerations of imaging.



12. Adjust the photomultiplier voltage and laser power so that the brightest signals produce slightly less than the maximum possible reading (i.e., 65,535, if a 16-bit scale), and the background values (between the arrayed spots) are consistently somewhat above zero, and then collect the data for the entire image. The aim in imaging the fluorescent signals is to use as much of the linear range of detection of the instrument as possible, and to scale both signals so that they occupy roughly the same range. Setting the device so that the assay background level is at the base of the range of values insures that the minimal detectable signal can be evaluated, and helps in the determination of whether one of the two signals is considerably stronger than the other.

13. Load the captured image into the data extraction software and examine images for overall quality of hybridization, noting the uniformity and level of backgrounds, the level and distribution of signals, the relative strengths of the signals in each channel, and the comparability of signals for most of the genes. A very good characterization of the performance of the system can be carried out with a series of four hybridizations. These basically ask whether the system accurately judges two identical samples to be identical, and checks the reproducibility of the detected differences when two samples are different. For a sample of this type of data, see Troubleshooting.

AGAROSE GEL ELECTROPHORESIS OF ESTs Gel imaging allows a rough quantitation of product while giving an excellent characterization of the product. Band size, as well as the number of bands observed in the PCR products, contribute to understanding the final results of the hybridization. The use of gel-well formats suitable for loading from 96-well plates, as well as programmable pipettors, make this form of analysis feasible on a large scale.

SUPPORT PROTOCOL 1

Materials 2% (w/v) agarose gel in 1× TAE buffer (see UNIT 2.5A) 50× TAE buffer (APPENDIX 2) Loading buffer (see recipe) 100-bp size standards (see recipe) Electrophoresis apparatus with capacity for four 50-well combs, (e.g., Owl Scientific) Disposable microtiter mixing trays (e.g., Becton Dickinson) Programmable, 12-channel pipettor with disposable tips (e.g., Matrix Technologies) Electrophoresis power supply Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5A) Perform gel electrophoresis 1. Cast a 2% agarose gel in 1× TAE buffer with 4 combs (i.e., 50-tooth) and submerge in an electrophoresis apparatus with sufficient 1× TAE buffer to just cover the surface of the gel (see UNIT 2.5A). 2. Prepare a reservoir of loading buffer, using 12 wells of a microtiter plate. 3. Program 12-channel pipettor to sequentially carry out the following steps: fill with 2 µl fill with 1 µl fill with 2 µl mix a volume of 5 µl five times expel 5 µl.

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–2400 bp –600 bp –100 bp –2400 bp –600 bp –100 bp

–2400 bp –600 bp –100 bp

–2400 bp –600 bp –100 bp

Figure 22.3.2 Ethidium bromide staining pattern of EST PCR products electrophoresed on a 2% agarose gel.

4. Place 12 disposable tips on the pipettor. 5. Load 2 µl of PCR product from wells A1 to A12 of the PCR plate. 6. Load 1 µl of air. 7. Load 2 µl loading buffer from the reservoir. 8. Place tips in clean wells of disposable microtiter mixing tray and allow pipettor to mix the sample and loading dye. 9. Place the pipettor in a 50-well row so that the tip containing the PCR product from well A1 is in the second well of the row, and the other tips are in every other succeeding well. 10. Repeat the process (changing tips each time), loading PCR plate row B starting in the third well, interleaved with the A row, the C row starting at well 26, and the D row at well 27, interleaved with the C row. 11. Place 5 µl of 100-bp size standard in wells 1 and 50. 12. Repeat this process, loading samples from rows E, F, G, and H in the second, 50-well row of gel wells, loading samples from two 96-well PCR plates per gel, or single-row samples from 16 PCR plates.


To reduce diffusion and mixing, apply voltage to the gel for a minute between loading each well strip. This will cause the DNA to enter the gel, and reduce band spreading and sample loss.



13. Apply voltage to the gel and run until the bromphenol blue (faster band) has nearly migrated to the next set of wells. For a gel that is 14 cm in the running dimension, and 3 cm between each row of wells, apply 200 V for 15 min.

14. Take a digital photo of gel and store image for future reference. The gels should show bands of fairly uniform brightness distributed in size between 600 to 2000 bp as in Figure 22.3.2. Further computer analysis of such images can be carried out with image-analysis packages to provide a list of the number and size of bands. Ideally this information can be made available during analysis of the data from hybridizations involving these PCR products.

FLUOROMETRIC DETERMINATION OF DNA CONCENTRATION While it would be ideal to be able to quantify exactly each EST PCR product and spot all DNA species at equivalent concentrations, it is impractical for most laboratories to do so when thousands of ESTs must be prepared. Fortunately, it is possible to use a strategy where excess DNA is spotted, so that the exact quantities used do not produce much variation in the observed results. When using this strategy, it is necessary to track the average productivity of the PCR reactions. Fluorometry provides a simple way to obtain an approximate concentration of the double-stranded PCR product in the PCR reaction mix.

SUPPORT PROTOCOL 2

Materials FluoReporter Blue dsDNA Quantitation Kit (Molecular Probes) Fluor buffer (see recipe) PCR product (see Basic Protocol 3) TE buffer, pH 8 (APPENDIX 2) 50, 100, 250, and 500 µg/ml dsDNA reference standards (e.g., see recipe) 96-well plates for fluorescent detection (e.g., Dynex) 12-channel multipipettor Fluorometer (e.g., PE Biosystems) Computer equipped with Microsoft Excel software Quantitate dsDNA 1. Label 96-well plates for fluorescence assay. With a 12-channel multipipettor, add 200 µl of fluor buffer to each well. 2. Add 1 µl PCR product from each well in a row of a PCR plate to a row of the fluorometry plate. Samples can be added to rows A through G of the fluorometry plate.

3. In the final row of the fluorometry plate, add 1 µl TE buffer to first well and add 1 µl of each of the series of dsDNA standards—50, 100, 250, and 500 µg/ml—dsDNA to the subsequent wells. Repeat this series twice in the final row. 4. Set the fluorometer for excitation at 346 nm and emission at 460 nm. Adjust as necessary to read the plate. If the fluorometer does not support automated analysis, export the data table to Excel.

5. Test to see that the response for the standards is linear and reproducible from the range of 0 to 500 µg/ml of dsDNA. Nucleic Acid Arrays


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6. Calculate the concentration of dsDNA in the PCR reactions using the following equation after subtracting the average 0 µg/ml value from all other sample and control values: [dsDNA (µg/ml)] = [(PCR sample value)/(average 100 µg/ml value)] × 100 Constantly tracking the yields of the PCRs makes it possible to rapidly detect many ways in which PCR can fail or perform poorly. This assay can also be applied after precipitation and resuspension of the PCR products to monitor overall recovery of product. SUPPORT PROTOCOL 3

COATING SLIDES WITH POLY-L-LYSINE Slides coated with poly-L-lysine have a surface that is both hydrophobic and positively charged. The hydrophobic character of the surface minimizes spreading of the printed spots, and the charge appears to help position the DNA on the surface in a way that makes cross-linking more efficient. Materials Cleaning solution (see recipe) Poly-L-lysine solution (see recipe) Gold Seal microscope slides (Becton Dickinson) 50-slide stainless steel rack and 50-slide glass tank (Wheaton) 25-slide plastic rack and 25-slide plastic box (Shandon Lipshaw) Plastic slide box with no paper or cork liners (e.g., PGC Scientific) Coat slides 1. Place slides into 50-slide racks and place racks in glass tanks with 500 ml of cleaning solution. Gold Seal slides are highly recommended, as they have been found to have consistently low levels of autofluorescence. It is important to wear powder-free gloves when handling the slides. Change gloves frequently, as random contact with skin and surfaces transfers grease to the gloves.

2. Place tanks on platform shaker for 2 hr at 60 rpm. 3. Pour out cleaning solution and wash in water for 3 min. Repeat wash four times. 4. Transfer slides to 25-slide plastic racks and place into small plastic boxes for coating. 5. Submerge slides in 200 ml poly-L-lysine solution per box and shake for 1 hr at 60 rpm. 6. Rinse slides three times with water and submerge slides in water for 1 min. 7. Centrifuge for 2 min at 400 × g and dry slide boxes used for coating. 8. Place slides back into slide box used for coating and let stand overnight before transferring to new slide box for storage. This allows the coating to dry before handling.

9. Allow slides to age for two weeks on the bench, in a new slide box, before printing on them. The coating dries slowly, becoming more hydrophobic with time. Profiling Human Gene Expression with cDNA Microarrays

Slide boxes used for long-term storage should be plastic and free of cork lining. The glue used to affix the cork will leach out over time and give slides stored in these types of boxes a greasy film that has a high degree of autofluorescence. Clean all glassware and racks used for slide cleaning and coating with highly purified water only. Do not use detergent.



REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps, unless otherwise noted. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

Carbenicillin stock solution, 100 mg/ml 1 g carbenicillin (Life Technologies) 10 ml sterile water Sterile filter with a 0.2-µm filter Store frozen at –20°C up to 2 months Cleaning solution 400 ml H2O 600 ml 100% ethanol 100 g NaOH Dissolve NaOH in water. Add ethanol and stir until the solution clears. If the solution does not clear, add water until it does. Store up to 24 hr at room temperature. dsDNA reference standards 50 ìg/ml: 90 µl TE buffer (APPENDIX 2) 10 µl 0.5 mg/ml dsDNA (Life Technologies) 100 ìg/ml: 80 µl TE buffer (APPENDIX 2) 10 µl 0.5 mg/ml dsDNA (Life Technologies) 250 ìg/ml: 50 µl TE buffer (APPENDIX 2) 50 µl 0.5 mg/ml dsDNA (Life Technologies) 500 ìg/ml: 0 µl TE buffer (APPENDIX 2) 100 µl 0.5 mg/ml dsDNA (Life Technologies) It is good practice to check both the integrity (i.e., on a agarose gel) and the concentration (i.e., absorbance) of the standard before use. The reference lambda dsDNA is available from Life Technologies.

Ethanol/acetate solution 950 ml 100% ethanol 50 ml 3 M sodium acetate, pH 6.0 (see recipe) H2O to 1000 ml Store up to 2 weeks at room temperature Fluor buffer 25 µl Hoechst 33258 solution (from FluoReporter Blue kit; Molecular Probes) 10 ml TNE buffer (from FluoReporter Blue kit; Molecular Probes) 10 ml water Hoechst 33258 solution contains the dye at an unspecified concentration in a 1:4 mixture of DMSO:H2O. TNE Buffer is 10 mM Tris⋅Cl (pH 7.4), 2 M NaCl, 1 mM EDTA.

Human C0t-1 DNA, 10 mg/ml Add 925 µl of 100% ethanol and 75 µl of 3 M sodium acetate (pH 5.2) to 500 µl of 1 µg/µl human C0t-1 DNA. Centrifuge at 14,000 × g. Aspirate supernatant and allow pellet to air dry for 5 min. Resuspend the pellet in 50 µl DEPC-treated water (UNIT 4.1). Store up to 6 months at −20°C.

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Low-T dNTP mix, 10× 25 µl 100 mM dGTP (0.5 mM in 1×) 25 µl 100 mM dATP (0.5 mM in 1×) 25 µl 100 mM dCTP (0.5 mM in 1×) 10 µl 100 mM dTTP (0.2 mM in 1×) 415 µl DEPC-treated H2O (UNIT 4.1) Total volume, 500 µl Store up to 2 months at −20°C Loading buffer 4.0 ml glycerol (enzyme grade) 0.9 ml DEPC-treated H2O (UNIT 4.1) 0.1 ml 0.25% (w/v) xylene cyanol FF/0.25% (w/v) bromphenol blue Store up to 1 month at room temperature Phosphate-buffered saline (PBS) 8.00 g/liter NaCl 0.20 g/liter KCl 1.44 g/liter Na2HPO4 (anhydrous) 0.24 g/liter KH2PO4 (anhydrous) Bring components to 1 liter volume with H2O Autoclave 20 min Cool to room temperature Pass through a 0.2-µm filter Store up to 6 months at room temperature Poly-L-lysine solution 35 ml poly-L-lysine (0.1% w/v; Sigma) 35 ml PBS (see recipe) 280 ml H2O Store up to 24 hr at room temperature Size standards, 100-bp 50 µl 1 mg/ml DNA ladder (Life Technologies) 5 µl 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2) 5 µl 0.5 M EDTA, pH 8.0 (APPENDIX 2) 440 µl loading buffer (see recipe) Store up to 1 month at 4°C Sodium acetate, 3 M (pH 6.0) Dissolve 408.24 g/liter sodium acetate trihydrate to prepare 3 M sodium acetate. Prepare 3 M acetic acid by diluting 172.4 ml glacial acetic acid to 1 liter with water. Titrate the pH of 3 M sodium acetate solution to pH 6.0 with the 3 M acetic acid solution. Filter sterilize using a 0.2-µm filter. Store up to 6 months at room temperature. Sodium borate, 1 M (pH 8.0) Dissolve 61.83 g boric acid in 900 ml DEPC-treated water (UNIT 4.1). Adjust the pH to 8.0 with 1 N NaOH. Bring volume up to 1 liter. Sterilize with a 0.2-µm filter and store up to 6 months at room temperature.


T low E buffer 10 ml 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2) 0.2 ml 0.5 M EDTA, pH 8.0 (APPENDIX 2) 900 ml DEPC-treated H2O (UNIT 4.1) Autoclave and store up to 6 months at room temperature.



Wash buffer, 0.5× SSC/0.01% (v/v) SDS Add 25 ml of 20× SSC to 974 ml DEPC water. Filter sterilize on a 0.5-µm filter device. Add 1 ml of 10% SDS, and mix well. Store up to 2 months at room temperature. Wash buffer (0.06× SSC) Add 3 ml of 20× SSC (APPENDIX 2) to 997 ml DEPC-treated water (UNIT 4.1). Filter sterilize on a 0.5-µm filter device. Store up to 2 months at room temperature. Yeast tRNA, 4 ìg/ml 1. Resuspend yeast tRNA at 10 mg/ml (based on the supplier’s quantitation of the RNA) in DEPC-treated water (UNIT 4.1) in a 1.5-ml polypropylene conical centrifuge tube. 2. Add 0.5 vol of buffered phenol (UNIT 2.1A), then vortex. 3. Add 0.5 vol chloroform, then vortex again. 4. Centrifuge 5 min at 10,000 × g. Transfer aqueous layer to a new 1.5-ml polypropylene conical centrifuge tube. 5. Add 1 vol chloroform, and vortex. Centrifuge 5 min at 10,000 × g. 6. Repeat chloroform extraction. 7. Transfer aqueous layer to a new 1.5-ml polypropylene conical centrifuge tube. Add 0.1 vol of 3 M sodium acetate, pH 5.2 (APPENDIX 2). Add 2 vol 100% ethanol. 8. Centrifuge 5 min at 10,000 × g. Aspirate supernatant, then add 1 vol of 70% ethanol. 9. Centrifuge 5 min at 10,000 × g. Aspirate supernatant again and allow pellet to dry. 10. Resuspend in DEPC-treated water (UNIT 4.1) at the original volume. 11. Determine the RNA concentration by spectrometry (APPENDIX 3D). 12. Dilute to 4 mg/ml and store frozen at –20°C. COMMENTARY Background Information The possibility of examining the expression patterns of many genes simultaneously has been more and more enthusiastically pursued as the sequences and clones of a greater fraction of genes from model organisms have become available. A large part of the enthusiasm derives from the growing recognition of the extraordinary levels of integration and interaction between genes in all cell systems. It is clear that the same cues can provoke a wide variety of cellular responses, dependent in large part upon the proteins currently expressed in the cell being examined. By observing changes in a particular gene’s expression against the backdrop of the patterns of change of other genes, it is possible to ask contextual questions about cellular function. Some of the simple but intriguing questions of this class that can be asked revolve around longstanding, basic issues. These would include questions about what activities are jointly regulated, what chains of events are involved in normal processes such

as cell division, and what expression patterns are correlated with particular processes or pathologies. Most of the large-scale expression profiling done today is carried out either on arrays of short oligonucleotides or on arrays of ESTs. A good single-source comparative review of the strengths and weakness of the techniques, as well as example applications of both can be found in a 1999 supplement of Nature Genetics (Phimister, 1999; also see overview in UNIT 22.1). Both of these technologies are still in their early phases, and it is reasonable to expect that the methodologies for obtaining and analyzing gene expression profiles will change at a very rapid pace.

Critical Parameters Fluorescent noise Assays based on fluorescent staining are vulnerable to many environmental sources of noise. Nearly all dust derived from paper or

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cloth fibers is intensely fluorescent across the visible spectrum. Many organic molecules are fluorescent in the spectral regions used for this type of study. It is necessary to keep exposure to dust to a minimum and to carefully monitor the purity of the reagents used to fabricate microarrays. All sources of grease in the preparation of slides must be carefully avoided, since greasy films will bind fluorescent molecules nonspecifically. For those steps where the array slides are handled, all glassware should be carefully cleaned and soaked in 1 N nitric acid before initial use, and then separately shelved and used only for arrays. Water should minimally be from either a glass still or prepared by reverse osmosis and then run through deionizing tanks and charcoal filtration tanks. RNA handling The routine precautions required for successful lab handling of RNA should be observed (see Chapter 4 introduction). For all DNA and RNA manipulations in these protocols use DEPC-treated water unless otherwise specified.


RNA purity The quality of the RNA coming into the labeling will have a marked effect on the quality of the labeling and hybridization. RNA preparations judged good by standard molecular biology criteria often produce poor results. Typical problems include disperse, fine-grain noise over the entire hybridized surface and nonspecific binding of fluorescent molecules to the zones of DNA immobilization on the slide. These problems seem likely to have some roots in contaminating carbohydrate, and, as would be expected with carbohydrate, the problems are exacerbated by ethanol precipitation before and after labeling. Very impure preparations will frequently produce visible aggregates if precipitated after labeling, which are essentially resistant to solubilization. It is well known that nucleic acids form strong aggregates with carbohydrate when either dried together or when coprecipitated. This interaction is the basis for nucleic acid immobilization onto chromatography supports such as cellulose. To minimize this sort of problem, the authors recommend preparative procedures that use few or preferably no ethanol precipitations during RNA preparation and labeling. Purity also exerts strong effects on the efficiency of labeling. Reverse transcription of RNA in the presence of nucleotides derivatized with fluorescent molecules is not an efficient

process at best, and is further impaired by impurities in the incoming template. The typical problems observed are total failure of the labeling or production of many very short reverse transcripts. The purification scheme described in this Basic Protocol 2 is designed to minimize carbohydrate and lipid carryover. It is a modification of the authors’ previous method combining chromatography and phase extraction, kindly suggested by Dr. Alvydas Mikulskis of New England Nuclear. Validation and verification A very important step in qualifying the reliability of an array assay system is to validate experimental results. An early experiment with two samples where there is an expectation that some of the genes on the array will be differentially expressed is strongly recommended. The expected differences should be observed, and verified by northern analysis with probes made from the EST segments immobilized as reporters. Additional observed differences beyond the initial expectations should also be observed. Testing should be carried out over a range of signal strengths and a range of differential expression. Clones and informatics The ability to collect and manage numerous human EST clones inevitably adds clone verification and database requirements that are not easily met by public databases and repositories. It is important that the complications associated with ESTs be understood before array experiments are designed. The major obstacles in human EST usage arise from three sources. The first problem is that the categorization of the structures of human genes is not complete. Human EST sequences and the clones from which these sequences are derived are continually being deposited in publicly available repositories. The deposited sequences are then examined for similarity to each other and to known genes, and are clustered into groups of sequences thought to represent multiple examples of a single type of transcript. Organizations such as NCBI carry out this operation on a periodic basis, reforming the clusters to account for new EST sequences and name assignments and make the results publicly available (http://www.ncbi.nlm.nih.gov/UniGene/ Hs.Home.html) as part of their UniGene project. The rate of change in assignment of an EST to a cluster remains fairly high, so that the assigned identities of many ESTs fluctuate. It is thus necessary to be able to update identity



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Figure 22.3.3 Scatter plots of Cy3 (green) and Cy5 (red) mean intensities observed in three hybridizations. The labeled cDNAs in the hybridizations were made from: (A) cell line ML1, Cy3 and Cy5, (B) cell line UACC903, Cy3 and Cy5, and (C) cell line UACC903, Cy5, and cell line ML1, Cy3. Nucleic Acid Arrays


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and cross-referenced data, which depends on that identity, for any EST inventory. The second obstacle derives from the fact that it is not currently possible to obtain such data for lists of ESTs from the public databases. The databases are set up to provide answers to queries on a gene by gene or EST by EST basis. Efficiently answering questions such as “What are the current chromosomal position assignments of the ESTs in a particular set?” or “Do any of the ESTs in my inventory correspond to a particular UniGene cluster?” requires a local relational database. A frequently updated minimal relational database for human ESTs based on UniGene, that can be set up and run on an

ordinary desktop computer, can be obtained to serve this purpose (http://www.nhgri.nih.gov/ DIR/LCG/15K/DATA/). The final serious difficulty is that many public and private EST inventories have a relatively high rate of inaccuracy. For instance, IMAGE clones available from public distributors have an ∼20% chance of either not having the 3′ sequence ascribed to them in the public databases or having multiple EST clones in a single well. For this reason, a number of EST suppliers are now supplying clone sets that have been clone purified and sequence verified.

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Figure 22.3.4 Scatter plots comparing the expression ratios of genes in cell line UACC903 and ML1 in three separate experiments. In panel A, the ratios in experiment A are compared to the ratios in experiment B. In panel B, the ratios in experiment A are compared to the ratios in experiment C.



Troubleshooting One of the most illuminating diagnostics for array performance is a hybridization that reveals how tightly an array estimates that two samples from the same source produce equivalent signals. An example of a panel of three experiments that can rigorously test this capability is shown in Figure 22.3.3. Panels A and B of Figure 22.3.3 are scatter plots of the normalized mean intensities of readings from 6782 array detectors, when a single sample source is used to generate both the Cy3 and Cy5 labeled cDNA hybridized to the array. Panel C is a scatter plot of the normalized mean intensities when the two very different sample types, a melanoma cell line and a myeloid cell line used in A and B, respectively, are the sources for the labeled cDNAs in the hybridization. From such plots, one can obtain an immediate qualitative sense of the level of error in judging identity and the extent of difference in excess of error that nonidentical samples display. These parameters may obviously be more rigorously defined by statistical analysis. After achieving satisfactory performance at the level of finding identity, it is worthwhile to examine the reproducibility with which differences are determined in multiple assays. In large measure, this is a test of the reproducibility of the printing process. A sample of this kind of testing is shown in Figure 22.3.4. Three repetitions of the experiment seen in Panel C of Figure 22.3.3 were carried out. Filtering for genes that had a minimum mean intensity of 50 in both channels developed a subset 3861 of well-detected genes. Scatter-plot comparisons of the ratios of the UACC903 (Cy5) to ML1 (Cy3) signal for genes that were well detected in all the experiments were then carried out. Panels A and B of Figure 22.3.4 show plots comparing the ratios observed in the first hybridization, experiment A, with the ratios observed in experiments B and C, respectively. Analysis at this level can easily show whether the assay is essentially working, failing in some areas of the array, or failing generally. Faults will show up as significant deviations from the diagonal. If only some portion of the genes are far from the diagonal, then their position on the array can be determined and the properties of local background and signal can be checked to see why the assay failed in those instances.

Anticipated Results Microarray experimentation should provide a means for determining whether there is a difference in the expression level between those

genes that are present at sufficient levels in the sample to be detected by hybridization to the EST PCR products represented on the array.

Time Considerations Time considerations for various operations described in this unit are as follows. Prepare replicates and cultures for template: 8 plates–5 hr. Prepare plasmid template from cultures: 4 plates–2.5 hr. Set up PCR using templates from cultures: 12 plates–2 hr. Prepare 200-well agarose gel: 2 gels–45 min. Load 2 plates of PCR samples: 1 gel– 30 min. Ethanol precipitate and resuspend PCR products: 12 plates–5.5 hr. (with overnight incubation) Extract RNA: 4 samples–4 hr. Label RNA: 4 samples–4 hr. Set up hybridization: 2 slides–0.5 hr.

Acknowledgements The authors would like to acknowledge the many excellent suggestions and kind support of our colleagues at NHGRI, Jeffrey Trent, Paul Meltzer, Yidong Chen, Javed Khan, Abdel Elkahloun, and Anthony Masiello. We would also like to thank the members of the Brown lab, especially Joe DeRisi for sharing the details of the printing methods that they developed, and the many members of the array research community who have generously shared their observations and insights with us.

Literature Cited Alwine, J.C., Kemp, D.J., and Stark, G.R. 1977. Method for detection of specific RNAs in agarose gels by transfer to diazobenzyloxymethyl-paper and hybridization with DNA probes. Proc. Natl. Acad. Sci. USA 74:53505354. Bishop, J.O., Morton, J.G., Rosbash, M., and Richardson, M. 1974. Three abundance classes in HeLa cell messenger RNA. Nature 250:199204. DeRisi, J.L., Iyer, V.R., and Brown, P.O. 1997. Exploring the metabolic and genetic control of gene expression on a genomic scale. Science 278:680686. Iyer, V.R., Eisen, M.B., Ross, D.T., Schuler, G., Moore, T., Lee, J.C.F., Trent, J.M., Staudt, L.M., Hudson, J. Jr., Boguski, M.S., Lashkari, D., Shalon, D., Botstein, D., and Brown, P.O. 1999. The

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transcriptional program in the response of human fibroblasts to serum. Science 283:83-87. Khan, J., Simon, R., Bittner, M., Chen, Y., Leighton, S.B., Pohida, T., Smith, P.D., Jiang, Y., Gooden, G.C., Trent, J.M., and Meltzer, P.S. 1998. Gene expression profiling of alveolar rhabdomyosarcoma with cDNA microarrays. Cancer Res. 58:5009-5013. Lockhart, D.J., Dong, H., Byrne, M.C., Follettie, M.T., Gallo, M.V., Chee, M.S., Mittmann, M., Wang, C., Kobayashi, M., Horton, H., and Brown, E.L. 1996. Expression monitoring by hybridization to high-density oligonucleotide arrays. Nat. Biotechnol. 14:1675-1680. Phimister, B.E. 1999. The chipping forecast. Nat. Genet. 21(1 Suppl).

Schena, M., Shalon, D., Davis, R.W., and Brown, P.O. 1995. Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science 270:467-470. Spellman, P.T., Sherlock, G., Zhang, M.Q., Iyer, V.R., Anders, K., Eisen, M.B., Brown, P.O., Botstein, D., and Futcher, B. 1998. Comprehensive identification of cell cycle-regulated genes of the yeast Saccharomyces cerevisiae by microarray hybridization. Mol. Biol. Cell 9:3273-3297.

Contributed by Yuan Jiang, John Lueders, Arthur Glatfelter, Chris Gooden, and Michael Bittner National Human Genome Research Institute, NIH Bethesda, Maryland




Overview of mRNA Expression Profiling Using Microarrays mRNA expression profiling using microarrays is a technology that allows simultaneous determination of the mRNA levels of many genes. Microarrays range from small custom arrays designed to monitor expression of a few hundred genes to very large arrays that represent tens of thousands of genes or entire genomes. Presently, there are three major applications for microarray data. One application treats microarray data as massively parallel expression assays. Typically, this data is used for identification of particular genes that undergo expression changes in response to particular treatments, in particular cell types, or in particular mutants. Such genes are often considered candidates for players in a biological process, such as response to a treatment. A second application involves treating expression profiles as descriptions of collective behaviors. The state of the cell from which the sample was prepared is collectively characterized by determining the expression levels of tens of thousands of genes. For example, an expression profile of a drug-treated cell describes the effect of the drug, and drugs with similar modes of action can be identified by comparing expression profiles of drug-treated cells. A third application which is becoming feasible is the mining of large expression profiling databases to characterize expression patterns of genes of interest over a wide range of tissue types, after various treatments, or in different mutants. Due to the complexity of microarray data, computational tools are required for analysis. These tools must be tailored according to the type of analysis being carried out. If the goal is to identify genes that show expression-level changes between different samples, statistical tools are needed to sort genes based on the degree of confidence that they are actually differentially expressed. If the goal is to identify patterns in expression profiles that are diagnostic of particular cell states, pattern-recognition tools are needed. Another aspect of handling complex data is that in many cases subjective decisions are applied. For example, subjective criteria are used to decide how two different types of data, such as microarray data analysis results and gene annotation, should be combined for selection of candidate genes for further study. In the following discussion, an

overview of issues to consider when designing microarray experiments and analyzing the data is presented.

STRENGTHS AND WEAKNESSES OF EXPRESSION PROFILING RNA and corresponding cDNA species have relatively homogenous chemical characteristics, and can be specifically detected based on hybridization of complementary strands. These advantages enable simultaneous monitoring of tens of thousands of genes with very good sensitivity and accuracy at reasonable cost. Another advantage of using cDNA species is that they can be amplified by PCR and/or in vitro transcription. Consequently, the amount of RNA required for expression profiling has been decreasing as methods for quantitative amplification of cDNA have improved (Wang et al., 2000; Baugh et al., 2001; Iscove et al., 2002). This feature allows expression profiling of very small amounts of tissue. Thus, microarray analysis could potentially have a very high spatial resolution (Ohyama et al., 2000). In terms of obtaining global quantitative information about a particular class of molecules, largescale profiling technologies for other classes of molecules, such as proteins and metabolites, cannot match the performance of expression profiling. On the other hand, the information obtained by expression profiling is simply the amount of each mRNA species present. mRNA levels do not necessarily correlate with levels of active, properly localized proteins or the amounts of metabolites that they produce. Consequently, the biological significance of observed changes in mRNA levels is open to question. This limitation must be kept in mind when interpreting the results of expression profiling experiments.

MICROARRAY TECHNOLOGIES In its first incarnation, microarray technology involved spotting random cDNA clones for the purpose of detecting genes with differential expression levels in different samples in organisms with very limited DNA sequence information. The cDNA clones corresponding to genes that were found to be differentially expressed were then sequenced. Such an approach is quite inefficient, and other methods are now available to identify differentially-expressed genes

Contributed by Fumiaki Katagiri and Jane Glazebrook Current Protocols in Molecular Biology (2004) 22.4.1-22.4.8 Copyright © 2004 by John Wiley & Sons, Inc.

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in organisms lacking extensive sequence information. An early example is differential display of mRNA (Liang and Pardee, 1992; UNIT 25B.3), followed more recently by methods including serial analysis of gene expression (SAGE; http://www.sagenet.org; UNIT 25B.6; Velculescu et al., 1995) and massively parallel signature sequencing (MPSS; www.lynxgen.com/wt/ tert.php3?page_name=mpss; Brenner et al., 2000). Due to their capacities for covering large numbers of genes and rapid short sequence tag determination, SAGE and MPSS are also used for discovery of new mRNA species in organisms where genome information is available. Generally, the cost per sample of these methods is fairly high.

Affymetrix Arrays

Overview of mRNA Expression Profiling Using Microarrays

These days the more common use of microarray technology is in monitoring expression of genes with known sequences. There are several different microarray technology platforms in use. Some platforms use arrays that must be purchased from commercial entities, while other types can be produced using widely available equipment. GeneChip arrays from Affymetrix (http://www.affymetrix.com) consist of a collection of short (typically 25-mer) oligonucleotides that are synthesized directly on the array (Lockhart et al., 1996). The oligonucleotides can be arrayed at extremely high density, so these arrays are usually used to represent entire genomes, or tens of thousands of different genes. Each gene is represented by eleven to sixteen pairs of exactly hybridizing oligonucleotides and oligonucleotides containing single-base mismatches. The signal from the mismatch oligos can be used for estimation of noise resulting from nonspecific hybridization. A single sample is hybridized to each array and the expression level for each gene is obtained by combining the signals from the multiple oligonucleotide probes. To compare data from different samples, the signals from each gene are normalized to the average signal for all the genes on the array, based on the assumption that most of the genes in the genome are not changing expression levels between any two experimental conditions. The main advantage of Affymetrix arrays is their excellent technical reproducibility due to the physical uniformity of the arrays themselves and the statistical power resulting from having multiple measurements for each gene. The disadvantages are that the design of the arrays cannot be altered to suit individual experiments, and the arrays are relatively expensive.

NimbleGen Arrays NimbleGen arrays (http://www.nimblegen. com) are also high density arrays of oligonucleotides synthesized directly on a substrate (Nuwaysir et al., 2002). In contrast to Affymetrix arrays, the composition of the arrays can be changed very easily, so arrays can be tailored to particular experiments. In addition, longer oligonucleotides can be used for NimbleGen arrays. NimbleGen arrays are not available for purchase; rather, RNA samples must be sent to the company, which carries out the experiments and returns the data. NimbleGen arrays have not yet been widely used so it is difficult to evaluate the quality of the data; however, the similarity to the Affymetrix technology and published validation experiments (Nuwaysir et al., 2002) suggest high accuracy. The main advantage is the flexibility of the system—the design of individual arrays can be customized using a simple computer interface. The disadvantage is that the total cost of obtaining expression data from an RNA sample, including array production, hybridization, and scanning, is, at the time of this writing, even more expensive for a NimbleGen array than it is for a standard Affymetrix array.

Spotted Arrays Rather than synthesizing oligonucleotides directly on a substrate, they can be spotted onto a glass slide. The required spotting robots are widely available, so this technology is suitable for production of custom arrays in academic laboratories. However, olignucleotides cannot presently be spotted at the density required for representing tens of thousands of genes with multiple oligonucleotides per gene. Therefore, each gene is usually represented by one or two spots of an oligonucleotide of 60 to 70 bp, which can be purchased commercially or synthesized in-house by automated oligonucleotide synthesizers. The advantages of this technology are its flexibility and relatively low cost. However, there are technical challenges associated with production of high-quality data. Spotted arrays are less uniform than Affymetrix or NimbleGen arrays, making it difficult to compare data obtained from different arrays. Consequently, two-color methods are often used. In these experiments, two samples are labeled with different fluorescent dyes, and hybridized to the same array. The ratio of the two samples is used for further analysis. This method compensates for variation in the amount of oligonucleotide present in each spot although it can also result in a high level of error



(see below). Arrays on which each oligonucleotide is spotted only once or twice have inherently less statistical power than those on which each gene is represented by multiple oligonucleotides. For specialized arrays representing only a few hundred genes, the entire array can be spotted multiple times. Multiple measurements for each gene increase statistical power and result in reduced technical error. Spotted arrays can also be produced using cDNA clones or PCR products rather than oligonucleotides. Such arrays can have high sensitivity due to the long lengths of the probes, but suffer from cross-hybridization of related genes. In addition, there is much more labor involved in the production, quality control, and tracking of the necessary PCR products or clones than in acquisition of a set of oligonucleotides. These arrays are also usually used with two-color methods.

Choosing an Array or an Alternative Technology Choosing an appropriate microarray technology platform requires consideration of a number of questions, including: Is a satisfactory commercial array available? Is a satisfactory array available from an academic cooperative? How many genes must be monitored? What is the budget for the project? If the number of genes to be monitored is low and the number of samples is large, it might be wise to consider alternatives to microarrays. One is high-throughput quantitative RT-PCR, which offers much better sensitivity than microarrays. There are also bead-based technologies in which gene-specific probes are attached to fluorescently-labeled beads that are then counted using an instrument similar to a flow cytometer (Yang et al., 2001).

COMMON APPLICATIONS OF EXPRESSION PROFILING BY MICROARRAYS Applications of microarray analysis fall into three major classes. One application is discovery of genes with different expression levels in particular samples. These might be genes induced in response to pathogen attack, genes expressed specifically in certain cell types, genes with altered expression after drug treatment, and so forth. Such genes might then be used as molecular markers for certain biological states, or investigated further to determine if they are causally associated with the biological process of interest. For example, one might test pathogen-induced genes for contributions

to disease resistance. To create accurate lists of differentially-expressed genes, experimental design and statistical analysis of the data are crucial. Generally, greater numbers of biological and technical replicates allow greater confidence in the accuracy of the lists of differentially-regulated genes. Another application is the use of expression profiles as descriptions of the collective behavior of many genes for each sample (i.e., description of the cell state) or of many samples for each gene (i.e., description of the expression behavior of each gene). By comparing expression profiles of various mutants, it is possible to recognize mutants with defects similar to those of known mutants. For example, Hughes et al. (2000) discovered a yeast gene involved in sterol metabolism based on the observation that a mutation in it resulted in an expression profile similar to those of other mutants with defects in sterol metabolism. For this sort of analysis, it is critical to have powerful methods for pattern recognition to allow the investigator to recognize similarities among profiles. In contrast to conclusions about differential expression of individual genes, patterns composed of the expression values of many genes are statistically robust. Indeed, Hughes et al. (2000) showed that even when genes with highmagnitude changes are excluded from an analysis, similarity relationships among expression profiles are still quite stable. By comparing expression levels of genes in many different samples, genes that are regulated very similarly in many different situations can be identified. These genes may be involved in a common biological process (e.g., Kim et al., 2001). Lastly, databases consisting of large amounts of expression profiling data can be used to rapidly obtain information about a gene of interest. For example, a database consisting of expression profiles from different tissues, mutants, and variously-treated samples can be mined to determine in what tissues a gene of interest is expressed, what signaling pathways control its expression, and what treatments alter its expression. The content of various microarray databases is rapidly increasing, so this sort of study will soon be possible for a variety of well-studied organisms. For this application, standardization of microarray data formats, data size and data quality are all important.

VARIATION IN MICROARRAY DATA Two kinds of variation affect microarray data: biological and technical variation. Bio-

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logical variation refers to real differences in the level of mRNA species between two samples that the investigator considers to be experimental replicates. They are caused by uncontrolled variables in experimental conditions as well as by stochastic biological events. For some samples, such as budding yeast growing in liquid culture, experimental conditions can be very tightly controlled. However, samples from two different mice, or worse yet, two different humans, are subject to many variations in experimental conditions that are beyond the investigator’s control. Biological variation is also dependent on the stability of the biological process of interest in the presence of small perturbations (Tao et al., 2003). Irrespective of the source of biological variation, greater variation results in lower statistical power. The adverse effects of relatively high system variation may be overcome by increasing the number of replicates. Technical variation results from variations in the mechanics of the microarray experiment itself (e.g., fabrication of the array, preparation of RNA from the sample, labeling, hybridization, washing, scanning). A simple way to assess the level of technical variation is to hybridize RNA from the same sample to two or more different arrays. Improvements in array technology and statistical procedures for dealing with background signal and normalization can reduce technical variation. In two-color methods, dye-swap technical replicates are highly desirable because the relative efficiencies of labeling with different dyes vary for different mRNA/cDNA species. While it is a good idea to assess technical variation for each sample in a microarray experiment, in the interests of economy, so-called “technical replicates” for experimental samples can be reduced (Churchill, 2002) or even omitted in favor of biological replicates, which include both the technical and biological variation in the system.

STATISTICS AND PRACTICAL DECISIONS


Statistics are important for many aspects of microarray data analysis. To begin with, microarray experiments should be designed based on statistical considerations, such as randomization (Churchill, 2002). For example, if mice cages for corresponding samples in biological replicates are kept in the same places, the correlation observed in gene expression might be due to the cage location rather than the intended experimental conditions. Data from a carefully designed experiment are particularly useful

when the data are shared for general use. However, for focused research, an experimental design that is not as rigorous may be used, dependent on the goals, bottlenecks, and budget of the research. For example, if the research goal is to identify a few good expression marker genes, running a microarray experiment with a small number of replicates, choosing a small number of candidate genes, and running a rigorous test for the candidate genes using quantitative RT-PCR could be more time- and costeffective than running a microarray experiment with a large number of replicates.

Calculating Expression Values For data analysis, there is the question of how to calculate a signal value for a probe spot based on the fluorescence intensity of a group of pixels in the scanned array image. Statistical principles are used to determine a small number of values (for example, the mean and standard deviation) that represent a larger group of values (such as the fluorescence intensity for a group of pixels). Microarray data must also be adjusted to compensate for nonspecific background signal. These calculations are usually performed using specialized software packages and require relatively little thought on the part of the typical user. For arrays on which each gene is represented by more than one probe spot, the values from each spot must be combined to yield a value for each gene. Normalization is required to allow comparisons between independent arrays with different overall signal intensities. There are multiple ways of doing this, which are often specific to particular microarray technology platforms. Comparison of different normalization methods is outside the scope of this overview. Interested readers may refer to other literature (e.g., Quackenbush, 2002; Bolstad et al., 2003).

Statistical Challenges in Comparing Expression Profiles Once data has been obtained in the form of normalized expression values, the next common issue is determination of which genes have different expression levels in one sample relative to another. This requires calculation of the probability that the observed differences in expression values merely represent random samples taken from an underlying single population of possible values. If this probability is small, it is likely that the expression level of the gene really is different between the two samples (the underlying populations are separate), while if it is large, the gene likely does not have a different expression level in the two samples.



Significance of gene expression level differences Determination of genes that are expressed at different levels in different samples from microarray data presents statistical challenges. Due to the high cost of carrying out the experiments, it is usually not practical to produce many independent replicates. If one assumes that the percent error (or coefficient of variation) of each gene is completely independent, analyses using small numbers of replicates have little statistical power, so only large expression differences are called statistically significant. One way to reduce the impact of this problem might be to assume the same percent errors for all genes. However, this is not a good assumption because weak signals (genes with low expression levels) typically have larger percent errors than strong signals (Jain et al., 2003). Commonly used tests for the significance of gene expression level differences, such as t-tests, assume that the underlying population of expression values is normally distributed; however, this assumption may not be valid, and its validity cannot be tested when the number of experimental replicates is small. It is possible to use nonparametric methods, in which no particular distribution is assumed, but these methods have less statistical power than parametric methods. Another problem arises from the fact that microarray experiments are massively parallel assays. If 1000 genes are monitored and a 5% rejection rate (i.e., cutoff at P = 0.05) for each gene is applied to detect genes that are differentially expressed between two samples, on average 50 genes are falsely called differentially expressed. Multiple-testing corrections are statistical methods that increase the stringency of P-value cutoffs according to the number of genes monitored to reduce the frequency of such false-positives. They also increase the number of false negatives, thereby excluding genes that really are differentially expressed. Permutation methods (SAM) Permutation methods have been developed to respond to the statistical challenges presented by microarray data. One such method that is often used is significance analysis of microarrays (SAM; Tusher et al., 2001). In an example of SAM, an experiment with four replicate control samples and four replicate experimental samples was performed (Tusher et al., 2001). The relative difference value was calculated for each gene. The relative difference value was the difference between the mean

expression level of each gene in the experimental versus control samples, divided by the genespecific scatter, defined as the standard deviation of repeated expression measurements. To assess the significance of this relative difference, the eight data sets were then randomly assigned to two groups of four data sets each (this is permutation), and the relative difference value for each gene was similarly calculated for the permuted groups. The expected relative difference for each gene was calculated by averaging the relative difference values of the gene from all the appropriate permuted group comparisons. The difference between the relative difference value of each gene determined in the control versus experiment comparison and the expected relative difference value of the gene was used as a measure of the likelihood that apparent expression differences were real.

Examples of Common Situations Since there are practical limitations on the level of confidence that can be obtained from microarray data, it is important to have a clear idea of the purpose of the experiment and the effects of statistical limitations, and to apply statistical methods that are appropriate to the data set under study. The following examples illustrate some common situations. Example 1: Identifying genes that may be important in a biological process If genes are selected using stringent criteria to reduce false positives, very few candidate genes are obtained. If the criteria are relaxed, more genes are obtained, but these certainly include false positives. The key question is capacity for testing candidate genes. If many candidates can be tested, then it is better to use less stringent criteria and rely on downstream testing to eliminate false positives. If downstream testing is onerous, it is better to use more stringent selection criteria. Example 2: Replicate data generated by two different individuals The variance among replicate samples generated by different individuals is large relative to the variance among replicates generated by the same individual. Data generated by the two individuals should not be simply combined in an analysis. Example 3: Non-normal distributions If the distributions of values among replicate samples clearly deviate substantially from the bell shape of a normal distribution, statistical

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tests that assume an underlying normal distribution should not be used. Either the values should be transformed first so that their distribution approximates a normal distribution, or a nonparametric test should be employed.

CHOICE OF STATISTICS FOR COMPARISON


When using array platforms with singlecolor detection methods (e.g., see Affymetrix Arrays), it is possible to compare expression values directly. This is convenient when many different combinations of data sets are used for comparison. When needed, by combining expression values from two appropriate experiments, expression ratio values can be calculated. With two-color methods commonly used with spotted arrays, only ratio values can be used. There is an advantage to using expression ratio values for comparisons. Unlike expression values, ratio values are independent of technology platform, so ratios obtained from one platform can be used for comparisons with ratios obtained from a different platform; however, when using ratio values, it is very important to consider the effect on error levels. The ratios are obtained from two measurements, each with an associated error, so the percent error in the ratio is larger than the percent error in either measurement. Ratios in which the expression level in one sample is very low will generally have large errors, since weak signals tend to have large percent errors. Deriving ratios from two different arrays rather than from a direct comparison on a single array, such as deriving the ratio of C/A from the ratios B/A obtained from one array and B/C obtained from another array, further increases errors. Therefore, it is ideal to have a direct comparison of each interesting pair of samples when using a two-color method. On the other hand, if many pair-wise comparisons are needed, using a common reference sample may be more practical despite the error problem discussed above. Factors affecting the choice of suitable reference samples have been discussed elsewhere (e.g., Churchill, 2002). For compiling standardized databases for general use, data from a single color method or data from a two-color method using the same reference sample for all the arrays is more valuable than other types of ratio data due to ease of comparison among various data sets. Expression values and ratio values are often log-transformed before comparison. This is appropriate if log-transformation brings the dataset closer to a normal distribution, improv-

ing the accuracy of statistical tests based on the assumption of a normal distribution. In addition, when expression ratios are log-transformed, the characteristics of the transformed values are more intuitive. X-fold induction and repression have the same absolute value with opposite signs. Also, the scale is compressed for larger numbers. On a log2 scale, the difference between a fold change of 2 (log = 1) and a fold change of 8 (log = 3) is 2, rather than 6 on a linear scale, while the difference between a fold change of 10 (log = 3.32) and a fold change of 16 (log = 4) is 0.68, rather than 6 on a linear scale.

THE PROBLEM OF CLASSIFYING GENES INTO A SMALL NUMBER OF GROUPS Genes are often classified into two groups such as those that are induced and those that are not. Such groups of genes are sometimes used to make arguments about how many genes are common to two groups that are induced under different conditions. Imagine that genes are divided into induced and uninduced genes using either a two-fold cutoff or a probability cutoff. Typically the cutoff values lie in the tail of the distribution of the values for all the genes. Consequently, many genes have values close to the cutoff value. For example, if the cutoff value is a two-fold change, there will be many genes with values of 1.9 or 2.1. Genes with a value of 1.9 will be judged as uninduced while those with a value of 2.1 will be judged as induced. Is there really a major difference between a 1.9-fold change and a 2.1-fold change? Among the genes that pass the cutoff, there will be large variations in the extent to which they pass, yet they are all classified in the same group. For example, a gene that shows a 2.1-fold change is grouped together with a gene that shows a 200-fold change. Are these fold-changes really similar? These problems become greater when two sets of induced genes are compared to determine the set of genes induced in common by two different treatments. The impact of these limitations on conclusions drawn from microarray data should be kept in mind.

ADVANTAGES OF COMMERCIAL SOFTWARE Several expensive software packages for microarray analysis are commercially available. Programs that will perform most of the analytical methods implemented in these packages are freely available somewhere on the web, so why should anyone spend a lot of money on the



commercial products? First, such products have the various methods put together nicely. It is quick and easy to apply different analytical methods and combine or compare the results. Second, they have integrated database management mechanisms, which can also take care of database security and sharing issues. Third, they are designed to be highly interactive. Wellorganized, highly interactive databases provide access to information that helps the investigator make subjective decisions about how to integrate different types of information. Fourth, they offer many ways to visualize the data and the results of analyses. People have excellent visual pattern recognition abilities, so visualization of data is a powerful tool aiding comprehension of large data sets. Fifth, they often offer ways to record a series of analyses applied to the data. Such a record is very important because various criteria used in analyses could be subjectively decided. In addition, such a recorded series of analyses can be used as a macro, so the same series of analyses are easily applied to different data sets. In using sophisticated software tools, it is necessary to understand the analytical tools being applied to avoid drawing faulty conclusions. For example, when using false color to display differences in gene expression levels, the eye is drawn to the boundaries between different colors, and one tends to conclude that genes in different color groups have very different expression levels, when they may actually be quite similar.

who produce data through carefully-designed and executed experiments thus have the opportunity to benefit the larger scientific community as well as to further their own research goals.

COMPARISON OF MULTIPLE PROFILES In an upcoming supplement, another aspect of expression profile data analysis will be considered: comparison of multiple profiles. Similarities among profiles can be used in a number of ways. For example, tumors that respond similarly to treatments can have similar expression profiles, and this can be used to determine which treatment may be most effective (Lapointe et al., 2004). Also, expression profiles of mutants can be used to predict the nature of the defects in the mutants (e.g., Hughes et al., 2000; Glazebrook et al., 2003). Detection of similarities among expression profiles is a problem of pattern-recognition among distributions of points in n-dimensional space, where each point represents an expression profile and n is the number of genes represented in each profile. While people have difficulty recognizing patterns in spaces of more than three dimensions, computer algorithms can handle as many dimensions as needed. Many computational methods for pattern recognition have developed, and several are routinely used for analysis of expression profiling data. These methods, together with methods under development, may prove to be even more useful in the future.

LITERATURE CITED SUMMARY Expression profile data obtained from microarrays can be enormously useful in addressing a variety of biological questions, including genome-scale questions that were previously unapproachable. Maximizing the potential of microarray experiments requires attention to aspects of experimental design and data analysis. Experiments should be designed so as to minimize systematic errors and errors arising in the process of data analysis. Good design can also make it possible to determine how much of the variation among replicates arises from various sources, such as technical variation or biological variation. When analyzing data, it is important to understand the limitations of the analytical methods used to avoid drawing erroneous conclusions. Public databases consisting of microarray data will be very useful for a variety of large-scale analyses, provided that the data sets are of high quality. Investigators

Baugh, L.R., Hill, A.A., Brown, E.L., and Hunter, C.P. 2001. Quantitative analysis of mRNA amplification by in vitro transcription. Nucl. Acids Res. 29:e29. Bolstad, B.M., Irizarry, R.A., Åstrand, M., and Speed, T.P. 2003. A comparison of normalization methods for high density oligonucleotide array data based on variance and bias. Bioinformatics 19:185-193. Brenner, S., Johnson, M., Bridgham, J., Golda, G., Lloyd, D.H., Johnson, D., Luo, S., McCurdy, S., Foy, M., Ewan, M., et al. 2000. Gene expression analysis by massively parallel signature sequencing (MPSS) on microbead arrays. Nat. Biotechnol. 18:630-634. Churchill, G.A. 2002. Fundamentals of experimental design for cDNA microarrays. Nat. Genet. 32:490-495. Glazebrook, J., Chen, W., Estes, B., Chang, H.-S., Nawrath, C., Métraux, J.-P., Zhu, T., and Katagiri, F. 2003. Topology of the network integrating salicylate and jasmonate signal transduction derived from global expression phenotyping. Plant J. 34:217-228.

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Hughes, T.R., Marton, M.J., Jones, A.R., Roberts, C.J., Stoughton, R., Armour, C.D., Bennett, H.A., Coffey, E., Dai, H., He, Y.D., et al. 2000. Functional discovery via a compendium of expression profiles. Cell 102:109-126. Iscove, N.N., Barbara, M., Gu, M., Gibson, M., Modi, C., and Winegarden, N. 2002. Representation is faithfully preserved in global cDNA amplified exponentially from sub-picogram quantities of mRNA. Nat. Biotechnol. 20:940943. Jain, N., Thatte, J., Braciale, T., Ley, K., O’Connell, M., and Lee, J.K. 2003. Local-pooled-error test for identifying differentially expressed genes with a small number of replicated microarrays. Bioinformatics 19:1945-1951. Kim, S.K., Lund, J., Kiraly, M., Duke, K., Jiang, M., Stuart, J.M., Eizinger, A., Wylie, B.N., and Davidson, G.S. 2001. A gene expression map for Caenorhabditis elegans. Science 293:20872092. Lapointe, J., Li, C., Higgins, J.P., van de Rijn, M., Bair, E., Montgomery, K., Ferrari, M., Egevad, L., Rayford, W., Bergerheim, U., et al. 2004. Gene expression profiling identifies clinically relevant subtypes of prostate cancer. Proc. Natl. Acad. Sci. U.S.A. 101:811-816. Liang, P. and Pardee, A.B. 1992. Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science 257:967971. Lockhart, D.J., Dong, H., Byrne, M.C., Follettie, M.T., Gallo, M.V., Chee, M.S., Mittmann, M., Wang, C., Kobayashi, M., Horton, H., and Brown, E.L. 1996. Expression monitoring by hybridization to high-density oligonucleotide arrays. Nat. Biotechnol. 14:1675-1680. Nuwaysir, E.F., Huang, W., Albert, T.J., Singh, J., Nuwaysir, K., Pitas, A., Richmond, T., Gorski, T., Berg, J.P., Ballin, J., et al. 2002. Gene expres-

sion analysis using oligonucleotide arrays produced by maskless photolithography. Genome Res. 12:1749-1755. Ohyama, H., Zhang, X., Kohno, Y., Alevizos, I., Posner, M., Wong, D.T., and Todd, R. 2000. Laser capture microdissection-generated target sample for high-density oligonucleotide array hybridization. Biotechniques 29:530-536. Quackenbush, J. 2002. Microarray data normalization and transformation. Nature Genet. 32:496501. Tao, Y., Xie, Z., Chen, W., Glazebrook, J., Chang, H.-S., Han, B., Zhu, T., Zou, G., and Katagiri, F. 2003. Quantitative nature of Arabidopsis responses during compatible and incompatible interactions with the bacterial pathogen Pseudomonas syringae. Plant Cell 15:317-330. Tusher, V.G., Tibshirani, R., and Chu, G. 2001. Significance analysis of microarrays applied to the ionizing radiation response. Proc. Natl. Acad. Sci. U.S.A. 98:5116-5121. Velculescu, V.E., Zhang, L., Vogelstein, B., and Kinzler, K.W. 1995. Serial analysis of gene expression. Science 270:484-487. Wang, E., Miller, L.D., Ohnmacht, G.A., Liu, E.T., and Marincola, F.M. 2000. High-fidelity mRNA amplification for gene profiling. Nat. Biotechnol. 18:457-459. Yang, L., Tran, D.K., and Wang, X. 2001. BADGE, BeadsArray for the Detection of Gene Expression, a high-throughput diagnostic bioassay. Genome Res. 11:1888-1898.

Contributed by Fumiaki Katagiri and Jane Glazebrook University of Minnesota St. Paul, Minnesota




CHAPTER 23 Manipulating the Mouse Genome INTRODUCTION

D

efining the role of a mammalian nucleic acid segment can be a difficult process. The first steps involve mRNA identification, gene cloning, and nucleic acid sequence determination. A variety of cell-free and cultured cell systems have been developed (see Chapters 8 to 22, 24) that can provide important clues to the function of a particular DNA sequence in regulating gene transcription or RNA translation. Many segments of DNA appear inert or function inappropriately in these model systems; thus, an important test of any functional model of a DNA segment is to characterize the phenotype of a viable organism carrying the sequence in its genome. Today the tools exist for manipulating DNA segments in a variety of mammals, from mice and rats to sheep and cows. However, the mouse, because of its small size (30 g) and short life cycle (about 8 weeks) from birth to the production of progeny, has been selected for the vast majority of laboratory experiments. Thus, this chapter focuses on the manipulation of the mouse genome. Two general methods are available for the introduction and modification of mouse genomic DNA sequence. Transgenic mice are produced by the injection of one or more transgenes (usually a DNA segment bearing its own promoter) into the pronucleus of a fertilized mouse oocyte, which, after reimplantation in a foster mother, gives birth to a transgenic mouse bearing one to several hundred copies of the transgene. Embryonic stem cell–derived mice are produced by the introduction of embryonic stem cells into the blastocyst, which is reimplanted in a foster mother, where it goes through normal mouse development, producing a mouse pup. Transgenic mice are useful for testing the overexpression of a gene segment while embryonic stem–cell derived mice are generally useful for defining the phenotype of mice lacking a gene segment. Both types of mice can be used to express altered protein sequences; transgenic mice will usually overproduce the altered protein, while embryonic stem cell–derived mice will usually express normal levels of protein. In this chapter we describe the production of embryonic stem cell derived–mice.

The production of embryonic stem cell–derived mice can be thought of as a three-stage process that normally takes about 6 months. These steps involve the production of a targeting vector (UNIT 23.1), the introduction of DNA sequences into the embryonic stem cell genome by homologous recombination (UNIT 23.5) and eventually to the production of genetically altered mice derived from embryonic stem cells. A critical part of this procedure is the maintenance of the cells. Embryonic stem cells are cultured on feeder cells and require constant attention; these procedures are described in UNITS 23.2 & 23.3. A procedure describing the derivation of embryonic stem cells is described in UNIT 23.4. After the genetically engineered embryonic stem cells are obtained they are injected into mouse blastocysts, which are then re-implanted into a foster mother that eventually gives birth to chimeric mice composed of wild-type cells and ES-derived cells (UNIT 23.7). Chimeric mice are then mated to wild-type mice and eventually mice derived from the ES cell genome are obtained. The genetically engineered mice are now available for physiologic and biochemical evaluation. The production and management of a colony of genetically engineered mice requires careful planning (UNIT 23.8) in order to have adequate numbers of mice of appropriate ages for study. Manipulating the Mouse Genome Contributed by J.G. Seidman Current Protocols in Molecular Biology (2003) 23.0.1-23.0.2 Copyright © 2003 by John Wiley & Sons, Inc.


The function of genes expressed in embryonic stem cells can be assessed directly in these cells. In UNIT 23.5, procedures for producing a single knockout or heterozygous cell line by homologous recombination are described. Homozygous (double-knockout) cells can be produced in tissue culture and these cells can be a useful tool for analyzing gene function in embryonic stem cells (UNIT 23.6). Taken together, these steps provide a powerful method for introducing a wide variety of specific changes into murine embryonic stem cells and eventually into the genome of a mouse. J.G. Seidman

Introduction



Overview of Gene Targeting by Homologous Recombination The analysis of mutant organisms and cell lines has been important in determining the function of specific proteins. Until recently, mutants were produced by mutagenesis followed by selection for a particular phenotypic change. Recent technological advances in gene targeting by homologous recombination in mammalian systems enable the production of mutants in any desired gene (Mansour, 1990; Robertson, 1991; Zimmer, 1992). This technology can be used to produce mutant mouse strains and mutant cell lines. Because most mammalian cells are diploid, they contain two copies, or alleles, of each gene encoded on an autosomal (nonsex) chromosome. In most cases, both alleles must be inactivated to produce a discernible phenotypic change in a mutant. The conversion from heterozygosity to homozygosity is accomplished by breeding in the case of mouse strains and by direct selection in cell lines. Bacteriophage recombinases such as Cre and its recognition sequence, loxP, have also allowed spatial control of knockouts. Another recombinase system, the yeast Flp/FRT system, can also be used (Fiering et al., 1993, 1999). The control can function along actual spatial coordinates when a viral gene transfer system is used, or in a cell type– or tissue-specific fashion when restricted promoters are employed. Adding temporal regulation of Cre, such as that achievable with the tetracycline regulatable system (UNIT 16.14), allows temporal control as well. To produce a mutant mouse strain by homologous recombination, two major elements are needed. An embryonic stem (ES) cell line capable of contributing to the germ line, and a targeting construct containing target-gene sequences with the desired mutation. Maintaining ES cells in their undifferentiated state is a major task during gene targeting (UNIT 23.3). This usually is accomplished by growing cells on a layer of feeder cells (UNIT 23.2). The targeting construct is then transfected into cultured ES cells (see UNIT 23.5). ES cell lines are derived from the inner cell mass of a blastocyst-stage embryo. Homologous recombination occurs in a small number of the transfected cells, resulting in introduction of the mutation present in the targeting construct into the target gene. Once identified, mutant ES cell clones can be Contributed by Richard Mortensen Current Protocols in Molecular Biology (2000) 23.1.1-23.1.11 Copyright © 2003 by John Wiley & Sons, Inc.

UNIT 23.1

microinjected into a normal blastocyst in order to produce a chimeric mouse. Because many ES cell lines retain the ability to differentiate into every cell type present in the mouse, the chimera can have tissues, including the germ line, with contribution from both the normal blastocyst and the mutant ES cells. Breeding germ-line chimeras yields animals that are heterozygous for the mutation introduced into the ES cell, and that can be interbred to produce homozygous mutant mice. Homologous recombination can also be used to produce homozygous mutant cell lines (see UNIT 23.6). Previously, inactivation of both alleles of a gene required two rounds of homologous recombination and selection (te Riele et al., 1990; Cruz et al., 1991; Mortensen et al., 1991). Now, however, inactivation of both alleles of many genes requires only a single round of homologous recombination using a single targeting construct (Mortensen et al., 1992). The homozygous mutant cells can then be analyzed for phenotypic changes to determine the function of the gene.

ANATOMY OF TARGETING CONSTRUCTS Two basic configurations of constructs are used for homologous recombination—insertion constructs and replacement constructs (Fig. 23.1.1). Each can be used for different purposes in specific situations, as discussed below. The insertion construct contains a region of homology to the target gene cloned as a single continuous sequence, and is linearized by cleavage of a unique restriction site within the region of homology. Homologous recombination introduces the insertion construct sequences into the homologous site of the target gene, interrupting normal target-gene structure by adding sequences. As a result, the normal gene can be regenerated from the mutated target gene by an intrachromosomal recombination event. The replacement construct is the second, more commonly used construct. It contains two regions of homology to the target gene located on either side of a mutation (usually a positive selectable marker; see below). Homologous recombination proceeds by a double cross-over event that replaces the target-gene sequences with the replacement-construct sequences. Be-

Manipulating the Mouse Genome


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Figure 23.1.1 Two configurations of constructs used for homologous recombination. Numbers indicate target-gene sequences in the genome. An asterisk indicates homologous target-gene sequences in the construct. Replacement constructs substitute their sequences (2*, neo, and 3*) for the endogenous target-gene sequences (2 and 3). Insertion constructs add their sequences (2*, neo, and 3*) to the endogenous target gene, resulting in tandem duplication and disruption of the normal gene structure.

cause no duplication of sequences occurs, the normal gene cannot be regenerated.

METHODS OF ENRICHMENT FOR HOMOLOGOUS RECOMBINANTS Positive Selection by Drug-Resistance Gene Nearly all constructs used for homologous recombination rely on the positive selection of a drug-resistance gene (e.g., neomycin or neo) that is also used to interrupt and mutate the target gene. When either insertion or replacement constructs are linearized, the drug-resistance gene is flanked by two regions of homology to the target gene. Selection of the cells using drugs (e.g., G418) eliminates the great majority of cells that have not stably incorporated the construct (see UNIT 9.5). However, in many of the surviving clones the construct has incorporated into the genome not by homologous recombination but rather through random integration. Therefore, methods to enrich for homologous recombinant clones have been developed. Overview of Gene Targeting by Homologous Recombination

Positive-Negative Selection The most commonly used method for eliminating cells in which the construct integrated

into the genome randomly, thus further enriching for homologous recombinants, is known as positive-negative selection. It is only applicable to replacement constructs (Fig. 23.1.2; Mansour et al., 1988). In these constructs, a negative selectable marker (e.g., herpes simplex virus thymidine kinase, HSV-TK) is included outside the region of homology to the target gene. In the presence of the TK gene, the cells are sensitive to acyclovir and its analogs (e.g., gancyclovir, GANC). The HSV-TK enzyme activates these drugs, resulting in their incorporation into growing DNA, causing chain termination and cell death. During homologous recombination, sequences outside the regions of homology to the target gene are lost due to crossing over. In contrast, during random integration all sequences in the construct tend to be retained because recombination usually occurs at the ends of the construct. The presence of the TK gene can be selected against by growing the cells in gancyclovir; the homologous recombinants will be G418-resistant and gancyclovirresistant, whereas clones in which the construct integrated randomly will be G418-resistant and gancyclovir-sensitive. In some cases, TK is inactivated without homologous recombination; thus, the gancyclovir-resistant clones



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Figure 23.1.2 Enrichment for homologous recombinants by positive-negative selection using the TK gene. Homologous recombination involving cross-overs on either side of the neo gene results in loss of the TK gene. Random integration tends to preserve the TK gene. The presence of TK can be selected against because any cell expressing the gene will be killed by gancyclovir (GANC). Although both homologous recombinants and clones in which the construct integrated randomly are G418-resistant, only homologous recombinants are gancyclovir-resistant. The construct is shown linearized so that the plasmid vector sequences remain attached to the TK gene. This configuration helps preserve the integrity of the TK gene. The superscript R denotes resistance and the superscript S denotes sensitivity.

must be screened to identify the true homologous recombinants. Other markers that are lethal to cells have also been used instead of TK and gancyclovir (e.g., diphtheria toxin; Yagi et al., 1990).

Endogenous Promoters Constructs that rely on an endogenous promoter to express the positive selectable marker can also give enrichment of homologous recombinants (Fig. 23.1.3), but can only be used if the gene of interest is expressed in the cell line. They contain the coding region of a selectable marker (e.g., neo) but lack a promoter for the marker. The coding sequence for the marker usually interrupts, and is in frame with, an exon of the target gene. Thus, when homologous recombination occurs, a fusion protein is produced driven by the endogenous target-gene

promoter. In contrast, when random integration occurs, the selectable-marker protein is not usually produced. Therefore, homologous recombinants are G418-resistant, whereas cells in which the construct integrated randomly are G418-sensitive. Constructs containing a promoterless selectable marker can be constructed in either replacement or insertion structure and can result in dramatic enrichment for homologous recombinants.

TYPES OF MUTATIONS Gene Inactivation Homologous recombination has most often been used to completely inactivate a gene (commonly termed “knockout”). Usually, an exon encoding an important region of the protein (or an exon 5′ to that region) is interrupted by a



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Figure 23.1.3 Enrichment for homologous recombinants using a positive selectable marker (neo) lacking a promoter. Clones in which integration of the construct provides an endogenous promoter to drive neo expression will be G418-resistant. The construct is designed so that homologous recombination will provide a promoter leading to neo expression, whereas random integration will most likely not provide a promoter, thus precluding neo expression.

Overview of Gene Targeting by Homologous Recombination

positive selectable marker (e.g., neo), preventing the production of normal mRNA from the target gene and resulting in inactivation of the gene. A gene may also be inactivated by creating a deletion in part of a gene, or by deleting the entire gene. By using a construct with two regions of homology to the target gene that are far apart in the genome, the sequences intervening the two regions can be deleted. Up to 15 kb have been deleted in this way; thus, many genes could be completely eliminated (Mombaerts et al., 1991). Gene inactivations may also be controlled using the Cre/loxP recombinase system either spatially, as in cell type– or tissue-specific knockout, or temporally, through control of the activity or expression of the recombinase (see Cre/loxP System, below). Mutations can be introduced that have mul-

tiple purposes. Homologous recombination has been used to introduce a replacement construct containing the coding sequence of β-galactosidase in frame with the 5′ end of the target gene. Downstream of the lacZ gene is a positive selectable marker driven by a heterologous promoter (Fig. 23.1.4). This construct not only disrupts target-gene function but also expresses a fusion protein with β-galactosidase activity, and thus can be used to monitor the activity of the endogenous gene’s promoter in various tissues during development (Mansour et al., 1990).

Subtle Gene Mutations Homologous recombination can also be used to introduce subtle mutations in a gene. One method is analogous to a method in yeast called transplacement or allele replacement (UNIT 13.10). It is called “hit and run” (Hasty et



lacZ

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Figure 23.1.4 LacZ reporter construct for gene targeting. This construct has two purposes: first, to disrupt the target gene and, second, to express the lacZ gene as a marker to monitor activity of the endogenous target gene’s promoter.

al., 1991) because duplications are introduced into the target gene and then removed. An insertion construct containing both positive and negative selectable markers (e.g., neo and TK) is used to introduce a duplication that contains a subtle mutation, such as a point mutation, into the target gene sequence (Fig. 23.1.5). After selection for integration of the construct using the positive selectable marker (e.g., G418), homologous recombinants are identified by screening. A homologous recombinant clone is cultured and then the presence of the negative selectable marker is selected against (e.g., selection against TK using gancyclovir). This selects for an intrachromosomal recombination that eliminates the target-gene duplications and the selectable markers but leaves the mutant target-gene sequences substituting for the nor-

mal target-gene sequences. Surviving clones are screened for the correct intrachromosonal rearrangements, leaving the desired mutation. A second method of introducing subtle mutations into a gene is to insert the mutation by homologous recombination and then use the Cre/loxP system to remove the selectable marker.

CRE/loxP SYSTEM The Cre/loxP system is derived from the bacteriophage P1. The recombinase Cre acts on the DNA site loxP. If there are two loxP sites in the same orientation near each other, Cre can act to loop out the sequence between the two sites, leaving a single loxP site in the original DNA and a second loxP in a circular piece of DNA containing the intervening sequence.



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Figure 23.1.5 Allele replacement “hit and run.” Cells are cultured in G418 to select for the integration of neo, then homologous recombinants are identified by screening and are cultured in gancyclovir (GANC) to select against the presence of the TK gene. This strategy may yield a reconstituted gene containing the subtle mutation present in the construct (indicated by the dark bar and †). Because intrachromosomal recombination may result in the loss of the subtle mutation, its presence must be verified (e.g., by a change in restriction site).


Therefore, a properly designed targeting construct containing loxP sites can be used for introducing subtle mutations or for a temporally or spatially controlled knockout (for a review of the control of transgenes, see Sauer, 1993). Other recombinase systems, such as the Flp/FRT system, can be similarly useful (Fiering et al., 1995; Vooijs et al., 1998).

Removing the Positive Selectable Marker Although many gene inactivation approaches involving homologous recombination still use constructs that leave the positive selectable marker in the genomic DNA, it has become increasingly clear that this can cause a number of unanticipated effects. For example, the presence of the neo gene, often with its own promoter, can alter the expression of neighboring loci (Olson et al., 1996; Pham et al., 1996).



∗ Cre expression

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Figure 23.1.6 Using the Cre/loxP system to introduce subtle mutations. The subtle mutation is introduced along with the selectable marker in the targeting vector. The selectable marker is then removed by transient expression of Cre, which leaves only the small loxP site in the genome in a silent location.

This can be a particular problem in gene clusters where neighboring genes are in the same family, since the genes affected may have similar or identical functions. As a result, slight differences in targeting constructs have led to marked differences in phenotype. If the targeting construct includes loxP sites flanking the neo gene, then neo can be removed after targeting by transient expression of the Cre recombinase (as discussed in UNIT 9.5 and Fig. 23.1.3). This will leave the small loxP site in the genomic DNA, but the construct can be engineered so that this is in an innocuous location, such as an intron. Although theoretically even a loxP site could cause alterations in the expression of neighboring genes, no such cases have yet been reported. The efficiency of Cre recombination from transient expression reported in the literature varies widely, from ∼2% to ∼15% (Sauer and Henderson, 1989; Abuin and Bradley, 1996). This rate should be distin-

guished from the efficiency of Cre recombination in vivo, where the expression of Cre is derived from sequences integrated into the genome and therefore will show longer-lasting expression in nearly all cases.

Introduction of Subtle Mutations Using Cre/loxP The strategy described in the previous section involves introducing subtle mutations by first duplicating sequences and then screening for intrachromosomal recombination that removes the redundant sequences and leaves the mutation. A limitation to this approach is that the second homologous recombination event occurs only infrequently. A more efficient method is to use a replacement construct containing the subtle mutation and then remove the positive selectable marker, which is flanked by loxP sites, using the Cre recombinase system (Fig. 23.1.6). This is identical in effect to re-



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Cre expression in culture

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Nonhomologous sequences positive selectable marker genomic sequences lox sequence vector sequences promoter sequence

Figure 23.1.7 Conditional gene targeting using the Cre/loxP system. The targeting vector contains three loxP sites that flank the regions of the gene to be removed and the positive selectable marker neo. After homologous recombination is obtained, the selectable marker is excised from the gene by transient expression of Cre. The correct recombination is identified by screening by Southern analysis or PCR. The mutant ES cells are then used to produce a transgenic animal, which is finally bred to an animal line expressing Cre under either temporal or spatial control. Cre can also be ubiquitously expressed to obtain a knockout in all tissues, including the germ line.

moving the neo locus after gene inactivation, except that instead of an inactive gene, the replaced sequences contain a subtly mutated version.

Spatial Control of Knockout Overview of Gene Targeting by Homologous Recombination

Spatially controlled targeted gene inactivations can be performed in two ways. The most common makes use of cell type–specific promoters (sometimes called tissue-specific promoters, even though tissues are actually made

of a number of different cell types). This approach begins with the creation of a transgenic animal that expresses Cre in only some cells using a cell type–restricted promoter. A second transgenic animal line is then created by homologous recombination that contains loxP sites flanking a portion of the gene that is critical for activity, typically important exons (Fig. 23.1.7). Initially there are three loxP sites flanking this important gene region and the selectable marker. After homologous recombi-



nation has been verified, Cre is transiently expressed, and loops out regions of DNA between pairs of loxP sites. The resultant colonies are screened for the desired recombination (loss of the selectable marker but retention of all regions of the gene). Depending on the frequency of recombination at the site, it may be useful to use a construct that contains a negative selectable marker (such as cytosine deaminase in the example shown in Fig. 23.1.2) between the loxP sites along with the positive selectable marker (also see UNIT 9.5). In this way cells that have lost the markers can be selected. The targeted line will have normal expression of the targeted gene, since its only modification is the presence of loxP sites in innocuous sites (e.g., introns). When the two lines are bred together, the Cre recombinase will loop out the DNA—inactivating the gene—only in those cells where it is expressed. In this way, tissue-specific knockouts of a number of genes have been generated (Gu et al., 1994; Agah et al., 1997). The method also has the advantage that, once a transgenic line is generated with the desired restricted expression of Cre, the approach can be applied to a number of targeted lines. In addition, it is not necessary to make separate constructs for a restricted and a complete knockout, since Cre-expressing lines have been made that will produce rearrangement in all tissues when bred to the targeted line (Schwenk et al., 1995). Another way of spatially controlling knockout is to use an expression system for Cre that can be applied to absolute location. In some cases, no restricted expression pattern is known for a gene that matches the desired spatial alteration; in others, the site may be particularly amenable to viral manipulation (as with an epithelial or endothelial surface) or accessible by direct injection (such as sterotactic injection of the central nervous system). By using a viral vector to express the Cre protein, it is possible to obtain knockouts that are spatially limited by the viral infection. This strategy has been applied to a number of tissues including the brain, liver, colon, and heart (Rohlmann et al., 1996; Wang et al., 1996; Agah et al., 1997; Shibata et al., 1997; van der Neut, 1997).

Temporal Control of Knockout In many cases the phenotype of interest is in the adult animal but, because the gene is necessary for development, no adult animals are obtained. Delaying the expression of Cre activity until the animal is an adult would allow normal development, and then the knockout

could be created in the adult (Rajewsky et al., 1996). This can be accomplished by using a conditional expression system (e.g., the tet-on, tet-off, or ecdysone systems; see UNIT 16.14 and St-Onge et al., 1996) or other inducible system (such as an interferon-inducible promoter; Kuhn et al., 1995) to express Cre at the proper time. This would, however, require the construction of animals containing three transgenes. Another approach that has been used is the creation of a fusion protein with either a modified estrogen receptor (Feil et al., 1996, 1997; Zhang et al., 1996; Brocard et al., 1997) or a modified glucocorticoid receptor (Brocard et al., 1998). These fusion proteins are inactive for recombination until the appropriate ligand is added, allowing temporal control in an animal with only transgenes. The Flp/FRT recombinase system can be used in an analogous way. Combination of the two systems can allow the production of complex schemes for gene mutation.

LITERATURE CITED Abuin, A. and Bradley, A. 1996. Recycling selectable markers in mouse embryonic stem cells. Mol. Cell. Biol. 16:1851-1856. Agah, R., Frenkel, P.A., French, B.A., Michael, L.H., Overbeek, P.A., and Schneider, M.D. 1997. Gene recombination in postmitotic cells. Targeted expression of Cre recombinase provokes cardiac-restricted, site-specific rearrangement in adult ventricular muscle in vivo. J. Clin. Invest. 100:169-179. Brocard, J., Warot, X., Wendling, O., Messaddeq, N., Vonesch, J.L., Chambon, P., and Metzger, D. 1997. Spatio-temporally controlled site-specific somatic mutagenesis in the mouse. Proc. Natl. Acad. Sci. U.S.A. 94:14559-14563. Brocard, J., Feil, R., Chambon, P., and Metzger, D. 1998. A chimeric Cre recombinase inducible by synthetic, but not by natural ligands of the glucocorticoid receptor. Nucl. Acids Res. 26:40864090. Cruz, A., Coburn, C.M., and Beverley, S.M. 1991. Double targeted gene replacement for creating null mutants. Proc. Natl. Acad. Sci. U.S.A. 88:7170-7174. Feil, R., Brocard, J., Mascrez, B., LeMeur, M., Metzger, D., and Chambon, P. 1996. Ligand-activated site-specific recombination in mice. Proc. Natl. Acad. Sci. U.S.A. 93:10887-10890. Feil, R., Wagner, J., Metzger, D., and Chambon, P. 1997. Regulation of Cre recombinase activity by mutated estrogen receptor ligand-binding domains. Biochem. Biophys. Res. Commun. 237:752-757.



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Fiering, S., Kim, C.G., Epner, E.M., and Groudine, M. 1993. An “in-out” strategy using gene targeting and FLP recombinase for the functional dissection of complex DNA regulatory elements: Analysis of the β-globin locus control region. Proc. Natl. Acad. Sci. U.S.A. 90:8469-8473. Fiering, S., Epner, E., Robinson, K., Zhuang, Y., Telling, A., Hu, M., Martin, D.I., Enver, T., Ley, T.J., and Groudine, M. 1995. Targeted deletion of 5′HS2 of the murine β-globin LCR reveals that it is not essential for proper regulation of the β-globin locus. Genes Dev. 9:2203-2213. Fiering S., Bender, M.A., and Groudine, M. 1999. Analysis of mammalian cis-regulatory DNA elements by homolgous recombination. Methods Enzymol. 306:42-66. Gu, H., Marth, J.D., Orban, P.C., Mossmann, H., and Rajewsky, K. 1994. Deletion of a DNA polymerase β gene segment in T cells using cell type– specific gene targeting. Science 265:103-106. Hasty, P., Ramirez, S.R., Krumlauf, R., and Bradley, A. 1991. Introduction of a subtle mutation into the Hox-2.6 locus in embryonic stem cells [published erratum appears in Nature, 1991, 353:94]. Nature 350:243-246. Kuhn, R., Schwenk, F., Aguet, M., and Rajewsky, K. 1995. Inducible gene targeting in mice. Science 269:1427-1429. Mansour, S.L. 1990. Gene targeting in murine embryonic stem cells: Introduction of specific alterations into the mammalian genome. Genet. Anal. Tech. Appl. 7:219-227. Mansour, S.L., Thomas, K.R., and Capecchi, M.R. 1988. Disruption of the proto-oncogene int-2 in mouse embryo-derived stem cells: A general strategy for targeting mutations to nonselectable genes. Nature 336:348-352. Mansour, S.L., Thomas, K.R., Deng, C.X., and Capecchi, M.R. 1990. Introduction of a lacZ reporter gene into the mouse int-2 locus by homologous recombination. Proc. Natl. Acad. Sci. U.S.A. 87:7688-7692. Mombaerts, P., Clarke, A.R., Hooper, M.L., and Tonegawa, S. 1991. Creation of a large genomic deletion at the T-cell antigen receptor betasubunit locus in mouse embryonic stem cells by gene targeting. Proc. Natl. Acad. Sci. U.S.A. 88:3084-3087. Mortensen, R.M., Zubiaur, M., Neer, E.J., and Seidman, J.G. 1991. Embryonic stem cells lacking a functional inhibitory G-protein subunit (alpha i2) produced by gene targeting of both alleles. Proc. Natl. Acad. Sci. U.S.A. 88:7036-7040. Mortensen, R.M., Conner, D.A., Chao, S., Geisterfer, L.A., and Seidman, J.G. 1992. Production of homozygous mutant ES cells with a single targeting construct. Mol. Cell. Biol. 12:2391-2395.


Olson, E.N., Arnold, H.H., Rigby, P.W., and Wold, B.J. 1996. Know your neighbors: Three phenotypes in null mutants of the myogenic bHLH gene MRF4. Cell 85:1-4.

Pham, C.T., MacIvor, D.M., Hug, B.A., Heusel, J.W., and Ley, T.J. 1996. Long-range disruption of gene expression by a selectable marker cassette. Proc. Natl. Acad. Sci. U.S.A. 93:1309013095. Rajewsky, K., Gu, H., Kuhn, R., Betz, U.A., Muller, W., Roes, J., and Schwenk, F. 1996. Conditional gene targeting. J. Clin. Invest. 98:600-603. Robertson, E.J. 1991. Using embryonic stem cells to introduce mutations into the mouse germ line. Biol. Reprod. 44:238-245. Rohlmann, A., Gotthardt, M., Willnow, T.E., Hammer, R.E., and Herz, J. 1996. Sustained somatic gene inactivation by viral transfer of Cre recombinase. Nat. Biotechnol. 14:1562-1565. Sauer, B. 1993. Manipulation of transgenes by sitespecific recombination: Use of Cre recombinase. Methods Enzymol. 225:890-900. Sauer, B. and Henderson, N. 1989. Cre-stimulated recombination at loxP-containing DNA sequences placed into the mammalian genome. Nucl. Acids Res. 17:147-161. Schwenk, F., Baron, U., and Rajewsky, K. 1995. A cre-transgenic mouse strain for the ubiquitous deletion of loxP-flanked gene segments including deletion in germ cells. Nucl. Acids Res. 23:5080-5081. Shibata, H., Toyama, K., Shioya, H., Ito, M., Hirota, M., Hasegawa, S., Matsumoto, H., Takano, H., Akiyama, T., Toyoshima, K., Kanamaru, R., Kanegae, Y., Saito, I., Nakamura, Y., Shiba, K., and Noda, T. 1997. Rapid colorectal adenoma formation initiated by conditional targeting of the Apc gene. Science 278:120-123. St-Onge, L., Furth, P.A., and Gruss, P. 1996. Temporal control of the Cre recombinase in transgenic mice by a tetracycline responsive promoter. Nucl. Acids Res. 24:3875-3877. te Riele, H., Maandag, E.R., Clarke, A., Hooper, M., and Berns, A. 1990. Consecutive inactivation of both alleles of the pim-1 proto-oncogene by homologous recombination in embryonic stem cells. Nature 348:649-651. van der Neut, R. 1997. Targeted gene disruption: Applications in neurobiology. J. Neurosci. Methods 71:19-27. Vooijs, M., van der Valk, M., te Riele, H., and Berns, A. 1998. Flp-mediated tissue-specific inactivation of the retinoblastoma tumor suppressor gene in the mouse. Oncogene 17:1-12. Wang, Y., Krushel, L.A., and Edelman, G.M. 1996. Targeted DNA recombination in vivo using an adenovirus carrying the cre recombinase gene. Proc. Natl. Acad. Sci. U.S.A. 93:3932-3936. Yagi, T., Ikawa, Y., Yoshida, K., Shigetani, Y., Takeda, N., Mabuchi, I., Yamamoto, T., and Aizawa, S. 1990. Homologous recombination at c-fyn locus of mouse embryonic stem cells with use of diphtheria toxin A-fragment gene in negative selection. Proc. Natl. Acad. Sci. U.S.A. 87:9918-9922.



Zhang, Y., Riesterer, C., Ayrall, A.M., Sablitzky, F., Littlewood, T.D., and Reth, M. 1996. Inducible site-directed recombination in mouse embryonic stem cells. Nucl. Acids Res. 24:543-548. Zimmer, A. 1992. Manipulating the genome by homologous recombination in embryonic stem cells. Annu. Rev. Neurosci. 15:115-137.

Contributed by Richard Mortensen University of Michigan Medical Center Ann Arbor, Michigan



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Mouse Embryo Fibroblast (MEF) Feeder Cell Preparation

UNIT 23.2

The production of mouse mutants using homologous recombination and blastocystmediated transgenesis requires the maintenance of mouse embryonic stem (ES) cells in an undifferentiated state. Many investigators rely on feeder layers to prevent ES cell differentiation; feeder layers prepared from mitotically inactivated primary mouse embryo fibroblasts (MEFs) are used most commonly. This unit describes a simple method to isolate and store MEFs (Basic Protocol 1 and Support Protocol) and two common techniques for mitotic inactivation: γ-irradiation (Basic Protocol 2) and mitomycin C treatment (Alternate Protocol). NOTE: All cell culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. ISOLATION OF PRIMARY MOUSE EMBRYO FIBROBLASTS MEFs are isolated from 12.5 to 13.5 postcoitum (p.c.) mouse embryos. The embryos are dissociated and then trypsinized to produce single-cell suspensions. After expansion, aliquots can be frozen and stored in liquid nitrogen indefinitely. Alternatively, MEFs suitable for ES cell culture may be obtained from commercial sources (see http://www.biosupplynet.com for a current list of suppliers). Commercial MEFs may be useful to researchers new to ES cell culture or to those who will grow ES cells on a limited scale.

BASIC PROTOCOL 1

Materials Mouse embryos, 12.5 to 13.5 days postcoitum (Hogan et al., 1994) DPBS (see recipe), sterile Trypsin/EDTA solution (see recipe) MEF medium (see recipe) with penicillin/streptomycin Laminar flow hood Inverted microscope 100-mm tissue culture dish Dissecting forceps and fine scissors, sterilized by autoclaving or ethanol flaming 10-ml syringe and 16-G needle 100-mm tissue culture plates or 75-cm2 flasks Additional reagents and equipment for passaging and freezing MEFs (see Support Protocol) 1. Dissect mouse embryos (12.5 to 13.5 days postcoitum) into 10 to 20 ml sterile DPBS in a 100-mm tissue culture dish. Process embryos from one mouse together. Remove embryonic internal organs from the abdominal cavity using dissecting forceps. Transfer the carcass to a clean dish with fresh DPBS. Organ removal can be done crudely. Initial dissection can be performed at the bench. Subsequent procedures should be performed in a laminar flow hood. Any mouse strain can be used as an embryo source. However, outbred mice or F1 hybrids will usually produce more embryos per mating than inbred mice. If possible, use mice maintained in a viral-antibody-free (VAF) facility to reduce the chance of contamination. If the feeder cells will be used during the selection of antibiotic-resistant ES cells, use embryos from a transgenic mouse expressing the appropriate selectable marker. Appropriate transgenic mice may be obtained from the ES cell community or from standard mouse vendors such as The Jackson Laboratory and Taconic. Contributed by David A. Conner Current Protocols in Molecular Biology (2000) 23.2.1-23.2.7 Copyright © 2000 by John Wiley & Sons, Inc.



2. Rinse embryos again in 10 to 20 ml DPBS. Transfer the embryos to a clean 100-mm tissue culture dish containing 3 to 5 ml trypsin/EDTA solution. 3. Dissociate embryos by aspirating into a 10-ml syringe through a 16-G needle and expelling the contents. Repeat two to four times. Too many repetitions will reduce the cell yield. The embryos should be dissociated to the extent that they can be easily aspirated into a 5- or 10-ml pipette.

4. Add trypsin/EDTA solution to 10 ml. Mix contents by trituration, return to dish, and incubate 5 to 10 min in a 37°C incubator. 5. Mix again by trituration and incubate for an additional 5 to 10 min at 37°C. 6. Transfer contents to a 50-ml conical tube and add an equal volume of MEF medium with penicillin/streptomycin. Let stand 3 to 5 min at room temperature to allow large tissue pieces to settle to the bottom. 7. Remove solution, avoiding large tissue pieces, and place in a fresh 50-ml tube. Centrifuge 5 min at 1000 × g, room temperature. 8. Remove supernatant and resuspend pellet in 10 to 50 ml fresh MEF medium with penicillin/streptomycin. Plate cells on 100-mm tissue culture plates or 75-cm2 flasks, using approximately one embryo per plate or flask. Add medium to a final volume of 10 to 15 ml/plate. 9. Grow cells until confluent (2 to 5 days). Monitor cell density using an inverted microscope. Change medium after the first day and every other day thereafter. 10. Passage cells by trypsinizing (see Support Protocol, steps 1 to 4), resuspending the cell pellet in 10 to 50 ml MEF medium with penicillin/streptomycin, and plating at a dilution of 1:5 to 1:10. Add medium to a final volume of 10 to 15 ml per 75-cm2 flask or 10-mm plate. Using this dilution, fibroblasts from each original embryo are now plated on five to ten 75-cm2 flasks or 100-mm plates.

11. Grow again until confluent (3 to 5 days) and freeze (see Support Protocol, steps 1 to 7) at ∼5 × 106 cells/ml. A representative vial should be thawed to check for viability. It is also a good habit to check each new batch for mycoplasma contamination using a commercial service or a PCR-based screening kit (e.g., Pan Vera, Stratagene; also see Coté, 2000)). In addition, MEFs made from previously untested transgenic mice should be grown in the appropriate antibiotic to ensure that they are resistant to the concentration of antibiotic used for selection. BASIC PROTOCOL 2


MITOTIC INACTIVATION OF MEFS WITH γ-IRRADIATION MEFs must be inactivated prior to use as a feeder layer for mouse ES cells. Mitotic inactivation prevents the dilution of ES cell lines with dividing fibroblasts. MEFs can be inactivated using γ-irradiation, as described here, or mitomycin C treatment (see Alternate Protocol). This procedure is faster and less labor intensive, but requires a convenient radiation source. Both methods produce feeder layers suitable for the maintenance of undifferentiated ES cells. Materials Frozen MEF culture (see Basic Protocol 1) MEF medium (see recipe) without penicillin/streptomycin Ca2+- and Mg2+-free HBSS (see recipe)



100-mm tissue culture plates or 75-cm2 flasks 150-cm2 tissue culture flasks 100-mm Petri dishes γ-Radiation source Additional reagents and equipment for passaging, freezing, and thawing MEFs (see Support Protocol) 1. Thaw a vial of frozen MEFs as described below (see Support Protocol, steps 8 to 10) and plate in a 75-cm2 tissue culture flask or 100-mm plate. Grow until confluent (3 to 5 days). Change medium after the first day and every other day thereafter. 2. Passage cells by trypsinizing (see Support Protocol, steps 1 to 4), resuspending the cell pellet in 10 to 50 ml MEF medium without penicillin/streptomycin, and plating at a 1:10 dilution. Add medium to a final volume of 10 to 15 ml per 75-cm2 flask or 100-mm plate. 3. Grow until confluent (3 to 5 days), and passage at a 1:5 to 1:10 dilution, using twenty-five to fifty 150-cm2 flasks. Primary fibroblasts undergo a limited and variable number of cell divisions. Further passaging may be possible, but the rate of cell division slows quickly. Thawing a fresh vial is usually the fastest way to generate more feeders.

4. Remove medium from confluent flasks, rinse with 15 ml Ca2+- and Mg2+-free HBSS, and trypsinize again (see Support Protocol, steps 1 to 4). MEFs from ten to fifteen 150-cm2 flasks can be processed together.

5. Resuspend pellet in 10 ml MEF medium without penicillin/streptomycin and transfer suspension in a 100-mm Petri dish. Use a constant volume of medium regardless of the number of cells to ensure consistent results. Use a Petri dish rather than a cell culture plate to prevent adherence of the MEFs to the surface during irradiation.

6. Expose cells to 4000 rads from a γ-radiation source. 7. Dilute suspension to 50 ml with MEF medium without penicillin/streptomycin. Count the number of cells and freeze as described below (see Support Protocol, steps 4 to 7). MITOTIC INACTIVATION OF MEFS WITH MITOMYCIN C MEFs may be inactivated by mitomycin C treatment if a γ-radiation source is not available. Although this method is more time and labor intensive, the inactivated feeders are equally suitable for ES cell culture.

ALTERNATE PROTOCOL

Additional Materials (also see Basic Protocol 2) 1 mg/ml mitomycin C (Sigma) stock solution, filter sterilized (store at 4°C protected from light) 1. Expand a vial of MEFs as described above (see Basic Protocol 2, steps 1 to 3). 2. Add 1 mg/ml mitomycin C stock solution to the medium to a final concentration of 10 µg/ml. Return plates to the incubator for 2 to 3 hr. 3. Rinse plates twice with 10 to 15 ml Ca2+- and Mg2+-free HBSS. 4. Trypsinize as if passaging (see Support Protocol, steps 1 to 4).



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5. Add an equal volume of MEF medium without penicillin/streptomycin. 6. Count and freeze cells (see Support Protocol, steps 4 to 7). SUPPORT PROTOCOL

FREEZING AND THAWING MEFS Freezing MEFs, particularly inactivated fibroblasts, is a great convenience. Large stocks of active or inactivated fibroblasts can be prepared at any time, obviating the need to coordinate MEF preparations with ES cell manipulations. Cells are frozen slowly in medium containing 10% (v/v) dimethyl sulfoxide and thawed rapidly. Materials Plates containing MEFs (see Basic Protocols 1 and 2; see Alternate Protocol) Ca2+- and Mg2+-free HBSS (see recipe) Trypsin/EDTA solution (see recipe) Freezing medium (see recipe) MEF medium (see recipe) with or without penicillin/streptomycin Cryovials Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3F) Freeze cells 1. Remove MEF medium and rinse plates with 10 to 15 ml Ca2+- and Mg2+-free HBSS. Washing the plates removes residual serum that will inhibit trypsin.

2. Remove HBSS and add trypsin/EDTA solution to cover the surface of the cells (e.g., 3 to 5 ml in a 75-cm2 flask). Incubate 3 to 5 min at 37°C and tap the flask to release the cells. When trypsinization is complete, tapping and rocking the plate should release the cells. Loose sheets of cells are visible to the naked eye. Incubate at 37°C for an additional 1 to 2 min if the cells do not slough off.

3. When the cell layer has loosened, add an equal volume of MEF medium and mix by trituration to produce a single-cell suspension. MEF medium should contain penicillin/streptomycin when preparing initial MEFs (i.e., Basic Protocol 1, step 10), but not for other protocols. Addition of medium containing serum will inhibit further trypsinization.

4. Count cells using a hemacytometer (APPENDIX 3F) and then pellet by centrifuging 3 to 5 min at 1000 × g, room temperature. 5. Resuspend pellet at 3 × 106 cells/ml in freezing medium and mix by trituration. 6. Dispense into cryovials in 1-ml aliquots, place cryovials in an insulated container at −80°C, and leave overnight. Under these conditions, the cells will freeze slowly enough to maintain viability.

7. Transfer cryovials to liquid nitrogen. Cells should not be quick-frozen in liquid nitrogen. Vials can be stored at −80°C for several months and in liquid nitrogen for years. Mouse Embryo Fibroblast (MEF) Feeder Cell Preparation

Thaw cells 8. Thaw cells rapidly by placing vials in a 37°C water bath.



9. Add cells to a tube with 10 ml MEF medium without penicillin/streptomycin. To remove DMSO, pellet by centrifuging 3 to 5 min at 1000 × g, room temperature. 10. Resuspend in a volume appropriate for the surface area of the plate (e.g., 10 to 15 ml for a 75-cm2 flask or 100-mm plate). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.

NOTE: Prepare all cell culture solutions from tissue culture–grade reagents. Use tissue culture–grade water (high resistance and endotoxin free). Sterilize all final solutions by filtration or prepare from sterile stocks. Use disposable sterile plasticware to prevent microbial and detergent contamination. Dulbecco’s phosphate-buffered saline (DPBS) 0.1 g/liter anhydrous CaCl2 0.1 g/liter MgCl2⋅6H2O 0.2 g/liter KCl 0.2 g/liter KH2PO4 8.0 g/liter NaCl 2.16 g/liter Na2HPO4⋅7H2O Adjust pH, if necessary, to 7.0 to 7.2 with 1 N HCl or 1 N NaOH Sterilize by filtration or by autoclaving Store at 4°C (stable indefinitely) Freezing medium Dulbecco’s modified Eagle medium (DMEM) with high glucose (4500 mg/liter), L-glutamine, sodium pyruvate, and pyridoxine hydrochloride or pyridoxal hydrochloride (e.g., Life Technologies, JRH Biosciences, or Sigma) containing: 20 mM HEPES, pH 7.3 20% (v/v) FBS, heat inactivated for 30 min at 56°C 10% (v/v) dimethyl sulfoxide (DMSO) Store up to 1 year at −20°C Freezing and thawing should be avoided. Commercial freezing media based on 10% DMSO can be used. Consult UNIT 23.3 regarding the selection of FBS.

Hank’s balanced salt solution (HBSS), Ca2+ and Mg2+ free 0.4 g/liter KCl 0.06 g/liter KH2PO4 8.0 g/liter NaCl 0.35 g/liter NaHCO3 0.048 g/liter Na2HPO4 1.g/liter D-glucose 0.01 g/liter phenol red Adjust pH, if necessary, to 7.0 to 7.4 with 1 N HCl or 1 N NaOH Store up to 6 months at 4°C



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Mouse embryo fibroblast (MEF) medium Dulbecco’s modified Eagle medium (DMEM) with high glucose (4500 mg/liter), L-glutamine, sodium pyruvate, and pyridoxine hydrochloride or pyridoxal hydrochloride (e.g., Life Technologies, JRH Biosciences, or Sigma) containing: 10% (v/v) FBS, heat inactivated for 30 min at 56°C 1× MEM nonessential amino acids (from 100× stock; Life Technologies) 2 mM L-glutamine (from 100× stock; Life Technologies) 0.1 mM 2-mercaptoethanol 20 mM HEPES, pH 7.3 1× penicillin/streptomycin when indicated (from 100× stock; Life Technologies) Store up to 2 weeks at 4°C Consult UNIT 23.3 regarding the selection of FBS. Penicillin/streptomycin should be included during the initial isolation of MEFs. It is not needed after Basic Protocol 1.

Trypsin/EDTA solution Ca2+- and Mg2+-free HBSS (see recipe) containing: 2.5 g/liter porcine trypsin (0.25%) 0.38 g/liter EDTA⋅4H2O Store aliquots up to 1 year at −20°C Aliquots can be thawed and stored at 4°C for up to 1 week. Repeated freezing and thawing should be avoided.



Historically, ES cells have been cultured under a variety of conditions to prevent differentiation (Wurst and Joyner, 1993; UNIT 23.3). The most common method utilizes feeder layers prepared from mouse embryo fibroblasts or from SIM mouse embryo fibroblasts resistant to thioguanine and oubain (STO cells; Hogan et al., 1994; Martin and Evans, 1975). Whereas MEFs are primary cultures with a limited mitotic potential, STO fibroblasts will divide indefinitely. STO cells resistant to G418 are readily available from the ES cell community. STO cells can be grown and inactivated using the protocols for MEFs. For more details see Robertson (1987), Wurst and Joyner (1993), or Hogan et al. (1994). The advantage of MEFs is that they provide a more consistent source of feeder cells. Early passage cells with reproducible characteristics must be used because they rapidly lose their ability to divide. Long-term propagation of STO cells can lead to changes that result in characteristics that are less favorable for ES cell growth. Because of these potential problems, researchers who are new to ES cell culture and who wish to generate ES cell clones capable of germ-line transmission will probably have more success with primary MEFs.

Critical Parameters and Troubleshooting The major problems encountered during the isolation and inactivation of MEFs are typically a result of poor aseptic technique, low cell yields during isolation, or inefficient mitotic inactivation. Special attention should be paid to the use of aseptic technique. After the initial primary cultures have been frozen, antibiotics should not be used; antibiotics can mask bad aseptic technique. Each new batch of MEFs should be checked for mycoplasma after growth without antibiotics (Coté, 2000). Inadvertent contamination will spread to ES cells and reduce their germ-line potential. Contaminated batches should be disposed of; it is best not to try to treat them. Contamination should be a rare occurrence. Aseptic technique is also important in the preparation of media and solutions. All solutions should be prepared using tissue culture– grade reagents and water (high resistance and endotoxin free). Although they are more expensive, solutions that have been screened for toxicity and mycoplasma are available from many companies. Laboratories with little cell culture experience will probably have the greatest success with commercial, prescreened solutions. Occasionally low cell yields are observed after the initial embryo dissociation. Plates from the initial dissociation should reach con-



fluence in 2 to 5 days. If this does not occur, it is likely due to low trypsin activity or cell lysis during dissociation with the needle and syringe. Some cells may be recovered by passaging without dilution or by concentrating the cells from several plates onto a single plate. Under these circumstances, the cells may still be used as long as they resume dividing and reach confluence. During future preparations, fresh trypsin solutions should be used and the number of times each embryo is passed through the needle should be reduced. Occasionally evidence of cell division is observed in cultures of mitotically inactivated MEFs. When preparing inactivated feeders for the first time, check to make sure that the treatment was effective. This can be accomplished by plating out an aliquot of the inactivated feeders at a density of ∼6 × 104 cells/cm2. The cells are then cultured for 10 to 14 days, with periodic changes of medium, and cultures are assessed for increases in cell density and foci of mitotic activity. Foci can be observed by eye as opaque splotches on the plate, but should be confirmed with a microscope. No growth should be seen on a 100-mm plate; however, feeders may still be used if there are only a few foci. If there is a dramatic change in cell density or there are many colonies of dividing cells, the preparation should be discarded and the procedure repeated with newly expanded cells. If mitomycin C was used, a fresh solution should be prepared. If γ-irradiation was used, the radiation source should be properly calibrated. Distance and shielding can affect the dose dramatically. In addition to these concerns, it is important to remember that MEFs are primary cells with limited mitotic potential. Expanding the cells more than suggested may work, but the rate of growth will decrease. Plates can still become confluent because the cell size will increase; however, the number of cells per plate will decrease and the time to reach confluence will increase. Feeder layers made under these conditions may not be ideal for ES cell culture.

Time Considerations Processing a single litter of embryos (six to ten) should take 1 to 2 hr. After plating the dissociated embryos, cultures should reach confluence within 2 to 5 days. Cells can be frozen at this point, but the total yield will be dramatically reduced. After passaging, the cells should be ready to freeze in 3 to 5 days. Thawed vials should reach confluence in 3 to 5 days. After each passage the cells should take 3 to 5 days to reach confluence. The cells can be inactivated at any stage, but the yield will be lower with less expansion.

Literature Cited Coté, R. 2000. Assessing and controlling microbial contamination in cell cultures. In Current Protocols in Cell Biology (J.S. Bonifacino, M. Dasso, J.B. Harford, J. Lippincott-Schwartz, and K.M. Yamada, eds.) pp. 1.5.1-1.5.18. John Wiley & Sons, New York. Hogan, B., Beddington, R., Constantini, F., and Lacy, E. 1994. Manipulating the Mouse Embryo: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Martin, G.R. and Evans, M.J. 1975. Differentiation of clonal lines of teratocarcinoma cells: Formation of embryoid bodies in vitro. Proc. Natl. Acad. Sci. U.S.A. 72:1441-1445. Robertson, E.J. 1987. Embryo-derived stem cell lines. In Teratocarcinomas and Embryonic Stem Cells: A Practical Approach (E.J. Robertson, ed.) pp. 71-112. IRL Press, Oxford. Wurst, W. and Joyner, A.L. 1993. Production of targeted embryonic stem cell clones. In Gene Targeting: A Practical Approach (A.L. Joyner, ed.) pp. 33-61. IRL Press, Oxford.

Key References Hogan et al., 1994. See above. Provides additional or alternative protocols and defines the context for use of MEFs during gene targeting in ES cells.

Internet Resources www.biosupplynet.com

Anticipated Results

Search this Web site for “embryonic stem cell reagents” to obtain a current list of suppliers that provide medium and MEFs suitable for ES cell culture.

Expect 10 to 30 vials of frozen cells from each embryo. Each vial can be expanded to produce 50 to 100 vials containing 1 ml of mitotically inactive feeders. Each vial should be sufficient for one targeting experiment.

Contributed by David A. Conner Harvard Medical School Boston, Massachusetts



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Mouse Embryonic Stem (ES) Cell Culture Culturing mouse embryonic stem (ES) cells has many similarities to the culture of any adherent cell line. However, there are some special concerns that warrant elaboration because ES cells must not be allowed to differentiate if they are to be used to generate germ-line chimeras. This unit describes a common method for ES cell culture utilizing gelatinized plates, feeder layers of mitotically inactive mouse embryo fibroblasts (MEFs; UNIT 23.2), and recombinant leukemia inhibitory factor (LIF). Special attention is paid to the timing of passaging to prevent differentiation during routine culture.


NOTE: All cell culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Materials 0.1% (w/v) gelatin solution (see recipe) Mitotically inactive mouse embryo fibroblasts (MEFs; UNIT 23.2) ES cell medium (see recipe) Embryonic stem (ES) cells (http://www.biosupplynet.com) Ca2+- and Mg2+-free HBSS (UNIT 23.2) DPBS/EDTA solution (see recipe) Trypsin/EDTA solution (see recipe) Freezing medium (UNIT 23.2) Cryovials Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3F) Prepare gelatin-coated plates 1. Pipet a sufficient volume of 0.1% gelatin solution onto plates to cover the bottom of the plate. 2. Leave the solution on the plate for ≥1 hr at room temperature or in a 37°C incubator. The temperature is not critical. Plates can be prepared in advance by adding the gelatin solution and storing the plate in the incubator (to prevent drying) for up to a week.

Prepare feeder plates 3. Aspirate gelatin solution just before adding feeder cells. 4. Plate mitotically inactive MEFs (feeders) in ES cell medium on gelatinized plates at ∼6 × 104 viable cells/cm2. Place cells in the incubator. Either freshly inactivated MEFs or frozen inactivated MEFs can be used. Frozen cells should be thawed rapidly at 37°C and DMSO should be removed (see UNIT 23.2). Note that some cell death occurs during the freezing and thawing procedure. Frozen MEFs should thus be plated at approximately twice the density (1.2 × 105 cells/cm2) of fresh MEFs (6 × 104 cells/cm2) to achieve a similar density of viable feeders. If the density is too high, the cells may lift off the plate. Feeders can be plated in either ES medium or MEF medium (see UNIT 23.2). Both have similar components, and it is often more convenient to prepare only one. The feeders attach to the plate within 30 min and begin to spread out over the next few hours.

5. Change medium to remove dead feeders the next day. The fibroblasts should spread to cover the entire surface of the plate by the day after plating. Feeder plates can be prepared in advance and used for routine ES cell culture within 7 to 10 days after plating. The medium should be changed every other day if they are not used immediately. Manipulating the Mouse Genome Contributed by David A. Conner Current Protocols in Molecular Biology (2000) 23.3.1-23.3.6 Copyright © 2000 by John Wiley & Sons, Inc.


Culture ES cells 6. Thaw a vial of ES cells rapidly at 37°C. Remove DMSO by diluting in 10 ml ES cell medium and centrifuging 3 to 5 min at 1000 × g, room temperature. 7. Resuspend pellet in ES cell medium and plate on feeder layers prepared at least one day earlier. The number of ES cells depends on the surface area of the feeder plate. Plate 1–3 × 106 cells on a 25-cm2 flask or a 60-mm plate. Plate cells frozen at a higher density on 75-cm2 flasks or 100-mm plates, and fewer cells in 6- or 24-well feeder plates.

8. Change medium daily and passage cells every 2 to 3 days depending on total cell density and colony size (see Critical Parameters and Troubleshooting for more details). Cells may require passaging the day after plating if the cell density is very high.

Passage ES cells 9. Change medium several hours before passaging to maximize plating efficiency. 10. Rinse plates with Ca2+- and Mg2+-free HBSS (e.g., 10 ml per 100-mm plate). Washing the plates removes residual serum that will inhibit trypsin.

11. Add sufficient DPBS/EDTA solution to cover the cells and incubate 3 min at 37°C. 12. Add an equal volume of Trypsin/EDTA solution. Rock the plate to mix and return to the incubator for 1 to 2 min. 13. Tap the plate to release the cells. If the cells do not slough off, return the plate to the incubator for another minute and tap again. Alternatively, the DPBS/EDTA step can be skipped, and the plate can be incubated for 3 to 7 min after addition of trypsin/EDTA, with periodic tapping to see if the cell layer has loosened. With either approach, the goal is to achieve a single-cell suspension with minimal trypsinization. Preincubation with DPBS/EDTA reduces the time necessary for trypsin treatment and yields the same degree of dispersion.

14. When the cell layer has loosened, add 2 to 3 vol ES cell medium and disperse the cells thoroughly by trituration. 15. Centrifuge 3 to 5 min at 1000 × g, room temperature. Resuspend the pellet in ES cell medium and plate on feeders at a 1:10 dilution. Freeze cells 16. Trypsinize cells (steps 9 to 13). 17. Add an equal volume of ES cell medium and mix by trituration to produce a single-cell suspension. Addition of medium containing serum will inhibit further trypsinization.

18. Count cells using a hemacytometer (APPENDIX 3F) and then pellet by centrifuging 3 to 5 min at 1000 × g, room temperature. 19. Resuspend pellet at 3 × 106 cells/ml in freezing medium and mix by trituration. 20. Dispense into cryovials in 1-ml aliquots, place the cryovials in an insulated container at −80°C, and leave overnight. Under these conditions, the cells will freeze slowly enough to maintain viability.

21. Transfer cryovials to liquid nitrogen. Mouse Embryonic Stem (ES) Cell Culture

Cells should not be quick-frozen in liquid nitrogen. Vials can be stored at −80°C for several months and in liquid nitrogen for years.




NOTE: Prepare all cell culture solutions from tissue culture–grade reagents. Use tissue culture–grade water (high resistance and endotoxin free). Sterilize all final solutions by filtration or prepare from sterile stocks. Use disposable sterile plasticware to prevent microbial and detergent contamination. DPBS/EDTA solution Prepare DPBS (UNIT 23.2) but without calcium and magnesium Add 0.2 g/liter EDTA⋅4H2O Sterilize by filtration or autoclaving Store at 4°C (stable indefinitely) ES cell medium Dulbecco’s modified Eagle medium (DMEM) with high glucose (4500 mg/liter), L-glutamine, sodium pyruvate, and pyridoxine hydrochloride or pyridoxal hydrochloride (e.g., Life Technologies, JRH Biosciences, or Sigma) containing: 15% (v/v) FBS, heat inactivated for 30 min at 56°C 1× MEM nonessential amino acids (from 100× stock; Life Technologies) 2 mM L-glutamine (from 100× stock; Life Technologies) 0.1 mM 2-mercaptoethanol 20 mM HEPES, pH 7.3 500 to 1000 U/ml murine leukemia inhibitory factor (LIF; e.g., ESGRO, Life Technologies; Chemicon) Store up to 1 week at 4°C The quality of FBS should be screened by assessing plating efficiency, morphology, and toxicity on ES cells plated at low density in 10%, 15%, and 30% heat-inactivated FBS. Plating efficiency should be ∼10% and should be similar for all FBS concentrations. The author has had good success with FBS from Hyclone. Several companies sell FBS that has been screened for ES cell growth using the same techniques. Although it is expensive, laboratories that are new to ES cell culture may benefit from prescreened FBS. In addition, prescreened FBS provides a standard of comparison for other FBS lots.

Gelatin solution, 0.1% (w/v) Prepare 0.1% (w/v) porcine skin gelatin (Sigma) in water and sterilize by autoclaving. Filter after cooling. Store up to 1 year at 4°C. Trypsin/EDTA solution Ca2+- and Mg2+-free HBSS (UNIT 23.2) containing: 0.5 g/liter porcine trypsin (0.05%) 0.2 g/liter EDTA⋅4H2O Store aliquots up to 1 year at −20°C Note that this recipe contains one-fifth the trypsin that is used for preparing MEFs (UNIT 23.2).

Aliquots can be thawed and stored at 4°C for up to 1 week. Repeated freezing and thawing should be avoided.



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COMMENTARY Background Information ES cells can be obtained from a number of investigators, from commercial sources, or by establishing new lines directly from mouse embryos (see http://www.biosupplynet.com for a current list of commercial suppliers). Historically, ES cells have been cultured under a variety of conditions to prevent differentiation. Lines have been successfully maintained in Buffalo rat liver (BRL) cell–conditioned medium (Smith and Hooper, 1987), in LIF-containing medium (Smith et al., 1988), and on STO or MEF feeder layers with or without LIF (Evans and Kaufman, 1981; Martin, 1981; Wurst and Joyner, 1993). ES cells may become accommodated to the culture conditions in which they were isolated; therefore, it may be advisable to modify the protocol outlined in this unit to match the growth conditions recommended for specific ES cell lines. However, most ES cell lines should do well on MEF feeder layers supplemented with LIF.


Mouse Embryonic Stem (ES) Cell Culture

The major problems encountered during ES cell culture are contamination and differentiation. Special attention should be paid to aseptic technique. ES cells should not be cultured routinely in antibiotics. Although penicillin and streptomycin do not reduce the germ-line potential of ES cells, the antibiotics can mask bad aseptic technique. Mycoplasma contamination can reduce germ-line potential and is not eliminated by penicillin and streptomycin. Cultures should be checked periodically for mycoplasma (Coté, 2000), and contaminated cultures should be discarded. It is best not to try to treat them. Contamination should be a rare occurrence. Aseptic technique is also important in the preparation of media and solutions. All solutions should be prepared using tissue culture– grade reagents and water (high resistance and endotoxin free). Although they are more expensive, solutions that have been screened for toxicity and mycoplasma are available from many companies. Laboratories with little cell culture experience will probably have the greatest success with commercial, prescreened solutions (http://www.biosupplynet.com). ES cells grow as three-dimensional colonies, not monolayers. Undifferentiated colonies have a rounded appearance with sharp refractal edges; individual cell borders within

a colony are difficult to discern (Fig. 23.3.1A). All cultures will contain some differentiated colonies (Fig. 23.3.1B). Typically these colonies grow as monolayers with ragged edges and easily distinguishable cell borders. The appearance of ES cell colonies can change depending on total cell density and the time since passaging. To assess the quality of the culture, the cells should be observed over several days. During this period most of the colonies should be undifferentiated. Assuming that quality reagents are used, differentiation of ES cells can be minimized by limiting the number of cell divisions and by passaging cultures at the appropriate density. It is advisable to keep track of the number of passages. New ES cell lines should be expanded with a minimum number of passages and frozen in many aliquots. A new low-passage vial should be thawed for each experiment. The passage number is relevant only because it is a rough estimate of the number of cell divisions the culture has undergone. Thus, one should not skip passaging at the appropriate density to avoid increasing the passage number. Cells should be viewed every day until their growth patterns become familiar. One day after passaging, ES cell colonies can be difficult to distinguish from cell debris on the surface of the feeder layer. By the second day, distinct colonies are apparent (Fig. 23.3.1C). Determining the appropriate density for passaging is the most difficult aspect of ES cell culture. Both the total plate density and the individual colony size should be considered. Cells will tend to differentiate within large colonies or at very high total plate densities. For example, the cells in Figure 23.3.1C can be cultured for another day; the plate density is low and the colonies are small. The cells in Figure 23.3.1D are ready to passage; the plate density is moderate but the individual colonies are large. Although the cells seem to grow best at relatively high densities, the cells should not be allowed to become confluent; colonies should remain separated. Passaging every 2 days at a dilution of 1:8 or 1:10, when the cells are growing well, will maintain an ideal plate density and colony size. Passaging at a lower frequency is necessary when the cells are seeded at a low density. Under these conditions, it is best to pay attention to individual colony size and to passage with little dilution to disperse colonies and increase plate density.



A

B

C

D

Figure 23.3.1 Morphology of ES cell colonies. Phase-contrast images of ES cell colonies on MEF feeders. (A) Appearance of an ideal ES cell colony that is ready to passage. (B) Appearance of a differentiating ES cell colony. Note that the colony has grown as a monolayer with distinct borders between cells. The edge of the colony is not yet ragged. (C) Appearance of ES cell colonies, at lower magnification, 2 days after plating at low density. Colonies can be distinguished from the feeder layer by their sharp refractal borders and their three-dimensional appearance. (D) Appearance of ES cell colonies 3 days after plating at an intermediate density. These cells are ready to passage.

Anticipated Results

Literature Cited

When plates are grown at maximum density, there should be 2.5–4.0 × 105 cells/cm2. For routine growth, the total density should be somewhat lower (e.g., 1.5–2 × 105 cells/cm2) and the cells should be passaged every 2 days.

Coté, R. 2000. Assessing and controlling microbial contamination in cell cultures. In Current Protocols in Cell Biology (J.S. Bonifacino, M. Dasso, J.B. Harford, J. Lippincott-Schwartz, and K.M. Yamada, eds.) pp. 1.5.1-1.5.18. John Wiley & Sons, New York.

Time Considerations

Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156.

A single vial (frozen at 3.0 × 106 cells/ml) can be expanded in ∼5 days to produce enough cells for a standard electroporation (2–4 × 107 cells). A single vial can be expanded to produce 50 to 100 vials of a low-passage stock in 7 to 8 days.

Martin, G.R. 1981. Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc. Natl. Acad. Sci. U.S.A. 78:7634-7636. Smith, A.G. and Hooper, M.L. 1987. Buffalo rat liver cells produce a diffusible activity which inhibits the differentiation of murine embryonal carcinoma and embryonic stem cells. Dev. Biol. 121:1-9.



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Smith, A.G., Heath, J.K., Donaldson, D.D., Wong, G.G., Moreau, J., Stahl, M., and Rogers, D. 1988. Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature 336:688-690.

Wurst and Joyner, 1993. See above.

Wurst, W. and Joyner, A.L. 1993. Production of targeted embryonic stem cell clones. In Gene Targeting: A Practical Approach (A.L. Joyner, ed.) pp. 33-61. IRL Press, Oxford.

Internet Resources

Key References Hogan, B., Beddington, R., Constantini, F., and Lacy, E. 1994. Manipulating the Mouse Embryo: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.

These three references, written by experts in the field, represent the best compilations of ES cell culture methods to date.

www.biosupplynet.com Search this Web site for “embryonic stem cell reagents” to obtain a current list of suppliers that provide medium and ES cells.


Robertson, E.J. 1987. Embryo-derived stem cell lines. In Teratocarcinomas and Embryonic Stem Cells: A Practical Approach (E.J. Robertson, ed.) pp. 71-112. IRL Press, Oxford.

Mouse Embryonic Stem (ES) Cell Culture



Mouse Embryonic Stem (ES) Cell Isolation

UNIT 23.4

Undifferentiated mouse embryonic stem (ES) cells are necessary for the production of mouse mutants using homologous recombination and blastocyst-mediated transgenesis. Suitable ES cell lines are available from commercial sources (UNIT 23.3) and from many investigators. However, investigators planning to use ES cells extensively may find that the isolation of new lines provides a simple and economical method for maintaining stocks of early passage ES cells. In addition, ES cells obtained from mutant mouse lines may facilitate the analysis of mutant phenotypes, particularly when the mutation causes early embryonic lethality. ISOLATION OF MOUSE ES CELLS ES cell isolation is straightforward, although success rates can be quite variable. ES cells are derived from the inner cell mass of blastocysts (i.e., 3.5-day-old embryos). Blastocysts are simply cultured for several days, during which time they attach to the surface of the tissue culture plate. Both trophoblast and inner cell mass cells divide after attachment. Inner cell mass outgrowths are picked, dispersed by trypsinization, and replated. Under appropriate conditions a percentage of the isolated outgrowths will continue to divide and maintain an undifferentiated ES cell morphology.

BASIC PROTOCOL

Materials Modified ES medium (see recipe) Blastocysts, 3.5-day-old post-coitum embryos (Hogan et al., 1994) Hanks’ balanced salt solution (HBSS), calcium- and magnesium-free (UNIT 23.2) DPBS-EDTA (UNIT 23.3) 0.25% (w/v) trypsin-EDTA (UNIT 23.2) Inverted microscope Gilson-style automatic pipettor with 20-µl pipet tips 96-well U-bottom plate Additional reagents and equipment for preparing gelatin-coated plates with MEF feeder layers for embryonic stem cell culture (UNIT 23.3) and for passaging and freezing embryonic stem cells (UNIT 23.3) NOTE: All cell culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Plate out embryos 1. Prepare gelatinized 24-well tissue culture plates with mitotically inactivated MEF (mouse embryo fibroblast) feeder layers as described in UNIT 23.3. Plates can be prepared a few hours in advance or the day before.

2. Replace medium in each well with modified ES medium prior to use. Drop individual blastocysts into separate wells and return plates to incubator. Aim for the center of the well, otherwise the blastocysts will be difficult to observe with the microscope because of distortion near the edge of wells.

3. Observe embryos daily with an inverted microscope. The majority of blastocysts will hatch from the zona pellucida and attach to the plate by the second or third day. Figure 23.4.1A shows an attached blastocyst approximately 48 hr after plating. The zona pellucida is not present. In this early stage, the embryo still retains its original shape. Contributed by David A. Conner Current Protocols in Molecular Biology (2000) 23.4.1-23.4.9 Copyright © 2000 by John Wiley & Sons, Inc.



A

B

C

D

Figure 23.4.1 Morphology of plated blastocysts as shown in phase-contrast images of blastocysts at various time points after plating on MEF feeder cells. (A) Appearance of a blastocyst shortly after attachment to the plate ~48 hr after plating. (B) Typical appearance of a blastocyst 3 to 4 days after plating. Note the expansion of inner cell mass and trophoblast cells spreading out beneath. (C) Typical appearance of blastocyst ready to pick. (D) Appearance of a blastocyst that has been left on the plate 1 day too long. Note the pigmentation on the top surface indicating substantial differentiation.

4. Change medium after embryos have attached to the plate. The trophoblast cells will spread out quickly under the embryo. Over the next several days the inner cell mass will begin to expand. Figure 23.4.1B shows the inner cell mass at an early stage of expansion. Note the large flattened trophoblast cells under the inner cell mass. The inner cell mass is expanding as a multilayered colony.


Pick inner cell mass outgrowths Approximately 5 or 6 days after plating, the inner cell mass outgrowths will be ready to disperse. Figure 23.4.1C shows an outgrowth ready to pick. Figure 23.4.1D shows an outgrowth that has been left too long. If the outgrowth is left on the plate for too long, the cells begin to differentiate. This is typically observed as an accumulation of pigmented cells on the top of the colony. Round, loosely attached cells may also appear on the top of the outgrowth as differentiation occurs. Outgrowths should be picked before significant pigment accumulates. Outgrowths with a small amount of differentiation should still be dispersed, because they can yield decent lines.



5. Prepare gelatinized 24-well plates with feeder cells in advance with enough wells for each of embryo (UNIT 23.3). 6. Replace medium in embryo cultures 2 to 4 hr prior to picking. This is thought to maximize plating efficiency.

7. Rinse wells with Hanks’ balanced salt solution (HBSS, calcium- and magnesiumfree). After aspirating, add a small volume of HBSS to wells to keep the cell layer from drying out. Process only a few wells at a time.

8. Pick up inner cell mass outgrowth using an automatic pipettor with a 20-µl tip and place the cell clump in a well of a 96-well U-bottom plate containing 25 µl DPBS-EDTA in each well. Try to transfer the cell mass in a minimal volume (i.e., ≤5 ìl). Some investigators find that picking is more easily performed using an inverted microscope with a low-power objective or a dissecting microscope. However, the cell mass can be seen by eye at this stage. With some practice, picking by eye is less tedious than picking with a microscope. In either case, the outgrowth can be dislodged by scraping the tip of the pipet around the cell mass to tear the trophoblast and MEF layer. Then, the inner cell mass can be picked up by simultaneously nudging the clump with the pipet tip and releasing the pipet plunger to aspirate the cells. It does not matter if some fibroblasts and trophoblasts are transferred with the inner cell mass. After placing the cells in the 96-well plate, check under the microscope to verify that the picking procedure was successful.

9. Add 50 µl of 0.25% trypsin-EDTA to each cell containing well of the 96-well plate and place in incubator for 5 to 10 min. Do not pick too many at one time. The first outgrowth picked should not sit in PBS-EDTA for longer than 10 min prior to addition of trypsin.

10. Add 100 µl modified ES medium to each well to inhibit trypsin. Triturate 3 to 10 times using a Gilson-style pipettor with 200-µl tips. Transfer cells to a new 24-well feeder plate. Try to disperse the outgrowth into several small clumps. Check with microscope to determine the degree of dispersal. Differentiation will occur if cells are not dispersed. However, do not try to achieve a single-cell suspension; trypsinization and trituration sufficient to achieve single-cell suspension usually kills cells at this stage.

Screen and expand putative ES cell lines 11. Monitor plates daily (within 2 days after plating, colonies should be visible). Mark the wells with ES cell–like colonies and observe their morphology on successive days. Look for ES cell colonies like those depicted in Figure 23.4.2A and B. These will appear as small rounded clumps of cells with sharp refractal edges; nuclei are usually distinct, but cell borders are not (see UNIT 23.3 for more detail and pictures). Most wells will contain some embryo-derived cells. Figure 23.4.2, panels C through E, shows common morphologies of non-ES cell colonies. 5% to 15% of the wells may contain ES cell–like colonies when using embryos from the 129 mouse strain (see Background Information and see Troubleshooting regarding the effects of mouse strain on ES cell yields).

12. After 3 to 6 days, passage ES cell colonies as follows: a. In wells with only a few ES cell colonies, pick cells by pipet as described for the initial inner cell mass outgrowths (steps 5 to 10) and replate on gelatinized 24-well feeder plates.



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A

B

C

D

E

Figure 23.4.2 Cell types observed after inner cell mass disaggregation. Phase-contrast images of some of the different cell morphologies seen 2 to 3 days after disaggregation. (A, B) Appearance of putative ES cells. (C,D,E) Appearance of common non-ES cell–like colonies. The cells in panel C resemble ES cells to some degree because of the distinct edge. However, unlike ES cells they are growing as a monolayer and the individual cells have distinct borders. Colonies of this type with more tightly packed cells look exactly like ES cells. After passage these misleading colonies will spread out and become easy to distinguish from true ES cells. The round, highly refractal cells in panel D look like the cells that often accumulate on the top of differentiating ES cell colonies. Note that the colony does not have a sharp, refractal edge. The colony in panel E has a diffuse border with giant trophoblast cells visible in the center.




b. In wells with many ES cell colonies, trypsinize the entire well and replate on gelatinized feeder plates with a larger surface area (e.g., 6-well or 25-cm2 plates) using the protocol described for passaging established ES cell lines (UNIT 23.3). When picking individual colonies, all colonies from a single well can be combined. Deciding when to disperse the colonies depends on colony size and total plate density as described in UNIT23.3. Passage all wells that have colonies that look remotely like ES cells and observe them daily afterwards; non-ES cells will become apparent after passage. In addition, colonies from wells with mixed morphologies should be split. ES cells will usually outgrow other cell types after several passages.

13. Expand each line that maintains an ES cell morphology until there are enough cells to freeze down 3 or more vials at a cell density per vial of 1–3 × 106 cells/ml (see UNIT 23.3). Typically, enough cells should be present on a 25-cm2 flask at medium density. The protocols for expansion and freezing are described in UNITS 23.2 & 23.3. Samples of new cell lines should be analyzed for mycoplasma contamination. Investigators may want to karyotype new lines; however, most early-passage lines will have a normal complement of 40 chromosomes.

ES CELL SEX DETERMINATION Male ES lines are used most commonly, primarily because of concerns of X chromosome instability in female ES lines (Robertson et al., 1983). This protocol describes a simple polymerase chain reaction (PCR) screen to determine the sex of an ES cell line.

SUPPORT PROTOCOL

Additional Materials (also see Basic Protocol) Digestion buffer (UNIT 23.5) Saturated NaCl 95% ethanol 10× amplification buffer (UNIT 15.1) 25 mM 4dNTP mix (UNIT 15.1) Primers (Kunieda et al., 1992): Set 1: SRY2: TCTTAAACTCTGAAGAAGAGAC SRY4: GTCTTGCCTGTATGTGATGG Set 2: NDS3: GAGTGCCTCATCTATACTTACAG NDS4: TCTAGTTCATTGTTGAGTTGC 5 U/µl Taq DNA polymerase 3% (w/v) agarose gel Molecular weight markers 1.5-ml microcentrifuge tubes 55°C incubator Additional reagents and equipment for PCR amplification (UNIT 15.1), culture of ES cells (UNITS 23.2 and 23.3), and agarose gel electrophoresis (UNIT 2.5) Prepare DNA 1. Plate ES cells onto gelatinized 24-well plates without feeder cells. Grow these cells for 3 to 5 passages without feeder cells. Dilute cells at least 1:10 at each split. The cells must be diluted out to prevent contamination of the PCR reactions from any residual feeder cells. ES cells can be grown in standard ES cell medium (UNIT 23.3) without antibiotics at this stage. Some differentiation will occur in the absence of the feeder layer.

2. Remove medium. Rinse well with HBSS. Manipulating the Mouse Genome


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3. Add 300 µl digestion buffer, transfer to a 1.5-ml microcentrifuge tube, and incubate overnight at 55°C. Plenty of DNA can be obtained from a single well of a 24-well plate, but it is often more convenient to use a larger surface area, such as that of a 25-cm2 flask. Use 1.5 ml of digestion buffer for a 25-cm2 flask and adjust the volume of subsequent solutions proportionally.

4. Add 150 µl saturated NaCl and vortex vigorously (the solution will turn milky white). Add 2 vol of 95% ethanol (the solution will turn clear except for precipitated DNA). Some investigators precipitate the DNA using 2 vol ethanol (or 1 vol isopropanol) without adding salt. However, the DNA pellet resuspends more easily if salt is added.

5. Resuspend DNA pellet in 50 µl water. Determine DNA concentration by measuring the absorbance at 260 nm (APPENDIX 3D). Perform PCR 6. Perform 2 PCR reactions with each sample: one with the SRY primers and the other with the NDS primers. Mix the following in a 0.5-ml thin-walled PCR tube for each primer set. 2.5 µl 10× PCR buffer (final MgCl2 concentration 1.5 mM) 0.2 µl 25 mM 4dNTP mix 0.5 µM primer 1 0.5 µM primer 2 0.2 to 1.0 µg DNA template 0.5 U Taq DNA polymerase up to 25 µl H2O 7. Using the following parameters, run the PCR. 35 cycles

30 sec 30 sec 60 sec

94°C 50°C 72°C

(denaturation) (annealing) (extension)

Analyze the product 8. Run 10 µl of the products on a 3% agarose gel with the appropriate molecular weight markers. The SRY primers are derived from the sex-determining region of the Y chromosome. Male cells will show the 404-bp product; female cells will have no product. The NDS primers span a microsatellite dinucleotide repeat on the X chromosome and serve as a positive control; both male and female cells should show the 244-bp product.


NOTE: Prepare all cell culture solutions from tissue culture-grade reagents. Use tissue culture-grade water (high resistance and endotoxin free). Sterilize all final solutions by filtration or prepare from sterile stocks. Use disposable sterile plasticware to prevent microbial and detergent contamination.




Modified ES medium Dulbecco’s modified Eagle medium (DMEM), with high glucose (4500 mg/liter), L-glutamine, sodium pyruvate, and pyridoxine (or pyridoxal) hydrochloride (e.g., Life Technologies, JRH Biosciences, or Sigma) containing: 20% (v/v) FBS, heat inactivated for 30 min at 56°C 1× MEM nonessential amino acids (Life Technologies) 2 mM glutamine (from 100× stock; Life Technologies) 0.1 mM 2-mercaptoethanol 20 mM HEPES, pH 7.3 500 to 1000 U/ml murine leukemia inhibitory factor (LIF; Life Technologies, Chemicon) 1× penicillin/streptomycin (from 100× stock; Life Technologies) 1× nucleoside stock (see recipe; optional) Store at 4°C for up to 1 week The quality of FBS should be screened by assessing plating efficiency, morphology, and toxicity on ES cells plated at low density in 10%, 15%, and 30% heat-inactivated FBS. Plating efficiency should be ~10% and should be similar for all FBS concentrations. The author has had good success with FBS from Hyclone. Several companies sell FBS that has been screened for ES cell growth using the same techniques. Although it is expensive, laboratories that are new to ES cell culture may benefit from prescreened FBS. In addition, prescreened FBS provides a standard of comparison for other FBS lots.

100× nucleoside stock In 100 ml of tissue culture grade water dissolve: 80 mg adenine 73 mg cytidine 85 mg guanosine 24 mg thymidine 73 mg uridine Dissolve by warming at 37°C and filter sterilize. Aliquot and freeze at −20°C. Thaw at 37°C and mix vigorously to redissolve nucleosides prior to use. The addition of a nucleoside stock to the modified ES medium is recommended by Robertson (1987).

COMMENTARY Background Information Pluripotent mouse embryonic stem cells were first isolated by Evans and Kaufman (1981) and Martin (1981). The protocol outlined in this unit is essentially that of Axelrod (1984) and Robertson (1987) with some modifications. The primary differences are related to the culturing conditions including the use of MEFs as feeder layers instead of STO (SIM mouse embryo fibroblasts resistant to thioguanine and oubain) cells and the addition of recombinant leukemia inhibitory factor (LIF) to inhibit differentiation. Evans and Kaufman (1981) used delayed blastocysts, reasoning that the increase in the number of cells in these embryos would increase the likelihood of successful stem cell isolation. Martin (1981) used immunosurgery to isolate the inner cell mass and cultured on STO feeder cell layers

with conditioned medium from embryonal carcinoma cells. Many of these ingenious tricks are no longer necessary because of a more refined understanding of the cell culture conditions required to maintain undifferentiated ES cells. Indeed, ES cell lines capable of contributing to the germ line have been isolated in media supplemented with LIF in the absence of feeder cell layers (Pease et al., 1990). In addition to methodology, genetic background can affect the efficiency of ES cell isolation. The majority of ES cells used for gene targeting are derived from 129 substrains, in part because of the ease with which ES cells can be established from this strain. The nomenclature of the 129 strain has been revised recently; refer to Festing et al. (1999) to clarify strain names. Lines have been isolated with more difficulty from some other common



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strains such as C57BL/6 (Ledermann and Burki, 1991), Balb/cJ (Noben-Trauth et al., 1996), and C3H/He (Kitani et al., 1996). The factors that influence strain permissiveness are not known.

Critical Parameters and Troubleshooting The isolation of ES cell lines is not a complicated procedure; however, this does not mean that the procedure is always successful. Several factors are important to consider. Highquality cell culture reagents and proper aseptic technique are necessary for success. Consult UNITS 23.2 & 23.3 for more detail on these issues. As indicated above, the strain of mouse from which the embryos are derived can have a dramatic effect on success. If a strain other than a standard 129 substrain must be used, first consult the literature for reports of successful ES cell derivation and for any strain-specific approaches. Second, increase the number of starting embryos, because the yield will probably be lower. Third, verify the technique and reagents used by isolating lines from a permissive strain. The procedure may not be successful even with these modifications. Consider more elaborate approaches such as that reported by McWhir et al. (1996). The most common problems during isolation of ES cell lines are encountered during the initial dispersion of the inner cell mass outgrowth. Under-trypsinization or very limited trituration will result in transfer of the whole cell clump without dispersion. The large colony will differentiate rapidly. At this stage, the well can sometimes be saved by immediately picking the outgrowth a second time, trypsinizing, and replating. Over-trypsinization or extensive trituration also creates a problem. Single cells do not clone well at this stage. Dispersing the outgrowth into a single-cell suspension will usually result in cell death or differentiation. Both of these problems can be avoided by working with only a few outgrowths at one time and by monitoring the procedure with a microscope. In addition, after dispersal, the wells should be scanned carefully for several days. Single ES cell–like colonies are easy to miss, especially if they sit near the edge of the well, where the image is distorted.

Anticipated Results Mouse Embryonic Stem (ES) Cell Isolation

With embryos derived from 129 strains (e.g., 129X1/SvJ or 129S6/SvEvTac), 5% to 15 % of the inner cell mass outgrowths should give rise

to ES cell lines. Half of the resultant lines should be male.

Time Considerations Initial inner cell mass outgrowths can be picked 5 to 6 days after plating the embryos. Within 3 to 5 days, ES cell–like clumps can be picked and dispersed; 3 to 5 days later wells containing ES cells can be expanded, and subsequently frozen 2 to 4 days later. For sex determination, aliquots of the cell lines should be expanded without feeder cells for 1 to 2 weeks (i.e., 3 to 5 passages).

Literature Cited Axelrod, H.R. 1984. Embryonic stem cell lines derived from blastocysts by a simplified technique. Devel. Biol. 101:225-228. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Festing, F.W., Simpson, E.M., Davisson, M.T., and Mobraaten, L.E. 1999. Revised nomenclature for strain 129 mice. Mamm. Genome 10:836. Hogan, B., Beddington, R., Constantini, F., and Lacy, E. 1994. Manipulating the Mouse Embryo: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Kitani, H., Takagi, N., Atsumi, T., Kawakura, K., Imamura, K., Goto, S., Kusakabe, M., and Fukuta, K. 1996. Isolation of a germline-transmissible embryonic stem (ES) cell line from C3H/He mice. Zool. Sci. 13:865-871. Kunieda, T., Xian, M., Kobayashi, E. Imamichi, T., Moriwaki, K., and Toyoda, Y. 1992. Sexing of mouse preimplantation embryos by detection of Y chromosome-specific sequences using polymerase chain reaction. Biol. Reprod. 46:692-697. Ledermann, B. and Burki, K. 1991. Establishment of a germ-line competent C57BL/6 embryonic stem cell line. Exp. Cell Res. 197:254-258. Martin, G.R. 1981. Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc. Natl. Acad. Sci. U.S.A. 78:7634-7636. McWhir, J., Schnieke, A.E., Ansell, R., Wallace, H., Colman, A., Scott, A.R., and Kind, A.J. 1996. Selective ablation of differentiated cells permits isolation of embryonic stem cell lines from murine embryos with a non-permissive genetic background. Nature Genet. 14:223-226. Noben-Trauth, N., Kohler, G., Burki, K., and Ledermann, B. 1996. Efficient targeting of the IL-4 gene in a BALB/c embryonic stem cell line. Transgenic Res. 5:487-491. Pease, S., Braghetta, P., Gearing, D., Grail, D., and Williams, R.L. 1990. Isolation of embryonic stem (ES) cells in media supplemented with recombinant leukemia inhibitory factor (LIF). Dev. Bio. 141:344-352.



Robertson, E.J. 1987. Embryo-derived stem cell lines. In Teratocarcinomas and Embryonic Stem Cells: A Practical Approach (E.J. Robertson, ed.) pp.71-112. IRL Press, Oxford. Robertson, E.J., Evans, M.J., and Kaufman, M.H. 1983. X-chromosome instability in pluripotential stem cell lines derived from parthenogenetic embryos. J. Embryol. Exp. Morph. 74:297-309.

Key References Hogan, B., et al., 1994. See above. Robertson, E.J. 1987. See above. Wurst, W. and Joyner, A.L. 1993. Production of targeted embryonic stem cell clones. In Gene Targeting: A Practical Approach (A.L. Joyner, ed.) pp. 33-61. IRL Press, Oxford. These three references, written by experts in the field, represent the best compilations of ES culture and isolation methods.

Internet Resources http://www.biosupplynet.com Search this web site for “embryonic stem cell reagents” to obtain a current list of suppliers that provide medium and ES cells.




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Production of a Heterozygous Mutant Cell Line by Homologous Recombination (Single Knockout)

UNIT 23.5

Gene targeting by homologous recombination allows the introduction of specific mutations into any cloned gene. In the method described here, the gene of interest is inactivated by interrupting its coding sequence with a positive selectable marker (e.g., neo). Expression of neo is obtained by including the phosphoglycerate kinase (PGK) promoter in the construct. To enrich for clones in which the target gene has undergone homologous recombination over those in which random integration of the construct has occurred, a negative selectable marker, herpes simplex virus thymidine kinase (HSV-TK), is included in the construct outside the region of homology to the target gene. Depending upon the target gene, it may be easier to assemble the construct by adding the neo and TK genes to the cloned target gene or by adding two fragments of the target gene to a plasmid containing the neo and TK genes (e.g., pNTK, Fig. 23.5.1). If the Cre-loxP system (UNIT 23.1) is to be used for removing the selectable marker, or for tissue-specific or temporally controlled knockout, then a construct already containing loxP sites flanking the marker is more convenient (e.g., pTKLNL, Fig. 23.5.2A). If selection for loss of selectable marker is desired, a construct that also contains a negative selectable marker can be used (e.g., pTKLNCL, Fig. 23.5.2, panel B).

f1 (+) ori

Ap r

Sac I Bst XI Sac II Eag I Not I Eco RI pPGK

ColE1 ori pNTK 7 kb

KpnI ApaI Dra II Xho I AccI SaI I Cla l Hin dlII

TK

KpnI

neo

pPGK

SpeI BamHl SmaI Pst I EcoRl

Figure 23.5.1 pNTK vector. Both the neo and TK genes are driven by a PGK promoter (pPGK) that is expressed in ES cells. Unique restriction enzyme sites that are useful are indicated in bold. One genomic fragment can be cloned into the BamHI site. A second genomic fragment can be cloned into the HindIII, ClaI, Sal I, XhoI sites. A site should be preserved that will linearize the construct, leaving the majority of plasmid vector sequences attached to the TK gene (e.g., XhoI). Contributed by Richard Mortensen Current Protocols in Molecular Biology (2000) 23.5.1-23.5.11 Copyright © 2000 by John Wiley & Sons, Inc.



Sac II (7205)

A

SacI (7198) ColE1 ori

Not I (1) XbaI (8) pPGK AccI (815) SnaBI (819) KpnI (1110) HSV-TK

HSV-TK construct

Apr ScaI (5427) f1 (+) ori KpnI (4339)

XmaI (1685) SmaI (1687) NcoI (1998)

pTKLNL 7210 bp

XbaI (2126) BamHI (2138)

XhoI (4320) AccI (4315) SalI (4314) loxP HindIII (4157) ScaI (4045) 3′ PGK Bgl II (3878) Bgl II (3870)

XmaI (2144) SmaI (2146) loxP PGK

neo construct

neo NcoI (3338) ClaI (3578)

poly(A) PGK HincII (9316)

B HindIII(9157) 3′ PGK

SalI (9314) loxP

Xho I (1) Asp718 (16) Kpn I (20) f1 (+) ori

poly(A) PGK

CD

PGK poly(A) PGK Nco I (6015) neo

Apr CD construct

pTKLNCL 9319 bp

ColE1 ori HSV-TK construct neo construct

PGK loxP BamHI (4815) XbaI (4803) Not I (4796) Sac II (4790) NcoI (4655)

Production of a Heterozygous Mutant Cell Line

pMC1 HincII (3166) Asp718 (3764) KpnI (3768) HSV-TK

Figure 23.5.2 Constructs containing loxP sites surrounding a positive selectable marker, neo (A), or both a positive and a negative selectable marker, neo and cytosine deaminase (CD; B). Constructs can be made by insertion of homologous sequences in unique restriction sites outside the loxP sites. If conditional targeting constructs are desired (as in Fig. 23.1.7), a third loxP site can be inserted into the region of homology and then the two regions of homology inserted into the vectors. Another version of these plasmids is also available with the TK and CD reversed (Milstone et al., 1999).



STRATEGIC PLANNING Figure 23.5.3 illustrates the production, selection, and identification of targeted gene disruption by homologous recombination. A replacement targeting construct requires the assembly of several different DNA sequences: 1. A genomic clone (preferably >10 kb) of the gene of interest, generally encoded on a bacteriophage or cosmid clone containing homologous sequences to be included in the construct. DNA isogenic to the ES cells (i.e., derived from the same animal strain) is preferred but not essential. An alternative for genes of sufficient size, and regions

create construct

culture ES cells

RE

neo

2∗ electroporate with DNA

RE

RE 2

1 homologous recombination

TK

3∗

3

4

probe E

culture and select with G418 and GANC

RE

RE

neo

2∗

1

3∗

4

HR pick clones and expand


isolate DNA Nonhomologous sequences analyze by Southern hybridization

HR E

positive selectable marker target gene negative selectable marker vector promoter

Figure 23.5.3 Production, selection, and identification of targeted gene disruption by homologous recombination. An example of a restriction enzyme site (RE) and hybridization probe that can be used to identify cells in which homologous recombination has occurred (shaded colony) is shown. The predicted size of the restriction fragment generated from an unaltered target gene (E) and a target gene that has undergone homologous recombination (HR) is shown. If equal amounts of DNA are present in the lanes of the Southern blot, the intensity of each of the two hybridizing fragments from the DNA of a homologous recombinant clone will be half of the intensity of the hybridizing fragment from unaltered clones.



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for which mouse sequences are known, is long PCR (Cheng et al., 1994). This approach works for many genes as long as they have large enough introns, and is particularly convenient when the intron-exon structure of the gene is known. Long PCR may introduce mutations, which may decrease homologous recombination rates just as nonisogenic DNA can. In addition, the mutations may occur in important parts of the gene; this risk makes the approach less suitable for conditional knockouts or subtle mutations. 2. Additional cloned target-gene DNA sequences not included in the construct, to be used as a hybridization probe to identify homologous recombinants. This probe can often be isolated from the same bacteriophage or cosmid clone that provided the homologous sequences included in the construct. The hybridization probe will hybridize with either an unaltered target gene or a target gene that has undergone homologous recombination, but will not hybridize with a construct that entered the genome by random integration. 3. A positive selectable marker, such as the gene encoding neomycin phosphotransferase (neo) or hygromycin-B-phosphotransferase (hyg), which is used to disrupt the target gene. If a homozygous mutant cell line is an ultimate goal, it is recommended that the neo coding sequence contain the point mutation that decreases the phosphotransferase activity (Yenofsky et al., 1990). Using the PGK promoter and the wild-type neo gene may result in cells containing a single neo gene that are resistant to >10 mg/ml G418, thus precluding the use of higher G418 concentration to isolate clones containing two neo genes (see UNIT 23.6). 4. A negative selectable marker such as HSV-TK, which is used to enrich for ES cell clones in which homologous recombination has occurred in the target gene over clones in which random integration of the construct has occurred. BASIC PROTOCOL


GENE TARGETING IN EMBRYONIC STEM CELLS The basic protocol is divided into three parts. First, it outlines the assembly of a replacement targeting construct and considerations in choosing its exact structure. Second, it briefly describes the culture of embryonic stem (ES) cells and the method for introducing the construct DNA into ES cells. A more detailed description of culturing ES cells and maintaining their undifferentiated state is found in UNITS 23.2 & 23.3. Third, it outlines the method for identifying clones in which the target gene has been altered by homologous recombination. The resulting homologous recombinants are heterozygous (one allele of the target gene is altered by homologous recombination and one allele is normal) and can be used to produce transgenic murine lines or to produce homozygous mutant cell lines (in which both alleles of the target gene are altered; UNIT 23.6). Materials Target gene from genomic library isogenic with ES cell line (e.g., 129 SV library; Stratagene) Plasmid vector (e.g., pNTK, available from R. Mortensen; see Fig. 23.5.1) 95% ethanol Sterile H2O Embryonic stem (ES) cells (UNITS 23.2 & 23.3; ATCC) ES/LIF medium (see recipe) Trypsin/EDTA: 0.25% (w/v) trypsin/1 mM EDTA (20 mM HEPES, pH 7.3, optional) ES medium (see recipe) Electroporation buffer (see recipe)



G418 (UNIT 9.5) Gancyclovir (GANC) Freezing medium (see recipe) Digestion buffer (see recipe) Saturated NaCl (see recipe) 1% agarose gel (UNIT 2.5A) Tissue culture hood Gelatin-coated tissue culture plates (UNIT 23.3): 100-mm plates and 24-well microtiter plates 4-mm electroporation cuvettes Pipet tips, sterilized by autoclaving Nylon membrane Additional reagents and equipment for subcloning DNA (UNIT 3.16), restriction enzyme digestion (UNIT 3.1), phenol/chloroform extraction of DNA (UNIT 2.1A), agarose gel electrophoresis (UNIT 2.5A), electroporation (UNIT 9.3), ES cell culture (UNITS 23.2 & 23.3 and APPENDIX 3F), stable transformation using selective medium (UNIT 9.5), and Southern blotting and hybridization (UNITS 2.9 & 2.10) NOTE: All tissue culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise noted. Create a replacement construct 1. Select a portion of the target gene to include in the construct and a separate portion of the target gene to use as a probe for hybridization of Southern blots to identify cells in which homologous recombination has occurred (see step 19). It should contain a rare restriction site within an exon encoding an important region of the protein (or an exon upstream of such a region) that is ideally flanked by >1 kb of target gene DNA on each side (most constructs are made with 2 kb). The rate of homologous recombination may increase with increasing lengths of homologous DNA up to 15 kb.

2. Construct a clone in a plasmid vector (UNIT 3.16) such that the neo gene interrupts the gene of interest, leaving regions of homology on either side of the neo gene. Include a thymidine kinase (TK) gene in the replacement construct outside the regions of homology (Fig. 23.5.4). This construction can be accomplished by either adding the neo and TK sequences to the subcloned homologous sequences or by adding regions of homologous sequences to a plasmid already containing the neo and TK genes (e.g., pNTK, see Fig. 23.5.1).

3. Digest the construct DNA with a restriction enzyme to linearize (UNIT 3.1). Linearize the construct DNA so that the plasmid vector sequences remain attached to the TK gene. This will help preserve the activity of TK gene if any loss of DNA sequence occurs during random insertion of the construct into the genome.

4. Purify and sterilize the digested construct DNA by phenol/chloroform extraction (UNIT 2.1A). 5. Precipitate the DNA by adding 2 vol of 95% ethanol and microcentrifuging 30 sec. Using sterile technique in a tissue culture hood, remove the supernatant and allow the pellet to air dry until only slightly moist. 6. Dissolve the pellet in 100 µl sterile water. Check for complete digestion and estimate DNA concentration by electrophoresis on an agarose gel (UNIT 2.5A). Manipulating the Mouse Genome


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A

B neo

target gene

TK

point to linearize

point to linearize

neo

TK

target gene

pNTK Homologous sequences construct and target gene

Nonhomologous sequences positive selectable marker target gene negative selectable marker vector promoter

Figure 23.5.4 Two strategies to create a replacement construct. In method A the target gene fragment is subcloned into a plasmid vector, then pPGK-neo is inserted into a rare restriction enzyme site in the target-gene fragment and pPGK-TK is inserted into the plasmid vector near the target gene. In method B the target-gene fragment is cleaved into two pieces that are subcloned into the polylinker sites of pNTK (see Fig. 23.5.1). Note that the relative orientation of homologous fragments in the construct must retain that found in the target gene.

Transfect construct and select ES cells 7. Culture ES cells in ES/LIF medium (UNITS 23.2 & 23.3). Passage cells every 2 to 3 days by seeding a 100-mm gelatin-coated tissue culture plate with 1–2 × 106 cells/plate. Leukemia inhibitory factor (LIF) prevents ES cells from differentiating. Some investigators suggest passaging cells at a higher density if blastocyst injection of the cells (UNIT 23.4) is planned (e.g., 1.5 × 106 cells per 25-cm2 flask). A detailed description of culture techniques for ES cells is found in UNITS 23.2 & 23.3.

8. Harvest ∼5 × 106 to 1 × 107 cells by adding trypsin/EDTA and incubating for ∼5 min until cells are freed from the plate surface. Dissociate to single cells by pipetting up and down five to ten times. Add 5 ml ES medium. Pellet cells and resuspend the cell pellet in 1 ml electroporation buffer in the same tube. Typically, 107 cells can be obtained from a near-confluent 100-mm tissue culture plate.

9. Add 1 pmol linearized, sterile construct DNA from step 6. 10. Electroporate the mixture at 450 V and 250 µF in a 4-mm electroporation cuvette (UNIT 9.3). Incubate 10 min at room temperature. Many electroporation conditions can be used with ES cells.

11. Plate cells in ES medium at ∼2 × 106 cells per 100-mm gelatin-coated tissue culture plate. Incubate 24 hr. Production of a Heterozygous Mutant Cell Line

12. Begin selection (UNIT 9.5) by changing ES medium to ES/LIF medium and adding G418 to 0.2 mg/ml and GANC to 2 µM (final).



13. Continue incubation, changing medium daily using ES/LIF medium and adding G418 (0.2 mg/ml final) and GANC (2 µm final), until single, isolated colonies are visible (typically 1 week after electroporation). Remove an individual colony from the plate using an autoclaved pipet tip, and place in a 35-µl drop of trypsin/EDTA for 5 min. Pipet up and down about five times to dissociate cells. Transfer cells to a well of a gelatin-coated 24-well microtiter plate containing 1 ml ES/LIF medium. 14. Incubate until colonies are visible, but the cells are not differentiating (typically 3 to 4 days). Passage half of the cells to a well of a clean gelatin-coated 24-well microtiter plate. Add the remaining cells to 0.5 ml freezing medium and place at −70°C. Freeze overnight, then transfer to liquid nitrogen. Undifferentiated cells grow in smooth, round colonies. Differentiated cells are flatter with distinct intercellular boundaries. Proceed immediately to step 15 after placing half the cells in the freezer.

Screen for homologous recombinants 15. Incubate ES cells in 24-well microtiter plate (step 14) to near confluence (usually 2 to 3 days). Because it is not critical to prevent differentiation of the ES cells at this stage, LIF can be omitted from the culture medium; however, the presence of LIF may help to maintain cell growth.

16. Add 300 µl digestion buffer to each well. Transfer well contents to a 1.5-ml microcentrifuge tube, and incubate overnight at 55°C. 17. Add 150 µl saturated NaCl and vortex vigorously (the solution will turn milky white). Add 2 vol of 95% ethanol (the solution will turn clear except for precipitated DNA). Some investigators precipitate the DNA using 2 vol ethanol (or 1 vol isopropanol) without adding salt. However, the DNA pellet resuspends more easily if salt is added.

18. Resuspend DNA pellet in 50 µl water. Determine DNA concentration by measuring the absorbance at 260 nm (APPENDIX 3D). 19. Digest 10 µg DNA (or 10 µl if DNA concentration was not determined) with the appropriate restriction enzyme (UNIT 3.16). 20. Fractionate the digested DNA on a 1% agarose gel (UNIT 2.5A). Transfer to a nylon membrane, and hybridize by Southern blotting (UNITS 2.9 & 2.10) to the target-gene hybridization probe chosen in step 1 to distinguish the unaltered target gene from a target gene that has undergone homologous recombination. 21. Select ES cell colonies that show two hybridizing fragments of approximately equal intensity—one fragment of the predicted size for the unaltered target gene and one fragment of the predicted size for a target gene that has undergone homologous recombination. If the two fragments are of unequal hybridization intensity, the cell population may not be clonal. Freeze cells and store in liquid nitrogen. 22. If desired remove selectable markers that are flanked by loxP sites by transient expression of Cre (see Support Protocol).



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SUPPORT PROTOCOL

TRANSIENT EXPRESSION OF CRE FOR RECOMBINATION Removal of sequences between the lox sites is accomplished by transient expression of Cre recombinase. If flanking lox sites are present in both alleles, Cre will recombine both alleles as efficiently as one. If selectable markers are between lox sites (as in Fig. 23.1.6), then sensitivity to selection media (e.g., containing G418) will be restored. Additional Materials (also see Basic Protocol) Cre expression plasmid using a promoter giving high expression levels in ES cells (e.g., pMC1 or pPGK) 12.5 mg/ml 5-fluorocytosine in PBS (if selecting against CD), sterile 1. Expand the homologously recombined clones obtained using the Basic Protocol by culturing and harvesting ES cells (see Basic Protocol, steps 7 and 8). 2. Prepare Cre expression plasmid DNA using the same procedure as for the original targeting vector (see Basic Protocol, steps 4 to 6). Do not linearize the DNA, as this will increase the probability of genomic integration.

3. Use 1 to 2 pmol of this DNA to transfect the expanded ES cell clones (see Basic Protocol, step 10). Transfecting more DNA will most likely increase expression; however, it also increases the probability of integration.

4. Plate cells at a lower density than for the original targeting (since survival is expected to be higher). The target is a number of clones per plate that will allow convenient colony picking. If no selection is to be performed, plating at a few hundred electroporated cells per 100-mm plate is a reasonable starting point. A range of dilutions should be plated (at least to a few thousand per plate), since the exact survival is not accurately predictable and plating at low density will decrease survival. If the negative selectable marker cytosine deaminase (CD) is used, then plate at 1,000 to 10,000 cells per 100-mm plate, since higher densities will give complete killing due to a neighbor selection effect. The number of surviving colonies will depend on the frequency of recombination.

5. Continue to culture cells, replacing medium daily with fresh ES/LIF medium. If selecting against CD, include 250 µg/ml 5-fluorocytosine (from 12.5 mg/ml stock) in the medium. 6. Screen colonies loss of the selectable marker (Cre-induced recombination) by Southern analysis (see Basic Protocol, steps 14 to 21). Although Cre-construct integration is an unlikely event, the clones for injection can also be screened for presence of Cre by reprobing the Southern blots or by PCR.



Digestion buffer 20 mM Tris⋅Cl, pH 8.0 10 mM NaCl 10 mM EDTA 0.5% (w/v) sodium dodecyl sulfate (SDS) Store indefinitely at room temperature Add 1 mg/ml proteinase K just before use



Electroporation buffer 20 mM HEPES, pH 7.3 137 mM NaCl 5 mM KCl 0.7 mM Na2HPO4 6 mM glucose 0.1 mM 2-mercaptoethanol (2-ME) Store indefinitely at 4°C ES medium Dulbecco’s minimum essential medium (DMEM), high-glucose + pyruvate formulation, containing: 15% FBS, heat-inactivated 1 hr at 56°C 0.1 mM 2-mercaptoethanol (2-ME) 20 mM HEPES, pH 7.3 (optional) DMEM containing 4500 mg/liter D-glucose and pyruvate can be obtained from GIBCO/BRL. A detailed description of culture conditions for embryo stem cells has been presented by Robertson (1987).

ES/LIF medium ES medium (see recipe) containing 1000 U/ml leukemia inhibitory factor (LIF; GIBCO/BRL). Store ≤1 week at 4°C. Conditioned medium from a CHO cell line overproducing LIF (Genetics Institute) can also be used at a dilution of 1:1000. An alternative to LIF for preventing differentiation of ES cells is to grow them on feeder layers of irradiated mouse embryo fibroblasts (MEF) in ES medium. Some investigators add 20 mM HEPES (pH 7.3) to culture medium.

Freezing medium DMEM, high-glucose + pyruvate formulation, containing: 10% FBS (Hyclone), heat-inactivated 1 hr at 56°C 10% (v/v) dimethylsulfoxide (DMSO) 20 mM HEPES, pH 7.3 (optional) Store at −20°C Saturated NaCl Add NaCl to distilled H2O until no more dissolves (∼6 M). Some solid NaCl should remain; decant solution for use. Store indefinitely at room temperature. COMMENTARY Background Information See UNIT 23.1 for an overview of gene targeting by homologous recombination. Although homologous recombination has been used by yeast geneticists for some time, it has only recently been shown to occur in somatic mammalian cells. It was first demonstrated between exogenously introduced DNA sequences (Folger et al., 1982) and later between an exogenously introduced DNA construct and an endogenous gene (Smithies et al., 1985). The mechanism of homologous recombination is not well understood but a number of characteristics are known. Homologous recom-

bination occurs more readily if the construct has free ends, rather than being circular (Wong and Capecchi, 1987). The rate of homologous recombination does not depend on the number of targets in the genome—at least when the target is present as tandem repeats of a dihydrofolate reductase (DHFR) amplified gene (Zheng and Wilson, 1990). Initially homologous recombination in mammalian cells was studied by introducing a mutated neo or TK gene, then restoring neo or TK activity by correcting the mutation through homologous recombination (reviewed by Capecchi, 1989). This approach provided an Manipulating the Mouse Genome


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easy method to detect homologous recombination. Embryonic stem (ES) cells and the related embryonic carcinoma (EC or EK) cells were first isolated and cultured in 1981 (Evans and Kaufman, 1981; Martin, 1981). They can give rise to a chimeric mouse when introduced into a normal blastocyst, which is then transferred into the uterus of a pseudopregnant foster mother. The ES cells contribute to all tissues of the chimeric mouse including the germ line (Bradley et al., 1984). Currently, ES cells, rather than EC cells, are used to produce chimeric mice, because the extent of chimerism and efficiency of germ line transmission is much higher with normal cells. Most ES cell lines used are derived from males because the karyotype of XY cells is more stable than that of XX cells and resulting chimeric male mice are easier to breed. Typically, the extent of the contribution of the ES cells to somatic tissues of the chimeric mouse is easily determined visually by choosing strains of mice for the sources of ES cells and blastocysts that have different coat colors.

Critical Parameters


The degree of homology between the construct and the target genome can have a dramatic effect on the rate of homologous recombination in two ways. First, homologous recombination requires stretches of exact DNA homology. The DNA used to construct the targeting vector must be from the same species as the cell in which the mutation is to be introduced. It should also be isogenic with the target cell (this is not absolutely required, but increases the probability of success). Because animal strains may differ just as individual outbred animals differ, there may be a mismatch of DNA on average every 500 bp. A single DNA mismatch is sufficient to dramatically decrease the rate of homologous recombination (Deng and Capecchi, 1992; teRiele et al., 1992). Mutations induced by making constructs using long PCR may similarly decrease homologous recombination rates. Second, the rate of homologous recombination increases with increasing length of the homologous DNA sequence (within limits). The exact length of homologous DNA that gives the maximum recombination rate is controversial but may be as high as 15 kb (Deng and Capecchi, 1992; Hasty et al., 1991). Homology should also be >1 kb for the shorter arm (most constructs have used >2 kb). Further, fidelity of recombination can be lower if the

length of homology is 90% by coat color) and sex conversion, there is a high likelihood of germline transmission. In this case, only good male chimeras need to be mated. When the level of chimerism is lower and there is no evidence of sex conversion, all reasonable male and female chimeras should be mated. The possibility of injecting other targeted clones should be considered. Low-level chimeras ( 5), where N equals the number of backcross generations (Silver, 1995). For the N10 congenic ∼20 cM surrounding the targeted locus remain in the congenic. The production of congenics can be accelerated using a newer approach known as speed congenics (Lander and Schork, 1994; Wakeland et al., 1997). The mating strategy is identical to the traditional approach, except that the mouse containing the least amount of the donor genome is selected at each generation for subsequent matings. The relative genome contributions are determined by screening for polymorphic marker loci spread throughout the genome at generation N2 and all subsequent generations. The contaminating donor genome contribution can be reduced to the same level seen in the N10 congenic in 5 or 6 generations,



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Mut

WT

129 chimera

X

outcross

WT

Mut

WT

WT

C57BL/6

WT

X

F1

Het

C57BL/6 back-cross

N2

Mut

WT

heterozygote

X

C57BL/6

Mut

WT

N10

heterozygote

Figure 23.8.3 Generation of a congenic strain. Congenic lines are generated by an outcross to the host strain (C57BL/6) followed by successive back-crosses of mice carrying the mutant allele to the host strain. The strain is considered congenic after the tenth back-cross generation. Most loci except those tightly linked to the mutant locus are derived from the host strain and are homozygous. At this stage, brother-to-sister matings are initiated to generate experimental animals. Note that if all matings are performed with host females and donor males, the host Y chromosome will not be transferred. At least one back-cross between a donor female and host male followed by mating of a mutant male to a host female in the next back-cross is required to fix the C57BL/6 Y chromosome in the congenic line.

Mouse Colony Management



reducing the time required to make a congenic strain to 1.5 to 2 years. This rapid method requires larger colonies to select the “best” mouse for subsequent matings. In addition, few laboratories have the resources to perform the marker-assisted screening in house, although there are companies and core facilities that will perform the screening.

Long-Term Strain Maintenance There are several options for maintaining mutant mouse lines when there is no immediate need to generate animals for experimentation. Most investigators do not choose to eliminate mouse lines, because of the time and effort required to regenerate them. A line can be maintained by keeping only a few mating pairs. Homozygous mutants, if viable and fertile, should be used, so that genotyping will not be necessary. Let the mating pairs breed actively. Most litters can be culled. Set up new mating pairs from the younger generations when the litter size becomes smaller or the litters become less frequent. The number of active mating pairs depends on the fecundity of the line: more pairs should be kept for lines that produce small litters or lines that have difficulty rearing pups. This may also be a good opportunity to perform back-crosses if a congenic line might be needed in the future. Alternatively, morula (2.5-day old embryos) can be frozen using a controlled-rate freezing protocol and stored in liquid nitrogen. Several of the major mouse suppliers provide morula-freezing services. There have been recent advances in sperm cryopreservation. However, this is not yet the method of choice because of the variable success of in vitro fertilization with frozen mouse sperm. Finally, some of the major mouse suppliers will freeze, maintain, and distribute interesting mutant mouse lines if the investigator is willing to relinquish control over their distribution.

MOUSE IDENTIFICATION Individual animals must be identifiable when maintaining colonies of mice with different genotypes and backgrounds. A variety of methods have been used to mark individual mice including ear tagging, ear punching, toe clipping, subcutaneous transponder implantation, and tail tattooing. To a large extent, the method that a laboratory will use depends on personal preferences. However, the method must be acceptable to the institutional animal care committee. Note that there may be some variation in protocol acceptance between different institutions, so that the resident animal resource center should be consulted before choosing a protocol. None of the procedures will cause chronic pain. There are several practical factors to consider when choosing a method of identification (Table 23.8.2). Subcutaneous transponders are the most expensive of the identification methods. The typical cost of each transponder is between $5 and $10. In most cases, they are not easily reused. Readers typically cost between $500 and $2000. Implanted animals can be identified easily and the method is permanent in most cases. However, because of the lack of standardization, transponders from different companies usually require specific readers. Subcutaneous insertion is not difficult, but may require more restraint than for the other methods: anesthesia may be necessary until the procedure becomes routine. The cost of ear punching and toe clipping is negligible beyond the purchase of a punch or scissors. Marking the mice by these methods is simple, but may become tedious when large, continuous numbering schemes are used (see Hogan et. al., 1994 for numbering schemes). Reading the number may also be difficult when using continuous numbering schemes. Both methods are very useful for identifying neo-

Table 23.8.2 Methods of Identification

Method

Cost

Ease

Anesthesia

Comments

Ear tag Transponder

Low High

Simple Simple

No Yes/No

Tags can tear out of older animals Lack of standardization requires transponder specific readers

Ear punch (toe clip)

Low

Simple

No

Good for identification of neonates Large numbering schemes can be difficult to read

Tail tattoo

Moderate

Requires training

Yes

Labeling with complex numbering schemes can be tedious


Supplement 57

nates in a single litter when the mice are too small for other forms of identification. Note that neonates may also be marked with India ink for short-term identification. Tail tattooing is permanent. Tattooing devices can be purchased for $500 to $1000. Marking of many animals this way can be tedious because of the technique and the requirement for anesthesia. Ear tags are inexpensive and easy to apply. Mice should be tagged at weaning or later: tags are more likely to tear out when placed in the small ear of a younger animal. Tags should be clipped to the base of the ear, not to the edge. This will reduce the chance of tear-out. Tags may eventually tear out in older animals. Place even-numbered tags in the right ear and odd numbers in the left ear to reduce confusion when tags fall out; the torn or punctured ear will indicate whether the original tag number was even or odd.

DNA PREPARATION FOR GENOTYPE ANALYSIS Typically, the mice are tagged for identification and tail biopsied for genotyping simultaneously at weaning (age 3 to 4 weeks). At this age, biopsies are thought to be less traumatic because the tip of the tail is still cartilaginous. Check institutional guidelines for recommended procedures: an inhalation anesthetic may be required for older animals. DNA can be isolated from tail samples in several ways depending on the method of genotype analysis. PCR screens should be used whenever possible for the sake of speed and convenience. The following are step-by-step instructions for the abovementioned procedures.

Tail Biopsy


1. Remove 0.5 cm or less of the tip of the tail from a 3 to 4 week old pup using clean, sharp scissors. The same scissors can be used for multiple mice. The mice will not get infections at the cut site and cross-contamination of DNA samples by the scissors should not be a problem. A styptic pen can be used to stop the bleeding. However, there is very little blood loss at the cut site in normal mice. 2. Place the tail in a labeled 1.5-ml microcentrifuge tube. Samples can be collected over 1 hr or more at room temperature without affecting the quality of the final DNA. The biopsies can be frozen at −20°C or processed immediately after collection. When Southern analysis is used to detect large bands (>15 kb), samples should be processed after collection

because freezing will decrease the DNA quality, making the identification of large bands more difficult. 3. Prepare DNA using the standard protocol for Southern analysis and PCR, or the rapid protocol for PCR (see steps for each procedure below). The standard method produces DNA that is suitable for Southern analysis (UNIT 2.9A) or PCR (UNIT 15.1). It is an overnight procedure. The rapid method produces template for PCR in less than 30 min.

Standard DNA Isolation for Southern Analysis and PCR 1. Add 0.5 ml of digestion solution consisting of 100 mM Tris⋅Cl, pH 8.5 (APPENDIX 2), 5 mM EDTA, 200 mM NaCl, 0.5% Tween 20, and 1 mg/ml proteinase K to each tail biopsy. The proteinase K should be added fresh just prior to digestion. 2. Incubate overnight at 55°C. The tissue should dissociated completely leaving some undigested debris and hair at the bottom of the tube. Alternatively tails may be digested for ∼4 hr at 65°C with intermittent agitation. 3. Mix the samples by gentle vortexing and centrifuge at top speed in a microcentrifuge at room temperature for 2 min. 4. Pour the supernatant into a new 1.5 ml microcentrifuge tube containing 0.5 ml isopropanol. Mix the tubes thoroughly by inversion and let stand at room temperature for 10 to 30 min. 5. Centrifuge at top speed in a microcentrifuge at room temperature for 2 min. Aspirate the supernatant and wash the pellet once with 70% ethanol. 8. Allow the DNA pellet to air dry and then resuspend in 100 µl TE buffer, pH 7.6 (APPENDIX 2). The DNA can be dissolved by gentle vortexing. If the DNA pellet is allowed to dry out completely it may become difficult to dissolve. The DNA concentration will range from 0.5 to 2 µg/µl, depending on the size of the tail biopsy. The DNA is suitable for restriction digestion or as a PCR template. Store the DNA at 4°C.

Rapid DNA Isolation for PCR Analysis 1. Add 0.5 ml of 0.05 M NaOH to each tail biopsy. Incubate for 10 to 20 min at 95°C. 2. Remove the samples from the heat and neutralize by adding 50 µl of a solution of 1 M Tris⋅Cl, pH 8 (APPENDIX 2), and 10 mM EDTA. Vortex to mix. The tail does not dissociate completely with this method, but DNA is released into solution. 1 or 2 µl can be used directly in a 20-µl PCR reaction (UNIT 15.1). The



protocol can be scaled to work in a 96-well format.

INFORMATION MANAGEMENT Mouse colonies have a tendency to expand rapidly. Good record-keeping practices should be established immediately. Records serve several purposes. Colony sizes and expenses can be kept to a minimum when an accurate tally of mice is maintained. Phenotypic variation attributable to differences in strain background and subtle difference in fecundity are more easily detected when complete records are maintained. Paper-based records will become unwieldy for all but the smallest colonies. Fortunately, there are many options for computerbased record keeping. Table 23.8.3 lists some examples of colony management software including commercial and freeware options. Demos of most of the commercial applications can be tested prior to purchase. Many investigators may find that simple, locally constructed databases are most convenient. These can be adapted to individual lab requirements and are usually less expensive than commercial colony management software. Whether using commercial, freeware, or locally constructed databases, it is a good practice

to record information both about individual animals and matings. Tables 23.8.4 and 23.8.5 each list a minimum set of data that should be collected for individual mice and matings, respectively. Mating data should be recorded even when there are no offspring. This type of database is easily created using Filemaker Pro. Filemaker Pro is relatively inexpensive, crossplatform (Macintosh and PC), and simple to program. Databases can be accessed through a network, allowing multiple users to log on simultaneously. Several Filemaker Pro colony databases can be downloaded from the Web site listed in Table 23.8.3 for MouSeek. These templates may be used directly or modified to individual lab requirements. In addition, handheld (Palm-compatible) versions of Filemaker Pro databases are easily created and can be synchronized with the desktop application. Handheld copies are convenient for data entry and review inside the animal facility. Regardless of the record-keeping system, it is much more common to regret recording too little information than too much. However, a database is only as useful as the data it contains; the database must remain simple enough so that a laboratory group will actually record the information.

Table 23.8.3

Examples of Colony Management Softwarea

Program

Source

Platform

Comments

Colony

Locus Technology, Inc. http://www.locustechnology.com

Windows

Very complete packages, good for large-scale operations Options for barcode scanning and transponder interface

Big Bench Mouse Progeny

Big Bench Software http://www.bigbenchsoftware.com Progeny http://www.progeny2000.com Topaz Technologies http://www.topaztracks.com

Macintosh Windows Windows

Basic management software that can be networked Pedigree software that can be used for colony management Primarily a subscription service to access remotely administered database with web browser

LAMS

Mark McKie www.hgu.mrc.ac.uk/Softdata/LAMS

Windows

MouSeek

Caleb F. Davis Macintosh http://Mickey.utmem.edu/main/databases.html Windows

Scion

Macintosh Windows

Freeware for basic colony management Freeware Filemaker Pro template; modifiable

aSeveral software options are listed ranging from extensive commercial applications (Colony and Scion) to freeware alternatives (LAMS and MouSeek). Colony has the most options including handheld devices for editing data on site and may be most useful for managing large colonies. Big Bench Mouse is new and appears best suited for medium and small colonies. Progeny requires the user to define the relational database. This increases both flexibility and complexity. LAMS and MouSeek are inexpensive alternatives for medium and small colonies. Both are freeware, but MouSeek requires the purchase of Filemaker Pro. The Web site cited for MouSeek also provides links to two other free Filemaker Pro colony management templates. Most of the databases can be run on a network allowing multiple users to access data simultaneously.


Supplement 57

Table 23.8.4

Mouse Dataa

Field

Type

Description

Unique ID

Number

Tag/ID

Text

A serial number that uniquely identifies the record; usually created automatically The mouse tag number

Date of death Cause of death

Date Text

Mating number

Number

Date of death Used to differentiate natural deaths from experimental deaths or those for colony culling Unique mating number from the mating table

Genotype Notes

Text Text

Genotype For any non-standard annotations

Sex

Text

Sex of the mouse

aMother, father, strain, date of birth, and category are all defined in the corresponding mating record (Table 23.8.5).

Table 23.8.5

Mating Dataa

Field

Type

Description

Mating number

Number

Unique serial number to identify the mating record usually created automatically

Start date Date of birth

Date Date

Date the mating was begun Date of birth of litter

Category Strain

Text Text

Differentiates multiple targeted or transgenic lines The background strain of the mouse

Mother

Number

ID that uniquely identifies the mother—the unique number from the mouse table

Father

Number

Notes

Text

ID that uniquely identifies the father—the unique number from the mouse table For any non-standard annotations

No. of Males No. of Females

Number Number

Number of male offspring Number of female offspring

aMouse (Table 23.8.4) and mating data tables list a minimum set of data that should be recorded in colony management

software. With this basic data individual animals and their mating behavior can be tracked and genealogies can be reconstructed. A simple database in Filemaker Pro can be constructed from scratch by creating these two tables and linking them using the mating number. Each mouse has a record in the mouse table. Each mating, whether productive or not, has a record in the mating table.

LITERATURE CITED Festing, M.F.W., Simpson, E.M., Davisson, M.T., and Mobraaten, L.E. 1999. Revised nomenclature for strain 129 mice. Mammalian Genome, 10:836. Hogan, B., Beddington, R., Constantini, F., and Lacy, E. 1994. Manipulating the Mouse Embryo: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.


Kolesnikov, Y., Jain, S., Wilson, R., and Pasternak, G.W. 1998. Lack of morphine tolerance in 129/SvEv mice: Evidence for a NMDA receptor defect. J. Pharmacol. Exp. Ther. 284:455-459.

Lander, E.S. and Schork, N.J. 1994. Genetic dissection of complex traits. Science. 265:2037-2048. Silver, L.M. 1995. Creation of a congenic strain In Mouse Genetics: Concepts and Applications, Section 3.3.3.2 Oxford University Press, New York. Simpson, E.M., Linder, C.C., Sargent, E.E., Davisson, M.T., Mobraaten, L.E., and Sharp, J.J. 1997. Genetic variation among 129 substrains and its importance for targeted mutagenesis in mice. Nature Genet. 16:19-27. Threadgill, D.W., Yee, D., Matin, A., Nadeau, J.H., and Magnuson, T. 1997. Genealogy of the 129



inbred strains: 129SvJ is a contaminated inbred strain. Mammalian Genome 8:390-393. Wakeland, E., Morel, L., Achey, K., Yui, M., and Longmate, J. 1997. Speed congenics: A classic technique in the fast lane (relatively speaking). Immunol. Today 18:472-477.

KEY REFERENCES Banbury Conference on Genetic Background in Mice. 1997. Mutant mice and neuroscience: Recommendations concerning genetic background. Neuron 19:755-759.

Electronic transponders http://www.bmds.com Web site for BioMedic Data Systems, Inc., supplier of electronic transponders (Tel. 302-628-4100). http://www.minimitter.com Web site for MiniMitter Co., Inc. (Tel., 800-6852999). http://www.stoeltingco.com Web site for Stoelting Co. (Tel., 630-860-9700).

This viewpoint article reviews many of the concerns regarding the effects of genetic heterogeneity on experimental interpretation. Suggestions for standardized strain backgrounds and mating strategies are presented.

Software

Silver, L.M. 1995. Mouse genetics: Concepts and applications. Oxford University Press, New York.

Mice

This is a useful and thorough review of the house mouse and its use in genetics. The text is available on line at http://www.princeton.edu/~lsilver/book/MGcontents.html.

INTERNET RESOURCES

http://www.filemaker.com Web site for Filemaker, Inc., supplier of Filemaker Pro Database Software (Tel., 800-325-2747).

http://www.criver.com Web site for Charles River Laboratories, supplier of numerous mouse strains (Tel., 978-658-6000). http://www.taconic.com Web site for Taconic Farms, supplier of numerous mouse strains (Tel., 518-537-6208).

http://www.biosupplynet.com

http://www.jax.org

Search this Web site for animal husbandry related items to obtain a current list of suppliers.

Web site for The Jackson Laboratory, supplier of numerous mouse strains (Tel., 207-288-6000).

Ear tags and applicators http://www.jorvet.com Web site for Jorgensen Laboratories, Inc., supplier for ear tags and applicators (Tel., 800-525-5614).

Contributed by David A. Conner Harvard Medical School Cambridge, Massachusetts

http://www.nationalband.com Web site for National Band and Tag Company (Tel., 859-261-2035).



Supplement 57

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Overview of Agents from Combinatorial Nucleic Acid and Protein Libraries

UNIT 24.1

UNIT 24.1

will be published in an upcoming supplement.

Generation and Use of Combinatorial Libraries Current Protocols in Molecular Biology (2000) 24.1.1 Copyright © 2000 by John Wiley & Sons, Inc.


Design, Synthesis, and Amplification of DNA Pools for Construction of Combinatorial Pools and Libraries

UNIT 24.2

This unit describes the design, synthesis, and amplification of a random sequence DNA pool. Functional nucleic acid–binding or catalytic species can be selected from these random sequence pools. In designing the DNA pool, careful consideration should be given both to the degree of randomization and the length of the random sequence region (see Strategic Planning). Following pool design, chemical synthesis on a commercial DNA synthesizer will yield a single-stranded DNA pool. The newly synthesized oligonucleotide pool can then be purified (see Basic Protocol 1). Prior to amplification, the initial complexity of the pool should be determined (see Support Protocol 1), the skewing of the pool should be determined (see Support Protocol 2), and amplification reaction conditions should be optimized (Support Protocol 3). If the nascent synthetic oligonucleotide is judged to be suitable for large-scale amplification, it can be enzymatically converted into a double-stranded DNA library (see Basic Protocol 2). Multiple copies of a singlestranded DNA pool can be derived from each double-stranded DNA library, or the library can be transcribed to yield a RNA pool or a modified RNA pool (see UNIT 24.3). Figure 24.2.1 outlines the procedure. NOTE: The above comments and the protocols in this chapter are most relevant to the design of pools from which nucleic acids with binding activities (aptamers) or other enzymatic activities can be selected. However, many of the concepts and procedures are relevant for the design and synthesis of nucleotide pools that encode peptides of variable sequence. Considerations specific to the construction of nucleotide pools designed to encode peptides of variable sequence are given in UNIT 23.4. STRATEGIC PLANNING Designing the Initial DNA Pool The nucleic acid pools used for in vitro selection experiments typically contain a randomized central core flanked by constant sequences that are required for enzymatic manipulations, such as PCR amplification, in vitro transcription, or restriction digestion (see also Fig. 24.2.2). Since a pool is relatively expensive to synthesize, both in terms of time and cost, some effort should be devoted to pool design. There are many subtle parameters to consider that can greatly influence the outcome of a selection experiment, including the degree of randomization, pool length, and pool modularity (see Table 24.2.1 for references to selection experiments that have previously been successfully executed with different types and sizes of pools). Type of selection and degree of randomization Most researchers who carry out in vitro selection experiments wish to either better define or optimize a known binding site (binding-site selection), or to identify a nucleic acid that binds a particular ligand or site (aptamer selection; UNIT 24.3). Each of these tasks in turn requires the synthesis of different types of pools. The sequences and structures that contribute to known binding sites are frequently best defined by selections that start from partially randomized pools. One example of binding-site definition that started from a partially randomized pool was a selection that defined critical residues of the Rev-responContributed by Jack Pollard, Sabine D. Bell, and Andrew D. Ellington Current Protocols in Molecular Biology (2000) 24.2.1-24.2.24 Copyright © 2000 by John Wiley & Sons, Inc.

Generation and Use of Combinatorial Libraries


sive element (RRE) of HIV-1 Rev (Bartel et al., 1991). This experiment is also described in more detail below. Biased pools can also be used for the optimization of a previously isolated motif. For example, aptamers that could bind to the Rex protein of HTLV-1 were selected from a partially randomized pool based on the wild-type Rex-binding element (XBE) but in the end bound Rex 9-fold better than the XBE (Baskerville et al., 1995).

design pool

novel binding site

known binding site

partial randomization

segmental randomization

complete randomization

degree of randomization

length of random sequence tract

primer design

RNA pool? then add promoter

synthesize

purify if too low, resynthesize

if unacceptable, resynthesize

determine extension efficiency

determine composition

optimize amplification

large-scale amplification

Design and Construction of Combinatorial DNA Pools and Libraries

storage

Figure 24.2.1 Flow chart outlining pool design, synthesis, and large-scale amplification.



T7 promotor 5' –GCTAATACGACTCACTATAGGGAGATCACT StyI AvaI 5' – GCTAATACGACTCACTATAGGGAGATCACTTACGGCACC ----- Nx ------- CCAAGGCTCGGGACAGCG – 3' BanI

5' – CGCTGTCCCGAGCCTTGG

T7 promotor 5' – GATAATACGACTCACTATAGGGAATGGATCCACATCTACGA PstI HindIII 5' –GGGAATGGATCCACATCTACGAATTC ------ N30 ------- TTCACTGCAGACTTGACGAAGCTT– 3' BamHI EcoRI

5' – AAGCTTCGTCAAGTCTGCAGTGAA

Figure 24.2.2 Two examples of pools used in in vitro selection. Primers are shown above and below the sequence of the pool. The T7 promoter is delineated in bold. Restriction sites are underlined, with their enzymes listed.

Table 24.2.1

Selection Experiments with Different Types and Sizes of Pools

Target

DNA/RNA Length of random region Reference

Bacteriophage T4 DNA polymerase

RNA

8

Tuerk and Gold (1990)

HIV-1 Rev

RNA

Ribozyme HIV-1 Rev

RNA RNA

HIV-1 Rev

RNA

4 and 6, segmental; 6-9 and 6-9, segmental

PKCβ HTLV-1 Rex

RNA RNA

120 Conrad et al. (1994) 43, doped (70% wild Baskerville et al. (1995) type, 30% non-wild type)

66, doped (65% wild Bartel et al. (1991) type, 30% non-wild type, 5% deleted) 120 30

Bartel and Szostak (1993) Tuerk and MacDougalWaugh (1993) Giver et al. (1993)

In contrast, completely random sequence pools explore a much wider swath of sequence space and are more useful for the isolation of novel binding species (aptamers) or catalytic species (Breaker, 1997; Jaeger, 1997). There are many examples of the selection of novel binding sites from completely random sequence pools (reviewed in Gold et al., 1995; Osborne and Ellington, 1997). Even when a natural binding site is known in advance, a completely different binding site may be selected from a random sequence pool; for example, Tuerk and MacDougal-Waugh (1993) isolated unique binders to Rev that bound better than the wild-type RBE sequence in vitro. Completely random sequence pools can also be used to extract aptamers that bind to proteins not normally thought to bind to nucleic acids; an example of this is the selection of an RNA aptamer that bound and inhibited the β isoform of protein kinase C (Conrad et al., 1994). Completely random sequence pools can also be used for the selection of novel nucleic acid catalysts. For example, starting from a pool with a 220-position random region, Bartel and Szostak (1993) isolated a novel ribozyme capable of RNA ligation. Generally, selections for catalysis require pools with a random region greater than 90 residues, while binding selections use pools with a random region of less than 70 residues.



Supplement 52

Intermediate between partially random and completely random sequence pools are segmentally random sequence pools. In a segmentally random pool, short tracts of sequence are completely randomized. Segmental randomization thus allows all possible sequences within a short region or set of residues to be examined. Thus, if a natural binding site is known, but a portion of that binding site is suspected to be particularly important for function, then a segmentally random pool can be used to identify all possible, functional sequences within the wild-type sequence context. For example, Tuerk and Gold (1990) selected aptamers that bound T4 DNA polymerase from a pool that contained 8 random sequence positions flanked by wild-type residues. Similarly, many binding sites are known to be presented within a particular structural context, such as a stem-loop or stem-bulge structure. In these cases, a portion of the structure can be completely randomized, and all possible functional stem-loops or stem-bulges can be identified. For example, the Rev-binding element was known to form a stem-internal loop-stem structure. Giver et al. (1993) segmentally randomized only the internal loop portion of the structure and selected Rev-binding species. Many of the anti-Rev aptamers had sequences that were significantly different than the wild-type, yet were still presented in the context of a stem-internal loop-stem structure. Partially random (doped) pool design (binding site selection) The most important issue in the synthesis of a doped pool is the level of randomization (the probability of sequence substitution/position). As a general rule, the substitution frequency of a doped pool should roughly correspond to the number of positions thought to be required for function. For example, if 10 residues within a nucleic acid binding site are thought to be functional, then the rate of substitution might be set to yield single mutants at least half the time. If the substitution frequency is set too low, there may be too few varying residues or combinations of residues to yield information about functional sequences or structures. In contrast, if the substitution frequency is set too high, the sequence space nearest the wild-type motif will only be sparsely sampled, and many of the highly mutated molecules may be nonfunctional because their sequences will have diverged too far from the wild-type. For example, an in vitro genetic analysis has been used to uncover the critical structural interactions between the HIV-1 Rev protein and its primary RNA binding site, the Rev-binding element (Bartel et al., 1991). The RBE had previously been mapped by deletion analysis to a short segment of HIV-1. Bartel and his co-workers assumed that the minimal RBE was smaller even than the region identified by deletion analysis, and thus decided to heavily dope a portion of a 66-nucleotide sequence at a frequency of 35% substitution/position. The initial RRE library contained ∼1013 molecules that had an average of 23 substitutions/template (0.35 probability substitution/position × 66 positions = ∼23 substitutions); less than 1 in 1012 molecules were completely wild-type. Following selection, a 20-nucleotide core-binding site within the 66-nucleotide pool was readily defined by sequence conservations and covarying residues. A lower substitution rate might not have precisely defined the relatively small binding site, while an even higher substitution rate might have created a mutational load that would have limited the selection of functional molecules or even have allowed the selection of novel, non-wildtype anti-Rev aptamers (Giver et al., 1993; Tuerk and MacDougal-Waugh, 1993). Conversely, if the binding site were larger than originally hypothesized, the relatively high rate of substitution might have meant that few functional molecules could have survived the selection unscathed. Design and Construction of Combinatorial DNA Pools and Libraries

The number and type of sequence substitutions, as opposed to the probable target size for mutation, can also be used to plan the synthesis of a doped sequence pool, as described by the following equations. Typically, a 1-µmol synthesis of a 100-residue template yields



14

Percent of pool containing a given number of substitutions

12

18% substitution/position

10

8

35% substitution/position

6

4

2

0 0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40 42 44 46 48 50 52 54 56 58 60 62 64 66

Number of substitutions

Figure 24.2.3 Comparison of substitution distributions for a 66-nucleotide pool doped to either 18% or 35%.

a pool of ∼1015 amplifiable molecules. Regardless of the degree of partial randomization or the precise doping strategy employed, the number of different mutational combinations is given by: 3n{L!/[n!(L − n)!]} where n is the number of sequence substitutions/template in a template of length L. For example, in the case of the 66-nucleotide RRE pool discussed earlier, there were ∼2.17 × 109 possible 5-residue substitutions and ∼1.25 × 1016 possible 10-residue substitutions. To calculate what fraction of a given set of substitutions are actually contained in a doped pool, the binomial probability distribution can be used: P(n,L,f) = {L!/[n!(L − n)!]}(fn )(1 − f)(L−n) where P is the fraction of the template population when f is the probability of substitution/position. If primarily single-base substitutions are desired, then f should be maximized for n = 1; if multiple mutations (e.g., double or triple substitutions) are desired, then f should be correspondingly higher. If the doping strategy is optimized for n substitutions, then this number of substitutions will occur most frequently, “n − 1” and “n + 1” substitutions will occur less frequently but in roughly equal numbers, and so forth. Higher levels of sequence substitution skew the mutant frequency distribution, allowing the sampling of some regions of sequence space at the exclusion of others (Fig. 24.2.3). Therefore, in the RRE example already cited, a pool of 1 × 1013 molecules doped at a frequency of 35% would contain few 5-residue substitutions [1 × 1013 × P(5,66,0.35) = ∼1.82 × 106 5-residue substitutions out of ∼2.17 × 109 possible 5-residue substitutions]. In contrast, if the pool were doped at a frequency of 18%, all 5-residue substitutions would almost certainly be included [1 × 1013 × P(5,66,0.18) = ∼9.3 ×1010 5-residue substitutions].



Supplement 52

Note that in a pool of only 1 × 1013 total molecules, neither doping scheme would yield all possible 10-residue substitutions. Completely random pool design (aptamer selection) Completely random sequence pools are used to initiate selection experiments when no functional nucleic acid sequence or structural motif is known in advance. There is really only one parameter to consider when designing a completely random pool: the length of the random region. While we will consider this parameter in detail below, we must first dismiss a frequent bogey of selection neophytes, the issue of complexity and representation. Random sequence space is a vast landscape of possibilities of which only a vanishingly small fraction can be sampled by either nature or man. Assuming a 4-monomer repertoire from which pools can be constructed, there are ∼1.6 × 1060 unique individual sequences in a sequence space bounded by a 100-residue template (4100 = ∼1.6 × 1060), a quantity of nucleic acid greater than an Avogadro’s number of Earth masses. While this grotesquely large value is clearly beyond the realm of experimental possibility, modern methods of chemical nucleic acid synthesis do allow the sampling of nearly as much sequence information as may be contained in the Earth’s biosphere. As a back-of-the-envelope calculation, consider that there are on the order of ∼1× 109 species in the biosphere, each with ∼1 × 105 genes. If each of these genes in turn is composed of ∼1 × 103 residues, then there are ∼1 × 1017 residues worth of information in a biosphere. In contrast, a typical 1-µmol synthesis of a 100-residue random sequence pool would contain 1 × 1015 molecules × ∼1 × 102 residues/molecule = ∼1 × 1017 unique residues or roughly 1 biosphere’s worth of information. Obviously, the connection and ordering of sequence information in organisms is important as well. Typically, a random sequence pool contains ∼1 × 1015 molecules, and thus can potentially sample on the order of all possible 25-mers (415 = ∼1.1 × 1015). In fact, since different 25-mers can be found in different “reading frames,” a slightly larger sequence space will likely be sampled. Because of this physical restriction, it is sometimes thought that random sequence pools should be no more than 25 residues in length—any longer, and only a fractional sampling would be possible, and many potential sequences would be lost. While this is true, it should be realized that longer pools do not lose any of the numerical complexity of smaller pools (except in those instances where long syntheses are extremely inefficient) and in fact gain access to some fraction of longer sequence and structural motifs as well. For example, tRNA molecules are roughly 76 nucleotides in length. It might prove more difficult to select tRNA mimics from a random sequence population containing 30 randomized residues than from a pool spanning 80 randomized residues. However, any short functional tRNA mimics present in the shorter population should also be present in equal or greater number in the longer population. In most instances, the relative completeness of the pool is not a consideration in the success of a selection. Indeed, it has been shown that functional nucleic acids are not extremely rare (for recent reviews see Gold et al., 1995; Fitzwater and Polisky, 1996) and can be isolated both from “complete” pools that span 20 random sequence positions and from very “incomplete” pools that span 90 random sequence positions.


Having dismissed considerations of complexity and representation, the one guiding principle that emerges from this analysis is that longer pools are more generally useful for selection experiments than shorter pools. However, this principle must be applied with appropriate caveats. First, aptamers derived from shorter pools are easier to analyze. Sequence and structural motifs embedded within a 30-nucleotide random sequence region are much more readily apparent than sequence and structural motifs embedded within a



90-nucleotide random sequence region, especially if the motifs are not colinear. Second, longer pools are more difficult to synthesize than shorter pools. Finally, longer pools are more likely to yield amplification or other selection artifacts than shorter pools. For example, pools that contain random regions greater than 90 nucleotides in length can form self-aggregates that precipitate from solution upon prolonged incubation, and thus require immobilization on a solid support prior to selection (Bartel and Szostak, 1993; Lorsch and Szostak, 1994). Because of these considerations, pools used for the in vitro selection of aptamers typically contain from 20 to 80 random sequence positions. Longer pools are not only desirable but are likely required in selections for complex functions, such as catalysis. Pools used for the selection of ribozymes typically contain from 50 to 220 random sequence positions (for recent reviews see Gold et al., 1995; Fitzwater and Polisky, 1996). The optimal length of the random region is an active area of research (Sabeti et al., 1997) where many of the fundamental parameters remain to be defined. Practically, though, longer pools must be synthesized as oligonucleotides of 150 residues or fewer in length because of the constraints of DNA synthetic chemistry. For this reason, pools longer than 150 bases are typically generated in a modular fashion by ligating together individual, synthetic oligonucleotides (Bartel and Szostak, 1993). Segments of shorter DNAs can be stitched together by the inclusion of unique restriction sites (Bartel and Szostak, 1993). Asymmetric restriction sites, such as AvaI (C|YCGRG), BanI (G|GYRCC), and StyI (C|CWWGG), are very useful for this task since they minimize intra-pool dimerization via self-ligation. Also, these enzymes are cost-effective for digesting large amounts of DNA. Alternatively, an overlapping region can be included at the 3′ end of each synthetic oligonucleotide and mutually primed synthesis (e.g., UNIT 8.2) of a longer template can be carried out. After assembling pool modules, the complexity (yield) of the new, aggregate pool will need to be freshly assessed. The upper bound of the complexity of an assembled pool (e.g., 1011 100-mer modules × 1011 100-mer modules) will likely be much larger than its actual complexity (e.g., 100 micrograms of ligated 200-mer, 9.12 × 1014 molecules). Segmentally random pool design (binding site and aptamer selection) In general, the rules governing the design of segmentally random pools are idiosyncratic, depending on experimental purpose. If the desire is to better define a known binding site, then relatively short sequence tracts (i.e., from four to ten residues) should be completely randomized. The randomization of longer sequence tracts may lead to the selection of novel binding sites rather than variants of a known binding site. The residues can either be colinear (as is the case for many DNA binding sites) or dispersed (as is the case for many RNA binding sites). If the desire is to identify a binding site within the context of a known structural element, then from four to twenty residues can be completely randomized. In this instance, the fewer the number of residues that are randomized, the more likely it will be that the selected sequences will resemble a wild-type binding site or retain an engineered structure. The greater the number of residues that are randomized, the more likely it will be that a novel aptamer sequence or structure will be discovered. Primer design Generally, the constant sequences at the 5′ and 3′ ends of a pool function as primer-binding sites and can be almost any sequence or length. Primers of 20 nucleotides in length are convenient because their melting temperatures are convenient for the PCR and they can easily be synthesized in high yields. In designing constant sequences and complementary primers, obvious artifacts associated with the PCR, such as secondary-structure formation or self-association that could lead to the production of primer dimers, should be avoided. Computer programs such as the Genetic Computer Group’s PRIME or the Whitehead’s PRIMER3 assist in designing constant regions. Other primer design programs include



Supplement 52

Amplify (Bill Engels, Dept. of Genetics, University of Wisconsin, Madison) and Oligo (National Biosciences). As a rule of thumb, one should try to avoid using the same triplet sequence more than once in either constant region. Beyond these basal considerations, there are two schools of thought regarding the sequence of the priming site itself. On the one hand, designing primers to possess a 3′ clamp of 5′-WSS-3′ (IUB codes: W = A or T, S = C or G), such as ACC, ensures good extension by polymerases. On the other hand, the inclusion of A/T-rich regions at the 3′ termini of primers reduces the frequency of mispriming and allows virtually “infinite” multiplication of DNA amplicons (Crameri and Stemmer, 1993). The inclusion of restriction sites within primer regions can facilitate cloning of selected nucleic acids, although palindromes adjacent to the 3′ ends can also facilitate the genesis of primer-dimers. Finally, primers for partially randomized pools should be designed so that they do not conflict with the folding or accessibility of a known DNA or RNA binding site. It is suggested that the secondary structure of the wild-type binding site with any appended primer-binding sites be determined using an algorithm such as Mulfold (Jaeger et al., 1989). If the native or wild-type structure of the binding site is not among the most common folds, then the primers should be redesigned. If an RNA pool is to be constructed, runoff RNA transcripts for in vitro selection are frequently made with T7 RNA polymerase. There are several known promoters for T7 RNA polymerase (Milligan et al., 1987), but the following minimal sequence gives good yields: −17 −1 5′-TAA-TAC-GAC-TCA-CTA-TA-3′ Addition of a G and C residue at the −18 and −19 positions of the minimal promoter helps to close the DNA duplex and stabilize the 5′ end of the promoter region, thereby increasing transcriptional yields. Transcription initiation is optimal when there are stretches of purines in the +1 and +2 positions, with GG being the best initiator (Milligan et al., 1987). Transcriptional yields also increase if uridine does not appear in the transcript before position 6. Typical pool designs incorporating all the elements described are shown in Figure 24.2.2. Chemically Synthesizing the Pool


While pools of genomic DNA sequences have been used for selection (Singer et al., 1997), partially or completely random sequence pools must be chemically synthesized. Modern DNA synthesizers utilize phosphoramidite chemistry (UNIT 2.11; Beaucage and Caruthers, 2000) or H-phosphonate chemistry (Stromberg and Stawinski, 2000) and can routinely produce usable amounts of DNA up to 150 nucleotides in length. Longer oligonucleotides can also be synthesized, but side reactions such as branching and depurination accumulate throughout the synthesis and the amount of final, usable product recovered can be vanishingly small. Since stepwise coupling efficiencies for a long oligonucleotide are on average ≥98%, the typical yield of a 100-base synthesis that starts with a 1-µmol column is 13.5%, or 13.5 nmol, or 1 × 1016 different molecules, of which ∼10% to 30% can be enzymatically elongated or amplified. Several strategies can be used to enhance the synthetic yield of oligonucleotides that are longer than 100 bases (see UNIT 2.11). Further, if a pool longer than ∼150 nucleotides is desired, smaller pools can be modularly synthesized and coupled by ligation or mutually-primed synthesis (see discussion of completely random pool design, above). During synthesis it is wise to prevent the cross-contamination of primers with their corresponding pool. It has recently been



Table 24.2.2 Representative Calculations Based on the Masses and Efficiencies for Couplings that Utilize the Canonical Tetrazole Activation Chemistry and Phosphoramidites Bearing Standard Protecting Groups

Phosphoramidite

Molecular mass (g/mol)

5′-CE-dA 5′-CE-dC 5′-CE-dG 5′-CE-dT

858 834 840 745

Mass Coupling efficiency correction correction 0.87 0.89 0.89 1.00

0.67 0.67 1.00 0.83

Overall correction 0.58 0.60 0.89 0.83

discovered (A. Friedman, pers. comm.) that when pools and primers are synthesized on identical ports of a DNA synthesizer, there is some mixing of the molecules. The contamination is sufficient to yield a positive signal following extensive (30 to 50 PCR cycles) amplification of a no-template negative control. The unprogrammed interleaving of pools and primers can lead to extreme skewing of amplified materials, such that only a few species from the original pool may comprise a significant fraction of a subsequent amplification reaction. Therefore, pools and their cognate primers should be synthesized on different synthesizer ports and/or the machine should be extensively flushed with acetonitrile between syntheses. Most synthesizers can be programmed for in-line, degenerate mixing of bases. While this method is useful when only a few positions must be randomized, because of the extremely fast reaction of the activated phosphoramidite with the newly deprotected 5′ hydroxyl, random sequences will be skewed towards the phosphoramidite that first enters the column. Therefore, for longer pools or pools that should contain a statistically random distribution of nucleotides, it is better to manually mix the phosphoramidites off-line and use this mixture for the synthesis of degenerate sequence positions. A more stochastic distribution can be obtained by including larger amounts of A and C phosphoramidites in the mix to compensate for the faster coupling times of G and T phosphoramidites (Zon et al., 1985). Suggested ratios include a 3:3:2:2.4 molar ratio of A:C:G:T phosphoramidites (D.P. Bartel, pers. comm.), and a 1.5:1.25:1.15:1 molar ratio of A:C:G:T (see User’s Manual for PE Biosystems Models 392 and 394 DNA/RNA Synthesis). Doped pools are perhaps the most difficult to synthesize (Hermes et al., 1989; Bartel et al., 1991). Doping can be accomplished by using phosphoramidite mixtures that have been adjusted to ensure the proper level of partial randomization of a given nucleotide. For example, if a doped pool is to be synthesized in which non-wild-type residues are included at a rate of 10%/position, then for the adenosine bottle a molar ratio of 33.43:1.50:1.00:1.21 of A:C:G:T phosphoramidites should be used. These ratios were derived by first adjusting for the relative molecular mass and coupling differentials of the individual phosphoramidites and then mixing the phosphoramidite solutions on a percent Table 24.2.3 Volumes of Acetonitrile Needed to Dissolve 1 g of Phosphoramidite

Phosphoramidite

Dissolved in X ml of acetonitrile

5′-CE-dA 5′-CE-dC 5′-CE-dG 5′-CE-dT

11.6 12.0 17.8 16.6



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Table 24.2.4 Amidite Mixtures for Synthesis of Doped Pool in Which Non-Wild-Type Residues Are Included at a Rate of 10%/Position

Phosphoramidite A C G T A C G T

Mutagenesis Total volume (%) (ml) 10 10 10 10 20 20 20 20

10 10 10 10 10 10 10 10

Volume each amidite to mix (ml) A

C

G

T

9.00 0.33 0.33 0.33 8.00 0.67 0.67 0.67

0.33 9.00 0.33 0.33 0.67 8.00 0.67 0.67

0.33 0.33 9.00 0.33 0.67 0.67 8.00 0.67

0.33 0.33 0.33 9.00 0.67 0.67 0.67 8.00

volume basis to yield the desired extent of doping. This process is described in greater detail below. To normalize the coupling of different phosphoramidites, relative correction factors that take into account different coupling efficiencies and molecular masses must be calculated. Multiplying together these correction factors gives an overall correction factor to provide equal molar coupling of each phosphoramidite. Table 24.2.2 displays representative calculations based on the masses and efficiencies for couplings that utilize the canonical tetrazole activation chemistry (UNIT 2.11; Beaucage and Caruthers, 2000) and phosphoramidites bearing standard protecting groups [cyanoethyl for the phosphates along either isobutyryl (N-2 of guanine) or benzoyl (N-6 of adenine and N-4 of cytidine) groups]. Other chemistries and protections may require the substitution of other correction factors. Most modern synthesizers require that ∼1 g of phosphoramidite be dissolved in ∼20 ml of acetonitrile to be used in the coupling reaction. Applying this constraint along with the combined mass-coupling (overall) correction factor gives the volumes shown in Table 24.2.3 to dissolve 1 g of each phosphoramidite. Therefore, if equal volumes of each of these solutions are mixed, equal molar coupling should occur since the molar concentrations have been adjusted to account for both the mass and coupling differentials. As in the example above, if a doped pool is to be synthesized in which non-wild-type residues are included at a rate of 10%/position, then the amidites should be mixed as in Table 24.2.4.


In addition to varying nucleotide composition, it is also possible to vary the length of random sequence that is synthesized. Deletions can be stochastically incorporated during a synthesis by replacing the capping step with an acetonitrile wash (Bartel et al., 1991). It is more difficult to stochastically incorporate insertions, but the lengths of segmental random sequences in a pool can be mixed. For example, in Giver et al. (1993), four columns were used to generate a pool with two random regions of 6 to 9 positions separated by a constant domain. The first column was synthesized with 6 random positions, the second with 7 random positions, etc. Following the addition of the intervening constant sequence, the synthesis was stopped, the four columns were opened, and the resins from the four columns were mixed. The mixed resins were then equally redivided into four new columns and the synthesis was resumed. The first column incorporated 6 positions, the second column 7 positions, etc. Thus, the first column



contained oligonucleotides in which the first random segment was 6, 7, 8, or 9 residues long and a second random segment that was uniformly 6 residues long. The second column contained oligonucleotides in which the first random segment was 6, 7, 8, or 9 residues long and a second random segment was uniformly 7 residues long, and so forth. Following the completion of all four syntheses, the reactions were combined to generate the final random sequence pool. PURIFICATION OF A RANDOM SEQUENCE POOL A newly synthesized oligonucleotide pool should be purified on a denaturing polyacrylamide gel (see e.g., UNIT 2.12) prior to amplification. Oligonucleotides can also be purified using an HPLC or commercially available spin columns, but HPLC purification is not recommended for ssDNA pools, due to concerns about cross-contamination. Since oligonucleotides of equivalent length but different sequence migrate at slightly varying rates (see User’s Guide for PE Biosystems Expedite Nucleic Acid Synthesis System), a pool should appear as a broader band than a homogeneous sequence. In fact, because of the presence of capped failure sequences and depurinated, cleaved fragments, it is likely that the oligonucleotide product will appear even more heterogeneous.

BASIC PROTOCOL 1

As a general note, since sequences exist as single copies prior to amplification, individual species can be easily lost. Therefore, it is important to wash and elute the various filters, tubes, and tips described below one or more times. The eluates can then be pooled for a final precipitation and eventual amplification. Contamination of primers or other solutions with a synthesized or isolated pool should be avoided by using aerosol barrier tips. Similarly, gel plates used during purification should be washed thoroughly to ensure that they are free of contamination with other pools or primers. Materials DNA pool Ammonium hydroxide n-butanol TE buffer, pH 8.0 (APPENDIX 2) Urea loading buffer, 2× (UNIT 2.12) 5 M NaCl Ethanol Fluorescent TLC plate (VWR), wrapped in plastic wrap UV lamp Razor blades Small-bore syringes 13-ml centrifuge tubes capable of withstanding temperature extremes (Sarstedt) 90°C water bath Rotary shaker Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (e.g., UNIT 2.12) 1. After synthesis, deprotection, and cleavage from the solid support, lyophilize the oligonucleotide solution (in concentrated ammonium hydroxide) to dryness or precipitate with a 10-fold volume of n-butanol. The n-butanol precipitation can occur quite quickly at room temperature for longer oligonucleotides. Shorter (5 hr at room temperature), followed by a brief elution with an aqueous buffer (∼1 hr). Isoamyl alcohol extraction (UNIT 2.12) can be used to bring the extracts to a convenient volume for subsequent precipitation.



6. Precipitate the eluted oligonucleotide pool by adjusting the salt concentration to 0.3 M using a 5 M NaCl stock solution, then adding 3 vol of ethanol. Keep at –20°C for 3 hr, then microcentrifuge at maximum speed 4°C. Lyophilize to dryness. Resuspend the synthetic pool in TE buffer, pH 8.0 (to protect against nuclease contamination or drastic pH changes). If the volume of the eluted oligonucleotide is too large to conveniently precipitate, concentrate the sample by extracting against an equal volume of n-butanol. Remove the upper butanol layer and repeat until the aqueous volume is convenient for precipitation. About 1/5 of the aqueous layer is extracted into the organic butanol layer for every volume of butanol used. If too much butanol is used, thereby completely extracting the aqueous layer into the butanol, add more water and repeat the concentration.

DETERMINING THE POOL COMPLEXITY The number of different molecules present in a population can affect the outcome of a selection experiment (see Troubleshooting). If the pool complexity is too low for a given application, the pool will have to be resynthesized.

SUPPORT PROTOCOL 1

Pool complexity is, in turn, a function of yield and of the number of molecules in the pool that can be fully extended by a polymerase. The overall yield of the synthesis can be calculated by determining the UV absorption of the pool. However, deletions, incompletely deprotected residues, or backbone lesions that arise during chemical synthesis decrease by 10% to 40% the fraction of molecules in a synthetic pool that can be fully extended by polymerases. For example, the rate of insertions (presumably due to DMT cleavage via tetrazole) has been measured to be as high as 0.4% per position, and the rate of deletions (presumably due to incomplete capping) has been found to be as high as 0.5% per position (A. Keefe and D. Wilson, pers. comm.). The number of usable DNA molecules that are actually present in a nascent pool can be calculated by determining the fraction of the pool that can be extended by Taq polymerase. Materials Purified ssDNA pool and labeled primers 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2) 10 mM MgCl2 5 mM DTT [γ-32P]ATP (>3000 Ci/mmol) T4 polynucleotide kinase 1:1 phenol/chloroform (UNIT 2.1A) Chloroform 4.0 M ammonium acetate Taq DNA polymerase TE buffer, pH 8.0 (APPENDIX 2) PCR amplification buffer (see recipe) 2× formamide loading buffer (see recipe) 15 × 17–cm denaturing polyacrylamide gel (UNIT 2.12) Thermal cycler Phosphor imager plate and phosphor imager Additional reagents and equipment for quantitation of DNA (e.g., APPENDIX 3D), end-labeling of DNA (e.g., UNIT 3.10), phenol/chloroform and chloroform extraction of DNA (UNIT 2.1A), PCR amplification (e.g., Chapter 15), and denaturing polyacrylamide gel electrophoresis (UNIT 2.12)



Supplement 52

1. Quantitate DNA by UV absorption assuming that A260 of 1.0 indicates ∼37 µg/ml of single stranded DNA. Also see APPENDIX 3D.

2. Label the 5′ end of the 3′ PCR primer with [γ-32P]ATP by preparing the following reaction mixture. For 30-ìl reaction (volume of reaction and concentration of DNA and [γP]ATP will vary depending on application):

32

50 mM Tris⋅Cl, pH 7.5 10 mM MgCl2 5 mM DTT 1 to 50 pmol dephosphorylated DNA, 5′ ends 50 pmol (150 µCi) [γ-32P]ATP 50 µg/ml BSA 20 U T4 polynucleotide kinase Incubate 60 min at 37°C, then stop reaction by adding 1 µl of 0.5 M EDTA. Phenol/chloroform and chloroform extract the labeled oligonucleotide (UNIT 2.1A), and precipitate by adding an equal volume of 4.0 M ammonium acetate and 2 vol ethanol. Microcentrifuge to collect the pellet, remove the supernatant, and redissolve the labeled DNA pellet in 10 µl of TE buffer, pH 8.0. This procedure ensures that most of the unincorporated label remains in the supernatant.

3. Incubate ∼50 pmol of labeled primer with a 2- to 5-fold molar excess of pool in a 50-µl extension reaction, under the same conditions that will be used in the final amplification, in a thermal cycler as follows (see UNIT 15.1). a. Denature and anneal the primer and template DNA in PCR amplification buffer (usually 94°C for the denaturation step and ∼50°C for the annealing step). b. Add Taq or other DNA polymerase (scaled to the anticipated enzyme concentration to be used in the large-scale amplification), then increase the temperature to 72°C for 20 min. It may be useful to take time points to determine whether the reaction has gone to completion.

c. Finally, terminate the reaction by the addition of 2× formamide loading buffer. 4. Heat the extension reaction to 90°C for 3 min and load the reaction on a 15 × 17–cm denaturing polyacrylamide gel with appropriate radiolabeled size markers. Electrophorese until the dye is at or near the bottom of the gel, but do not let the radiolabeled primers run off. It is also useful to load a separate well with an aliquot of the primer alone. Choose an acrylamide percentage that allows efficient separation of small primers from larger extended products.

5. Dry and expose the gel to a phosphor imager plate. Using a phosphor imager, quantify the control primer band and the extended product band.


There may be a smear leading up to the extended band. One should use one’s best judgment in determining how much near-full-length material will be included in the quantitation. Calculate the percent extension by dividing counts of labeled, extended product by counts of labeled primer. Percent extension for a gel-purified ssDNA pool can range from 10% to 30%. The complexity of the pool is then the yield (determined in step 1) multiplied by the extension efficiency (percent extension determined above). If the complexity of the pool is insufficient for planned experiments, then the pool must be resynthesized.



DETERMINING THE POOL BIAS Following extension, the reaction should be repeated using a cold primer and the nonradioactive double-stranded DNA pool should be amplified in a PCR reaction, cloned (e.g., using a TA cloning kit from Invitrogen), and individual members sequenced to determine the degree of partial or completely randomness. The cloning step could also be carried out following PCR optimization (see Support Protocol 3). From 20 to 30 clones should be sequenced to determine the base composition of the starting pool. The random region should be composed of roughly 25% of each base. A pool with the random region skewed toward one or more bases (>30%) should be resynthesized. SMALL-SCALE PCR OPTIMIZATION OF POOL AMPLIFICATION To enhance yield and further avoid bias, the amplification conditions for a pool should be optimized prior to carrying out a large-scale amplification. Moreover, since amplifying a pool is costly in terms of both time and money, any optimization of the PCR should first take place on a small scale. The more involved large-scale amplification can then be carried out with confidence.

SUPPORT PROTOCOL 2

SUPPORT PROTOCOL 3

Materials dNTPs (UNIT 3.4) Taq DNA polymerase (e.g., Boehringer Mannheim) PCR amplification buffer containing 1.5 mM Mg2+ (see recipe) dsDNA mass markers (e.g., Life Technologies) 4% Nu Sieve agarose gel (FMC Bioproducts) Thermal cycler Densitometer Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5) 1. Carry out a 0.1 ml PCR reaction using 2 nM of synthetic pool oligonucleotide as template, 2 µM primers, and PCR buffer with 1.5 mM magnesium. Use the manufacturer’s suggested quantity of Taq (e.g., 2.5 U of Boehringer Mannheim Taq) in a reaction containing 200 µM dNTPs. A suggested temperature regime is: 10 to 15 cycles: 2 min 1 min

95°C 55°C

3 min

72°C

(denaturation) (dependent on primer composition) (annealing) (extension).

After 10 to 15 cycles of amplification, check the length and purity of the amplified DNA on a 4% Nu Sieve agarose gel in 1× TBE buffer (UNIT 2.5). A 0.1 ml reaction typically yields ∼1 ìg, but the amount can vary from 0.1 to 10 ìg. A fuzzy band may indicate that too many cycles of PCR have been carried out. In this case, set up the reaction again and perform fewer cycles.

2. Dilute the double-stranded PCR DNA product 1:128, and repeat the PCR reaction, removing a 5- to 10-µl aliquot during the last 10 sec of the cycle-7 extension step. Serially dilute the amplified product 1:2, 1:4, ... 1:128. Electrophorese all of the samples on a large agarose gel. Note that it is quite difficult to accurately pipet solutions at 72°C. It may therefore be desirable to pipet an amount slightly larger than that intended for use in the serial dilution. Generation and Use of Combinatorial Libraries


Supplement 52

3. Calculate the average PCR efficiency by identifying to what extent the cycle-7 PCR reaction is the result of progressive doublings of the original synthetic DNA. Determine which dilution lanes lack detectable DNA. The largest dilution that lacks detectable DNA is also the dilution that is a minimum estimate of the number of doublings. For example, if the 1/64 dilution is the largest dilution without detectable DNA, this implies that 6 “doublings” of the synthetic DNA yielded at least 64-fold more DNA. This is expressed as follows:

(average efficiency)no. of theoretical doublings (i.e., PCR cycles) = fold increase in DNA Thus, if 7 cycles of PCR were performed, then the average number of doublings per cycle is ∼1.81 [from (∼1.81)7 = 64].

4. Modulate PCR conditions to enhance PCR efficiency. If the pool’s average number of doublings per cycle is 5 ìM are generally not helpful). It may be useful to scan both above and below 2.5 ìM in 0.5-ìM increments. Magnesium concentration affects both primer annealing and the fidelity of Taq (which decreases with increasing magnesium concentration). Starting at the magnesium supplied in the PCR buffer (usually 1.5 mM), scan in 1-mM increments toward 5 mM as a maximal concentration. DNA denaturation at temperatures above 95°C is usually impractical since this greatly reduces Taq’s half-life. While other thermostable polymerases can be more resistant to higher temperatures, they usually have a lower extension efficiency and are more expensive than Taq. Annealing temperatures are dependent upon both primer sequence and length. The primer annealing temperatures should already be known from the primer design p rocess, o r may be calculated via an algorithm that can be found at http://paris.chem.yale.edu/extinct.html. This algorithm takes into account nucleotide composition, stacking energies (according to Turner’s rules), and empirical data. An annealing temperature ∼5°C less than the calculated annealing temperature is a good place to begin optimization. The amplification is more efficient at a lower annealing temperature, but mispriming and secondary structural problems are more pronounced. Higher temperatures improve the specificity, but decrease the overall yield of the reaction. To determine the optimum annealing temperature for a given primer and magnesium concentration, one should scan in both directions around the annealing temperature in 5°C increments. Finally, extension temperatures are modulated by the properties of Taq, which will extend (although inefficiently) at temperatures as low as 65°C. When extending at temperatures above Taq’s optimum temperature (70° to 75°C) somewhat more polymerase may be required; scanning of the enzyme quantity should be done in 2.5-U increments. However, too much Taq may be harmful to structured single-stranded nucleic acids (Lyamichev et al., 1993).


5. Confirm the results of the extension reaction described in Support Protocol 1 by the optimization method as follows. After optimizing pool PCR conditions for >1.8 average number of doublings per cycle, determine the pool complexity by performing another 0.1-ml PCR reaction with 2 nM of the original, synthetic pool oligonucleotide under the now optimized reaction conditions. After 7 or more cycles of PCR, perform agarose gel electrophoresis on serial dilutions of the PCR reaction adjacent to serial



dilutions of dsDNA mass markers. Calculate the amount of amplified DNA using either a densitometer or by estimating which dilutions are most similar. Calculate the approximate pool complexity as follows: g of PCR DNA after N cycles of PCR = g of starting extendable ssDNA g avg no. of doublings per cycle (see step 4) g of starting extendable ssDNA = mol of starting extendable ssDNA 330 g/mol × (no. of bases in full-length product) mol starting extendable ssDNA × (6.02 × 1023 ) = molecules of starting extendabless DNA molecules of starting extendable ssDNA = fraction of extendable ssDNA starting molecules fraction of extendable ssDNA × no. of synthetic pool molecules = pool complexity PCR efficiency should be optimized to balance the average number of doublings per cycle against the total reaction volume. A pool of 1 × 1015 molecules (∼1.7 × 109 mol) at a starting template concentration of 2 nM will require 0.85 L for amplification. Therefore, it is greatly desirable to amplify the pool at the highest template concentration that still gives a reasonable number of doublings per cycle. The amplification should generate at least 8 copies of pool DNA if the pool complexity is to be archived and preserved (see Basic Protocol 2).

LARGE-SCALE PCR AMPLIFICATION OF POOL DNA Very long and complex pools often require PCR amplification on a multiple-milliliter scale. Large-scale PCR differs from conventional PCR in that it is typically conducted in water baths using 15 ml, 17 × 120–mm, screw-capped (Sarstedt) thermostable tubes to accommodate the larger volumes. Amplification reactions of up to 2.5 L have been carried out in this way. Medium-scale amplifications can sometimes be carried out in thermal cyclers that can accommodate multiple samples (e.g., 96-well PCR plates).

BASIC PROTOCOL 2

Materials Purified ssDNA pool and primers EDTA 1:1 phenol/chloroform (UNIT 2.1A) Chloroform 4 M ammonium acetate Ethanol TE buffer, pH 8.0, containing 50 mM of a salt such as potassium chloride Thermal cycler or three water baths (one must be a circulating water bath) 96-well PCR plate or 13-ml thermostable tubes (Sarstedt) Thermometer Styrofoam racks Spectrophotometer or fluorometer Additional reagents and equipment for PCR amplification (UNIT 15.1; see Support Protocol 3 for determination of conditions on a small scale) and phenol/chloroform and chloroform extraction of DNA (UNIT 2.1A)



Supplement 52

Plan the reaction Since large-scale reactions are quite expensive in terms of nucleotides and enzyme, preparedness and planning for the large-scale amplification cannot be overemphasized. Primers

Current Protocols in Molecular Biology

Current protocols Molecular Biology

Current Protocols in Molecular Biology