Genes VIII

Molecular Biology GENES ARE DNA 1.1.1 Introduction Key Terms The genome is the complete set of sequences in the genet...

Author: Benjamin Lewin

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Molecular Biology

GENES ARE DNA

1.1.1 Introduction Key Terms The genome is the complete set of sequences in the genetic material of an organism. It includes the sequence of each chromosome plus any DNA in organelles. Nucleic acids are molecules that encode genetic information. They consist of a series of nitrogenous bases connected to ribose molecules that are linked by phosphodiester bonds. DNA is deoxyribonucleic acid, and RNA is ribonucleic acid. A gene (cistron) is the segment of DNA specifying production of a polypeptide chain; it includes regions preceding and following the coding region (leader and trailer) as well as intervening sequences (introns) between individual coding segments (exons). An allele is one of several alternative forms of a gene occupying a given locus on a chromosome. A locus is the position on a chromosome at which the gene for a particular trait resides; a locus may be occupied by any one of the alleles for the gene. Linkage describes the tendency of genes to be inherited together as a result of their location on the same chromosome; measured by percent recombination between loci.

The hereditary nature of every living organism is defined by its genome, which consists of a long sequence of nucleic acid that provides the information need to construct the organism. We use the term "information" because the genome does not itself perform any active role in building the organism; rather it is the sequence of the individual subunits (bases) of the nucleic acid that determines hereditary features. By a complex series of interactions, this sequence is used to produce all the proteins of the organism in the appropriate time and place. The proteins either form part of the structure of the organism, or have the capacity to build the structures or to perform the metabolic reactions necessary for life. The genome contains the complete set of hereditary information for any organism. Physically the genome may be divided into a number of different nucleic acid molecules. Functionally it may be divided into genes. Each gene is a sequence within the nucleic acid that represents a single protein. Each of the discrete nucleic acid molecules comprising the genome may contain a large number of genes. Genomes for living organisms may contain as few as 40,000 for Man. In this Chapter, we analyze the properties of the gene in terms of its basic molecular construction. Figure 1.1 summarizes the stages in the transition from the historical concept of the gene to the modern definition of the genome.

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Figure 1.1 A brief history of genetics.

The basic behavior of the gene was defined by Mendel more than a century ago. Summarized in his two laws, the gene was recognized as a "particulate factor" that passes unchanged from parent to progeny. A gene may exist in alternative forms. These forms are called alleles. In diploid organisms, which have two sets of chromosomes, one copy of each chromosome is inherited from each parent. This is the same behavior that is displayed by genes. One of the two copies of each gene is the paternal allele (inherited from the father), the other is the maternal allele (inherited from the mother). The equivalence led to the discovery that chromosomes in fact carry the genes. Each chromosome consists of a linear array of genes. Each gene resides at a particular location on the chromosome. This is more formally called a genetic locus. We can then define the alleles of this gene as the different forms that are found at this locus. The key to understanding the organization of genes into chromosomes was the discovery of genetic linkage. This describes the observation that alleles on the same chromosome tend to remain together in the progeny instead of assorting independently as predicted by Mendel's laws (see Molecular Biology Supplement 32.3 Linkage and mapping). Once the unit of recombination (reassortment) was introduced as the measure of linkage, the construction of genetic maps became possible. Introduction | SECTION 1.1.1 © 2004. Virtual Text / www.ergito.com

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On the genetic maps of higher organisms established during the first half of this century, the genes are arranged like beads on a string. They occur in a fixed order, and genetic recombination involves transfer of corresponding portions of the string between homologous chromosomes. The gene is to all intents and purposes a mysterious object (the bead), whose relationship to its surroundings (the string) is unclear. The resolution of the recombination map of a higher eukaryote is restricted by the small number of progeny that can be obtained from each mating. Recombination occurs so infrequently between nearby points that it is rarely observed between different mutations in the same gene. By moving to a microbial system in which a very large number of progeny can be obtained from each genetic cross, it became possible to demonstrate that recombination occurs within genes. It follows the same rules that were previously deduced for recombination between genes. Mutations within a gene can be arranged into a linear order, showing that the gene itself has the same linear construction as the array of genes on a chromosome. So the genetic map is linear within as well as between loci: it consists of an unbroken sequence within which the genes reside. This conclusion leads naturally into the modern view that the genetic material of a chromosome consists of an uninterrupted length of DNA representing many genes. A genome consists of the entire set of chromosomes for any particular organism. It therefore comprises a series of DNA molecules (one for each chromosome), each of which contains many genes. The ultimate definition of a genome is to determine the sequence of the DNA of each chromosome. The first definition of the gene as a functional unit followed from the discovery that individual genes are responsible for the production of specific proteins. The difference in chemical nature between the DNA of the gene and its protein product led to the concept that a gene codes for a protein. This in turn led to the discovery of the complex apparatus that allows the DNA sequence of gene to generate the amino acid sequence of a protein. Understanding the process by which a gene is expressed allows us to make a more rigorous definition of its nature. Figure 1.2 shows the basic theme of this book. A gene is a sequence of DNA that produces another nucleic acid, RNA. The DNA has two strands of nucleic acid, and the RNA has only one strand. The sequence of the RNA is determined by the sequence of the DNA (in fact, it is identical to one of the DNA strands). In many, but not in all cases, the RNA is in turn used to direct production of a protein. Thus a gene is a sequence of DNA that codes for an RNA; in protein-coding genes, the RNA in turn codes for a protein.


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Figure 1.2 A gene codes for an RNA, which may code for protein.

From the demonstration that a gene consists of DNA, and that a chromosome consists of a long stretch of DNA representing many genes, we move to the overall organization of the genome in terms of its DNA sequence. In Molecular Biology 1.2 The interrupted gene we take up in more detail the organization of the gene and its representation in proteins. In Molecular Biology 1.3 The content of the genome we consider the total number of genes, and in Molecular Biology 1.4 Clusters and repeats we discuss other components of the genome and the maintenance of its organization (for review see 1; 2; 5). Last updated on July 18, 2002


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Reviews 1.

Cairns, J., Stent, G., and Watson, J. D. (1966). Phage and the origins of molecular biology. Cold Spring Harbor Symp. Quant. Biol..

2.

Olby, R. (1974). . The Path to the Double Helix.

5.

Judson, H. (1978). . The Eighth Day of Creation.

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GENES ARE DNA

1.1.2 DNA is the genetic material of bacteria Key Terms Transformation of bacteria is the acquisition of new genetic material by incorporation of added DNA. Avirulent mutants of a bacterium or virus have lost the capacity to infect a host productively, that is, to make more bacterium or virus. The transforming principle is DNA that is taken up by a bacterium and whose expression then changes the properties of the recipient cell. Deoxyribonucleic acid (DNA) is a nucleic acid molecule consisting of long chains of polymerized (deoxyribo)nucleotides. In double-stranded DNA the two strands are held together by hydrogen bonds between complementary nucleotide base pairs. Key Concepts

• Bacterial transformation provided the first proof that DNA is the genetic material. Genetic properties can be transferred from one bacterial strain to another by extracting DNA from the first strain and adding it to the second strain.

The idea that genetic material is nucleic acid had its roots in the discovery of transformation in 1928. The bacterium Pneumococcus kills mice by causing pneumonia. The virulence of the bacterium is determined by its capsular polysaccharide. This is a component of the surface that allows the bacterium to escape destruction by the host. Several types (I, II, III) of Pneumococcus have different capsular polysaccharides. They have a smooth (S) appearance. Each of the smooth Pneumococcal types can give rise to variants that fail to produce the capsular polysaccharide. These bacteria have a rough (R) surface (consisting of the material that was beneath the capsular polysaccharide). They are avirulent. They do not kill the mice, because the absence of the polysaccharide allows the animal to destroy the bacteria. When smooth bacteria are killed by heat treatment, they lose their ability to harm the animal. But inactive heat-killed S bacteria and the ineffectual variant R bacteria together have a quite different effect from either bacterium by itself. Figure 1.3 shows that when they are jointly injected into an animal, the mouse dies as the result of a Pneumococcal infection. Virulent S bacteria can be recovered from the mouse postmortem.

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Figure 1.3 Neither heat-killed S-type nor live R-type bacteria can kill mice, but simultaneous injection of both can kill mice just as effectively as the live S-type.

In this experiment, the dead S bacteria were of type III. The live R bacteria had been derived from type II. The virulent bacteria recovered from the mixed infection had the smooth coat of type III. So some property of the dead type III S bacteria can transform the live R bacteria so that they make the type III capsular polysaccharide, and as a result become virulent (371). Figure 1.4 shows the identification of the component of the dead bacteria responsible for transformation. This was called the transforming principle. It was purified by developing a cell-free system, in which extracts of the dead S bacteria could be added to the live R bacteria before injection into the animal. Purification of the transforming principle in 1944 showed that it is deoxyribonucleic acid (DNA) (372).

Figure 1.4 The DNA of S-type bacteria can transform R-type bacteria into the same S-type.


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References 371. Griffith, F. (1928). The significance of pneumococcal types. J. Hyg. 27, 113-159. 372. Avery, O. T., MacLeod, C. M., and McCarty, M. (1944). Studies on the chemical nature of the substance inducing transformation of pneumococcal types. J. Exp. Med. 98, 451-460.



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1.1.3 DNA is the genetic material of viruses Key Concepts

• Phage infection proved that DNA is the genetic material of viruses. When the DNA and protein components of bacteriophages are labeled with different radioactive isotopes, only the DNA is transmitted to the progeny phages produced by infecting bacteria.

Having shown that DNA is the genetic material of bacteria, the next step was to demonstrate that DNA provides the genetic material in a quite different system. Phage T2 is a virus that infects the bacterium E. coli. When phage particles are added to bacteria, they adsorb to the outside surface, some material enters the bacterium, and then ~20 minutes later each bacterium bursts open (lyses) to release a large number of progeny phage. Figure 1.5 illustrates the results of an experiment in 1952 in which bacteria were infected with T2 phages that had been radioactively labeled either in their DNA component (with 32P) or in their protein component (with 35S). The infected bacteria were agitated in a blender, and two fractions were separated by centrifugation. One contained the empty phage coats that were released from the surface of the bacteria. The other fraction consisted of the infected bacteria themselves.

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Figure 1.5 The genetic material of phage T2 is DNA.

Most of the 32P label was present in the infected bacteria. The progeny phage particles produced by the infection contained ~30% of the original 32P label. The progeny received very little – less than 1% – of the protein contained in the original phage population. The phage coats consist of protein and therefore carried the 35S radioactive label. This experiment therefore showed directly that only the DNA of the parent phages enters the bacteria and then becomes part of the progeny phages, exactly the pattern of inheritance expected of genetic material (373). A phage (virus) reproduces by commandeering the machinery of an infected host cell to manufacture more copies of itself. The phage possesses genetic material whose behavior is analogous to that of cellular genomes: its traits are faithfully reproduced, and they are subject to the same rules that govern inheritance. The case of T2 reinforces the general conclusion that the genetic material is DNA, whether part of the genome of a cell or virus.


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References 373. Hershey, A. D. and Chase, M. (1952). Independent functions of viral protein and nucleic acid in growth of bacteriophage. J. Gen. Physiol. 36, 39-56.



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GENES ARE DNA

1.1.4 DNA is the genetic material of animal cells Key Terms Transfection of eukaryotic cells is the acquisition of new genetic markers by incorporation of added DNA. Key Concepts

• DNA can be used to introduce new genetic features into animal cells or whole animals.

• In some viruses, the genetic material is RNA.

When DNA is added to populations of single eukaryotic cells growing in culture, the nucleic acid enters the cells, and in some of them results in the production of new proteins. When a purified DNA is used, its incorporation leads to the production of a particular protein (2486). Figure 1.6 depicts one of the standard systems.

Figure 1.6 Eukaryotic cells can acquire a new phenotype as the result of transfection by added DNA.

Although for historical reasons these experiments are described as transfection DNA is the genetic material of animal cells | SECTION 1.1.4 © 2004. Virtual Text / www.ergito.com

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when performed with eukaryotic cells, they are a direct counterpart to bacterial transformation. The DNA that is introduced into the recipient cell becomes part of its genetic material, and is inherited in the same way as any other part. Its expression confers a new trait upon the cells (synthesis of thymidine kinase in the example of the figure). At first, these experiments were successful only with individual cells adapted to grow in a culture medium. Since then, however, DNA has been introduced into mouse eggs by microinjection; and it may become a stable part of the genetic material of the mouse (see Molecular Biology 4.18.18 Genes can be injected into animal eggs). Such experiments show directly not only that DNA is the genetic material in eukaryotes, but also that it can be transferred between different species and yet remain functional. The genetic material of all known organisms and many viruses is DNA. However, some viruses use an alternative type of nucleic acid, ribonucleic acid (RNA), as the genetic material. The general principle of the nature of the genetic material, then, is that it is always nucleic acid; in fact, it is DNA except in the RNA viruses.

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References 2486. Pellicer, A., Wigler, M., Axel, R., and Silverstein, S. (1978). The transfer and stable integration of the HSV thymidine kinase gene into mouse cells. Cell 14, 133-141.


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GENES ARE DNA

1.1.5 Polynucleotide chains have nitrogenous bases linked to a sugar-phosphate backbone Key Concepts

• A nucleoside consists of a purine or pyrimidine base linked to position 1 of a pentose sugar.

• Positions on the ribose ring are described with a prime ( ′ ) to distinguish them. • The difference between DNA and RNA is in the group at the 2 ′ position of the

sugar. DNA has a deoxyribose sugar (2 ′ –H); RNA has a ribose sugar (2 ′ –OH).

• A nucleotide consists of a nucleoside linked to a phosphate group on either the 5 ′ or 3 ′ position of the (deoxy)ribose.

• Successive (deoxy)ribose residues of a polynucleotide chain are joined by a

phosphate group between the 3 ′ position of one sugar and the 5 ′ position of the next sugar.

• One end of the chain (conventionally the left) has a free 5 ′ end and the other end has a free 3 ′ end.

• DNA contains the four bases adenine, guanine, cytosine, and thymine; RNA has uracil instead of thymine.

The basic building block of nucleic acids is the nucleotide. This has three components: • a nitrogenous base; • a sugar; • and a phosphate. The nitrogenous base is a purine or pyrimidine ring. The base is linked to position 1 on a pentose sugar by a glycosidic bond from N1 of pyrimidines or N9 of purines. To avoid ambiguity between the numbering systems of the heterocyclic rings and the sugar, positions on the pentose are given a prime ( ′ ). Nucleic acids are named for the type of sugar; DNA has 2 ′ –deoxyribose, whereas RNA has ribose. The difference is that the sugar in RNA has an OH group at the 2 ′ position of the pentose ring. The sugar can be linked by its 5 ′ or 3 ′ position to a phosphate group. A nucleic acid consists of a long chain of nucleotides. Figure 1.7 shows that the backbone of the polynucleotide chain consists of an alternating series of pentose Polynucleotide chains have nitrogenous bases linked to a sugar-phosphate backbone | SECTION 1.1.5 © 2004. Virtual Text / www.ergito.com

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Figure 1.7 A polynucleotide chain consists of a series of 5 ′ -3 ′ sugar-phosphate links that form a backbone from which the bases protrude.

Each nucleic acid contains 4 types of base. The same two purines, adenine and guanine, are present in both DNA and RNA. The two pyrimidines in DNA are cytosine and thymine; in RNA uracil is found instead of thymine. The only difference between uracil and thymine is the presence of a methyl substituent at position C5. The bases are usually referred to by their initial letters. DNA contains A, G, C, T, while RNA contains A, G, C, U. The terminal nucleotide at one end of the chain has a free 5 ′ group; the terminal nucleotide at the other end has a free 3 ′ group. It is conventional to write nucleic acid sequences in the 5 ′ → 3 ′ direction – that is, from the 5 ′ terminus at the left to the 3 ′ terminus at the right. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.5

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GENES ARE DNA

1.1.6 DNA is a double helix Key Terms Base pairing describes the specific (complementary) interactions of adenine with thymine or of guanine with cytosine in a DNA double helix (thymine is replaced by uracil in double helical RNA). Complementary base pairs are defined by the pairing reactions in double helical nucleic acids (A with T in DNA or with U in RNA, and C with G). Antiparallel strands of the double helix are organized in opposite orientation, so that the 5 ′ end of one strand is aligned with the 3 ′ end of the other strand. The minor groove of DNA is 12Å across. The major groove of DNA is 22Å across. A helix is said to be right-handed if the turns runs clockwise along the helical axis. B-form DNA is a right-handed double helix with 10 base pairs per complete turn (360°) of the helix. This is the form found under physiological conditions whose structure was proposed by Crick and Watson. A stretch of overwound DNA has more base pairs per turn than the usual average (10 bp = 1 turn). This means that the two strands of DNA are more tightly wound around each other, creating tension. A stretch of underwound DNA has fewer base pairs per turn than the usual average (10 bp = 1 turn). This means that the two strands of DNA are less tightly wound around each other; ultimately this can lead to strand separation. Key Concepts

• The B-form of DNA is a double helix consisting of two polynucleotide chains that run antiparallel.

• The nitrogenous bases of each chain are flat purine or pyrimidine rings that face

inwards and pair with one another by hydrogen bonding to form A-T or G-C pairs only.

• The diameter of the double helix is 20 Å, and there is a complete turn every 34 Å, with 10 base pairs per turn.

• The double helix forms a major (wide) groove and a minor (narrow) groove.

The observation that the bases are present in different amounts in the DNAs of different species led to the concept that the sequence of bases is the form in which genetic information is carried. By the 1950s, the concept of genetic information was common: the twin problems it posed were working out the structure of the nucleic acid, and explaining how a sequence of bases in DNA could represent the sequence of amino acids in a protein.

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Three notions converged in the construction of the double helix model for DNA by Watson and Crick in 1953: • X-ray diffraction data showed that DNA has the form of a regular helix, making a complete turn every 34 Å (3.4 nm), with a diameter of ~20 Å (2 nm). Since the distance between adjacent nucleotides is 3.4 Å, there must be 10 nucleotides per turn. • The density of DNA suggests that the helix must contain two polynucleotide chains. The constant diameter of the helix can be explained if the bases in each chain face inward and are restricted so that a purine is always opposite a pyrimidine, avoiding partnerships of purine-purine (too wide) or pyrimidine-pyrimidine (too narrow). • Irrespective of the absolute amounts of each base, the proportion of G is always the same as the proportion of C in DNA, and the proportion of A is always the same as that of T. So the composition of any DNA can be described by the proportion of its bases that is G + C. This ranges from 26% to 74% for different species. Watson and Crick proposed that the two polynucleotide chains in the double helix associate by hydrogen bonding between the nitrogenous bases. G can hydrogen bond specifically only with C, while A can bond specifically only with T. These reactions are described as base pairing, and the paired bases (G with C, or A with T) are said to be complementary. The model proposed that the two polynucleotide chains to run in opposite directions (antiparallel), as illustrated in Figure 1.8. Looking along the helix, one strand runs in the 5 ′ → 3 ′ direction, while its partner runs 3 ′ → 5 ′ (374; 376; 375).


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Figure 1.8 The double helix maintains a constant width because purines always face pyrimidines in the complementary A-T and G-C base pairs. The sequence in the figure is T-A, C-G, A-T, G-C.

The sugar-phosphate backbone is on the outside and carries negative charges on the phosphate groups. When DNA is in solution in vitro, the charges are neutralized by the binding of metal ions, typically by Na+. In the cell, positively charged proteins provide some of the neutralizing force. These proteins play an important role in determining the organization of DNA in the cell. The bases lie on the inside. They are flat structures, lying in pairs perpendicular to the axis of the helix. Consider the double helix in terms of a spiral staircase: the base pairs form the treads, as illustrated schematically in Figure 1.9. Proceeding along the helix, bases are stacked above one another, in a sense like a pile of plates.


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Figure 1.9 Flat base pairs lie perpendicular to the sugar-phosphate backbone.

Each base pair is rotated ~36° around the axis of the helix relative to the next base pair. So ~10 base pairs make a complete turn of 360°. The twisting of the two strands around one another forms a double helix with a minor groove (~12 Å across) and a major groove (~22 Å across), as can be seen from the scale model of Figure 1.10. The double helix is right-handed; the turns run clockwise looking along the helical axis. These features represent the accepted model for what is known as the B-form of DNA.


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Figure 1.10 The two strands of DNA form a double helix.

It is important to realize that the B-form represents an average, not a precisely specified structure. DNA structure can change locally. If it has more base pairs per turn it is said to be overwound; if it has fewer base pairs per turn it is underwound. Local winding can be affected by the overall conformation of the DNA double helix in space or by the binding of proteins to specific sites. Last updated on February 9, 2004


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References 374. Watson, J. D., and Crick, F. H. C. (1953). A structure for DNA. Nature 171, 737-738. 375. Watson, J. D., and Crick, F. H. C. (1953). Genetic implications of the structure of DNA. Nature 171, 964-967. 376. Wilkins, M. F. H., Stokes, A. R., and Wilson, H. R. (1953). Molecular structure of DNA. Nature 171, 738-740.



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1.1.7 DNA replication is semiconservative Key Terms A parental strand or duplex of DNA refers to the DNA that will be replicated. The antisense strand (Template strand) of DNA is complementary to the sense strand, and is the one that acts as the template for synthesis of mRNA. A daughter strand or duplex of DNA refers to the newly synthesized DNA. Semiconservative replication is accomplished by separation of the strands of a parental duplex, each then acting as a template for synthesis of a complementary strand. Key Concepts

• The Meselson-Stahl experiment used density labeling to prove that the single

polynucleotide strand is the unit of DNA that is conserved during replication.

• Each strand of a DNA duplex acts as a template to synthesize a daughter strand. • The sequences of the daughter strands are determined by complementary base pairing with the separated parental strands.

It is crucial that the genetic material is reproduced accurately. Because the two polynucleotide strands are joined only by hydrogen bonds, they are able to separate without requiring breakage of covalent bonds. The specificity of base pairing suggests that each of the separated parental strands could act as a template strand for the synthesis of a complementary daughter strand. Figure 1.11 shows the principle that a new daughter strand is assembled on each parental strand. The sequence of the daughter strand is dictated by the parental strand; an A in the parental strand causes a T to be placed in the daughter strand, a parental G directs incorporation of a daughter C, and so on.

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Figure 1.11 Base pairing provides the mechanism for replicating DNA.

The top part of the figure shows a parental (unreplicated) duplex that consists of the original two parental strands. The lower part shows the two daughter duplexes that are being produced by complementary base pairing. Each of the daughter duplexes is identical in sequence with the original parent, and contains one parental strand and one newly synthesized strand. The structure of DNA carries the information needed to perpetuate its sequence. The consequences of this mode of replication are illustrated in Figure 1.12. The parental duplex is replicated to form two daughter duplexes, each of which consists of one parental strand and one (newly synthesized) daughter strand. The unit conserved from one generation to the next is one of the two individual strands comprising the parental duplex. This behavior is called semiconservative replication.


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Figure 1.12 Replication of DNA is semiconservative.

The figure illustrates a prediction of this model. If the parental DNA carries a "heavy" density label because the organism has been grown in medium containing a suitable isotope (such as 15N), its strands can be distinguished from those that are synthesized when the organism is transferred to a medium containing normal "light" isotopes. The parental DNA consists of a duplex of two heavy strands (red). After one generation of growth in light medium, the duplex DNA is "hybrid" in density – it consists of one heavy parental strand (red) and one light daughter strand (blue). After a second generation, the two strands of each hybrid duplex have separated; each gains a light partner, so that now half of the duplex DNA remains hybrid while half is entirely light (both strands are blue). The individual strands of these duplexes are entirely heavy or entirely light. This pattern was confirmed experimentally in the Meselson-Stahl experiment of 1958, which followed the semiconservative replication of DNA through three generations of growth of E. coli. When DNA was extracted from bacteria and its density measured by centrifugation, the DNA formed bands corresponding to its density – heavy for parental, hybrid for the first generation, and half hybrid and half light in the second generation (377; for review see 2524).


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Reviews 2524. Holmes, F. (2001). . Meselson, Stahl, and the Replication of DNA: A History of The Most Beautiful Experiment in Biology.

References 377. Meselson, M. and Stahl, F. W. (1958). The replication of DNA in E. coli. Proc. Natl. Acad. Sci. USA 44, 671-682.



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1.1.8 DNA strands separate at the replication fork Key Terms A replication fork (Growing point) is the point at which strands of parental duplex DNA are separated so that replication can proceed. A complex of proteins including DNA polymerase is found at the fork. A DNA polymerase is an enzyme that synthesizes a daughter strand(s) of DNA (under direction from a DNA template). Any particular enzyme may be involved in repair or replication (or both). RNA polymerases are enzymes that synthesize RNA using a DNA template (formally described as DNA-dependent RNA polymerases). A deoxyribonuclease (DNAase) is an enzyme that attacks bonds in DNA. It may cut only one strand or both strands. Ribonucleases (RNAase) are enzymes that cleave RNA. They may be specific for single-stranded or for double-stranded RNA, and may be either endonucleases or exonucleases. Exonucleases cleave nucleotides one at a time from the end of a polynucleotide chain; they may be specific for either the 5 ′ or 3 ′ end of DNA or RNA. Endonucleases cleave bonds within a nucleic acid chain; they may be specific for RNA or for single-stranded or double-stranded DNA. Key Concepts

• Replication of DNA is undertaken by a complex of enzymes that separate the parental strands and synthesize the daughter strands.

• The replication fork is the point at which the parental strands are separated. • The enzymes that synthesize DNA are called DNA polymerases; the enzymes that synthesize RNA are RNA polymerases.

• Nucleases are enzymes that degrade nucleic acids; they include DNAases and RNAases, and can be divided into endonucleases and exonucleases.

Replication requires the two strands of the parental duplex to separate. However, the disruption of structure is only transient and is reversed as the daughter duplex is formed. Only a small stretch of the duplex DNA is separated into single strands at any moment. The helical structure of a molecule of DNA engaged in replication is illustrated in Figure 1.10. The nonreplicated region consists of the parental duplex, opening into the replicated region where the two daughter duplexes have formed. The double helical structure is disrupted at the junction between the two regions, which is called the replication fork. Replication involves movement of the replication fork along the parental DNA, so there is a continuous unwinding of the parental strands and DNA strands separate at the replication fork | SECTION 1.1.8 © 2004. Virtual Text / www.ergito.com

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rewinding into daughter duplexes.

Figure 1.10 The replication fork is the region of DNA in which there is a transition from the unwound parental duplex to the newly replicated daughter duplexes.

The synthesis of nucleic acids is catalyzed by specific enzymes, which recognize the template and undertake the task of catalyzing the addition of subunits to the polynucleotide chain that is being synthesized. The enzymes are named according to the type of chain that is synthesized: DNA polymerases synthesize DNA, and RNA polymerases synthesize RNA. Degradation of nucleic acids also requires specific enzymes: deoxyribonucleases (DNAases) degrade DNA, and ribonucleases (RNAases) degrade RNA. The nucleases fall into the general classes of exonucleases and endonucleases: • Endonucleases cut individual bonds within RNA or DNA molecules, generating discrete fragments. Some DNAases cleave both strands of a duplex DNA at the target site, while others cleave only one of the two strands. Endonucleases are involved in cutting reactions, as shown in Figure 1.11. • Exonucleases remove residues one at a time from the end of the molecule, generating mononucleotides. They always function on a single nucleic acid strand, and each exonuclease proceeds in a specific direction, that is, starting at either a 5 ′ or at a 3 ′ end and proceeding toward the other end. They are involved in trimming reactions, as shown in Figure 1.12.

Figure 1.11 An endonuclease cleaves a bond within a nucleic acid. This example shows an enzyme that attacks one strand of a DNA duplex.

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Figure 1.12 An exonuclease removes bases one at a time by cleaving the last bond in a polynucleotide chain.

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GENES ARE DNA

1.1.9 Nucleic acids hybridize by base pairing Key Terms Denaturation of protein describes its conversion from the physiological conformation to some other (inactive) conformation. Renaturation describes the reassociation of denatured complementary single strands of a DNA double helix. Annealing of DNA describes the renaturation of a duplex structure from single strands that were obtained by denaturing duplex DNA. Hybridization describes the pairing of complementary RNA and DNA strands to give an RNA-DNA hybrid. Key Concepts

• Heating causes the two strands of a DNA duplex to separate. • The Tm is the midpoint of the temperature range for denaturation. • Complementary single strands can renature when the temperature is reduced. • Denaturation and renaturation/hybridization can occur with DNA-DNA, DNA-RNA, or RNA-RNA combinations, and can be intermolecular or intramolecular.

• The ability of two single-stranded nucleic acid preparations to hybridize is a measure of their complementarity.

A crucial property of the double helix is the ability to separate the two strands without disrupting covalent bonds. This makes it possible for the strands to separate and reform under physiological conditions at the (very rapid) rates needed to sustain genetic functions. The specificity of the process is determined by complementary base pairing. The concept of base pairing is central to all processes involving nucleic acids. Disruption of the base pairs is a crucial aspect of the function of a double-stranded molecule, while the ability to form base pairs is essential for the activity of a single-stranded nucleic acid.Figure 1.16 shows that base pairing enables complementary single-stranded nucleic acids to form a duplex structure.

Nucleic acids hybridize by base pairing | SECTION 1.1.9 © 2004. Virtual Text / www.ergito.com

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Figure 1.16 Base pairing occurs in duplex DNA and also in intra- and inter-molecular interactions in single-stranded RNA (or DNA).

• An intramolecular duplex region can form by base pairing between two complementary sequences that are part of a single-stranded molecule. • A single-stranded molecule may base pair with an independent, complementary single-stranded molecule to form an intermolecular duplex. Formation of duplex regions from single-stranded nucleic acids is most important for RNA, but single-stranded DNA also exists (in the form of viral genomes). Base pairing between independent complementary single strands is not restricted to DNA-DNA or RNA-RNA, but can also occur between a DNA molecule and an RNA molecule. The lack of covalent links between complementary strands makes it possible to manipulate DNA in vitro. The noncovalent forces that stabilize the double helix are disrupted by heating or by exposure to low salt concentration. The two strands of a double helix separate entirely when all the hydrogen bonds between them are broken. The process of strand separation is called denaturation or (more colloquially) melting. ("Denaturation" is also used to describe loss of authentic protein structure; it is a general term implying that the natural conformation of a macromolecule has been converted to some other form.) Denaturation of DNA occurs over a narrow temperature range and results in striking changes in many of its physical properties. The midpoint of the temperature range over which the strands of DNA separate is called the melting temperature (Tm). It depends on the proportion of G·C base pairs. Because each G·C base pair has three hydrogen bonds, it is more stable than an A·T base pair, which has only two hydrogen bonds. The more G·C base pairs are contained in a DNA, the greater the energy that is needed to separate the two strands. In solution under physiological conditions, a DNA that is 40% G·C – a value typical of mammalian genomes – denatures with a Tm of about 87°C. So duplex DNA is stable at the temperature prevailing in the cell.


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The denaturation of DNA is reversible under appropriate conditions. The ability of the two separated complementary strands to reform into a double helix is called renaturation. Renaturation depends on specific base pairing between the complementary strands. Figure 1.17 shows that the reaction takes place in two stages. First, single strands of DNA in the solution encounter one another by chance; if their sequences are complementary, the two strands base pair to generate a short double-helical region. Then the region of base pairing extends along the molecule by a zipper-like effect to form a lengthy duplex molecule. Renaturation of the double helix restores the original properties that were lost when the DNA was denatured.

Figure 1.17 Denatured single strands of DNA can renature to give the duplex form.

Renaturation describes the reaction between two complementary sequences that were separated by denaturation. However, the technique can be extended to allow any two complementary nucleic acid sequences to react with each other to form a duplex structure. This is sometimes called annealing, but the reaction is more generally described as hybridization whenever nucleic acids of different sources are involved, as in the case when one preparation consists of DNA and the other consists of RNA. The ability of two nucleic acid preparations to hybridize constitutes a precise test for their complementarity since only complementary sequences can form a duplex structure. The principle of the hybridization reaction is to expose two single-stranded nucleic acid preparations to each other and then to measure the amount of double-stranded material that forms. Figure 1.18 illustrates a procedure in which a DNA preparation is denatured and the single strands are adsorbed to a filter. Then a second denatured DNA (or RNA) preparation is added. The filter is treated so that the second preparation can adsorb to it only if it is able to base pair with the DNA that was originally adsorbed. Usually the second preparation is radioactively labeled, so that the reaction can be measured as the amount of radioactive label retained by the filter.


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Figure 1.18 Filter hybridization establishes whether a solution of denatured DNA (or RNA) contains sequences complementary to the strands immobilized on the filter.

The extent of hybridization between two single-stranded nucleic acids is determined by their complementarity. Two sequences need not be perfectly complementary to hybridize. If they are closely related but not identical, an imperfect duplex is formed in which base pairing is interrupted at positions where the two single strands do not correspond. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.9


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GENES ARE DNA

1.1.10 Mutations change the sequence of DNA Key Terms Spontaneous mutations occur in the absence of any added reagent to increase the mutation rate, as the result of errors in replication (or other events involved in the reproduction of DNA) or by environmental damage. The background level of mutation describes the rate at which sequence changes accumulate in the genome of an organism. It reflects the balance between the occurrence of spontaneous mutations and their removal by repair systems, and is characteristic for any species. Mutagens increase the rate of mutation by inducing changes in DNA sequence, directly or indirectly. Induced mutations result from the action of a mutagen. The mutagen may act directly on the bases in DNA or it may act indirectly to trigger a pathway that leads to a change in DNA sequence. Key Concepts

• All mutations consist of changes in the sequence of DNA. • Mutations may occur spontaneously or may be induced by mutagens.

Mutations provide decisive evidence that DNA is the genetic material. When a change in the sequence of DNA causes an alteration in the sequence of a protein, we may conclude that the DNA codes for that protein. Furthermore, a change in the phenotype of the organism may allow us to identify the function of the protein. The existence of many mutations in a gene may allow many variant forms of a protein to be compared, and a detailed analysis can be used to identify regions of the protein responsible for individual enzymatic or other functions. All organisms suffer a certain number of mutations as the result of normal cellular operations or random interactions with the environment. These are called spontaneous mutations; the rate at which they occur is characteristic for any particular organism and is sometimes called the background level. Mutations are rare events, and of course those that damage a gene are selected against during evolution. It is therefore difficult to obtain large numbers of spontaneous mutants to study from natural populations. The occurrence of mutations can be increased by treatment with certain compounds. These are called mutagens, and the changes they cause are referred to as induced mutations. Most mutagens act directly by virtue of an ability either to modify a particular base of DNA or to become incorporated into the nucleic acid. The effectiveness of a mutagen is judged by how much it increases the rate of mutation above background. By using mutagens, it becomes possible to induce many changes in any gene (for review see 3). Mutations change the sequence of DNA | SECTION 1.1.10 © 2004. Virtual Text / www.ergito.com

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Spontaneous mutations that inactivate gene function occur in bacteriophages and bacteria at a relatively constant rate of 3-4 × 10–3 per genome per generation (2221). Given the large variation in genome sizes between bacteriophages and bacteria, this corresponds to wide differences in the mutation rate per base pair. This suggests that the overall rate of mutation has been subject to selective forces that have balanced the deleterious effects of most mutations against the advantageous effects of some mutations. This conclusion is strengthened by the observation that an archaeal microbe that lives under harsh conditions of high temperature and acidity (which are expected to damage DNA) does not show an elevated mutation rate, but in fact has an overall mutation rate just below the average range (2203). Figure 1.19 shows that in bacteria, the mutation rate corresponds to ~10–6 events per locus per generation or to an average rate of change per base pair of 10–9-10–10 per generation. The rate at individual base pairs varies very widely, over a 10,000 fold range. We have no accurate measurement of the rate of mutation in eukaryotes, although usually it is thought to be somewhat similar to that of bacteria on a per-locus per-generation basis (2487).

Figure 1.19 A base pair is mutated at a rate of 10-9 - 10-10 per generation, a gene of 1000 bp is mutated at ~10-6 per generation, and a bacterial genome is mutated at 3 × 10-3 per generation.

Last updated on 8-15-2002

Mutations change the sequence of DNA | SECTION 1.1.10 © 2004. Virtual Text / www.ergito.com

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Reviews 3.

Drake, J. W. and Balz, R. H. (1976). The biochemistry of mutagenesis. Annu. Rev. Biochem. 45, 11-37.

2487. Drake, J. W., Charlesworth, B., Charlesworth, D., and Crow, J. F. (1998). Rates of spontaneous mutation. Genetics 148, 1667-1686.

References 2203. Grogan, D. W., Carver, G. T., and Drake, J. W. (2001). Genetic fidelity under harsh conditions: analysis of spontaneous mutation in the thermoacidophilic archaeon Sulfolobus acidocaldarius. Proc. Natl. Acad. Sci. USA 98, 7928-7933. 2221. Drake, J. W. (1991). A constant rate of spontaneous mutation in DNA-based microbes. Proc. Natl. Acad. Sci. USA 88, 7160-7164.


Mutations change the sequence of DNA | SECTION 1.1.10 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.11 Mutations may affect single base pairs or longer sequences Key Terms A point mutation is a change in the sequence of DNA involving a single base pair. A transition is a mutation in which one pyrimidine is replaced by the other and/or in which one purine is replaced by the other. A transversion is a mutation in which a purine is replaced by a pyrimidine or vice versa. Base mispairing is a coupling between two bases that does not conform to the Watson-Crick rule, e.g., adenine with cytosine, thymine with guanine. An insertion is the addition of a stretch of base pairs in DNA. Duplications are a special class of insertions. A transposon (transposable element) is a DNA sequence able to insert itself (or a copy of itself) at a new location in the genome, without having any sequence relationship with the target locus. A deletion is the removal of a sequence of DNA, the regions on either side being joined together except in the case of a terminal deletion at the end of a chromosome. Key Concepts

• A point mutation changes a single base pair. • Point mutations can be caused by the chemical conversion of one base into another or by mistakes that occur during replication.

• A transition replaces a G·C base pair with an A·T base pair or vice-versa. • A transversion replaces a purine with a pyrimidine, such as changing A·T to T·A. • Insertions are the most common type of mutation, and result from the movement of transposable elements.

Any base pair of DNA can be mutated. A point mutation changes only a single base pair, and can be caused by either of two types of event (for review see 3238): • Chemical modification of DNA directly changes one base into a different base. • A malfunction during the replication of DNA causes the wrong base to be inserted into a polynucleotide chain during DNA synthesis. Point mutations can be divided into two types, depending on the nature of the change when one base is substituted for another: Mutations may affect single base pairs or longer sequences | SECTION 1.1.11 © 2004. Virtual Text / www.ergito.com

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• The most common class is the transition, comprising the substitution of one pyrimidine by the other, or of one purine by the other. This replaces a G·C pair with an A·T pair or vice versa. • The less common class is the transversion, in which a purine is replaced by a pyrimidine or vice versa, so that an A·T pair becomes a T·A or C·G pair. The effects of nitrous acid provide a classic example of a transition caused by the chemical conversion of one base into another. Figure 1.20 shows that nitrous acid performs an oxidative deamination that converts cytosine into uracil. In the replication cycle following the transition, the U pairs with an A, instead of with the G with which the original C would have paired. So the C·G pair is replaced by a T·A pair when the A pairs with the T in the next replication cycle. (Nitrous acid also deaminates adenine, causing the reverse transition from A·T to G·C.)

Figure 1.20 Mutations can be induced by chemical modification of a base.

Transitions are also caused by base mispairing, when unusual partners pair in defiance of the usual restriction to Watson-Crick pairs. Base mispairing usually occurs as an aberration resulting from the incorporation into DNA of an abnormal base that has ambiguous pairing properties. Figure 1.21 shows the example of bromouracil (BrdU), an analog of thymine that contains a bromine atom in place of the methyl group of thymine. BrdU is incorporated into DNA in place of thymine. But it has ambiguous pairing properties, because the presence of the bromine atom allows a shift to occur in which the base changes structure from a keto (=O) form to an enol (–OH) form. The enol form can base pair with guanine, which leads to substitution of the original A·T pair by a G·C pair.

Mutations may affect single base pairs or longer sequences | SECTION 1.1.11 © 2004. Virtual Text / www.ergito.com

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Figure 1.21 Mutations can be induced by the incorporation of base analogs into DNA.

The mistaken pairing can occur either during the original incorporation of the base or in a subsequent replication cycle. The transition is induced with a certain probability in each replication cycle, so the incorporation of BrdU has continuing effects on the sequence of DNA. Point mutations were thought for a long time to be the principal means of change in individual genes. However, we now know that insertions of stretches of additional material are quite frequent. The source of the inserted material lies with transposable elements, sequences of DNA with the ability to move from one site to another (see Molecular Biology 4.16 Transposons and Molecular Biology 4.17 Retroviruses and retroposons.) An insertion usually abolishes the activity of a gene. Where such insertions have occurred, deletions of part or all of the inserted material, and sometimes of the adjacent regions, may subsequently occur. A significant difference between point mutations and the insertions/deletions is that the frequency of point mutation can be increased by mutagens, whereas the occurrence of changes caused by transposable elements is not affected. However, insertions and deletions can also occur by other mechanisms – for example, involving mistakes made during replication or recombination – although probably Mutations may affect single base pairs or longer sequences | SECTION 1.1.11 © 2004. Virtual Text / www.ergito.com

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these are less common. And a class of mutagens called the acridines introduce (very small) insertions and deletions.


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Reviews 3238. Maki, H. (2002). Origins of Spontaneous Mutations: Specificity and Directionality of Base-Substitution, Frameshift, and Sequence-Substitution Mutageneses. Annu. Rev. Genet. 36, 279-303.



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GENES ARE DNA

1.1.12 The effects of mutations can be reversed Key Terms Revertants are derived by reversion of a mutant cell or organism to the wild-type phenotype. Forward mutations inactivate a wild-type gene. A back mutation reverses the effect of a mutation that had inactivated a gene; thus it restores wild type. A true reversion is a mutation that restores the original sequence of the DNA. Second-site reversion occurs when a second mutation suppresses the effect of a first mutation. Suppression occurs when a second event eliminates the effects of a mutation without reversing the original change in DNA. A suppressor is a second mutation that compensates for or alters the effects of a primary mutation. Key Concepts

• Forward mutations inactivate a gene, and back mutations (or revertants) reverse their effects.

• Insertions can revert by deletion of the inserted material, but deletions cannot revert.

• Suppression occurs when a mutation in a second gene bypasses the effect of mutation in the first gene.

Figure 1.22 shows that the isolation of revertants is an important characteristic that distinguishes point mutations and insertions from deletions:

The effects of mutations can be reversed | SECTION 1.1.12 © 2004. Virtual Text / www.ergito.com

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Figure 1.22 Point mutations and insertions can revert, but deletions cannot revert.

• A point mutation can revert by restoring the original sequence or by gaining a compensatory mutation elsewhere in the gene. • An insertion of additional material can revert by deletion of the inserted material. • A deletion of part of a gene cannot revert. Mutations that inactivate a gene are called forward mutations. Their effects are reversed by back mutations, which are of two types. An exact reversal of the original mutation is called true reversion. So if an A·T pair has been replaced by a G·C pair, another mutation to restore the A·T pair will exactly regenerate the wild-type sequence. Alternatively, another mutation may occur elsewhere in the gene, and its effects The effects of mutations can be reversed | SECTION 1.1.12 © 2004. Virtual Text / www.ergito.com

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compensate for the first mutation. This is called second-site reversion. For example, one amino acid change in a protein may abolish gene function, but a second alteration may compensate for the first and restore protein activity. A forward mutation results from any change that inactivates a gene, whereas a back mutation must restore function to a protein damaged by a particular forward mutation. So the demands for back mutation are much more specific than those for forward mutation. The rate of back mutation is correspondingly lower than that of forward mutation, typically by a factor of ~10. Mutations can also occur in other genes to circumvent the effects of mutation in the original gene. This effect is called suppression. A locus in which a mutation suppresses the effect of a mutation in another locus is called a suppressor. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.12

The effects of mutations can be reversed | SECTION 1.1.12 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.13 Mutations are concentrated at hotspots Key Terms A hotspot is a site in the genome at which the frequency of mutation (or recombination) is very much increased, usually by at least an order of magnitude relative to neighboring sites. Key Concepts

• The frequency of mutation at any particular base pair is determined by statistical fluctuation, except for hotspots, where the frequency is increased by at least an order of magnitude.

So far we have dealt with mutations in terms of individual changes in the sequence of DNA that influence the activity of the genetic unit in which they occur. When we consider mutations in terms of the inactivation of the gene, most genes within a species show more or less similar rates of mutation relative to their size. This suggests that the gene can be regarded as a target for mutation, and that damage to any part of it can abolish its function. As a result, susceptibility to mutation is roughly proportional to the size of the gene. But consider the sites of mutation within the sequence of DNA; are all base pairs in a gene equally susceptible or are some more likely to be mutated than others? What happens when we isolate a large number of independent mutations in the same gene? Many mutants are obtained. Each is the result of an individual mutational event. Then the site of each mutation is determined. Most mutations will lie at different sites, but some will lie at the same position. Two independently isolated mutations at the same site may constitute exactly the same change in DNA (in which case the same mutational event has happened on more than one occasion), or they may constitute different changes (three different point mutations are possible at each base pair). The histogram of Figure 1.23 shows the frequency with which mutations are found at each base pair in the lacI gene of E. coli. The statistical probability that more than one mutation occurs at a particular site is given by random-hit kinetics (as seen in the Poisson distribution). So some sites will gain one, two, or three mutations, while others will not gain any. But some sites gain far more than the number of mutations expected from a random distribution; they may have 10× or even 100× more mutations than predicted by random hits. These sites are called hotspots. Spontaneous mutations may occur at hotspots; and different mutagens may have different hotspots.

Mutations are concentrated at hotspots | SECTION 1.1.13 © 2004. Virtual Text / www.ergito.com

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Figure 1.23 Spontaneous mutations occur throughout the lacI gene of E. coli, but are concentrated at a hotspot. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.13

Mutations are concentrated at hotspots | SECTION 1.1.13 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.14 Many hotspots result from modified bases Key Terms Modified bases are all those except the usual four from which DNA (T, C, A, G) or RNA (U, C, A, G) are synthesized; they result from postsynthetic changes in the nucleic acid. A mismatch describes a site in DNA where the pair of bases does not conform to the usual G-C or A-T pairs. It may be caused by incorporation of the wrong base during replication or by mutation of a base. Key Concepts

• A common cause of hotspots is the modified base 5-methylcytosine, which is spontaneously deaminated to thymine.

A major cause of spontaneous mutation results from the presence of an unusual base in the DNA. In addition to the four bases that are inserted into DNA when it is synthesized, modified bases are sometimes found. The name reflects their origin; they are produced by chemically modifying one of the four bases already present in DNA. The most common modified base is 5-methylcytosine, generated by a methylase enzyme that adds a methyl group to certain cytosine residues at specific sites in the DNA. Sites containing 5-methylcytosine provide hotspots for spontaneous point mutation in E. coli. In each case, the mutation takes the form of a G·C to A·T transition. The hotspots are not found in strains of E. coli that cannot methylate cytosine. The reason for the existence of the hotspots is that cytosine bases suffer spontaneous deamination at an appreciable frequency. In this reaction, the amino group is replaced by a keto group. Recall that deamination of cytosine generates uracil (see Figure 1.20). Figure 1.24 compares this reaction with the deamination of 5-methylcytosine where deamination generates thymine. The effect in DNA is to generate the base pairs G·U and G·T, respectively, where there is a mismatch between the partners.

Many hotspots result from modified bases | SECTION 1.1.14 © 2004. Virtual Text / www.ergito.com

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Figure 1.24 Deamination of cytosine produces uracil, whereas deamination of 5-methyl-cytosine produces thymine.

All organisms have repair systems that correct mismatched base pairs by removing and replacing one of the bases. The operation of these systems determines whether mismatched pairs such as G·U and G·T result in mutations. Figure 1.25 shows that the consequences of deamination are different for 5-methylcytosine and cytosine. Deaminating the (rare) 5-methylcytosine causes a mutation, whereas deamination of the more common cytosine does not have this effect (382). This happens because the repair systems are much more effective in recognizing G·U than G·T.


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Figure 1.25 The deamination of 5-methylcytosine produces thymine (by C·G to T·A transitions), while the deamination of cytosine produces uracil (which usually is removed and then replaced by cytosine).

E. coli contains an enzyme, uracil-DNA-glycosidase, that removes uracil residues from DNA (see Molecular Biology 4.15.22 Base flipping is used by methylases and glycosylases). This action leaves an unpaired G residue, and a "repair system" then inserts a C base to partner it. The net result of these reactions is to restore the original sequence of the DNA. This system protects DNA against the consequences of spontaneous deamination of cytosine (although it is not active enough to prevent the effects of the increased level of deamination caused by nitrous acid; see Figure 1.20). But the deamination of 5-methylcytosine leaves thymine. This creates a mismatched base pair, G·T. If the mismatch is not corrected before the next replication cycle, a mutation results. At the next replication, the bases in the mispaired G·T partnership separate, and then they pair with new partners to produce one wild-type G·C pair and one mutant A·T pair. Deamination of 5-methylcytosine is the most common cause of production of G·T mismatched pairs in DNA. Repair systems that act on G·T mismatches have a bias toward replacing the T with a C (rather than the alternative of replacing the G with an A), which helps to reduce the rate of mutation (see Molecular Biology 4.15.24 Controlling the direction of mismatch repair ). However, these Many hotspots result from modified bases | SECTION 1.1.14 © 2004. Virtual Text / www.ergito.com

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systems are not as effective as the removal of U from G·U mismatches. As a result, deamination of 5-methylcytosine leads to mutation much more often than does deamination of cytosine. 5-methylcytosine also creates hotspots in eukaryotic DNA. It is common at CpG dinucleotides that are concentrated in regions called CpG islands (see Molecular Biology 5.21.19 CpG islands are regulatory targets). Although 5-methylcytosine accounts for ~1% of the bases in human DNA, sites containing the modified base account for ~30% of all point mutations. This makes the state of 5-methylcytosine a particularly important determinant of mutation in animal cells. The importance of repair systems in reducing the rate of mutation is emphasized by the effects of eliminating the mouse enzyme MBD4, a glycosylase that can remove T (or U) from mismatches with G. The result is to increase the mutation rate at CpG sites by a factor of 3× (2845). (The reason the effect is not greater is that MBD4 is only one of several systems that act on G·T mismatches; we can imagine that elimination of all the systems would increase the mutation rate much more.) The operation of these systems casts an interesting light on the use of T in DNA compared with U in RNA. Perhaps it relates to the need of DNA for stability of sequence; the use of T means that any deaminations of C are immediately recognized, because they generate a base (U) not usually present in the DNA. This greatly increases the efficiency with which repair systems can function (compared with the situation when they have to recognize G·T mismatches, which can be produced also by situations where removing the T would not be the appropriate response). Also, the phosphodiester bond of the backbone is more labile when the base is U. Last updated on 8-15-2002


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References 382. Coulondre, C. et al. (1978). Molecular basis of base substitution hotspots in E. coli. Nature 274, 775-780. 2845. Millar, C. B., Guy, J., Sansom, O. J., Selfridge, J., MacDougall, E., Hendrich, B., Keightley, P. D., Bishop, S. M., Clarke, A. R., and Bird, A. (2002). Enhanced CpG mutability and tumorigenesis in MBD4-deficient mice. Science 297, 403-405.



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GENES ARE DNA

1.1.15 A gene codes for a single polypeptide Key Terms A homomultimer is a protein composed of identical subunits. A heteromultimer is a protein that is composed of nonidentical subunits (coded by different genes). Key Concepts

• The one gene : one enzyme hypothesis summarizes the basis of modern genetics: that a gene is a stretch of DNA coding for a single polypeptide chain.

• Most mutations damage gene function.

The first systematic attempt to associate genes with enzymes showed that each stage in a metabolic pathway is catalyzed by a single enzyme and can be blocked by mutation in a different gene. This led to the one gene : one enzyme hypothesis. Each metabolic step is catalyzed by a particular enzyme, whose production is the responsibility of a single gene. A mutation in the gene alters the activity of the protein for which it is responsible. A modification in the hypothesis is needed to accommodate proteins that consist of more than one subunit. If the subunits are all the same, the protein is a homomultimer, represented by a single gene. If the subunits are different, the protein is a heteromultimer. Stated as a more general rule applicable to any heteromultimeric protein, the one gene : one enzyme hypothesis becomes more precisely expressed as one gene : one polypeptide chain. Identifying which protein represents a particular gene can be a protracted task. The mutation responsible for creating Mendel's wrinkled-pea mutant was identified only in 1990 as an alteration that inactivates the gene for a starch branching enzyme! It is important to remember that a gene does not directly generate a protein. As shown previously in Figure 1.2, a gene codes for an RNA, which may in turn code for a protein. Most genes code for proteins, but some genes code for RNAs that do not give rise to proteins. These RNAs may be structural components of the apparatus responsible for synthesizing proteins or may have roles in regulating gene expression. The basic principle is that the gene is a sequence of DNA that specifies the sequence of an independent product. The process of gene expression may terminate in a product that is either RNA or protein. A mutation is a random event with regard to the structure of the gene, so the greatest probability is that it will damage or even abolish gene function. Most mutations that affect gene function are recessive: they represent an absence of function, because the mutant gene has been prevented from producing its usual protein.Figure 1.26 illustrates the relationship between recessive and wild-type alleles. When a A gene codes for a single polypeptide | SECTION 1.1.15 © 2004. Virtual Text / www.ergito.com

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heterozygote contains one wild-type allele and one mutant allele, the wild-type allele is able to direct production of the enzyme. The wild-type allele is therefore dominant. (This assumes that an adequate amount of protein is made by the single wild-type allele. When this is not true, the smaller amount made by one allele as compared to two alleles results in the intermediate phenotype of a partially dominant allele in a heterozygote.)

Figure 1.26 Genes code for proteins; dominance is explained by the properties of mutant proteins. A recessive allele does not contribute to the phenotype because it produces no protein (or protein that is nonfunctional).

Last updated on 7-18-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.15

A gene codes for a single polypeptide | SECTION 1.1.15 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.16 Mutations in the same gene cannot complement Key Terms A complementation test determines whether two mutations are alleles of the same gene. It is accomplished by crossing two different recessive mutations that have the same phenotype and determining whether the wild-type phenotype can be produced. If so, the mutations are said to complement each other and are probably not mutations in the same gene. Two mutants are said to complement each other when a diploid that is heterozygous for each mutation produces the wild type phenotype. A complementation group is a series of mutations unable to complement when tested in pairwise combinations in trans; defines a genetic unit (the cistron). A cistron is the genetic unit defined by the complementation test; it is equivalent to the gene. A gene (cistron) is the segment of DNA specifying production of a polypeptide chain; it includes regions preceding and following the coding region (leader and trailer) as well as intervening sequences (introns) between individual coding segments (exons). Key Concepts

• A mutation in a gene affects only the protein coded by the mutant copy of the gene, and does not affect the protein coded by any other allele.

• Failure of two mutations to complement (produce wild-phenotype) when they are present in trans configuration in a heterozygote means that they are part of the same gene.

How do we determine whether two mutations that cause a similar phenotype lie in the same gene? If they map close together, they may be alleles. However, they could also represent mutations in two different genes whose proteins are involved in the same function. The complementation test is used to determine whether two mutations lie in the same gene or in different genes. The test consists of making a heterozygote for the two mutations (by mating parents homozygous for each mutation). If the mutations lie in the same gene, the parental genotypes can be represented as:

The first parent provides an m1 mutant allele and the second parent provides an m2 Mutations in the same gene cannot complement | SECTION 1.1.16 © 2004. Virtual Text / www.ergito.com

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allele, so that the heterozygote has the constitution:

No wild-type gene is present, so the heterozygote has mutant phenotype. If the mutations lie in different genes, the parental genotypes can be represented as:

Each chromosome has a wild-type copy of one gene (represented by the plus sign) and a mutant copy of the other. Then the heterozygote has the constitution:

in which the two parents between them have provided a wild-type copy of each gene. The heterozygote has wild phenotype; the two genes are said to complement. The complementation test is shown in more detail in Figure 1.27. The basic test consists of the comparison shown in the top part of the figure. If two mutations lie in the same gene, we see a difference in the phenotypes of the trans configuration and the cis configuration. The trans configuration is mutant, because each allele has a (different) mutation. But the cis configuration is wild-type, because one allele has two mutations but the other allele has no mutations. The lower part of the figure shows that if the two mutations lie in different genes, we always see a wild phenotype. There is always one wild-type and one mutant allele of each gene, and the configuration is irrelevant. The basic test and some exceptions to it are discussed in Molecular Biology Supplement 32.9 Complementation.

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Figure 1.27 The cistron is defined by the complementation test. Genes are represented by bars; red stars identify sites of mutation.

Failure to complement means that two mutations are part of the same genetic unit. Mutations that do not complement one another are said to comprise part of the same complementation group. Another term that is used to describe the unit defined by the complementation test is the cistron. This is the same as the gene. Basically these three terms all describe a stretch of DNA that functions as a unit to give rise to an RNA or protein product. The properties of the gene with regards to complementation are explained by the fact that this product is a single molecule that behaves as a functional unit. Last updated on 7-18-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.16

Mutations in the same gene cannot complement | SECTION 1.1.16 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.17 Mutations may cause loss-of-function or gain-of-function Key Terms A null mutation completely eliminates the function of a gene. Leaky mutations leave some residual function, for instance when the mutant protein is partially active (in the case of a missense mutation), or when read-through produces a small amount of wild-type protein (in the case of a nonsense mutation). A loss-of-function mutation eliminates or reduces the activity of a gene. It is often, but not always, recessive. A gain-of-function mutation usually refers to a mutation that causes an increase in the normal gene activity. It sometimes represents acquisition of certain abnormal properties. It is often, but not always, dominant. Silent mutations do not change the sequence of a protein because they produce synonymous codons. Neutral substitutions in a protein cause changes in amino acids that do not affect activity. Key Concepts

• Recessive mutations are due to loss-of-function by the protein product. • Dominant mutations result from a gain-of-function. • Testing whether a gene is essential requires a null mutation (one that completely eliminates its function).

• Silent mutations have no effect, either because the base change does not change the sequence or amount of protein, or because the change in protein sequence has no effect.

• Leaky mutations do affect the function of the gene product, but are not revealed in the phenotype because sufficient activity remains.

The various possible effects of mutation in a gene are summarized in Figure 1.28.

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Figure 1.28 Mutations that do not affect protein sequence or function are silent. Mutations that abolish all protein activity are null. Point mutations that cause loss-of-function are recessive; those that cause gain-of-function are dominant.

When a gene has been identified, insight into its function in principle can be gained by generating a mutant organism that entirely lacks the gene. A mutation that completely eliminates gene function, usually because the gene has been deleted, is called a null mutation. If a gene is essential, a null mutation is lethal. To determine what effect a gene has upon the phenotype, it is essential to characterize a null mutant. When a mutation fails to affect the phenotype, it is always possible that this is because it is a leaky mutation – enough active product is made to fulfill its function, even though the activity is quantitatively reduced or qualitatively different from the wild type. But if a null mutant fails to affect a phenotype, we may safely conclude that the gene function is not necessary. Null mutations, or other mutations that impede gene function (but do not necessarily abolish it entirely) are called loss-of-function mutations. A loss-of-function mutation is recessive (as in the example of Figure 1.26). Sometimes a mutation has the opposite effect and causes a protein to acquire a new function; such a change is called a gain-of-function mutation. A gain-of-function mutation is dominant. Not all mutations in DNA lead to a detectable change in the phenotype. Mutations without apparent effect are called silent mutations. They fall into two types. Some involve base changes in DNA that do not cause any change in the amino acid present in the corresponding protein. Others change the amino acid, but the replacement in Mutations may cause loss-of-function or gain-of-function | SECTION 1.1.17 © 2004. Virtual Text / www.ergito.com

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the protein does not affect its activity; these are called neutral substitutions. Last updated on 10-2-2003 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.17

Mutations may cause loss-of-function or gain-of-function | SECTION 1.1.17 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.18 A locus may have many different mutant alleles Key Terms A locus is said to have multiple alleles when more than two allelic forms have been found. Each allele may cause a different phenotype. Key Concepts

• The existence of multiple alleles allows heterozygotes to occur representing any pairwise combination of alleles.

If a recessive mutation is produced by every change in a gene that prevents the production of an active protein, there should be a large number of such mutations in any one gene. Many amino acid replacements may change the structure of the protein sufficiently to impede its function. Different variants of the same gene are called multiple alleles, and their existence makes it possible to create a heterozygote between mutant alleles. The relationship between these multiple alleles takes various forms. In the simplest case, a wild-type gene codes for a protein product that is functional. Mutant allele(s) code for proteins that are nonfunctional. But there are often cases in which a series of mutant alleles have different phenotypes. For example, wild-type function of the white locus of D. melanogaster is required for development of the normal red color of the eye. The locus is named for the effect of extreme (null) mutations, which cause the fly to have a white eye in mutant homozygotes. To describe wild-type and mutant alleles, wild genotype is indicated by a plus superscript after the name of the locus (w+ is the wild-type allele for [red] eye color in D. melanogaster). Sometimes + is used by itself to describe the wild-type allele, and only the mutant alleles are indicated by the name of the locus. An entirely defective form of the gene (or absence of phenotype) may be indicated by a minus superscript. To distinguish among a variety of mutant alleles with different effects, other superscripts may be introduced, such as wi or wa. The w+ allele is dominant over any other allele in heterozygotes. There are many different mutant alleles. Figure 1.29 shows a (small) sample. Although some alleles have no eye color, many alleles produce some color. Each of these mutant alleles must therefore represent a different mutation of the gene, which does not eliminate its function entirely, but leaves a residual activity that produces a characteristic phenotype. These alleles are named for the color of the eye in a homozygote. (Most A locus may have many different mutant alleles | SECTION 1.1.18 © 2004. Virtual Text / www.ergito.com

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w alleles affect the quantity of pigment in the eye, and the examples in the Figure are arranged in [roughly] declining amount of color, but others, such as wsp, affect the pattern in which it is deposited.)

Figure 1.29 The w locus has an extensive series of alleles, whose phenotypes extend from wild-type (red) color to complete lack of pigment.

When multiple alleles exist, an animal may be a heterozygote that carries two different mutant alleles. The phenotype of such a heterozygote depends on the nature of the residual activity of each allele. The relationship between two mutant alleles is in principle no different from that between wild-type and mutant alleles: one allele may be dominant, there may be partial dominance, or there may be codominance. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.18

A locus may have many different mutant alleles | SECTION 1.1.18 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.19 A locus may have more than one wild-type allele Key Terms Polymorphism (more fully genetic polymorphism) refers to the simultaneous occurrence in the population of genomes showing variations at a given position. The original definition applied to alleles producing different phenotypes. Now it is also used to describe changes in DNA affecting the restriction pattern or even the sequence. For practical purposes, to be considered as an example of a polymorphism, an allele should be found at a frequency > 1% in the population. Key Concepts

• A locus may have a polymorphic distribution of alleles, with no individual allele that can be considered to be the sole wild-type.

There is not necessarily a unique wild-type allele at any particular locus. Control of the human blood group system provides an example. Lack of function is represented by the null type, O group. But the functional alleles A and B provide activities that are codominant with one another and dominant over O group. The basis for this relationship is illustrated in Figure 1.30.

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Figure 1.30 The ABO blood group locus codes for a galactosyltransferase whose specificity determines the blood group.

The O (or H) antigen is generated in all individuals, and consists of a particular carbohydrate group that is added to proteins. The ABO locus codes for a galactosyltransferase enzyme that adds a further sugar group to the O antigen. The specificity of this enzyme determines the blood group. The A allele produces an enzyme that uses the cofactor UDP-N-acetylgalactose, creating the A antigen. The B allele produces an enzyme that uses the cofactor UDP-galactose, creating the B antigen. The A and B versions of the transferase protein differ in 4 amino acids that presumably affect its recognition of the type of cofactor. The O allele has a mutation (a small deletion) that eliminates activity, so no modification of the O antigen occurs. This explains why A and B alleles are dominant in the AO and BO heterozygotes: the corresponding transferase activity creates the A or B antigen. The A and B alleles are codominant in AB heterozygotes, because both transferase activities are expressed. The OO homozygote is a null that has neither activity, and therefore lacks both antigens. Neither A nor B can be regarded as uniquely wild type, since they represent alternative activities rather than loss or gain of function. A situation such as this, in which there are multiple functional alleles in a population, is described as a polymorphism (see Molecular Biology 1.3.3 Individual genomes show extensive A locus may have more than one wild-type allele | SECTION 1.1.19 © 2004. Virtual Text / www.ergito.com

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variation). This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.19

A locus may have more than one wild-type allele | SECTION 1.1.19 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.20 Recombination occurs by physical exchange of DNA Key Terms Crossing-over describes the reciprocal exchange of material between chromosomes that occurs during prophase I of meiosis and is responsible for genetic recombination. A bivalent is the structure containing all four chromatids (two representing each homologue) at the start of meiosis. Chromatids are the copies of a chromosome produced by replication. The name is usually used to describe the copies in the period before they separate at the subsequent cell division. A chiasma (pl. chiasmata) is a site at which two homologous chromosomes appear to have exchanged material during meiosis. Breakage and reunion describes the mode of genetic recombination, in which two DNA duplex molecules are broken at corresponding points and then rejoined crosswise (involving formation of a length of heteroduplex DNA around the site of joining). Heteroduplex DNA (Hybrid DNA) is generated by base pairing between complementary single strands derived from the different parental duplex molecules; it occurs during genetic recombination. Key Concepts

• Recombination is the result of crossing-over that occurs at chiasmata and involves two of the four chromatids.

• Recombination occurs by a breakage and reunion that proceeds via an intermediate of hybrid DNA.

Genetic recombination describes the generation of new combinations of alleles that occurs at each generation in diploid organisms. The two copies of each chromosome may have different alleles at some loci. By exchanging corresponding parts between the chromosomes, recombinant chromosomes can be generated that are different from the parental chromosomes. Recombination results from a physical exchange of chromosomal material. This is visible in the form of the crossing-over that occurs during meiosis (the specialized division that produces haploid germ cells). Meiosis starts with a cell that has duplicated its chromosomes, so that it has four copies of each chromosome. Early in meiosis, all four copies are closely associated (synapsed) in a structure called a bivalent. Each individual chromosomal unit is called a chromatid at this stage. Pairwise exchanges of material occur between the chromatids.

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The visible result of a crossing-over event is called a chiasma, and is illustrated diagrammatically in Figure 1.31. A chiasma represents a site at which two of the chromatids in a bivalent have been broken at corresponding points. The broken ends have been rejoined crosswise, generating new chromatids. Each new chromatid consists of material derived from one chromatid on one side of the junction point, with material from the other chromatid on the opposite side. The two recombinant chromatids have reciprocal structures. The event is described as a breakage and reunion. Its nature explains why a single recombination event can produce only 50% recombinants: each individual recombination event involves only two of the four associated chromatids.

Figure 1.31 Chiasma formation is responsible for generating recombinants.

The complementarity of the two strands of DNA is essential for the recombination process. Each of the chromatids shown in Figure 1.31 consists of a very long duplex of DNA. For them to be broken and reconnected without any loss of material requires a mechanism to recognize exactly corresponding positions. This is provided by complementary base pairing. Recombination involves a process in which the single strands in the region of the crossover exchange their partners. Figure 1.32 shows that this creates a stretch of hybrid DNA in which the single strand of one duplex is paired with its complement from the other duplex. The mechanism of course involves other stages (strands must be broken and resealed), and we discuss this in more detail in Molecular Biology 4.15 Recombination and repair, but the crucial feature that makes precise recombination possible is the complementarity of DNA strands. The figure shows only some stages of the reaction, but we see that a stretch of hybrid DNA forms in the recombination intermediate when a single strand crosses over from one duplex to the other. Each recombinant consists of one parental duplex DNA at the left, connected by a stretch of hybrid DNA to the other parental duplex at the right. Each duplex DNA corresponds to one of the chromatids involved in recombination in Figure 1.31. Recombination occurs by physical exchange of DNA | SECTION 1.1.20 © 2004. Virtual Text / www.ergito.com

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Figure 1.32 Recombination involves pairing between complementary strands of the two parental duplex DNAs.

The formation of hybrid DNA requires the sequences of the two recombining duplexes to be close enough to allow pairing between the complementary strands. If there are no differences between the two parental genomes in this region, formation of hybrid DNA will be perfect. But the reaction can be tolerated even when there are small differences. In this case, the hybrid DNA has points of mismatch, at which a base in one strand faces a base in the other strand that is not complementary to it. The correction of such mismatches is another feature of genetic recombination (see Molecular Biology 4.15 Recombination and repair). This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.20

Recombination occurs by physical exchange of DNA | SECTION 1.1.20 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.21 The genetic code is triplet Key Terms The genetic code is the correspondence between triplets in DNA (or RNA) and amino acids in protein. A codon is a triplet of nucleotides that represents an amino acid or a termination signal. Frameshift mutations arise by deletions or insertions that are not a multiple of 3 base pairs and change the frame in which triplets are translated into protein. The term is inappropriate outside of coding sequences. Acridines are mutagens that act on DNA to cause the insertion or deletion of a single base pair. They were useful in defining the triplet nature of the genetic code. A suppressor is a second mutation that compensates for or alters the effects of a primary mutation. A frameshift suppressor is an insertion or deletion of a base that restores the original reading frame in a gene that has had a base deletion or insertion. Key Concepts

• The genetic code is read in triplet nucleotides called codons. • The triplets are nonoverlapping and are read from a fixed starting point. • Mutations that insert or delete individual bases cause a shift in the triplet sets after the site of mutation.

• Combinations of mutations that together insert or delete 3 bases (or multiples of three) insert or delete amino acids but do not change the reading of the triplets beyond the last site of mutation.

Each gene represents a particular protein chain. The concept that each protein consists of a particular series of amino acids dates from Sanger's characterization of insulin in the 1950s. The discovery that a gene consists of DNA faces us with the issue of how a sequence of nucleotides in DNA represents a sequence of amino acids in protein. A crucial feature of the general structure of DNA is that it is independent of the particular sequence of its component nucleotides. The sequence of nucleotides in DNA is important not because of its structure per se, but because it codes for the sequence of amino acids that constitutes the corresponding polypeptide. The relationship between a sequence of DNA and the sequence of the corresponding protein is called the genetic code. The structure and/or enzymatic activity of each protein follows from its primary sequence of amino acids. By determining the sequence of amino acids in each The genetic code is triplet | SECTION 1.1.21 © 2004. Virtual Text / www.ergito.com

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protein, the gene is able to carry all the information needed to specify an active polypeptide chain. In this way, a single type of structure – the gene – is able to represent itself in innumerable polypeptide forms. Together the various protein products of a cell undertake the catalytic and structural activities that are responsible for establishing its phenotype. Of course, in addition to sequences that code for proteins, DNA also contains certain sequences whose function is to be recognized by regulator molecules, usually proteins. Here the function of the DNA is determined by its sequence directly, not via any intermediary code. Both types of region, genes expressed as proteins and sequences recognized as such, constitute genetic information. The genetic code is deciphered by a complex apparatus that interprets the nucleic acid sequence. This apparatus is essential if the information carried in DNA is to have meaning. In any given region, only one of the two strands of DNA codes for protein, so we write the genetic code as a sequence of bases (rather than base pairs). The genetic code is read in groups of three nucleotides, each group representing one amino acid. Each trinucleotide sequence is called a codon. A gene includes a series of codons that is read sequentially from a starting point at one end to a termination point at the other end. Written in the conventional 5 ′ → 3 ′ direction, the nucleotide sequence of the DNA strand that codes for protein corresponds to the amino acid sequence of the protein written in the direction from N-terminus to C-terminus. The genetic code is read in nonoverlapping triplets from a fixed starting point: • Nonoverlapping implies that each codon consists of three nucleotides and that successive codons are represented by successive trinucleotides. • The use of a fixed starting point means that assembly of a protein must start at one end and work to the other, so that different parts of the coding sequence cannot be read independently. The nature of the code predicts that two types of mutations will have different effects. If a particular sequence is read sequentially, such as: UUU AAA GGG CCC (codons) aa1 aa2 aa3 aa4 (amino acids) then a point mutation will affect only one amino acid. For example, the substitution of an A by some other base (X) causes aa2 to be replaced by aa5: UUU AAX GGG CCC aa1 aa5 aa3 aa4 because only the second codon has been changed.

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But a mutation that inserts or deletes a single base will change the triplet sets for the entire subsequent sequence. A change of this sort is called a frameshift. An insertion might take the form: UUU AAX AGG GCC C aa1 aa5 aa6 aa7 Because the new sequence of triplets is completely different from the old one, the entire amino acid sequence of the protein is altered beyond the site of mutation. So the function of the protein is likely to be lost completely. Frameshift mutations are induced by the acridines, compounds that bind to DNA and distort the structure of the double helix, causing additional bases to be incorporated or omitted during replication. Each mutagenic event sponsored by an acridine results in the addition or removal of a single base pair (for review see 4). If an acridine mutant is produced by, say, addition of a nucleotide, it should revert to wild type by deletion of the nucleotide. But reversion can also be caused by deletion of a different base, at a site close to the first. Combinations of such mutations provided revealing evidence about the nature of the genetic code. Figure 1.33 illustrates the properties of frameshift mutations. An insertion or a deletion changes the entire protein sequence following the site of mutation. But the combination of an insertion and a deletion causes the code to be read incorrectly only between the two sites of mutation; correct reading resumes after the second site.


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Figure 1.33 Frameshift mutations show that the genetic code is read in triplets from a fixed starting point.

Genetic analysis of acridine mutations in the rII region of the phage T6 in 1961 showed that all the mutations could be classified into one of two sets, described as (+) and (–). Either type of mutation by itself causes a frameshift, the (+) type by virtue of a base addition, the (–) type by virtue of a base deletion. Double mutant combinations of the types (+ +) and (––) continue to show mutant behavior. But combinations of the types (+ –) or (– +) suppress one another, giving rise to a description in which one mutation is described as a supressor of the other. (In the context of this work, "suppressor" is used in an unusual sense, because the second mutation is in the same gene as the first.) These results show that the genetic code must be read as a sequence that is fixed by the starting point, so additions or deletions compensate for each other, whereas double additions or double deletions remain mutant. But this does not reveal how many nucleotides make up each codon. When triple mutants are constructed, only (+ + +) and (––– ) combinations show the wild phenotype, while other combinations remain mutant. If we take three additions or three deletions to correspond respectively to the addition or omission overall of a single amino acid, this implies that the code is read in triplets. An incorrect amino acid sequence is found between the two outside sites of mutation, and the sequence on either side remains wild type, as indicated in Figure 1.33 (378; 379). Last updated on January 27, 2004


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Reviews 4.

Roth, J. R. (1974). Frameshift mutations. Annu. Rev. Genet. 8, 319-346.

References 378. Benzer, S. and Champe, S. P. (1961). Ambivalent rII mutants of phage T4. Proc. Natl. Acad. Sci. USA 47, 403-416. 379. Crick, F. H. C., Barnett, L., Brenner, S., and Watts-Tobin, R. J. (1961). General nature of the genetic code for proteins. Nature 192, 1227-1232.



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GENES ARE DNA

1.1.22 Every sequence has three possible reading frames Key Terms A reading frame is one of the three possible ways of reading a nucleotide sequence. Each reading frame divides the sequence into a series of successive triplets. There are three possible reading frames in any sequence, depending on the starting point. If the first frame starts at position 1, the second frame starts at position 2, and the third frame starts at position 3. An open reading frame (ORF) is a sequence of DNA consisting of triplets that can be translated into amino acids starting with an initiation codon and ending with a termination codon. The initiation codon is a special codon (usually AUG) used to start synthesis of a protein. A stop codon (Termination codon) is one of three triplets (UAG, UAA, UGA) that causes protein synthesis to terminate. They are also known historically as nonsense codons. The UAA codon is called ochre, and the UAA codon is called amber, after the names of the nonsense mutations by which they were originally identified. A blocked reading frame cannot be translated into protein because of the occurrence of termination codons. Key Concepts

• Usually only one reading frame is translated and the other two are blocked by frequent termination signals.

If the genetic code is read in nonoverlapping triplets, there are three possible ways of translating any nucleotide sequence into protein, depending on the starting point. These called reading frames. For the sequence ACGACGACGACGACGACG the three possible reading frames are ACG ACG ACG ACG ACG ACG ACG CGA CGA CGA CGA CGA CGA CGA GAC GAC GAC GAC GAC GAC GAC A reading frame that consists exclusively of triplets representing amino acids is called an open reading frame or ORF. A sequence that is translated into protein has Every sequence has three possible reading frames | SECTION 1.1.22 © 2004. Virtual Text / www.ergito.com

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a reading frame that starts with a special initiation codon (AUG) and that extends through a series of triplets representing amino acids until it ends at one of three types of termination codon (see Molecular Biology 2.5 Messenger RNA). A reading frame that cannot be read into protein because termination codons occur frequently is said to be blocked. If a sequence is blocked in all three reading frames, it cannot have the function of coding for protein. When the sequence of a DNA region of unknown function is obtained, each possible reading frame is analyzed to determine whether it is open or blocked. Usually no more than one of the three possible frames of reading is open in any single stretch of DNA. Figure 1.34 shows an example of a sequence that can be read in only one reading frame, because the alternative reading frames are blocked by frequent termination codons. A long open reading frame is unlikely to exist by chance; if it were not translated into protein, there would have been no selective pressure to prevent the accumulation of termination codons. So the identification of a lengthy open reading frame is taken to be prima facie evidence that the sequence is translated into protein in that frame. An open reading frame (ORF) for which no protein product has been identified is sometimes called an unidentified reading frame (URF).

Figure 1.34 An open reading frame starts with AUG and continues in triplets to a termination codon. Blocked reading frames may be interrupted frequently by termination codons. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.22

Every sequence has three possible reading frames | SECTION 1.1.22 © 2004. Virtual Text / www.ergito.com

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GENES ARE DNA

1.1.23 Prokaryotic genes are colinear with their proteins Key Terms A colinear relationship describes the 1:1 representation of a sequence of triplet nucleotides in a sequence of amino acids. Key Concepts

• A prokaryotic gene consists of a continuous length of 3N nucleotides that codes for N amino acids.

• The gene, mRNA, and protein are all colinear.

By comparing the nucleotide sequence of a gene with the amino acid sequence of a protein, we can determine directly whether the gene and the protein are colinear: whether the sequence of nucleotides in the gene corresponds exactly with the sequence of amino acids in the protein. In bacteria and their viruses, there is an exact equivalence. Each gene contains a continuous stretch of DNA whose length is directly related to the number of amino acids in the protein that it represents. A gene of 3N bp is required to code for a protein of N amino acids, according to the genetic code. The equivalence of the bacterial gene and its product means that a physical map of DNA will exactly match an amino acid map of the protein. How well do these maps fit with the recombination map? The colinearity of gene and protein was originally investigated in the tryptophan synthetase gene of E. coli (see Great Experiments 1.2 Gene-protein colinearity). Genetic distance was measured by the percent recombination between mutations; protein distance was measured by the number of amino acids separating sites of replacement. Figure 1.35 compares the two maps. The order of seven sites of mutation is the same as the order of the corresponding sites of amino acid replacement. And the recombination distances are relatively similar to the actual distances in the protein. The recombination map expands the distances between some mutations, but otherwise there is little distortion of the recombination map relative to the physical map (380; 1225).

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Figure 1.35 The recombination map of the tryptophan synthetase gene corresponds with the amino acid sequence of the protein.

The recombination map makes two further general points about the organization of the gene. Different mutations may cause a wild-type amino acid to be replaced with different substituents. If two such mutations cannot recombine, they must involve different point mutations at the same position in DNA. If the mutations can be separated on the genetic map, but affect the same amino acid on the upper map (the connecting lines converge in the figure), they must involve point mutations at different positions that affect the same amino acid. This happens because the unit of genetic recombination (actually 1 bp) is smaller than the unit coding for the amino acid (actually 3 bp).


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References 380. Yanofsky, C. et al. (1964). On the colinearity of gene structure and protein structure. Proc. Natl. Acad. Sci. USA 51, 266-272. 1225. Yanofsky, C., Drapeau, G. R., Guest, J. R., and Carlton, B. C. (1967). The complete amino acid sequence of the tryptophan synthetase A protein ( µ subunit) and its colinear relationship with the genetic map of the A gene. Proc. Natl. Acad. Sci. USA 57, 2966-2968.



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GENES ARE DNA

1.1.24 Several processes are required to express the protein product of a gene Key Terms Messenger RNA (mRNA) is the intermediate that represents one strand of a gene coding for protein. Its coding region is related to the protein sequence by the triplet genetic code. Transcription describes synthesis of RNA on a DNA template. Translation is synthesis of protein on the mRNA template. A coding region is a part of the gene that represents a protein sequence. The leader of a protein is a short N-terminal sequence responsible for initiating passage into or through a membrane. A trailer (3 ′ UTR) is a nontranslated sequence at the 3 ′ end of an mRNA following the termination codon. Pre-mRNA is used to describe the nuclear transcript that is processed by modification and splicing to give an mRNA. Processing of RNA describes changes that occur after its transcription, including modification of the 5 ′ and 3 ′ ends, internal methylation, splicing, or cleavage. RNA splicing is the process of excising the sequences in RNA that correspond to introns, so that the sequences corresponding to exons are connected into a continuous mRNA. Key Concepts

• A prokaryotic gene is expressed by transcription into mRNA and then by translation of the mRNA into protein.

• In eukaryotes, a gene may contain internal regions that are not represented in protein.

• Internal regions are removed from the RNA transcript by RNA splicing to give an mRNA that is colinear with the protein product.

• Each mRNA consists of a nontranslated 5 ′ leader, a coding region, and a nontranslated 3 ′ trailer.

In comparing gene and protein, we are restricted to dealing with the sequence of DNA stretching between the points corresponding to the ends of the protein. However, a gene is not directly translated into protein, but is expressed via the production of a messenger RNA (abbreviated to mRNA), a nucleic acid intermediate actually used to synthesize a protein (as we see in detail in Molecular Biology 2.5 Messenger RNA).

Several processes are required to express the protein product of a gene | SECTION 1.1.24 © 2004. Virtual Text / www.ergito.com

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Messenger RNA is synthesized by the same process of complementary base pairing used to replicate DNA, with the important difference that it corresponds to only one strand of the DNA double helix. Figure 1.36 shows that the sequence of messenger RNA is complementary with the sequence of one strand of DNA and is identical (apart from the replacement of T with U) with the other strand of DNA. The convention for writing DNA sequences is that the top strand runs 5 ′ → 3 ′ , with the sequence that is the same as RNA.

Figure 1.36 RNA is synthesized by using one strand of DNA as a template for complementary base pairing.

The process by which a gene gives rise to a protein is called gene expression. In bacteria, it consists of two stages. The first stage is transcription, when an mRNA copy of one strand of the DNA is produced. The second stage is translation of the mRNA into protein. This is the process by which the sequence of an mRNA is read in triplets to give the series of amino acids that make the corresponding protein. A messenger RNA includes a sequence of nucleotides that corresponds with the sequence of amino acids in the protein. This part of the nucleic acid is called the coding region. But the messenger RNA includes additional sequences on either end; these sequences do not directly represent protein. The 5 ′ nontranslated region is called the leader, and the 3 ′ nontranslated region is called the trailer. The gene includes the entire sequence represented in messenger RNA. Sometimes mutations impeding gene function are found in the additional, noncoding regions, confirming the view that these comprise a legitimate part of the genetic unit. Figure 1.37 illustrates this situation, in which the gene is considered to comprise a continuous stretch of DNA, needed to produce a particular protein. It includes the sequence coding for that protein, but also includes sequences on either side of the coding region.


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Figure 1.37 The gene may be longer than the sequence coding for protein.

A bacterium consists of only a single compartment, so transcription and translation occur in the same place, as illustrated in Figure 1.38.

Figure 1.38 Transcription and translation take place in the same compartment in bacteria.

In eukaryotes transcription occurs in the nucleus, but the RNA product must be transported to the cytoplasm in order to be translated. For the simplest eukaryotic genes (just like in bacteria) the transcript RNA is in fact the mRNA. But for more complex genes, the immediate transcript of the gene is a pre-mRNA that requires processing to generate the mature mRNA. The basic stages of gene expression in a eukaryote are outlined in Figure 1.39. This results in a spatial separation between transcription (in the nucleus) and translation (in the cytoplasm).


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Figure 1.39 Gene expression is a multistage process.

The most important stage in processing is RNA splicing. Many genes in eukaryotes (and a majority in higher eukaryotes) contain internal regions that do not code for protein. The process of splicing removes these regions from the pre-mRNA to generate an RNA that has a continuous open reading frame (see Figure 2.1). Other processing events that occur at this stage involve the modification of the 5 ′ and 3 ′ ends of the pre-mRNA (see Figure 5.16). Translation is accomplished by a complex apparatus that includes both protein and RNA components. The actual "machine" that undertakes the process is the ribosome, a large complex that includes some large RNAs (ribosomal RNAs, abbreviated to rRNAs) and many small proteins. The process of recognizing which amino acid corresponds to a particular nucleotide triplet requires an intermediate transfer RNA (abbreviated to tRNA); there is at least one tRNA species for every amino acid. Many ancillary proteins are involved. We describe translation in Molecular Biology 2.5 Messenger RNA, but note for now that the ribosomes are the large structures in Figure 1.38 that move along the mRNA. The important point to note at this stage is that the process of gene expression involves RNA not only as the essential substrate, but also in providing components of the apparatus. The rRNA and tRNA components are coded by genes and are generated by the process of transcription (just like mRNA, except that there is no subsequent stage of translation). This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.24


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GENES ARE DNA

1.1.25 Proteins are trans-acting but sites on DNA are cis-acting Key Terms cis configuration describes two sites on the same molecule of DNA. trans configuration of two sites refers to their presence on two different molecules of DNA (chromosomes). A cis-acting site affects the activity only of sequences on its own molecule of DNA (or RNA); this property usually implies that the site does not code for protein. Key Concepts

• All gene products (RNA or proteins) are trans-acting. They can act on any copy of a gene in the cell.

• cis-acting mutations identify sequences of DNA that are targets for recognition by trans-acting products. They are not expressed as RNA or protein and affect only the contiguous stretch of DNA.

A crucial step in the definition of the gene was the realization that all its parts must be present on one contiguous stretch of DNA. In genetic terminology, sites that are located on the same DNA are said to be in cis. Sites that are located on two different molecules of DNA are described as being in trans. So two mutations may be in cis (on the same DNA) or in trans (on different DNAs). The complementation test uses this concept to determine whether two mutations are in the same gene (see Figure 1.27 in Molecular Biology 1.1.16 Mutations in the same gene cannot complement). We may now extend the concept of the difference between cis and trans effects from defining the coding region of a gene to describing the interaction between regulatory elements and a gene. Suppose that the ability of a gene to be expressed is controlled by a protein that binds to the DNA close to the coding region. In the example depicted in Figure 1.40, messenger RNA can be synthesized only when the protein is bound to the DNA. Now suppose that a mutation occurs in the DNA sequence to which this protein binds, so that the protein can no longer recognize the DNA. As a result, the DNA can no longer be expressed.

Proteins are trans-acting but sites on DNA are cis-acting | SECTION 1.1.25 © 2004. Virtual Text / www.ergito.com

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Figure 1.40 Control sites in DNA provide binding sites for proteins; coding regions are expressed via the synthesis of RNA.

So a gene can be inactivated either by a mutation in a control site or by a mutation in a coding region. The mutations cannot be distinguished genetically, because both have the property of acting only on the DNA sequence of the single allele in which they occur. They have identical properties in the complementation test, and a mutation in a control region is therefore defined as comprising part of the gene in the same way as a mutation in the coding region. Figure 1.41 shows that a deficiency in the control site affects only the coding region to which it is connected; it does not affect the ability of the other allele to be expressed. A mutation that acts solely by affecting the properties of the contiguous sequence of DNA is called cis-acting.


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Figure 1.41 A cis-acting site controls the adjacent DNA but does not influence the other allele.

We may contrast the behavior of the cis-acting mutation shown in Figure 1.41 with the result of a mutation in the gene coding for the regulator protein. Figure 1.42 shows that the absence of regulator protein would prevent both alleles from being expressed. A mutation of this sort is said to be trans-acting.


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Figure 1.42 A trans-acting mutation in a protein affects both alleles of a gene that it controls.

Reversing the argument, if a mutation is trans-acting, we know that its effects must be exerted through some diffusible product (typically a protein) that acts on multiple targets within a cell. But if a mutation is cis-acting, it must function via affecting directly the properties of the contiguous DNA, which means that it is not expressed in the form of RNA or protein. Last updated on January 15, 2004 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.25


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GENES ARE DNA

1.1.26 Genetic information can be provided by DNA or RNA Key Terms The central dogma describes the basic nature of genetic information: sequences of nucleic acid can be perpetuated and interconverted by replication, transcription, and reverse transcription, but translation from nucleic acid to protein is unidirectional, because nucleic acid sequences cannot be retrieved from protein sequences. A retrovirus is an RNA virus with the ability to convert its sequence into DNA by reverse transcription. Reverse transcription is synthesis of DNA on a template of RNA. It is accomplished by the enzyme reverse transcriptase. Key Concepts

• Cellular genes are DNA, but viruses and viroids may have genes of RNA. • DNA is converted into RNA by transcription, and RNA may be converted into DNA by reverse transcription.

• The translation of RNA into protein is unidirectional.

The central dogma defines the paradigm of molecular biology. Genes are perpetuated as sequences of nucleic acid, but function by being expressed in the form of proteins. Replication is responsible for the inheritance of genetic information. Transcription and translation are responsible for its conversion from one form to another. Figure 1.43 illustrates the roles of replication, transcription, and translation, viewed from the perspective of the central dogma:

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Figure 1.43 The central dogma states that information in nucleic acid can be perpetuated or transferred, but the transfer of information into protein is irreversible.

• The perpetuation of nucleic acid may involve either DNA or RNA as the genetic material. Cells use only DNA. Some viruses use RNA, and replication of viral RNA occurs in the infected cell. • The expression of cellular genetic information usually is unidirectional. Transcription of DNA generates RNA molecules that can be used further only to generate protein sequences; generally they cannot be retrieved for use as genetic information. Translation of RNA into protein is always irreversible. These mechanisms are equally effective for the cellular genetic information of prokaryotes or eukaryotes, and for the information carried by viruses. The genomes of all living organisms consist of duplex DNA. Viruses have genomes that consist of DNA or RNA; and there are examples of each type that are double-stranded (ds) or single-stranded (ss). Details of the mechanism used to replicate the nucleic acid vary among the viral systems, but the principle of replication via synthesis of complementary strands remains the same, as illustrated in Figure 1.44.


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Figure 1.44 Double-stranded and single-stranded nucleic acids both replicate by synthesis of complementary strands governed by the rules of base pairing.

Cellular genomes reproduce DNA by the mechanism of semi-conservative replication. Double-stranded virus genomes, whether DNA or RNA, also replicate by using the individual strands of the duplex as templates to synthesize partner strands. Viruses with single-stranded genomes use the single strand as template to synthesize a complementary strand; and this complementary strand in turn is used to synthesize its complement, which is, of course, identical with the original starting strand. Replication may involve the formation of stable double-stranded intermediates or use double-stranded nucleic acid only as a transient stage. The restriction to unidirectional transfer from DNA to RNA is not absolute. It is overcome by the retroviruses, whose genomes consist of single-stranded RNA molecules. During the infective cycle, the RNA is converted by the process of reverse transcription into a single-stranded DNA, which in turn is converted into a double-stranded DNA. This duplex DNA becomes part of the genome of the cell, and is inherited like any other gene. So reverse transcription allows a sequence of RNA to be retrieved and used as genetic information. The existence of RNA replication and reverse transcription establishes the general principle that information in the form of either type of nucleic acid sequence can be converted into the other type. In the usual course of events, however, the cell relies on the processes of DNA replication, transcription, and translation. But on rare occasions (possibly mediated by an RNA virus), information from a cellular RNA is converted into DNA and inserted into the genome. Although reverse transcription plays no role in the regular operations of the cell, it becomes a mechanism of potential importance when we consider the evolution of the genome. The same principles are followed to perpetuate genetic information from the massive genomes of plants or amphibians to the tiny genomes of mycoplasma and the yet smaller genetic information of DNA or RNA viruses. Figure 1.45 summarizes some Genetic information can be provided by DNA or RNA | SECTION 1.1.26 © 2004. Virtual Text / www.ergito.com

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examples that illustrate the range of genome types and sizes.

Figure 1.45 The amount of nucleic acid in the genome varies over an enormous range.

Throughout the range of organisms, with genomes varying in total content over a 100,000 fold range, a common principle prevails. The DNA codes for all the proteins that the cell(s) of the organism must synthesize; and the proteins in turn (directly or indirectly) provide the functions needed for survival. A similar principle describes the function of the genetic information of viruses, whether DNA or RNA. The nucleic acid codes for the protein(s) needed to package the genome and also for any functions additional to those provided by the host cell that are needed to reproduce the virus during its infective cycle. (The smallest virus, the satellite tobacco necrosis virus [STNV], cannot replicate independently, but requires the simultaneous presence of a "helper" virus [tobacco necrosis virus, TNV], which is itself a normally infectious virus.) This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.1.26


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GENES ARE DNA

1.1.27 Some hereditary agents are extremely small Key Terms A viroid is a small infectious nucleic acid that does not have a protein coat. Virion is the physical virus particle (irrespective of its ability to infect cells and reproduce). A subviral pathogen is an infectious agent that is smaller than a virus, such as a virusoid. Scrapie is a infective agent made of protein. A prion is a proteinaceous infectious agent, which behaves as an inheritable trait, although it contains no nucleic acid. Examples are PrPSc, the agent of scrapie in sheep and bovine spongiform encephalopathy, and Psi, which confers an inherited state in yeast. PrP is the protein that is the active component of the prion that causes scrapie and related diseases. The form involved in the disease is called PrPSc. Key Concepts

• Some very small hereditary agents do not code for protein but consist of RNA or of protein that has hereditary properties.

Viroids are infectious agents that cause diseases in higher plants (for review see 2525). They are very small circular molecules of RNA. Unlike viruses, where the infectious agent consists of a virion, a genome encapsulated in a protein coat, the viroid RNA is itself the infectious agent. The viroid consists solely of the RNA, which is extensively but imperfectly base paired, forming a characteristic rod like the example shown in Figure 1.46. Mutations that interfere with the structure of the rod reduce infectivity.

Figure 1.46 PSTV RNA is a circular molecule that forms an extensive double-stranded structure, interrupted by many interior loops. The severe and mild forms differ at three sites.

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in infected cells. Its sequence is faithfully perpetuated in its descendants. Viroids fall into several groups. A given viroid is identified with a group by its similarity of sequence with other members of the group. For example, four viroids related to PSTV (potato spindle tuber viroid) have 70-83% similarity of sequence with it. Different isolates of a particular viroid strain vary from one another, and the change may affect the phenotype of infected cells. For example, the mild and severe strains of PSTV differ by three nucleotide substitutions. Viroids resemble viruses in having heritable nucleic acid genomes. They fulfill the criteria for genetic information. Yet viroids differ from viruses in both structure and function. They are sometimes called subviral pathogens. Viroid RNA does not appear to be translated into protein. So it cannot itself code for the functions needed for its survival. This situation poses two questions. How does viroid RNA replicate? And how does it affect the phenotype of the infected plant cell? Replication must be carried out by enzymes of the host cell, subverted from their normal function. The heritability of the viroid sequence indicates that viroid RNA provides the template. Viroids are presumably pathogenic because they interfere with normal cellular processes. They might do this in a relatively random way, for example, by sequestering an essential enzyme for their own replication or by interfering with the production of necessary cellular RNAs. Alternatively, they might behave as abnormal regulatory molecules, with particular effects upon the expression of individual genes (for review see 12). An even more unusual agent is scrapie, the cause of a degenerative neurological disease of sheep and goats. The disease is related to the human diseases of kuru and Creutzfeldt-Jakob syndrome, which affect brain function. The infectious agent of scrapie does not contain nucleic acid. This extraordinary agent is called a prion (proteinaceous infectious agent) (for review see 2523). It is a 28 kD hydrophobic glycoprotein, PrP. PrP is coded by a cellular gene (conserved among the mammals) that is expressed in normal brain. The protein exists in two forms. The product found in normal brain is called PrPc. It is entirely degraded by proteases. The protein found in infected brains is called PrPsc. It is extremely resistant to degradation by proteases. PrPc is converted to PrPsc by a modification or conformational change that confers protease-resistance, and which has yet to be fully defined (383). As the infectious agent of scrapie, PrPsc must in some way modify the synthesis of its normal cellular counterpart so that it becomes infectious instead of harmless (see Molecular Biology 5.23.24 Prions cause diseases in mammals). Mice that lack a PrP gene cannot be infected to develop scrapie, which demonstrates that PrP is essential for development of the disease (386).

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Reviews 12.

Diener, T. O. (1986). Viroid processing: a model involving the central conserved region and hairpin. Proc. Natl. Acad. Sci. USA 83, 58-62.

2523. Prusiner, S. B. (1998). Prions. Proc. Natl. Acad. Sci. USA 95, 13363-13383. 2525. Diener, T. O. (1999). Viroids and the nature of viroid diseases. Arch. Virol. Suppl. 15, 203-220.

References 383. McKinley, M. P., Bolton, D. C., and Prusiner, S. B. (1983). A protease-resistant protein is a structural component of the scrapie prion. Cell 35, 57-62. 386. Bueler, H. et al. (1993). Mice devoid of PrP are resistant to scrapie. Cell 73, 1339-1347.


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GENES ARE DNA

1.1.28 Summary Two classic experiments proved that DNA is the genetic material. DNA isolated from one strain of Pneumococcus bacteria can confer properties of that strain upon another strain. And DNA is the only component that is inherited by progeny phages from the parental phages. DNA can be used to transfect new properties into eukaryotic cells. DNA is a double helix consisting of antiparallel strands in which the nucleotide units are linked by 5 ′ –3 ′ phosphodiester bonds. The backbone provides the exterior; purine and pyrimidine bases are stacked in the interior in pairs in which A is complementary to T while G is complementary to C. The strands separate and use complementary base pairing to assemble daughter strands in semiconservative replication. Complementary base pairing is also used to transcribe an RNA representing one strand of a DNA duplex. A stretch of DNA may code for protein. The genetic code describes the relationship between the sequence of DNA and the sequence of the protein. Only one of the two strands of DNA codes for protein. A codon consists of three nucleotides that represent a single amino acid. A coding sequence of DNA consists of a series of codons, read from a fixed starting point. Usually only one of the three possible reading frames can be translated into protein. A chromosome consists of an uninterrupted length of duplex DNA that contains many genes. Each gene (or cistron) is transcribed into an RNA product, which in turn is translated into a polypeptide sequence if the gene codes for protein. An RNA or protein product of a gene is said to be trans-acting. A gene is defined as a unit on a single stretch of DNA by the complementation test. A site on DNA that regulates the activity of an adjacent gene is said to be cis-acting. A gene may have multiple alleles. Recessive alleles are caused by a loss-of-function. A null allele has total loss-of-function. Dominant alleles are caused by gain-of-function. A mutation consists of a change in the sequence of A·T and G·C base pairs in DNA. A mutation in a coding sequence may change the sequence of amino acids in the corresponding protein. A frameshift mutation alters the subsequent reading frame by inserting or deleting a base; this causes an entirely new series of amino acids to be coded after the site of mutation. A point mutation changes only the amino acid represented by the codon in which the mutation occurs. Point mutations may be reverted by back mutation of the original mutation. Insertions may revert by loss of the inserted material, but deletions cannot revert. Mutations may also be suppressed indirectly when a mutation in a different gene counters the original defect. The natural incidence of mutations is increased by mutagens. Mutations may be concentrated at hotspots. A type of hotspot responsible for some point mutations is caused by deamination of the modified base 5-methylcytosine. Summary | SECTION 1.1.28 © 2004. Virtual Text / www.ergito.com

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Summary | SECTION 1.1.28 © 2004. Virtual Text / www.ergito.com

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THE INTERRUPTED GENE

1.2.1 Introduction Key Terms An exon is any segment of an interrupted gene that is represented in the mature RNA product. An intron (Intervening sequence) is a segment of DNA that is transcribed, but removed from within the transcript by splicing together the sequences (exons) on either side of it. A transcript is the RNA product produced by copying one strand of DNA. It may require processing to generate a mature RNA. RNA splicing is the process of excising the sequences in RNA that correspond to introns, so that the sequences corresponding to exons are connected into a continuous mRNA. A structural gene codes for any RNA or protein product other than a regulator. Key Concepts

• Eukaryotic genomes contain interrupted genes in which exons (represented in the final RNA product) alternate with introns (removed from the initial transcript).

• The exon sequences occur in the same order in the gene and in the RNA, but an

interrupted gene is longer than its final RNA product because of the presence of the introns.

Until eukaryotic genes were characterized by molecular mapping, we assumed that they would have the same organization as prokaryotic genes. We expected the gene to consist of a length of DNA that is colinear with the protein. But a comparison between the structure of DNA and the corresponding mRNA shows a discrepancy in many cases. The mRNA always includes a nucleotide sequence that corresponds exactly with the protein product according to the rules of the genetic code. But the gene includes additional sequences that lie within the coding region, interrupting the sequence that represents the protein. (For a description of the discovery see Great Experiments 4.1 The discovery of RNA splicing and Great Experiments 4.2 The discovery of split genes and RNA splicing.) The sequences of DNA comprising an interrupted gene are divided into the two categories depicted in Figure 2.1:


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Figure 2.1 Interrupted genes are expressed via a precursor RNA. Introns are removed when the exons are spliced together. The mRNA has only the sequences of the exons.

• The exons are the sequences represented in the mature RNA. By definition, a gene starts and ends with exons, corresponding to the 5 ′ and 3 ′ ends of the RNA. • The introns are the intervening sequences that are removed when the primary transcript is processed to give the mature RNA. The expression of interrupted genes requires an additional step that does not occur for uninterrupted genes. The DNA gives rise to an RNA copy (a transcript) that exactly represents the genome sequence. But this RNA is only a precursor; it cannot be used for producing protein. First the introns must be removed from the RNA to give a messenger RNA that consists only of the series of exons. This process is called RNA splicing. It involves a precise deletion of an intron from the primary transcript; the ends of the RNA on either side are joined to form a covalently intact molecule (see Molecular Biology 5.24 RNA splicing and processing). The structural gene comprises the region in the genome between points corresponding to the 5 ′ and 3 ′ terminal bases of mature mRNA. We know that transcription starts at the 5 ′ end of the mRNA, but usually it extends beyond the 3 ′ end, which is generated by cleavage of the RNA (see Molecular Biology 5.24.19 The 3 ′ ends of mRNAs are generated by cleavage and polyadenylation). The gene is considered to include the regulatory regions on both sides of the gene that are required for initiating and (sometimes) terminating gene expression. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.2.1


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1.2.2 An interrupted gene consists of exons and introns Key Concepts

• Introns are removed by the process of RNA splicing, which occurs only in cis on an individual RNA molecule.

• Only mutations in exons can affect protein sequence, but mutations in introns can affect processing of the RNA and therefore prevent production of protein.

How does the existence of introns change our view of the gene? Following splicing, the exons are always joined together in the same order in which they lie in DNA. So the colinearity of gene and protein is maintained between the individual exons and the corresponding parts of the protein chain. Figure 2.2 shows that the order of mutations in the gene remains the same as the order of amino acid replacements in the protein. But the distances in the gene do not correspond at all with the distances in the protein. Genetic distances, as seen on a recombination map, have no relationship to the distances between the corresponding points in the protein. The length of the gene is defined by the length of the initial (precursor) RNA instead of by the length of the messenger RNA.

Figure 2.2 Exons remain in the same order in mRNA as in DNA, but distances along the gene do not correspond to distances along the mRNA or protein products. The distance from A-B in the gene is smaller than the distance from B-C; but the distance from A-B in the mRNA (and protein) is greater than the distance from B-C.

All the exons are represented on the same molecule of RNA, and their splicing together occurs only as an intramolecular reaction. There is usually no joining of exons carried by different RNA molecules, so the mechanism excludes any splicing together of sequences representing different alleles. Mutations located in different exons of a gene cannot complement one another; thus they continue to be defined as members of the same complementation group. Mutations that directly affect the sequence of a protein must lie in exons. What are An interrupted gene consists of exons and introns | SECTION 1.2.2 © 2004. Virtual Text / www.ergito.com

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the effects of mutations in the introns? Since the introns are not part of the messenger RNA, mutations in them cannot directly affect protein structure. However, they can prevent the production of the messenger RNA – for example, by inhibiting the splicing together of exons. A mutation of this sort acts only on the allele that carries it. So it fails to complement any other mutation in that allele, and constitutes part of the same complementation group as the exons. Mutations that affect splicing are usually deleterious. The majority are single base substitutions at the junctions between introns and exons. They may cause an exon to be left out of the product, cause an intron to be included, or make splicing occur at an aberrant site. The most common result is to introduce a termination codon that results in truncation of the protein sequence. About 15% of the point mutations that cause human diseases are caused by disruption of splicing (for review see 3645). Eukaryotic genes are not necessarily interrupted. Some correspond directly with the protein product in the same manner as prokaryotic genes. In yeast, most genes are uninterrupted. In higher eukaryotes, most genes are interrupted; and the introns are usually much longer than exons, creating genes that are very much larger than their coding regions (for review see 8). Last updated on 3-10-2003

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Reviews 8.

Breathnach, R. and Chambon, P. (1981). Organization and expression of eukaryotic split genes coding for proteins. Annu. Rev. Biochem. 50, 349-383.

3645. Faustino, N. A. and Cooper, T. A. (2003). Pre-mRNA splicing and human disease. Genes Dev. 17, 419-437.


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1.2.3 Restriction endonucleases are a key tool in mapping DNA Key Terms Restriction endonucleases recognize specific short sequences of DNA and cleave the duplex (sometimes at target site, sometimes elsewhere, depending on type). A restriction map is a linear array of sites on DNA cleaved by various restriction enzymes. A kilobase (kb) is a measure of length and may be used to refer to DNA (1000 base pairs) or to RNA (1000 bases). A megabase (Mb) is 1 million base pairs of DNA. Key Concepts

• Restriction endonucleases can be used to cleave DNA into defined fragments. • A map can be generated by using the overlaps between the fragments generated by different restriction enzymes.

The characterization of eukaryotic genes was made possible by the development of techniques for physically mapping DNA. The techniques can be extended to (single-stranded) RNA by making a (double-stranded) DNA copy of the RNA. A physical map of any DNA molecule can be obtained by breaking it at defined points whose distance apart can be accurately determined. Specific breaks are made possible by the ability of restriction endonucleases to recognize rather short sequences of double-stranded DNA as targets for cleavage. Each restriction enzyme has a particular target in duplex DNA, usually a specific sequence of 4-6 base pairs. The enzyme cuts the DNA at every point at which its target sequence occurs. Different restriction enzymes have different target sequences, and a large range of these activities (obtained from a wide variety of bacteria) now is available. A restriction map represents a linear sequence of the sites at which particular restriction enzymes find their targets. Distance along such maps is measured directly in base pairs (abbreviated bp) for short distances; longer distances are given in kb, corresponding to kilobase (103) pairs in DNA or to kilobases in RNA. At the level of the chromosome, a map is described in megabase pairs (1 Mb = 106 bp). When a DNA molecule is cut with a suitable restriction enzyme, it is cleaved into distinct fragments. These fragments can be separated on the basis of their size by gel electrophoresis, as shown in Figure 2.3. The cleaved DNA is placed on top of a gel made of agarose or polyacrylamide. When an electric current is passed through the gel, each fragment moves down at a rate that is inversely related to the log of its molecular weight. This movement produces a series of bands. Each band corresponds Restriction endonucleases are a key tool in mapping DNA | SECTION 1.2.3 © 2004. Virtual Text / www.ergito.com

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to a fragment of particular size, decreasing down the gel.

Figure 2.3 Fragments generated by cleaving DNA with a restriction endonuclease can be separated according to their sizes.

By analyzing the restriction fragments of DNA, we can generate a map of the original molecule in the form shown in Figure 2.4. The method is explained in detail in Molecular Biology Supplement 32.11 Restriction mapping. The map shows the positions at which particular restriction enzymes cut DNA; the distances between the sites of cutting are measured in base pairs. So the DNA is divided into a series of regions of defined lengths that lie between sites recognized by the restriction enzymes. An important feature is that a restriction map can be obtained for any sequence of DNA, irrespective of whether mutations have been identified in it, or, indeed, whether we have any knowledge of its function (392) (for review see 6; 10).

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Figure 2.4 A restriction map is a linear sequence of sites separated by defined distances on DNA. The map identifies the sites cleaved by enzymes A and B, as defined by the individual fragments produced by the single and double digests.


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Reviews 6.

Nathans, D. and Smith, H. O. (1975). Restriction endonucleases in the analysis and restructuring of DNA molecules. Annu. Rev. Biochem. 44, 273-293.

10.

Wu, R. (1978). DNA sequence analysis. Annu. Rev. Biochem. 47, 607-734.

References 392. Danna, K. J., Sack, G. H., and Nathans, D. (1973). Studies of SV40 DNA VII A cleavage map of the SV40 genome. J. Mol. Biol. 78, 363-376.



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1.2.4 Organization of interrupted genes may be conserved Key Concepts

• Introns can be detected by the presence of additional regions when genes are

compared with their RNA products by restriction mapping or electron microscopy, but the ultimate definition is based on comparison of sequences.

• The positions of introns are usually conserved when homologous genes are

compared between different organisms, but the lengths of the corresponding introns may vary greatly.

• Introns usually do not code for proteins.

When a gene is uninterrupted, the restriction map of its DNA corresponds exactly with the map of its mRNA. When a gene possesses an intron, the map at each end of the gene corresponds with the map at each end of the message sequence. But within the gene, the maps diverge, because additional regions are found in the gene, but are not represented in the message. Each such region corresponds to an intron. The example of Figure 2.5 compares the restriction maps of a β-globin gene and mRNA. There are two introns. Each intron contains a series of restriction sites that are absent from the cDNA. But the pattern of restriction sites in the exons is the same in both the cDNA and the gene (387; 388; 389; 390; 391).

Figure 2.5 Comparison of the restriction maps of cDNA and genomic DNA for mouse β -globin shows that the gene has two introns that are not present in the cDNA. The exons can be aligned exactly between cDNA and gene.

Ultimately a comparison of the nucleotide sequences of the genomic and mRNA sequences precisely defines the introns. As indicated in Figure 2.6, an intron usually has no open reading frame. An intact reading frame is created in the mRNA sequence by the removal of the introns. Organization of interrupted genes may be conserved | SECTION 1.2.4 © 2004. Virtual Text / www.ergito.com

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Figure 2.6 An intron is a sequence present in the gene but absent from the mRNA (here shown in terms of the cDNA sequence). The reading frame is indicated by the alternating open and shaded blocks; note that all three possible reading frames are blocked by termination codons in the intron.

The structures of eukaryotic genes show extensive variation. Some genes are uninterrupted, so that the genomic sequence is colinear with that of the mRNA. Most higher eukaryotic genes are interrupted, but the introns vary enormously in both number and size. All classes of genes may be interrupted: nuclear genes coding for proteins, nucleolar genes coding for rRNA, and genes coding for tRNA. Interruptions also are found in mitochondrial genes in lower eukaryotes, and in chloroplast genes. Interrupted genes do not appear to be excluded from any class of eukaryotes, and have been found in bacteria and bacteriophages, although they are extremely rare in prokaryotic genomes. Some interrupted genes possess only one or a few introns. The globin genes provide an extensively studied example (see Molecular Biology 1.2.11 The members of a gene family have a common organization). The two general types of globin gene, α and β, share a common type of structure. The consistency of the organization of mammalian globin genes is evident from the structure of the "generic" globin gene summarized in Figure 2.7.

Figure 2.7 All functional globin genes have an interrupted structure with three exons. The lengths indicated in the figure apply to the mammalian β -globin genes.

Interruptions occur at homologous positions (relative to the coding sequence) in all known active globin genes, including those of mammals, birds, and frogs. The first intron is always fairly short, and the second usually is longer, but the actual lengths Organization of interrupted genes may be conserved | SECTION 1.2.4 © 2004. Virtual Text / www.ergito.com

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can vary. Most of the variation in overall lengths between different globin genes results from the variation in the second intron. In the mouse, the second intron in the α-globin gene is only 150 bp long, so the overall length of the gene is 850 bp, compared with the major β-globin gene where the intron length of 585 bp gives the gene a total length of 1382 bp. The variation in length of the genes is much greater than the range of lengths of the mRNAs ( α-globin mRNA = 585 bases, β-globin mRNA = 620 bases). The example of DHFR, a somewhat larger gene, is shown in Figure 2.8. The mammalian DHFR (dihydrofolate reductase) gene is organized into 6 exons that correspond to the 2000 base mRNA. But they extend over a much greater length of DNA because the introns are very long. In three mammals the exons remain essentially the same, and the relative positions of the introns are unaltered, but the lengths of individual introns vary extensively, resulting in a variation in the length of the gene from 25-31 kb.

Figure 2.8 Mammalian genes for DHFR have the same relative organization of rather short exons and very long introns, but vary extensively in the lengths of corresponding introns.

The globin and DHFR genes present examples of a general phenomenon: genes that are related by evolution have related organizations, with conservation of the positions of (at least some) of the introns. Variations in the lengths of the genes are primarily determined by the lengths of the introns.

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References 387. Wenskink, P. et al. (1974). A system for mapping DNA sequences in the chromosomes ofD. melanogaster. Cell 3, 315-325. 388. Berget, S. M., Moore, C., and Sharp, P. (1977). Spliced segments at the 5 ′ terminus of adenovirus 2 late mRNA. Proc. Natl. Acad. Sci. USA 74, 3171-3175. 389. Chow, L. T., Gelinas, R. E., Broker, T. R., and Roberts, R. J. (1977). An amazing sequence arrangement at the 5 ′ ends of adenovirus 2 mRNA. Cell 12, 1-8. 390. Glover, D. M. and Hogness, D. S. (1977). A novel arrangement of the 8S and 28S sequences in a repeating unit of D. melanogaster rDNA. Cell 10, 167-176. 391. Jeffreys, A. J. and Flavell, R. A. (1977). The rabbit β -globin gene contains a large insert in the coding sequence. Cell 12, 1097-1108.


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1.2.5 Exon sequences are conserved but introns vary Key Concepts

• Comparisons of related genes in different species show that the sequences of the corresponding exons are usually conserved but the sequences of the introns are much less well related.

• Introns evolve much more rapidly than exons because of the lack of selective pressure to produce a protein with a useful sequence.

Is a structural gene unique in its genome? The answer can be ambiguous. The entire length of the gene is unique as such, but its exons often are related to those of other genes. As a general rule, when two genes are related, the relationship between their exons is closer than the relationship between the introns. In an extreme case, the exons of two genes may code for the same protein sequence, but the introns may be different. This implies that the two genes originated by a duplication of some common ancestral gene. Then differences accumulated between the copies, but they were restricted in the exons by the need to code for protein functions. As we see later when we consider the evolution of the gene, exons can be considered as basic building blocks that are assembled in various combinations. A gene may have some exons that are related to exons of another gene, but the other exons may be unrelated. Usually the introns are not related at all in such cases. Such genes may arise by duplication and translocation of individual exons. The relationship between two genes can be plotted in the form of the dot matrix comparison of Figure 2.9. A dot is placed to indicate each position at which the same sequence is found in each gene. The dots form a line at an angle of 45° if two sequences are identical. The line is broken by regions that lack similarity, and it is displaced laterally or vertically by deletions or insertions in one sequence relative to the other.

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Figure 2.9 The sequences of the mouse α maj and α min globin genes are closely related in coding regions, but differ in the flanking regions and large intron. Data kindly provided by Philip Leder.

When the two β-globin genes of the mouse are compared, such a line extends through the three exons and through the small intron. The line peters out in the flanking regions and in the large intron. This is a typical pattern, in which coding sequences are well related, the relationship can extend beyond the boundaries of the exons, but it is lost in longer introns and the regions on either side of the gene. The overall degree of divergence between two exons is related to the differences between the proteins. It is caused mostly by base substitutions. In the translated regions, the exons are under the constraint of needing to code for amino acid sequences, so they are limited in their potential to change sequence. Many of the changes do not affect codon meanings, because they change one codon into another that represents the same amino acid. Changes occur more freely in nontranslated regions (corresponding to the 5 ′ leader and 3 ′ trailer of the mRNA). In corresponding introns, the pattern of divergence involves both changes in size (due to deletions and insertions) and base substitutions. Introns evolve much more rapidly than exons. When a gene is compared in different species, sometimes the exons are homologous, while the introns have diverged so much that corresponding sequences cannot be recognized. Mutations occur at the same rate in both exons and introns, but are removed more effectively from the exons by adverse selection. However, in the absence of the constraints imposed by a coding function, an intron is able quite freely to accumulate point substitutions and other changes. These changes imply that the intron does not have a sequence-specific function. Whether its presence is at all necessary for gene function is not clear. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.1.2.5

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1.2.6 Genes can be isolated by the conservation of exons Key Terms A zoo blot describes the use of Southern blotting to test the ability of a DNA probe from one species to hybridize with the DNA from the genomes of a variety of other species. Exon trapping inserts a genomic fragment into a vector whose function depends on the provision of splicing junctions by the fragment. Key Concepts

• Conservation of exons can be used as the basis for identifying coding regions by identifying fragments whose sequences are present in multiple organisms.

Some major approaches to identifying genes are based on the contrast between the conservation of exons and the variation of introns. In a region containing a gene whose function has been conserved among a range of species, the sequence representing the protein should have two distinctive properties: • it must have an open reading frame; • and it is likely to have a related sequence in other species. These features can be used to isolate genes. Suppose we know by genetic data that a particular genetic trait is located in a given chromosomal region. If we lack knowledge about the nature of the gene product, how are we to identify the gene in a region that may be (for example) >1 Mb? A heroic approach that has proved successful with some genes of medical importance is to screen relatively short fragments from the region for the two properties expected of a conserved gene. First we seek to identify fragments that cross-hybridize with the genomes of other species. Then we examine these fragments for open reading frames. The first criterion is applied by performing a zoo blot. We use short fragments from the region as (radioactive) probes to test for related DNA from a variety of species by Southern blotting. If we find hybridizing fragments in several species related to that of the probe – the probe is usually human – the probe becomes a candidate for an exon of the gene. The candidates are sequenced, and if they contain open reading frames, are used to isolate surrounding genomic regions. If these appear to be part of an exon, we may Genes can be isolated by the conservation of exons | SECTION 1.2.6 © 2004. Virtual Text / www.ergito.com

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then use them to identify the entire gene, to isolate the corresponding cDNA or mRNA, and ultimately to identify the protein. This approach is especially important when the target gene is spread out because it has many large introns. This proved to be the case with Duchenne muscular dystrophy (DMD), a degenerative disorder of muscle, which is X-linked and affects 1 in 3500 of human male births. The steps in identifying the gene are summarized in Figure 2.10.

Figure 2.10 The gene involved in Duchenne muscular dystrophy was tracked down by chromosome mapping and walking to a region in which deletions can be identified with the occurrence of the disease.

Linkage analysis localized the DMD locus to chromosomal band Xp21. Patients with the disease often have chromosomal rearrangements involving this band. By comparing the ability of X-linked DNA probes to hybridize with DNA from patients and with normal DNA, cloned fragments were obtained that correspond to the region that was rearranged or deleted in patients' DNA. Once some DNA in the general vicinity of the target gene has been obtained, it is Genes can be isolated by the conservation of exons | SECTION 1.2.6 © 2004. Virtual Text / www.ergito.com

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possible to "walk" along the chromosome until the gene is reached (see Molecular Biology Supplement 32.12 Genome mapping). A chromosomal walk was used to construct a restriction map of the region on either side of the probe, covering a region of >100 kb. Analysis of the DNA from a series of patients identified large deletions in this region, extending in either direction. The most telling deletion is one contained entirely within the region, since this delineates a segment that must be important in gene function and indicates that the gene, or at least part of it, lies in this region (3032; 3033). Having now come into the region of the gene, we need to identify its exons and introns. A zoo blot identified fragments that cross-hybridize with the mouse X chromosome and with other mammalian DNAs. As summarized in Figure 2.11, these were scrutinized for open reading frames and the sequences typical of exon-intron junctions. Fragments that met these criteria were used as probes to identify homologous sequences in a cDNA library prepared from muscle mRNA.

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Figure 2.11 The Duchenne muscular dystrophy gene was characterized by zoo blotting, cDNA hybridization, genomic hybridization, and identification of the protein.

The cDNA corresponding to the gene identifies an unusually large mRNA, ~14 kb. Hybridization back to the genome shows that the mRNA is represented in >60 exons, which are spread over ~2000 kb of DNA. This makes DMD the longest gene identified; in fact, it is 10× longer than any other known gene (3034; 3035). The gene codes for a protein of ~500 kD, called dystrophin, which is a component of muscle, present in rather low amounts. All patients with the disease have deletions at this locus, and lack (or have defective) dystrophin.


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Muscle also has the distinction of having the largest known protein, titin, with almost 27,000 amino acids. Its gene has the largest number of exons (178) and the longest single exon in the human genome (17,000 bp). Another technique that allows genomic fragments to be scanned rapidly for the presence of exons is called exon trapping (3030). Figure 2.12 shows that it starts with a vector that contains a strong promoter, and has a single intron between two exons. When this vector is transfected into cells, its transcription generates large amounts of an RNA containing the sequences of the two exons. A restriction cloning site lies within the intron, and is used to insert genomic fragments from a region of interest. If a fragment does not contain an exon, there is no change in the splicing pattern, and the RNA contains only the same sequences as the parental vector. But if the genomic fragment contains an exon flanked by two partial intron sequences, the splicing sites on either side of this exon are recognized, and the sequence of the exon is inserted into the RNA between the two exons of the vector. This can be detected readily by reverse transcribing the cytoplasmic RNA into cDNA, and using PCR to amplify the sequences between the two exons of the vector. So the appearance in the amplified population of sequences from the genomic fragment indicates that an exon has been trapped. Because introns are usually large and exons are small in animal cells, there is a high probability that a random piece of genomic DNA will contain the required structure of an exon surrounded by partial introns. In fact, exon trapping may mimic the events that have occurred naturally during evolution of genes (see Molecular Biology 1.2.9 How did interrupted genes evolve?).


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Figure 2.12 A special splicing vector is used for exon trapping. If an exon is present in the genomic fragment, its sequence will be recovered in the cytoplasmic RNA, but if the genomic fragment consists solely of sequences from within intron, splicing does not occur, and the mRNA is not exported to the cytoplasm.



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References 3030. Buckler, A. J., Chang, D. D., Graw, S. L., Brook, J. D., Haber, D. A., Sharp, P. A., and Housman, D. E. (1991). Exon amplification: a strategy to isolate mammalian genes based on RNA splicing. Proc. Natl. Acad. Sci. USA 88, 4005-4009. 3032. Kunkel, L. M., Monaco, A. P., Middlesworth, W., Ochs, H. D., and Latt, S. A. (1985). Specific cloning of DNA fragments absent from the DNA of a male patient with an X chromosome deletion. Proc. Natl. Acad. Sci. USA 82, 4778-4782. 3033. Monaco, A.P., Bertelson, C. J., Middlesworth, W., Colletti, C. A., Aldridge, J., Fischbeck, K. H., Bartlett, R., Pericak-Vance, M. A., Roses, A. D., and Kunkel, L. M. (1985). Detection of deletions spanning the Duchenne muscular dystrophy locus using a tightly linked DNA segment. Nature 316, 842-845. 3034. van Ommen, G. J., Verkerk, J. M., Hofker, M. H., Monaco, A. P., Kunkel, L. M., Ray, P., Worton, R., Wieringa, B., Bakker, E., and Pearson, P. L. (1986). A physical map of 4 million bp around the Duchenne muscular dystrophy gene on the human X-chromosome. Cell 47, 499-504. 3035. Koenig, M., Hoffman, E. P., Bertelson, C. J., Monaco, A. P., Feener, C., and Kunkel, L. M. (1987). Complete cloning of the Duchenne muscular dystrophy (DMD) cDNA and preliminary genomic organization of the DMD gene in normal and affected individuals. Cell 50, 509-517.



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1.2.7 Genes show a wide distribution of sizes Key Concepts

• Most genes are uninterrupted in yeasts, but are interrupted in higher eukaryotes. • Exons are usually short, typically coding for 96%) are not interrupted, and those that have exons usually remain reasonably compact. There are virtually no S. cerevisiae genes with more than 4 exons.

Figure 2.13 Most genes are uninterrupted in yeast, but most genes are interrupted in flies and mammals. (Uninterrupted genes have only 1 exon, and are totaled in the leftmost column.)

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In insects and mammals, the situation is reversed. Only a few genes have uninterrupted coding sequences (6% in mammals). Insect genes tend to have a fairly small number of exons, typically fewer than 10. Mammalian genes are split into more pieces, and some have several 10s of exons. ~50% of mammalian genes have >10 introns. Examining the consequences of this type of organization for the overall size of the gene, we see in Figure 2.14 that there is a striking difference between yeast and the higher eukaryotes. The average yeast gene is 1.4 kb long, and very few are longer than 5 kb. The predominance of interrupted genes in high eukaryotes, however, means that the gene can be much larger than the unit that codes for protein. Relatively few genes in flies or mammals are shorter than 2 kb, and many have lengths between 5 kb and 100 kb. The average human gene is 27 kb long (see Figure 3.22).

Figure 2.14 Yeast genes are small, but genes in flies and mammals have a dispersed distribution extending to very large sizes.

The switch from largely uninterrupted to largely interrupted genes occurs in the lower eukaryotes. In fungi (excepting the yeasts), the majority of genes are interrupted, but they have a relatively small number of exons (50 modified bases. • Modification usually involves direct alteration of the primary bases in tRNA, but there are some exceptions in which a base is removed and replaced by another base.

Transfer RNA is unique among nucleic acids in its content of "unusual" bases. An unusual base is any purine or pyrimidine ring except the usual A, G, C, and U from which all RNAs are synthesized. All other bases are produced by modification of one of the four bases after it has been incorporated into the polyribonucleotide chain. All classes of RNA display some degree of modification, but in all cases except tRNA this is confined to rather simple events, such as the addition of methyl groups. In tRNA, there is a vast range of modifications, ranging from simple methylation to wholesale restructuring of the purine ring. Modifications occur in all parts of the tRNA molecule There are >50 different types of modified bases in tRNA. Figure 7.7 shows some of the more common modified bases. Modifications of pyrimidines (C and U) are less complex than those of purines (A and G). In addition to the modifications of the bases themselves, methylation at the 2 ′ –O position of the ribose ring also occurs.

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Figure 7.7 All of the four bases in tRNA can be modified.

The most common modifications of uridine are straightforward. Methylation at position 5 creates ribothymidine (T). The base is the same commonly found in DNA; but here it is attached to ribose, not deoxyribose. In RNA, thymine constitutes an unusual base, originating by modification of U. Dihydrouridine (D) is generated by the saturation of a double bond, changing the ring structure. Pseudouridine ( ψ) interchanges the positions of N and C atoms (see Figure 24.40). And 4-thiouridine has sulfur substituted for oxygen. The nucleoside inosine is found normally in the cell as an intermediate in the purine biosynthetic pathway. However, it is not incorporated directly into RNA, where instead its existence depends on modification of A to create I. Other modifications of A include the addition of complex groups. Two complex series of nucleotides depend on modification of G. The Q bases, such as queuosine, have an additional pentenyl ring added via an NH linkage to the methyl group of 7-methylguanosine. The pentenyl ring may carry various further groups. tRNA contains modified bases | SECTION 2.7.5 © 2004. Virtual Text / www.ergito.com

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The Y bases, such as wyosine, have an additional ring fused with the purine ring itself; the extra ring carries a long carbon chain, again to which further groups are added in different cases. The modification reaction usually involves the alteration of, or addition to, existing bases in the tRNA. An exception is the synthesis of Q bases, where a special enzyme exchanges free queuosine with a guanosine residue in the tRNA. The reaction involves breaking and remaking bonds on either side of the nucleoside. The modified nucleosides are synthesized by specific tRNA-modifying enzymes. The original nucleoside present at each position can be determined either by comparing the sequence of tRNA with that of its gene or (less efficiently) by isolating precursor molecules that lack some or all of the modifications. The sequences of precursors show that different modifications are introduced at different stages during the maturation of tRNA. Some modifications are constant features of all tRNA molecules – for example, the D residues that give rise to the name of the D arm, and the ψ found in the T ψC sequence. On the 3 ′ side of the anticodon there is always a modified purine, although the modification varies widely. Other modifications are specific for particular tRNAs or groups of tRNAs. For example, wyosine bases are characteristic of tRNAPhe in bacteria, yeast, and mammals. There are also some species-specific patterns. The many tRNA-modifying enzymes (~60 in yeast) vary greatly in specificity (for review see 3461). In some cases, a single enzyme acts to make a particular modification at a single position. In other cases, an enzyme can modify bases at several different target positions. Some enzymes undertake single reactions with individual tRNAs; others have a range of substrate molecules. The features recognized by the tRNA-modifying enzymes are unknown, but probably involve recognition of structural features surrounding the site of modification. Some modifications require the successive actions of more than one enzyme. Last updated on 1-30-2003


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Reviews 3461. Hopper, A. K. and Phizicky, E. M. (2003). tRNA transfers to the limelight. Genes Dev. 17, 162-180.



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USING THE GENETIC CODE

2.7.6 Modified bases affect anticodon-codon pairing Key Concepts

• Modifications in the anticodon affect the pattern of wobble pairing and therefore are important in determining tRNA specificity.

The most direct effect of modification is seen in the anticodon, where change of sequence influences the ability to pair with the codon, thus determining the meaning of the tRNA. Modifications elsewhere in the vicinity of the anticodon also influence its pairing. When bases in the anticodon are modified, further pairing patterns become possible in addition to those predicted by the regular and wobble pairing involving A, C, U, and G. Figure 7.8 shows the use of inosine (I), which is often present at the first position of the anticodon. Inosine can pair with any one of three bases, U, C, and A.

Figure 7.8 Inosine can pair with any of U, C, and A.

This ability is especially important in the isoleucine codons, where AUA codes for isoleucine, while AUG codes for methionine. Because with the usual bases it is not possible to recognize A alone in the third position, any tRNA with U starting its anticodon would have to recognize AUG as well as AUA. So AUA must be read together with AUU and AUC, a problem that is solved by the existence of tRNA Modified bases affect anticodon-codon pairing | SECTION 2.7.6 © 2004. Virtual Text / www.ergito.com

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with I in the anticodon. Actually, some of the predicted regular combinations do not occur, because some bases are always modified. There seems to be an absolute ban on the employment of A; usually it is converted to I. And U at the first position of the anticodon is usually converted to a modified form that has altered pairing properties. Some modifications create preferential readings of some codons with respect to others. Anticodons with uridine-5-oxyacetic acid and 5-methoxyuridine in the first position recognize A and G efficiently as third bases of the codon, but recognize U less efficiently. Another case in which multiple pairings can occur, but with some preferred to others, is provided by the series of queuosine and its derivatives. These modified G bases continue to recognize both C and U, but pair with U more readily. A restriction not allowed by the usual rules can be achieved by the employment of 2-thiouridine in the anticodon. Figure 7.9 shows that its modification allows the base to continue to pair with A, but prevents it from indulging in wobble pairing with G (for review see 32).

Figure 7.9 Modification to 2-thiouridine restricts pairing to A alone because only one H-bond can form with G.

These and other pairing relationships make the general point that there are multiple ways to construct a set of tRNAs able to recognize all the 61 codons representing amino acids. No particular pattern predominates in any given organism, although the absence of a certain pathway for modification can prevent the use of some recognition patterns. So a particular codon family is read by tRNAs with different anticodons in different organisms. Often the tRNAs will have overlapping responses, so that a particular codon is read by more than one tRNA. In such cases there may be differences in the efficiencies of Modified bases affect anticodon-codon pairing | SECTION 2.7.6 © 2004. Virtual Text / www.ergito.com

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the alternative recognition reactions. (As a general rule, codons that are commonly used tend to be more efficiently read.) And in addition to the construction of a set of tRNAs able to recognize all the codons, there may be multiple tRNAs that respond to the same codons. The predictions of wobble pairing accord very well with the observed abilities of almost all tRNAs. But there are exceptions in which the codons recognized by a tRNA differ from those predicted by the wobble rules. Such effects probably result from the influence of neighboring bases and/or the conformation of the anticodon loop in the overall tertiary structure of the tRNA. Indeed, the importance of the structure of the anticodon loop is inherent in the idea of the wobble hypothesis itself. Further support for the influence of the surrounding structure is provided by the isolation of occasional mutants in which a change in a base in some other region of the molecule alters the ability of the anticodon to recognize codons. Another unexpected pairing reaction is presented by the ability of the bacterial initiator, fMet-tRNAf, to recognize both AUG and GUG. This misbehavior involves the third base of the anticodon.

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Reviews 32.

Bjork, G. R (1987). Transfer RNA modification. Annu. Rev. Biochem. 56, 263-287.


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2.7.7 There are sporadic alterations of the universal code Key Concepts

• Changes in the universal genetic code have occurred in some species. • They are more common in mitochondrial genomes, where a phylogenetic tree can be constructed for the changes.

• In nuclear genomes, they are sporadic and usually affect only termination codons.

The universality of the genetic code is striking, but some exceptions exist. They tend to affect the codons involved in initiation or termination and result from the production (or absence) of tRNAs representing certain codons. The changes found in principal (bacterial or nuclear) genomes are summarized in Figure 7.10.

Figure 7.10 Changes in the genetic code in bacterial or eukaryotic nuclear genomes usually assign amino acids to stop codons or change a codon so that it no longer specifies an amino acid. A change in meaning from one amino acid to another is unusual.

Almost all of the changes that allow a codon to represent an amino acid affect termination codons: • In the prokaryote Mycoplasma capricolum, UGA is not used for termination, but instead codes for tryptophan. In fact, it is the predominant Trp codon, and UGG is used only rarely. Two Trp-tRNA species exist, with the anticodons UCA ← There are sporadic alterations of the universal code | SECTION 2.7.7 © 2004. Virtual Text / www.ergito.com

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(reads UGA and UGG) and CCA ← (reads only UGG). • Some ciliates (unicellular protozoa) read UAA and UAG as glutamine instead of termination signals. Tetrahymena thermophila, one of the ciliates, contains three tRNAGlu species. One recognizes the usual codons CAA and CAG for glutamine, one recognizes both UAA and UAG (in accordance with the wobble hypothesis), and the last recognizes only UAG. We assume that a further change is that the release factor eRF has a restricted specificity, compared with that of other eukaryotes. • In another ciliate (Euplotes octacarinatus), UGA codes for cysteine. Only UAA is used as a termination codon, and UAG is not found. The change in meaning of UGA might be accomplished by a modification in the anticodon of tRNACys to allow it to read UGA with the usual codons UGU and UGC. • The only substitution in coding for amino acids occurs in a yeast (Candida), where CUG means serine instead of leucine (and UAG is used as a sense codon). Acquisition of a coding function by a termination codon requires two types of change: a tRNA must be mutated so as to recognize the codon; and the class 1 release factor must be mutated so that it does not terminate at this codon. The other common type of change is loss of the tRNA that responds to a codon, so that the codon no longer specifies any amino acid. What happens at such a codon will depend on whether the termination factor evolves to recognize it. All of these changes are sporadic, which is to say that they appear to have occurred independently in specific lines of evolution. They may be concentrated on termination codons, because these changes do not involve substitution of one amino acid for another. Once the genetic code was established, early in evolution, any general change in the meaning of a codon would cause a substitution in all the proteins that contain that amino acid. It seems likely that the change would be deleterious in at least some of these proteins, with the result that it would be strongly selected against. The divergent uses of the termination codons could represent their "capture" for normal coding purposes. If some termination codons were used only rarely, they could be recruited to coding purposes by changes that allowed tRNAs to recognize them. Exceptions to the universal genetic code also occur in the mitochondria from several species. Figure 7.11 constructs a phylogeny for the changes. It suggests that there was a universal code that was changed at various points in mitochondrial evolution. The earliest change was the employment of UGA to code for tryptophan, which is common to all (non-plant) mitochondria (for review see 41).

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Figure 7.11 Changes in the genetic code in mitochondria can be traced in phylogeny. The minimum number of independent changes is generated by supposing that the AUA=Met and the AAA=Asn changes each occurred independently twice, and that the early AUA=Met change was reversed in echinoderms.

Some of these changes make the code simpler, by replacing two codons that had different meanings with a pair that has a single meaning. Pairs treated like this include UGG and UGA (both Trp instead of one Trp and one termination) and AUG and AUA (both Met instead of one Met and the other Ile). Why have changes been able to evolve in the mitochondrial code? Because the mitochondrion synthesizes only a small number of proteins (~10), the problem of disruption by changes in meaning is much less severe. Probably the codons that are altered were not used extensively in locations where amino acid substitutions would have been deleterious. The variety of changes found in mitochondria of different species suggests that they have evolved separately, and not by common descent from an ancestral mitochondrial code. According to the wobble hypothesis, a minimum of 31 tRNAs (excluding the initiator) are required to recognize all 61 codons (at least 2 tRNAs are required for each codon family and 1 tRNA is needed per codon pair or single codon). But an unusual situation exists in (at least) mammalian mitochondria in which there are only 22 different tRNAs. How does this limited set of tRNAs accommodate all the codons? The critical feature lies in a simplification of codon-anticodon pairing, in which one tRNA recognizes all four members of a codon family. This reduces to 23 the minimum number of tRNAs required to respond to all usual codons. The use of AGAG for termination reduces the requirement by one further tRNA, to 22. In all eight codon families, the sequence of the tRNA contains an unmodified U at the first position of the anticodon. The remaining codons are grouped into pairs in which all the codons ending in pyrimidines are read by G in the anticodon, and all the codons ending in purines are read by a modified U in the anticodon, as predicted by the wobble hypothesis. The complication of the single UGG codon is avoided by the change in the code to read UGA with UGG as tryptophan; and in mammals, AUA ceases to represent isoleucine and instead is read with AUG as methionine. There are sporadic alterations of the universal code | SECTION 2.7.7 © 2004. Virtual Text / www.ergito.com

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This allows all the nonfamily codons to be read as 14 pairs. The 22 identified tRNA genes therefore code for 14 tRNAs representing pairs, and 8 tRNAs representing families. This leaves the two usual termination codons UAG and UAA unrecognized by tRNA, together with the codon pair AGAG. Similar rules are followed in the mitochondria of fungi (for review see 33). Last updated on 12-17-2001


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Reviews 33.

Fox, T. D. (1987). Natural variation in the genetic code. Annu. Rev. Genet. 21, 67-91.

41.

Osawa, S. et al. (1992). Recent evidence for evolution of the genetic code. Microbiol. Rev. 56, 229-264.



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2.7.8 Novel amino acids can be inserted at certain stop codons Key Concepts

• Changes in the reading of specific codons can occur in individual genes. • The insertion of seleno-Cys-tRNA at certain UGA codons requires several proteins to modify the Cys-tRNA and insert it into the ribosome.

• Pyrrolysine can be inserted at certain UAG codons.

Specific changes in reading the code occur in individual genes. The specificity of such changes implies that the reading of the particular codon must be influenced by the surrounding bases. A striking example is the incorporation of the modified amino acid seleno-cysteine at certain UGA codons within the genes that code for selenoproteins in both prokaryotes and eukaryotes. Usually these proteins catalyze oxidation-reduction reactions, and contain a single seleno-cysteine residue, which forms part of the active site. The most is known about the use of the UGA codons in three E. coli genes coding for formate dehydrogenase isozymes. The internal UGA codon is read by a seleno-Cys-tRNA. This unusual reaction is determined by the local secondary structure of mRNA, in particular by the presence of a hairpin loop downstream of the UGA. Mutations in 4 sel genes create a deficiency in selenoprotein synthesis. selC codes for tRNA (with the anticodon ACU ← ) that is charged with serine. selA and selD are required to modify the serine to seleno-cysteine. SelB is an alternative elongation factor. It is a guanine nucleotide-binding protein that acts as a specific translation factor for entry of seleno-Cys-tRNA into the A site; it thus provides (for this single tRNA) a replacement for factor EF-Tu. The sequence of SelB is related to both EF-Tu and IF-2 (for review see 39). Why is seleno-Cys-tRNA inserted only at certain UGA codons? These codons are followed by a stem-loop structure in the mRNA. Figure 7.12 shows that the stem of this structure is recognized by an additional domain in SelB (one that is not present in EF-Tu or IF-2). A similar mechanism interprets some UGA codons in mammalian cells, except that two proteins are required to identify the appropriate UGA codons. One protein (SBP2) binds a stem-loop structure far downstream from the UGA codon, while the counterpart of SelB (called SECIS) binds to SBP2 and simultaneously binds the tRNA to the UGA codon (1186).

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Figure 7.12 SelB is an elongation factor that specifically binds Seleno-Cys-tRNA to a UGA codon that is followed by a stem-loop structure in mRNA.

Another example of the insertion of a special amino acid is the placement of pyrrolysine at a UAG codon. This happens in both an archaea and a bacterium (2492; 2493). The mechanism is probably similar to the insertion of seleno-cysteine. An unusual tRNA is charged with lysine, which is presumably then modified. The tRNA has a CUA anticodon, which responds to UAG. There must be other components of the system that restricts its response to the appropriate UAG codons. Last updated on 5-28-2002


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Reviews 39.

Bock, A. (1991). Selenoprotein synthesis: an expansion of the genetic code. Trends Biochem. Sci. 16, 463-467.

References 1186. Fagegaltier, D., Hubert, N., Yamada, K., Mizutani, T., Carbon, P., and Krol, A. (2000). Characterization of mSelB, a novel mammalian elongation factor for selenoprotein translation. EMBO J. 19, 4796-4805. 2492. Srinivasan, G., James, C. M., and Krzycki, J. A. (2002). Pyrrolysine encoded by UAG in Archaea: charging of a UAG-decoding specialized tRNA. Science 296, 1459-1462. 2493. Hao, B., Gong, W., Ferguson, T. K., James, C. M., Krzycki, J. A., and Chan, M. K. (2002). A new UAG-encoded residue in the structure of a methanogen methyltransferase. Science 296, 1462-1466.



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2.7.9 tRNAs are charged with amino acids by synthetases Key Terms Cognate tRNAs (Isoaccepting tRNA) are those recognized by a particular aminoacyl-tRNA synthetase. They all are charged with the same amino acid. Key Concepts

• Aminoacyl-tRNA synthetases are enzymes that charge tRNA with an amino acid to generate aminoacyl-tRNA in a two-stage reaction that uses energy from ATP.

• There are 20 aminoacyl-tRNA synthetases in each cell. Each charges all the tRNAs that represent a particular amino acid.

• Recognition of a tRNA is based on a small number of points of contact in the tRNA sequence.

It is necessary for tRNAs to have certain characteristics in common, yet be distinguished by others. The crucial feature that confers this capacity is the ability of tRNA to fold into a specific tertiary structure. Changes in the details of this structure, such as the angle of the two arms of the "L" or the protrusion of individual bases, may distinguish the individual tRNAs. All tRNAs can fit in the P and A sites of the ribosome, where at one end they are associated with mRNA via codon-anticodon pairing, while at the other end the polypeptide is being transferred. Similarly, all tRNAs (except the initiator) share the ability to be recognized by the translation factors (EF-Tu or eEF1) for binding to the ribosome. The initiator tRNA is recognized instead by IF-2 or eIF2. So the tRNA set must possess common features for interaction with elongation factors, but the initiator tRNA can be distinguished. Amino acids enter the protein synthesis pathway through the aminoacyl-tRNA synthetases, which provide the interface for connection with nucleic acid. All synthetases function by the two-step mechanism depicted in Figure 7.13:

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Figure 7.13 An aminoacyl-tRNA synthetase charges tRNA with an amino acid.

• First, the amino acid reacts with ATP to form aminoacyl~adenylate, releasing pyrophosphate. Energy for the reaction is provided by cleaving the high energy bond of the ATP. • Then the activated amino acid is transferred to the tRNA, releasing AMP. The synthetases sort the tRNAs and amino acids into corresponding sets. Each synthetase recognizes a single amino acid and all the tRNAs that should be charged with it. Usually, each amino acid is represented by more than one tRNA. Several tRNAs may be needed to respond to synonym codons, and sometimes there are multiple species of tRNA reacting with the same codon. Multiple tRNAs representing the same amino acid are called isoaccepting tRNAs; because they are all recognized by the same synthetase, they are also described as its cognate tRNAs. Many attempts to deduce similarities in sequence between cognate tRNAs, or to tRNAs are charged with amino acids by synthetases | SECTION 2.7.9 © 2004. Virtual Text / www.ergito.com

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induce chemical alterations that affect their charging, have shown that the basis for recognition is different for different tRNAs, and does not necessarily lie in some feature of primary or secondary structure alone. We know from the crystal structure that the acceptor stem and the anticodon stem make tight contacts with the synthetase, and mutations that alter recognition of a tRNA are found in these two regions. (The anticodon itself is not necessarily recognized as such; for example, the "suppressor" mutations discussed later in this chapter change a base in the anticodon, and therefore the codons to which a tRNA responds, without altering its charging with amino acids.) A group of isoaccepting tRNAs must be charged only by the single aminoacyl-tRNA synthetase specific for their amino acid. So isoaccepting tRNAs must share some common feature(s) enabling the enzyme to distinguish them from the other tRNAs. The entire complement of tRNAs is divided into 20 isoaccepting groups; each group is able to identify itself to its particular synthetase. tRNAs are identified by their synthetases by contacts that recognize a small number of bases, typically from 1-5. Three types of feature commonly are used: • Usually (but not always), at least one base of the anticodon is recognized. Sometimes all the positions of the anticodon are important. • Often one of the last three base pairs in the acceptor stem is recognized. An extreme case is represented by alanine tRNA, which is identified by a single unique base pair in the acceptor stem. • The so-called discriminator base, which lies between the acceptor stem and the CCA terminus, is always invariant among isoacceptor tRNAs. No one of these features constitutes a unique means of distinguishing 20 sets of tRNAs, or provides sufficient specificity, so it appears that recognition of tRNAs is idiosyncratic, each following its own rules. Several synthetases can specifically charge a "minihelix" consisting only of the acceptor and T ψ C arms (equivalent to one arm of the L-shaped molecule) with the correct amino acid. For certain tRNAs, specificity depends exclusively upon the acceptor stem. However, it is clear that there are significant variations between tRNAs, and in some cases the anticodon region is important. Mutations in the anticodon can affect recognition by the class II Phe-tRNA synthetase. Multiple features may be involved; minihelices from the tRNAVal and tRNAMet (where we know that the anticodon is important in vivo) can react specifically with their synthetases. So recognition depends on an interaction between a few points of contact in the tRNA, concentrated at the extremities, and a few amino acids constituting the active site in the protein. The relative importance of the roles played by the acceptor stem and anticodon is different for each tRNA·synthetase interaction (for review see 35).


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Reviews 35.

Schimmel, P. (1989). Parameters for the molecular recognition of tRNAs. Biochemistry 28, 2747-2759.



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2.7.10 Aminoacyl-tRNA synthetases fall into two groups Key Concepts

• Aminoacyl-tRNA synthetases are divided into the class I and class II groups by sequence and structural similarities.

In spite of their common function, synthetases are a rather diverse group of proteins. The individual subunits vary from 40-110 kD, and the enzymes may be monomeric, dimeric, or tetrameric. Homologies between them are rare. Of course, the active site that recognizes tRNA comprises a rather small part of the molecule. It is interesting to compare the active sites of different synthetases. Synthetases have been divided into two general groups, each containing 10 enzymes, on the basis of the structure of the domain that contains the active site. A general type of organization that applies to both groups is represented in Figure 7.14. The catalytic domain includes the binding sites for ATP and amino acid. It can be recognized as a large region that is interrupted by an insertion of the domain that binds the acceptor helix of the tRNA. This places the terminus of the tRNA in proximity to the catalytic site. A separate domain binds the anticodon region of tRNA. Those synthetases that are multimeric also possess an oligomerization domain (for review see 34).

Figure 7.14 An aminoacyl-tRNA synthetase contains three or four regions with different functions. (Only multimeric synthetases possess an oligomerization domain.)

Class I synthetases have an N-terminal catalytic domain that is identified by the presence of two short, partly conserved sequences of amino acids, sometimes called "signature sequences." The catalytic domain takes the form of a motif called a nucleotide-binding fold (which is also found in other classes of enzymes that bind nucleotides). The nucleotide fold consists of alternating parallel β-strands and Aminoacyl-tRNA synthetases fall into two groups | SECTION 2.7.10 © 2004. Virtual Text / www.ergito.com

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-helices; the signature sequence forms part of the ATP-binding site. The insertion that contacts the acceptor helix of tRNA differs widely between different class I enzymes. The C-terminal domains of the class I synthetases, which include the tRNA anticodon-binding domain and any oligomerization domain, also are quite different from one another. Class II enzymes share three rather general similarities of sequence in their catalytic domains. The active site contains a large antiparallel β-sheet surrounded by α-helices. Again, the acceptor helix-binding domain that interrupts the catalytic domain has a structure that depends on the individual enzyme. The anticodon-binding domain tends to be N-terminal. The location of any oligomerization domain is widely variable. The lack of any apparent relationship between the two groups of synthetases is a puzzle. Perhaps they evolved independently of one another. This makes it seem possible even that an early form of life could have existed with proteins that were made up of just the 10 amino acids coded by one type or the other. A general model for synthetase·tRNA binding suggests that the protein binds the tRNA along the "side" of the L-shaped molecule. The same general principle applies for all synthetase·tRNA binding: the tRNA is bound principally at its two extremities, and most of the tRNA sequence is not involved in recognition by a synthetase. However, the detailed nature of the interaction is different between class I and class II enzymes, as can be seen from the models of Figure 7.15, which are based on crystal structures. The two types of enzyme approach the tRNA from opposite sides, with the result that the tRNA-protein models look almost like mirror images of one another.

Figure 7.15 Crystal structures show that class I and class II aminoacyl-tRNA synthetases bind the opposite faces of their tRNA substrates. The tRNA is shown in red, and the protein in blue. Photographs kindly provided by Dino Moras.

A class I enzyme (Gln-tRNA synthetase) approaches the D-loop side of the tRNA. It recognizes the minor groove of the acceptor stem at one end of the binding site, and interacts with the anticodon loop at the other end. Figure 7.16 is a diagrammatic representation of the crystal structure of the tRNAGln·synthetase complex. A revealing feature of the structure is that contacts with the enzyme change the structure of the tRNA at two important points. These can be seen by comparing the Aminoacyl-tRNA synthetases fall into two groups | SECTION 2.7.10 © 2004. Virtual Text / www.ergito.com

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dotted and solid lines in the anticodon loop and acceptor stem:

Figure 7.16 A class I tRNA synthetase contacts tRNA at the minor groove of the acceptor stem and at the anticodon.

• Bases U35 and U36 in the anticodon loop are pulled farther out of the tRNA into the protein. • The end of the acceptor stem is seriously distorted, with the result that base pairing between U1 and A72 is disrupted. The single-stranded end of the stem pokes into a deep pocket in the synthetase protein, which also contains the binding site for ATP. This structure explains why changes in U35, G73, or the U1-A72 base pair affect the recognition of the tRNA by its synthetase. At all of these positions, hydrogen bonding occurs between the protein and tRNA (447). A class II enzyme (Asp-tRNA synthetase) approaches the tRNA from the other side, and recognizes the variable loop, and the major groove of the acceptor stem, as drawn in Figure 7.17. The acceptor stem remains in its regular helical conformation. ATP is probably bound near to the terminal adenine. At the other end of the binding site, there is a tight contact with the anticodon loop, which has a change in conformation that allows the anticodon to be in close contact with the protein (448).

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Figure 7.17 A class II aminoacyl-tRNA synthetase contacts tRNA at the major groove of the acceptor helix and at the anticodon loop.


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Reviews 34.

Schimmel, P. (1987). Aminoacyl-tRNA synthetases: general scheme of structure-function relationships on the polypeptides and recognition of tRNAs. Annu. Rev. Biochem. 56, 125-158.

References 447. Rould, M. A. et al. (1989). Structure of E. coli glutaminyl-tRNA synthetase complexed with tRNAGln and ATP at 28Å resolution. Science 246, 1135-1142. 448. Ruff, M. et al. (1991). Class II aminoacyl tRNA synthetases: crystal structure of yeast aspartyl-tRNA synthetase complexes with tRNAAsp. Science 252, 1682-1689.



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2.7.11 Synthetases use proofreading to improve accuracy Key Terms Proofreading refers to any mechanism for correcting errors in protein or nucleic acid synthesis that involves scrutiny of individual units after they have been added to the chain. Kinetic proofreading describes a proofreading mechanism that depends on incorrect events proceeding more slowly than correct events, so that incorrect events are reversed before a subunit is added to a polymeric chain. Chemical proofreading describes a proofreading mechanism in which the correction event occurs after addition of an incorrect subunit to a polymeric chain, by reversing the addition reaction. Key Concepts

• Specificity of recognition of both amino acid and tRNA is controlled by

aminoacyl-tRNA synthetases by proofreading reactions that reverse the catalytic reaction if the wrong component has been incorporated.

Aminoacyl-tRNA synthetases have a difficult job. Each synthetase must distinguish 1 out of 20 amino acids, and and must differentiate cognate tRNAs (typically 1-3) from the total set (perhaps 100 in all). Many amino acids are closely related to one another, and all amino acids are related to the metabolic intermediates in their particular synthetic pathway. It is especially difficult to distinguish between two amino acids that differ only in the length of the carbon backbone (that is, by one CH2 group). Intrinsic discrimination based on relative energies of binding two such amino acids would be only ~1/5. The synthetase enzymes improve this ratio ~1000 fold. Intrinsic discrimination between tRNAs is better, because the tRNA offers a larger surface with which to make more contacts, but it is still true that all tRNAs conform to the same general structure, and there may be a quite limited set of features that distinguish the cognate tRNAs from the noncognate tRNAs. We can imagine two general ways in which the enzyme might select its substrate: • The cycle of admittance, scrutiny, rejection/acceptance could represent a single binding step that precedes all other stages of whatever reaction is involved. This is tantamount to saying that the affinity of the binding site is sufficient to control the entry of substrate. In the case of synthetases, this would mean that only the correct amino acids and cognate tRNAs could form a stable attachment at the site. Synthetases use proofreading to improve accuracy | SECTION 2.7.11 © 2004. Virtual Text / www.ergito.com

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• Alternatively, the reaction proceeds through some of its stages, after which a decision is reached on whether the correct species is present. If it is not present, the reaction is reversed, or a bypass route is taken, and the wrong member is expelled. This sort of postbinding scrutiny is generally described as proofreading. In the example of synthetases, it would require that the charging reaction proceeds through certain stages even if the wrong tRNA or amino acid is present. Synthetases use proofreading mechanisms to control the recognition of both types of substrates. They improve significantly on the intrinsic differences among amino acids or among tRNAs, but, consistent with the intrinsic differences in each group, make more mistakes in selecting amino acids (error rates are 10–4 - 10–5) than in selecting tRNAs (error rates are ~10-6) (see Figure 6.8). Transfer RNA binds to synthetase by the two stage reaction depicted in Figure 7.18. Cognate tRNAs have a greater intrinsic affinity for the binding site, so they are bound more rapidly and dissociate more slowly. Following binding, the enzyme scrutinizes the tRNA that has been bound. If the correct tRNA is present, binding is stabilized by a conformational change in the enzyme. This allows aminoacylation to occur rapidly. If the wrong tRNA is present, the conformational change does not occur. As a result, the reaction proceeds much more slowly; this increases the chance that the tRNA will dissociate from the enzyme before it is charged. This type of control is called kinetic proofreading (450).

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Figure 7.18 Recognition of the correct tRNA by synthetase is controlled at two steps. First, the enzyme has a greater affinity for its cognate tRNA. Second, the aminoacylation of the incorrect tRNA is very slow.

Specificity for amino acids varies among the synthetases. Some are highly specific for initially binding a single amino acid, but others can also activate amino acids closely related to the proper substrate. Although the analog amino acid can sometimes be converted to the adenylate form, in none of these cases is an incorrectly activated amino acid actually used to form a stable aminoacyl-tRNA. The presence of the cognate tRNA usually is needed to trigger proofreading, even if the reaction occurs at the stage before formation of aminoacyl-adenylate. (An exception is provided by Met-tRNA synthetase, which can reject noncognate aminoacyl-adenylate complexes even in the absence of tRNA.) There are two stages at which proofreading of an incorrect aminoacyl-adenylate may Synthetases use proofreading to improve accuracy | SECTION 2.7.11 © 2004. Virtual Text / www.ergito.com

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occur during formation of aminoacyl-tRNA. Figure 7.19 shows that both use chemical proofreading, in which the catalytic reaction is reversed. The extent to which one pathway or the other predominates varies with the individual synthetase:

Figure 7.19 When a synthetase binds the incorrect amino acid, proofreading requires binding of the cognate tRNA. It may take place either by a conformation change that causes hydrolysis of the incorrect aminoacyl-adenylate, or by transfer of the amino acid to tRNA, followed by hydrolysis.

• The noncognate aminoacyl-adenylate may be hydrolyzed when the cognate tRNA binds. This mechanism is used predominantly by several synthetases, including those for methionine, isoleucine, and valine. (Usually, the reaction cannot be seen in vivo, but it can be followed for Met-tRNA synthetase when the incorrectly activated amino acid is homocysteine, which lacks the methyl group of methionine). Proofreading releases the amino acid in an altered form, as homocysteine thiolactone. In fact, homocysteine thiolactone is produced in E. coli as a by-product of the charging reaction of Met-tRNA synthetase. This shows that continuous proofreading is part of the process of charging a tRNA with its amino acid (451). • Some synthetases use chemical proofreading at a later stage. The wrong amino acid is actually transferred to tRNA, is then recognized as incorrect by its structure in the tRNA binding site, and so is hydrolyzed and released. The Synthetases use proofreading to improve accuracy | SECTION 2.7.11 © 2004. Virtual Text / www.ergito.com

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process requires a continual cycle of linkage and hydrolysis until the correct amino acid is transferred to the tRNA. A classic example in which discrimination between amino acids depends on the presence of tRNA is provided by the Ile-tRNA synthetase of E. coli. The enzyme can charge valine with AMP, but hydrolyzes the valyl-adenylate when tRNAIle is added. The overall error rate depends on the specificities of the individual steps, as summarized in Figure 7.20. The overall error rate of 1.5 × 10–5 is less than the measured rate at which valine is substituted for isoleucine (in rabbit globin), which is 2-5 × 10–4. So mischarging probably provides only a small fraction of the errors that actually occur in protein synthesis.

Figure 7.20 The accuracy of charging tRNAIle by its synthetase depends on error control at two stages.

Ile-tRNA synthetase uses size as a basis for discrimination among amino acids. Figure 7.21 shows that it has two active sites: the synthetic (or activation) site and the editing (or hydrolytic) site. The crystal structure of the enzyme shows that the synthetic site is too small to allow leucine (a close analog of isoleucine) to enter. All amino acids large than isoleucine are excluded from activation because they cannot enter the synthetic site. An amino acid that can enter the synthetic site is placed on tRNA. Then the enzyme tries to transfer it to the editing site. Isoleucine is safe from editing because it is too large to enter the editing site. However, valine can enter this site, and as a result an incorrect Val-tRNAIle is hydrolyzed. Essentially the enzyme provides a double molecular sieve, in which size of the amino acid is used to discriminate between closely related species (452; for review see 40).


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Figure 7.21 Ile-tRNA synthetase has two active sites. Amino acids larger than Ile cannot be activated because they do not fit in the synthetic site. Amino acids smaller than Ile are removed because they are able to enter the editing site.

One interesting feature of Ile-tRNA synthetase is that the synthetic and editing sites are a considerable distance apart, ~34Å. A crystal structure of the enzyme complexed with an edited analog of isoleucine shows that the amino acid is transported from the synthetic site to the editing site (2164). Figure 7.22 shows that this involves a change in the conformation of the tRNA. The amino acid acceptor stem of tRNAIle can exist in alternative conformations. It adopts an unusual hairpin in order to be aminoacylated by an amino acid in the synthetic site. Then it returns to the more common helical structure in order to move the amino acid to the editing site. The translocation between sites is the rate-limiting step in proofreading (2165). Ile-tRNA synthetase is a class I synthetase, but the double sieve mechanism is used also by class II synthetases (2166).


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Figure 7.22 An amino acid is transported from the synthetic site to the editing site of Ile-tRNA synthetase by a change in the conformation of the amino acceptor stem of tRNA.



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Reviews 40.

Jakubowski, H. and Goldman, E. (1992). Editing of errors in selection of amino acids for protein synthesis. Microbiol. Rev. 56, 412-429.

References 450. Hopfield, J. J. (1974). Kinetic proofreading: a new mechanism for reducing errors in biosynthetic processes requiring high specificity. Proc. Natl. Acad. Sci. USA 71, 4135-4139. 451. Jakubowski, H. (1990). Proofreading in vivo: editing of homocysteine by methionyl-tRNA synthetase in E. coli. Proc. Natl. Acad. Sci. USA 87, 4504-4508. 452. Nureki, O. et al. (1998). Enzyme structure with two catalytic sites for double sieve selection of substrate. Science 280, 578-581. 2164. Silvian, L. F., Wang, J., and Steitz, T. A. (1999). Insights into editing from an ile-tRNA synthetase structure with tRNAIle and mupirocin. Science 285, 1074-1077. 2165. Nomanbhoy, T. K., Hendrickson, T. L., and Schimmel, P. (1999). Transfer RNA-dependent translocation of misactivated amino acids to prevent errors in protein synthesis. Mol. Cell 4, 519-528. 2166. Dock-Bregeon, A., Sankaranarayanan, R., Romby, P., Caillet, J., Springer, M., Rees, B., Francklyn, C. S., Ehresmann, C., and Moras, D. (2000). Transfer RNA-mediated editing in threonyl-tRNA synthetase. The class II solution to the double discrimination problem. Cell 103, 877-884.



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2.7.12 Suppressor tRNAs have mutated anticodons that read new codons Key Terms A suppressor is a second mutation that compensates for or alters the effects of a primary mutation. A nonsense suppressor is a gene coding for a mutant tRNA able to respond to one or more of the termination codons and insert an amino acid at that site. A missense suppressor codes for a tRNA that has been mutated so as to recognize a different codon. By inserting a different amino acid at a mutant codon, the tRNA suppresses the effect of the original mutation. Key Concepts

• A suppressor tRNA typically has a mutation in the anticodon that changes the codons to which it responds.

• When the new anticodon corresponds to a termination codon, an amino acid is

inserted and the polypeptide chain is extended beyond the termination codon. This results in nonsense suppression at a site of nonsense mutation or in readthrough at a natural termination codon.

• Missense suppression occurs when the tRNA recognizes a different codon from usual, so that one amino acid is substituted for another.

Isolation of mutant tRNAs has been one of the most potent tools for analyzing the ability of a tRNA to respond to its codon(s) in mRNA, and for determining the effects that different parts of the tRNA molecule have on codon-anticodon recognition. Mutant tRNAs are isolated by virtue of their ability to overcome the effects of mutations in genes coding for proteins. In general genetic terminology, a mutation that is able to overcome the effects of another mutation is called a suppressor. In tRNA suppressor systems, the primary mutation changes a codon in an mRNA so that the protein product is no longer functional. The secondary, suppressor mutation changes the anticodon of a tRNA, so that it recognizes the mutant codon instead of (or as well as) its original target codon. The amino acid that is now inserted restores protein function. The suppressors are described as nonsense suppressors or missense suppressors, depending on the nature of the original mutation. In a wild-type cell, a nonsense mutation is recognized only by a release factor, terminating protein synthesis. The suppressor mutation creates an aminoacyl-tRNA that can recognize the termination codon; by inserting an amino acid, it allows protein synthesis to continue beyond the site of nonsense mutation. This new Suppressor tRNAs have mutated anticodons that read new codons | SECTION 2.7.12 © 2004. Virtual Text / www.ergito.com

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capacity of the translation system allows a full-length protein to be synthesized, as illustrated in Figure 7.23. If the amino acid inserted by suppression is different from the amino acid that was originally present at this site in the wild-type protein, the activity of the protein may be altered.

Figure 7.23 Nonsense mutations can be suppressed by a tRNA with a mutant anticodon, which inserts an amino acid at the mutant codon, producing a full length protein in which the original Leu residue has been replaced by Tyr.

Missense mutations change a codon representing one amino acid into a codon representing another amino acid, one that cannot function in the protein in place of the original residue. (Formally, any substitution of amino acids constitutes a missense mutation, but in practice it is detected only if it changes the activity of the protein.) The mutation can be suppressed by the insertion either of the original amino acid or of some other amino acid that is acceptable to the protein. Figure 7.24 demonstrates that missense suppression can be accomplished in the Suppressor tRNAs have mutated anticodons that read new codons | SECTION 2.7.12 © 2004. Virtual Text / www.ergito.com

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same way as nonsense suppression, by mutating the anticodon of a tRNA carrying an acceptable amino acid so that it responds to the mutant codon. So missense suppression involves a change in the meaning of the codon from one amino acid to another.

Figure 7.24 Missense suppression occurs when the anticodon of tRNA is mutated so that it responds to the wrong codon. The suppression is only partial because both the wild-type tRNA and the suppressor tRNA can respond to AGA. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.2.7.12

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2.7.13 There are nonsense suppressors for each termination codon Key Concepts

• Each type of nonsense codon is suppressed by tRNAs with mutant anticodons. • Some rare suppressor tRNAs have mutations in other parts of the molecule.

Nonsense suppressors fall into three classes, one for each type of termination codon. Figure 7.25 describes the properties of some of the best characterized suppressors.

Figure 7.25 Nonsense suppressor tRNAs are generated by mutations in the anticodon.

The easiest to characterize have been amber suppressors. In E. coli, at least 6 tRNAs have been mutated to recognize UAG codons. All of the amber suppressor tRNAs have the anticodon CUA ← , in each case derived from wild type by a single base change. The site of mutation can be any one of the three bases of the anticodon, as seen from supD, supE, and supF. Each suppressor tRNA recognizes only the UAG codon, instead of its former codon(s). The amino acids inserted are serine, glutamine, or tyrosine, the same as those carried by the corresponding wild-type tRNAs. Ochre suppressors also arise by mutations in the anticodon. The best known are supC and supG, which insert tyrosine or lysine in response to both ochre (UAA) and amber (UAG) codons. This conforms with the prediction of the wobble hypothesis that UAA cannot be recognized alone. A UGA suppressor has an unexpected property. It is derived from tRNATrp, but its only mutation is the substitution of A in place of G at position 24. This change replaces a G·U pair in the D stem with an A·U pair, increasing the stability of the helix. The sequence of the anticodon remains the same as the wild type, CCA ← . So the mutation in the D stem must in some way alter the conformation of the anticodon loop, allowing CCA ← to pair with UGA in an unusual wobble pairing of C with A. There are nonsense suppressors for each termination codon | SECTION 2.7.13 © 2004. Virtual Text / www.ergito.com

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The suppressor tRNA continues to recognize its usual codon, UGG. A related response is seen with a eukaryotic tRNA. Bovine liver contains a tRNASer with the anticodon mCCA ← . The wobble rules predict that this tRNA should respond to the tryptophan codon UGG; but in fact it responds to the termination codon UGA. So it is possible that UGA is suppressed naturally in this situation. The general importance of these observations lies in the demonstration that codon-anticodon recognition of either wild-type or mutant tRNA cannot be predicted entirely from the relevant triplet sequences, but is influenced by other features of the molecule. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.2.7.13

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2.7.14 Suppressors may compete with wild-type reading of the code Key Terms The context of a codon in mRNA refers to the fact that neighboring sequences may change the efficiency with which a codon is recognized by its aminoacyl-tRNA or is used to terminate protein synthesis. Readthrough at transcription or translation occurs when RNA polymerase or the ribosome, respectively, ignores a termination signal because of a mutation of the template or the behavior of an accessory factor. Key Concepts

• Suppressor tRNAs compete with wild-type tRNAs that have the same anticodon to read the corresponding codon(s).

• Efficient suppression is deleterious because it results in readthrough past normal termination codons.

• The UGA codon is leaky and is misread by Trp-tRNA at 1-3% frequency.

There is an interesting difference between the usual recognition of a codon by its proper aminoacyl-tRNA and the situation in which mutation allows a suppressor tRNA to recognize a new codon. In the wild-type cell, only one meaning can be attributed to a given codon, which represents either a particular amino acid or a signal for termination. But in a cell carrying a suppressor mutation, the mutant codon has the alternatives of being recognized by the suppressor tRNA or of being read with its usual meaning. A nonsense suppressor tRNA must compete with the release factors that recognize the termination codon(s). A missense suppressor tRNA must compete with the tRNAs that respond properly to its new codon. The extent of competition influences the efficiency of suppression; so the effectiveness of a particular suppressor depends not only on the affinity between its anticodon and the target codon, but also on its concentration in the cell, and on the parameters governing the competing termination or insertion reactions. The efficiency with which any particular codon is read is influenced by its location. So the extent of nonsense suppression by a given tRNA can vary quite widely, depending on the context of the codon. We do not understand the effect that neighboring bases in mRNA have on codon-anticodon recognition, but the context can change the frequency with which a codon is recognized by a particular tRNA by more than an order of magnitude. The base on the 3 ′ side of a codon appears to have a particularly strong effect. A nonsense suppressor is isolated by its ability to respond to a mutant nonsense Suppressors may compete with wild-type reading of the code | SECTION 2.7.14 © 2004. Virtual Text / www.ergito.com

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codon. But the same triplet sequence constitutes one of the normal termination signals of the cell! The mutant tRNA that suppresses the nonsense mutation must in principle be able to suppress natural termination at the end of any gene that uses this codon. Figure 7.26 shows that this readthrough results in the synthesis of a longer protein, with additional C-terminal material. The extended protein will end at the next termination triplet sequence found in the phase of the reading frame. Any extensive suppression of termination is likely to be deleterious to the cell by producing extended proteins whose functions are thereby altered.

Figure 7.26 Nonsense suppressors also read through natural termination codons, synthesizing proteins that are longer than wild-type.

Amber suppressors tend to be relatively efficient, usually in the range of 10-50%, depending on the system. This efficiency is possible because amber codons are used relatively infrequently to terminate protein synthesis in E. coli. Ochre suppressors are difficult to isolate. They are always much less efficient, usually with activities below 10%. All ochre suppressors grow rather poorly, which indicates that suppression of both UAA and UAG is damaging to E. coli, probably because the ochre codon is used most frequently as a natural termination signal. UGA is the least efficient of the termination codons in its natural function; it is misread by Trp-tRNA as frequently as 1-3% in wild-type situations. In spite of this deficiency, however, it is used more commonly than the amber triplet to terminate bacterial genes. One gene's missense suppressor is likely to be another gene's mutator. A suppressor corrects a mutation by substituting one amino acid for another at the mutant site. But in other locations, the same substitution will replace the wild-type amino acid with a Suppressors may compete with wild-type reading of the code | SECTION 2.7.14 © 2004. Virtual Text / www.ergito.com

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new amino acid. The change may inhibit normal protein function. This poses a dilemma for the cell: it must suppress what is a mutant codon at one location, while failing to change too extensively its normal meaning at other locations. The absence of any strong missense suppressors is therefore explained by the damaging effects that would be caused by a general and efficient substitution of amino acids. A mutation that creates a suppressor tRNA can have two consequences. First, it allows the tRNA to recognize a new codon. Second, sometimes it prevents the tRNA from recognizing the codons to which it previously responded. It is significant that all the high-efficiency amber suppressors are derived by mutation of one copy of a redundant tRNA set. In these cases, the cell has several tRNAs able to respond to the codon originally recognized by the wild-type tRNA. So the mutation does not abolish recognition of the old codons, which continue to be served adequately by the tRNAs of the set. In the unusual situation in which there is only a single tRNA that responds to a particular codon, any mutation that prevents the response is lethal (for review see 31; 36; 37; 38). Suppression is most often considered in the context of a mutation that changes the reading of a codon. However, there are some situations in which a stop codon is read as an amino acid at a low frequency in the wild-type situation. The first example to be discovered was the coat protein gene of the RNA phage Q β. The formation of infective Q β particles requires that the stop codon at the end of this gene is suppressed at a low frequency to generate a small proportion of coat proteins with a C-terminal extension. In effect, this stop codon is leaky. The reason is that Trp-tRNA recognizes the codon at a low frequency (3062; 3063). Readthrough past stop codons occurs also in eukaryotes, where it is employed most often by RNA viruses. This may involve the suppression of UAG/UAA by Tyr-tRNA, Gln-tRNA, or Leu-tRNA, or the suppression of UGA by Trp-tRNA or Arg-tRNA. The extent of partial suppression is dictated by the context surrounding the codon (for review see 3061). Last updated on 11-2-2002

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Reviews 31.

Murgola, E. J. (1985). tRNA, suppression, and the code. Annu. Rev. Genet. 19, 57-80.

36.

Eggertsson, G. and Soll, D. (1988). Transfer RNA-mediated suppression of termination codons in E. coli. Microbiol. Rev. 52, 354-374.

37.

Normanly, J. and Abelson, J. (1989). Transfer RNA identity. Annu. Rev. Biochem. 58, 1029-1049.

38.

Atkins, J. F (1991). Towards a genetic dissection of the basis of triplet decoding, and its natural subversion: programmed reading frameshifts and hops. Annu. Rev. Genet. 25, 201-228.

3061. Beier, H. and Grimm, M. (2001). Misreading of termination codons in eukaryotes by natural nonsense suppressor tRNAs. Nucleic Acids Res. 29, 4767-4782.

References 3062. Hirsh, D. (1971). Tryptophan transfer RNA as the UGA suppressor. J. Mol. Biol. 58, 439-458. 3063. Weiner, A. M. and Weber, K. (1973). A single UGA codon functions as a natural termination signal in the coliphage q beta coat protein cistron. J. Mol. Biol. 80, 837-855.


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2.7.15 The ribosome influences the accuracy of translation Key Concepts

• The structure of the 16S rRNA at the P and A sites of the ribosome influences the accuracy of translation.

The lack of detectable variation when the sequence of a protein is analyzed demonstrates that protein synthesis must be extremely accurate. Very few mistakes are apparent in the form of substitutions of one amino acid for another. There are two general stages in protein synthesis at which errors might be made (see Figure 6.8 in Molecular Biology 2.6.3 Special mechanisms control the accuracy of protein synthesis): • Charging a tRNA only with its correct amino acid clearly is critical. This is a function of the aminoacyl-tRNA synthetase. Probably the error rate varies with the particular enzyme, but generally mistakes occur in 15 human diseases have been linked to disorders in peroxisome function. All of the components of the peroxisome are imported from the cytosol. Proteins that are required for peroxisome formation are called peroxins. 23 genes coding for peroxins have been identified, and human peroxisomal diseases have been mapped to 12 complementation groups, most identified with specific genes. Peroxisomes appear to be absent from cells that have null mutations in some of these genes. In some of these cases, introduction of a wild-type gene leads to the reappearance of peroxisomes (2317; 2318). It has generally been assumed that, like other membrane-bounded organelles, peroxisomes can arise only by duplication of pre-existing peroxisomes. But these results raised the question of whether it might be possible to assemble them de novo from their components. In at least some cases, however, the absence of peroxins leaves the cells with peroxisomal ghosts – empty membrane bodies. Even when they cannot be easily seen, it is hard to exclude the possibility that there is some remnant that serves to regenerate the peroxisomes (for review see 2305). Transport of proteins to peroxisomes occurs post-translationally. Proteins that are imported into the matrix have either of two short sequences, called PTS1 and PTS2. Peroxisomes employ another type of translocation system | SECTION 2.8.20 © 2004. Virtual Text / www.ergito.com

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The PTS1 signal is a tri- or tetrapeptide at the C-terminus. It was originally characterized as the sequence SKL (Ser-Lys-Leu) (2319), but now a large variety of sequences have been shown to act as a PTS1 signal (2320). The addition of a suitable sequence to the C-terminus of cytosolic proteins is sufficient to ensure their import into the organelle. The PTS2 signal is a sequence of 9 amino acids, again with much diversity, and this can be located near the N-terminus or internally (for review see 1962). It is possible there may be a third type of sequence called PTS3. Several peroxisomal proteins are necessary for the import of proteins from the cytosol. The peroxisomal receptors that bind the two types of signals are called Pex5p and Pex7p, respectively. The other proteins are part of membrane-associated complexes concerned with the translocation reaction. Transport into the peroxisome has unusual features that mark important differences from the system used for transport into other organelles. Proteins can be imported into the peroxisome in their mature, fully-folded state (1961). This contrasts with the requirement to unfold a protein for passage into the ER or mitochondrion, where it passes through a channel in the membrane into the organelle in something akin to an unfolded thread of amino acids. It is not clear how the structure of a preexisting channel could expand to permit this. One possibility is to resurrect an old idea and to suppose that the channel assembles around the substrate protein when it associates with the membrane. The Pex5p and Pex7p receptors are not integral membrane proteins, but are largely cytosolic, with only a small proportion associated with peroxisomes (2321; 2322). They behave in the same way, cycling between the peroxisome and the cytosol. Figure 8.47 shows that the receptor binds a substrate protein in the cytosol, takes it to the peroxisome, moves with it through the membrane into the interior, and then returns to the cytosol to undertake another cycle. This shuttling behavior resembles the carrier system for import into the nucleus (see Molecular Biology 2.8.28 Transport receptors carry cargo proteins through the pore).

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Figure 8.47 The Pex5p receptor binds a substrate protein in the cytosol, carries it across the membrane into the peroxisome, and then returns to the cytosol.

The import pathways converge at the perixosomal membrane, where Pex5p and Pex7p both interact with the same membrane protein complex, consisting of Pex14p and Pex13p. The receptors dock with this complex, and then several other peroxins are involved with the process of transport into the lumen. The details of the transport process are not yet clear. Proteins that are incorporated into the peroxisomal membrane have a sequence called the mPTS, but little is known about the process of integration. Pex3p may be a key protein, because in its absence other proteins are not found in peroxisomal membranes. Pex3p has its own mPTS, which raises the question of how it enters the membrane. Perhaps it interacts with Pex3p that is already in the membrane (for review see 2305). This bears on the question of whether peroxisomes can ever assemble de novo. Last updated on 1-22-2002


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Reviews 1962. Subramani, S., Koller, A., and Snyder, W. B. (2000). Import of peroxisomal matrix and membrane proteins. Annu. Rev. Biochem. 69, 399-418. 2305. Purdue, P. E. and Lazarow, P. B. (2001). Peroxisome biogenesis. Annu. Rev. Cell Dev. Biol. 17, 701-752.

References 1961. Walton, P. A., Hill, P. E., and Hill, S. (1995). Import of stably folded proteins into peroxisomes. Mol. Biol. Cell 6, 675-683. 2316. Goldfischer, S., Moore, C. L., Johnson, A. B., Spiro, A. J., Valsamis, M. P., Wisniewski, H. K., Ritch, R. H., Norton, W. T., Rapin, I., and Gartner, L. M. (1973). Peroxisomal and mitochondrial defects in the cerebro-hepato-renal syndrome. Science 182, 62-64. 2317. Matsuzono, Y., Kinoshita, N., Tamura, S., Shimozawa, N., Hamasaki, M., Ghaedi, K., Wanders, R. J., Suzuki, Y., Kondo, N., and Fujiki, Y. (1999). Human PEX19: cDNA cloning by functional complementation, mutation analysis in a patient with Zellweger syndrome, and potential role in peroxisomal membrane assembly. Proc. Natl. Acad. Sci. USA 96, 2116-2121. 2318. South, S. T. and Gould, S. J. (1999). Peroxisome synthesis in the absence of preexisting peroxisomes. J. Cell Biol. 144, 255-266. 2319. Gould, S. J., Keller, G. A., Hosken, N., Wilkinson, J., and Subramani, S. (1989). A conserved tripeptide sorts proteins to peroxisomes. J. Cell Biol. 108, 1657-1664. 2320. Elgersma, Y., Vos, A., van den Berg, M., van Roermund, C. W., van der Sluijs, P., Distel, B., and Tabak, H. F. (1996). Analysis of the carboxyl-terminal peroxisomal targeting signal 1 in a homologous context in S. cerevisiae. J. Biol. Chem. 271, 26375-26382. 2321. Dodt, G. and Gould, S. J. (1996). Multiple PEX genes are required for proper subcellular distribution and stability of Pex5p, the PTS1 receptor: evidence that PTS1 protein import is mediated by a cycling receptor. J. Cell Biol. 135, 1763-1774. 2322. Elgersma, Y., Elgersma-Hooisma, M., Wenzel, T., McCaffery, J. M., Farquhar, M. G., and Subramani, S. (1998). A mobile PTS2 receptor for peroxisomal protein import in Pichia pastoris. J. Cell Biol. 140, 807-820.



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PROTEIN LOCALIZATION

2.8.21 Bacteria use both co-translational and post-translational translocation Key Terms The periplasm (or periplasmic space) is the region between the inner and outer membranes in the bacterial envelope. Signal peptidase is an enzyme within the membrane of the ER that specifically removes the signal sequences from proteins as they are translocated. Analogous activities are present in bacteria, archaebacteria, and in each organelle in a eukaryotic cell into which proteins are targeted and translocated by means of removable targeting sequences. Signal peptidase is one component of a larger protein complex. Key Concepts

• Bacterial proteins that are exported to or through membranes use both post-translational and co-translational mechanisms.

The bacterial envelope consists of two membrane layers. The space between them is called the periplasm. Proteins are exported from the cytoplasm to reside in the envelope or to be secreted from the cell. The mechanisms of secretion from bacteria are similar to those characterized for eukaryotic cells, and we can recognize some related components. Figure 8.48 shows that proteins that are exported from the cytoplasm have one of four fates:

Figure 8.48 Bacterial proteins may be exported either post-translationally or co-translationally, and may be located within either membrane or the periplasmic space, or may be secreted.

Bacteria use both co-translational and post-translational translocation | SECTION 2.8.21 © 2004. Virtual Text / www.ergito.com

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• to be inserted into the inner membrane • to be translocated through the inner membrane to rest in the periplasm • to be inserted into the outer membrane • to be translocated through the outer membrane into the medium. Different protein complexes in the inner membrane are responsible for transport of proteins depending on whether their fate is to pass through or stay within the inner membrane. This resembles the situation in mitochondria, where different complexes in each of the inner and outer membranes handle different subsets of protein substrates depending on their destinations (see Molecular Biology 2.8.17 Post-translational membrane insertion depends on leader sequences) A difference from import into organelles is that transfer in E. coli may be either coor post-translational. Some proteins are secreted both co-translationally and post-translationally, and the relative kinetics of translation versus secretion through the membrane could determine the balance. Exported bacterial proteins have N-terminal leader sequences, with a hydrophilic N-terminus and an adjacent hydrophobic core. The leader is cleaved by a signal peptidase that recognizes precursor forms of several exported proteins. The signal peptidase is an integral membrane protein, located in the inner membrane. Mutations in N-terminal leaders prevent secretion; they are suppressed by mutations in other genes, which are thus defined as components of the protein export apparatus. Several genes given the general description sec are implicated in coding for components of the secretory apparatus by the occurrence of mutations that block secretion of many or all exported proteins. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.2.8.21

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2.8.22 The Sec system transport proteins into and through the inner membrane Key Concepts

• The bacterial SecYEG translocon in the inner membrane is related to the eukaryotic Sec61 translocon.

• Various chaperones are involved in directing secreted proteins to the translocon.

There are several systems for transport through the inner membrane. The best characterized is the Sec system, whose components are shown in Figure 8.49. The translocon that is embedded in the membrane consists of three subunits that are related to the components of mammalian/yeast Sec61 (for review see 45; 47). Each of the subunits is an integral transmembrane protein. (SecY has 10 transmembrane segments and SecE has 3 transmembrane segments.) The functional translocon is a trimer with one copy of each subunit (1066). The major pathway for directing proteins to the translocon consists of SecB and SecA. SecB is a chaperone that binds to the nascent protein to control its folding. It transfers the protein to SecA, which in turn transfers it to the translocon.

Figure 8.49 The Sec system has the SecYEG translocon embedded in the membrane, the SecA associated protein that pushes proteins through the channel, the SecB chaperone that transfers nascent proteins to SecA, and the signal peptidase that cleaves the N-terminal signal from the translocated protein.

Figure 8.50 shows that there are two predominant ways of directing proteins to the Sec channel: The Sec system transport proteins into and through the inner membrane | SECTION 2.8.22 © 2004. Virtual Text / www.ergito.com

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Figure 8.50 SecB/SecA transfer proteins to the translocon in order for them to pass through the membrane. 4.4S RNA transfers proteins that enter the membrane.

• the SecB chaperone; • and the 4.5S RNA-based SRP (2357). Several chaperones can increase the efficiency of bacterial protein export by preventing premature folding; they include "trigger factor" (characterized as a chaperone that assists export), GroEL (see earlier), and SecB (identified as the product of one of the sec mutants). Although SecB is the least abundant of these proteins, it has the major role in promoting export. It has two functions. First, it behaves as a chaperone and binds to a nascent protein to retard folding. It cannot reverse the change in structure of a folded protein, so it does not function as an unfolding factor. Its role is therefore to inhibit improper folding of the newly synthesized protein. Second, it has an affinity for the protein SecA. This allows it to target a precursor protein to the membrane (456; 457). The SecB-SecYEG pathway is used for translocation of proteins that are secreted into the periplasm and is summarized inFigure 8.51.

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Figure 8.51 SecB transfers a nascent protein to SecA, which inserts the protein into the channel. Translocation requires hydrolysis of ATP and a protonmotive force. SecA undergoes cycles of association and dissociation with the channel and provides the motive force to push the protein through.

SecA is a large peripheral membrane protein that has alternative ways to associate with the membrane. As a peripheral membrane protein, it associates with the membrane by virtue of its affinity for acidic lipids and for the SecY component of the translocon., which are part of a multisubunit complex that provides the translocase function. However, in the presence of other proteins (SecD and SecF), SecA can be found as a membrane-spanning protein. It probably provides the motor that pushes the substrate protein through the SecYEG translocon. SecA recognizes both SecB and the precursor protein that it chaperones; probably features of the mature protein sequence as well as its leader are required for recognition. SecA has an ATPase activity that depends upon binding to lipids, SecY, and a precursor protein. The ATPase functions in a cyclical manner during translocation. After SecA binds a precursor protein, it binds ATP, and ~20 amino acids are translocated through the membrane. Hydrolysis of ATP is required to The Sec system transport proteins into and through the inner membrane | SECTION 2.8.22 © 2004. Virtual Text / www.ergito.com

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release the precursor from SecA. Then the cycle may be repeated. Precursor protein is bound again to provide the spur to bind more ATP, translocate another segment of protein, and release the precursor. SecA may alternate between the peripheral and integral membrane forms during translocation; with each cycle, a 30 kD domain of SecA may insert into the membrane and then retract (471). Another process can also undertake translocation. When a precursor is released by SecA, it can be driven through the membrane by a protonmotive force (that is, an electrical potential across the membrane). This process cannot initiate transfer through the membrane, but can continue the process initiated by a cycle of SecA ATPase action. So after or between cycles of the SecA-ATP driven reaction, the protonmotive force can drive translocation of the precursor. The E. coli ribonucleoprotein complex of 4.5S RNA with Ffh and FtsY proteins is a counterpart to the eukaryotic SRP (see Molecular Biology 2.8.10 The SRP interacts with the SRP receptor). It probably plays the role of keeping the nascent protein in an appropriate conformation until it interacts with other components of the secretory apparatus. It is needed for the secretion of some, but not all, proteins. As we see in Figure 8.50, its substrates are integral membrane proteins (1065). The basis for differential selection of substrates is that the E. coli SRP recognizes an anchor sequence in the protein (anchor sequences by definition are present only in integral membrane proteins). Chloroplasts have counterparts to the Ffh and FtsY proteins, but do not require an RNA component. Last updated on 2-22-2002


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Reviews 45.

Oliver, D. (1985). Protein secretion in E. coli. Annu. Rev. Immunol. 39, 615-648.

47.

Lee, C. and Beckwith, J. (1986). Cotranslational and posttranslational protein translocation in prokaryotic systems. Annu. Rev. Cell Biol. 2, 315-336.

References 456. Collier, D. N. et al. (1988). The antifolding activity of SecB promotes the export of the E. coli maltose-binding protein. Cell 53, 273-283. 457. Crooke, E. et al. (1988). ProOmpA is stabilized for membrane translocation by either purified E. coli trigger factor or canine signal recognition particle. Cell 54, 1003-1011. 471. Brundage, L. et al. (1990). The purified E. coli integral membrane protein SecY/E is sufficient for reconstitution of SecA-dependent precursor protein translocation. Cell 62, 649-657. 1065. Beck, K., Wu, L. F., Brunner, J., and Muller, M. (2000). Discrimination between SRP- and SecA/SecB-dependent substrates involves selective recognition of nascent chains by SRP and trigger factor. EMBO J. 19, 134-143. 1066. Yahr, T. L. and Wickner, W. T. (2000). Evaluating the oligomeric state of SecYEG in preprotein translocase. EMBO J. 19, 4393-4401. 2357. Valent, Q. A., Scotti, P. A., High, S., von Heijne, G., Lentzen, G., Wintermeyer, W., Oudega, B., and Luirink, J. (1998). The E. coli SRP and SecB targeting pathways converge at the translocon. EMBO J. 17, 2504-2512.



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2.8.23 Sec-independent translocation systems in E. coli Key Concepts

• E. coli and organelles have related systems for protein translocation. • One system allows certain proteins to insert into membranes without a translocation apparatus.

• YidC is homologous to a mitochondrial system for transferring proteins into the inner membrane.

• The tat system transfers proteins with a twin arginine motif into the periplasmic space.

The most striking alternative system for protein translocation in E. coli is revealed by the coat protein of phage M13. Figure 8.52 shows that this does not appear to require any translocation apparatus! It can insert post-translationally into protein-free liposomes (2358). Targeting the protein to the membrane requires specific sequences (comprising basic residues) in the N- and C-terminal regions of the protein. They may interact with negatively charged heads of phospholipids. Then the protein enters the membrane by using hydrophobic groups in its N-terminal leader sequence and an internal anchor sequence. Hydrophobicity is the main driving force for translocation, but it can be assisted by a protonmotive force that is generated between the positively charged periplasmic side of the membrane and an acidic region in the protein. This drives the protein through the membrane, and leader peptidase can then cleave the N-terminal sequence. The generality of this mechanism in bacteria is unclear; it may apply only to the special case of bacteriophage coat proteins (for review see 2353). Some chloroplast proteins may insert into the thylakoid membrane by a similar pathway.

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Figure 8.52 M13 coat protein inserts into the inner membrane by making an initial electrostatic contact, followed by insertion of hydrophobic sequences. Translocation is driven by hydrophobic interactions and a protonmotive force until the anchor sequence enters the membrane.

Mutations in the gene yidC block insertion of proteins into the inner membrane (1065; 1088). YidC is homologous to the protein Oxa1p that is required when proteins are inserted into the inner mitochondrial membrane from the matrix. It can function either independently of SecYEG or in conjunction with it. The insertion of some of the YidC-dependent proteins requires SecYEG, suggesting that YidC acts in conjunction with the translocon to divert the substrate into membrane insertion as opposed to secretion (2360). Other proteins whose insertion depends on YidC do not require SecYEG: it seems likely that some other (unidentified) functions are required instead of the translocon. The Tat system is named for its ability to transport proteins bearing a twin arginine Sec-independent translocation systems in E. coli | SECTION 2.8.23 © 2004. Virtual Text / www.ergito.com

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targeting motif. It is responsible for translocation of proteins that have tightly bound cofactors. This may mean that they have limitations on their ability to unfold for passage through the membrane. This would be contrary to the principle of most translocation systems, where the protein passes through the membrane in an unfolded state, and then must be folded into its mature conformation after passage (for review see 1064). This system is related to a system in the chloroplast thylakoid lumen called Hcf106 (for review see 1064). Both of these systems transport proteins into the periplasm. Last updated on 2-22-2002


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Reviews 1064. Dalbey, R. E. and Robinson, C. (1999). Protein translocation into and across the bacterial plasma membrane and the plant thylakoid membrane. Trends Biochem. Sci. 24, 17-22. 2353. Dalbey, R. E. and Kuhn, A. (2000). Evolutionarily related insertion pathways of bacterial, mitochondrial, and thylakoid membrane proteins. Annu. Rev. Cell Dev. Biol. 16, 51-87.

References 1065. Beck, K., Wu, L. F., Brunner, J., and Muller, M. (2000). Discrimination between SRP- and SecA/SecB-dependent substrates involves selective recognition of nascent chains by SRP and trigger factor. EMBO J. 19, 134-143. 1088. Samuelson, J. C., Chen, M., Jiang, F., Moller, I., Wiedmann, M., Kuhn, A., Phillips, G. J., and Dalbey, R. E. (2000). YidC mediates membrane protein insertion in bacteria. Nature 406, 637-641. 2358. Soekarjo, M., Eisenhawer, M., Kuhn, A., and Vogel, H. (1996). Thermodynamics of the membrane insertion process of the M13 procoat protein, a lipid bilayer traversing protein containing a leader sequence. Biochemistry 35, 1232-1241. 2360. Scotti, P. A., Urbanus, M. L., Brunner, J., de Gier, J. W., von Heijne, G., van der Does, C., Driessen, A. J., Oudega, B., and Luirink, J. (2000). YidC, theE. coli homologue of mitochondrial Oxa1p, is a component of the Sec translocase. EMBO J. 19, 542-549.



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2.8.24 Pores are used for nuclear import and export Key Terms The nuclear envelope is a layer of two concentric membranes (inner and outer nuclear membranes) that surrounds the nucleus and its underlying intermediate filament lattice, the nuclear lamina. The nuclear envelope is penetrated by nuclear pores. The outer membrane is continuous with the membrane of the rough endoplasmic reticulum. A nuclear pore complex (NPC) is a very large, proteinaceous structure that extends through the nuclear envelope, providing a channel for bidirectional transport of molecules and macromolecules between the nucleus and the cytosol. Key Concepts

• The same nuclear pores are used for importing proteins into the nucleus and for exporting proteins and RNA from the nucleus.

The nucleus is segregated from the cytoplasm by a layer of two membranes that constitute the nuclear envelope. The inner membrane contacts the nuclear lamina, providing in effect a surface layer for the nucleus. The outer membrane is continuous with the endoplasmic reticulum in the cytosol. The space between the two membranes is continuous with the lumen of the endoplasmic reticulum. The two membranes come into contact at openings called nuclear pore complexes. At the center of each complex is a pore that provides a water-soluble channel between nucleus and cytoplasm. This means that the nucleus and cytosol have the same ionic milieu. There are ~3000 pore complexes on the nuclear envelope of an animal cell. Transport between nucleus and cytoplasm proceeds in both directions. Since all proteins are synthesized in the cytosol, any proteins required in the nucleus must be transported there. Since all RNA is synthesized in the nucleus, the entire cytoplasmic complement of RNA (mRNA, rRNA, tRNA, and other small RNAs) must be derived by export from the nucleus. The nuclear pores are used for both import and export of material. Figure 8.53 summarizes the frequency with which the pores are used for some of the more prominent substrates.

Figure 8.53 Nuclear pores are used for import and export. Pores are used for nuclear import and export | SECTION 2.8.24 © 2004. Virtual Text / www.ergito.com

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We can form an impression of the magnitude of import by considering the histones, the major protein components of chromatin. In a dividing cell, enough histones must be imported into the nucleus during the period of DNA synthesis to associate with a diploid complement of chromosomes. Since histones form about half the protein mass of chromatin, we may conclude that overall about 200 chromosomal protein molecules must be imported through each pore per minute. Uncertainties about the processing and stability of mRNA make it more difficult to calculate the number of mRNA molecules exported, but to account for the ~250,000 molecules of mRNA per cell probably requires ~1 event per pore per minute. The major RNA synthetic activity of the nucleus is of course the production of rRNA, which is exported in the form of assembled ribosomal subunits. Just to double the number of ribosomes during one cell cycle would require the export of ~5 ribosomal subunits (60S and 40S) through each pore per minute. For ribosomal proteins to assemble with the rRNA, they must first be imported into the nucleus. So ribosomal proteins must shuttle into the nucleus as free proteins and out again as assembled ribosomal subunits. Given ~80 proteins per ribosome, their import must be comparable in magnitude to that of the chromosomal proteins. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.2.8.24

Pores are used for nuclear import and export | SECTION 2.8.24 © 2004. Virtual Text / www.ergito.com

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2.8.25 Nuclear pores are large symmetrical structures Key Concepts

• The nuclear pore is an annular structure with 8-fold symmetry.

How does a nuclear pore accommodate the transit of material of varied sizes and characteristics in either direction? Nuclear pore complexes have a uniform appearance when examined by microscopy. The pores can be released from the nuclear envelope by detergent, and Figure 8.54 shows that they appear as annular structures, consisting of rosettes made of 8 spokes. Figure 8.55 shows a model for the pore based on three-dimensional reconstruction of electron microscopic images. It consists of an upper ring and a lower ring, connected by a lattice of 8 structures.

Figure 8.54 Nuclear pores appear as annular structures by electron microscopy. The circle around one pore has a diameter of 120 nm. Photograph kindly provided by Ronald Milligan.

Nuclear pores are large symmetrical structures | SECTION 2.8.25 © 2004. Virtual Text / www.ergito.com

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Figure 8.55 A model for the nuclear pore shows 8-fold symmetry. Two rings form the upper and lower surfaces (shown in yellow); they are connected by the spokes (shown in green on the inside and blue on the outside). Photograph kindly provided by Ronald Milligan.

The basis for the 8-fold symmetry is explained in terms of individual components in the schematic view from above shown in Figure 8.56. This includes the central structure of Figure 8.55, and extends it with an internal transporter and surrounding radial arms. The outside of the pore complex as such consists of a ring of diameter ~120 nm. The ring itself consists of 8 subunits. The 8 radial arms outside the ring may be responsible for anchoring the pore complex in the nuclear envelope; they penetrate the membrane. The 8 interior spokes project from the ring, closing the opening to a diameter of ~48 nm. Within this region is the transporter, which contains a pore that approximates a cylinder –d mutation.


Multimeric proteins have special genetic properties | SECTION 3.10.9 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.10 Repressor protein binds to the operator Key Terms A palindrome is a DNA sequence that reads the same on each strand of DNA when the strand is read in the 5 ′ to 3 ′ direction. It consists of adjacent inverted repeats. Key Concepts

• Repressor protein binds to the double-stranded DNA sequence of the operator. • The operator is a palindromic sequence of 26 bp. • Each inverted repeat of the operator binds to the DNA-binding site of one repressor subunit.

The repressor was isolated originally by purifying the component able to bind the gratuitous inducer IPTG (509; see Great Experiments 2.1 Isolation of repressor). (Because the amount of repressor in the cell is so small, in order to obtain enough material it was necessary to use a promoter up mutation to increase lacI transcription, and to place this lacI locus on a DNA molecule present in many copies per cell. This results in an overall overproduction of 100-1000-fold.) The repressor binds to double-stranded DNA containing the sequence of the wild-type lac operator. The repressor does not bind DNA from an Oc mutant. The addition of IPTG releases the repressor from operator DNA in vitro. The in vitro reaction between repressor protein and operator DNA therefore displays the characteristics of control inferred in vivo; so it can be used to establish the basis for repression (510). How does the repressor recognize the specific sequence of operator DNA? The operator has a feature common to many recognition sites for bacterial regulator proteins: it is a palindrome. The inverted repeats are highlighted in Figure 10.12. Each repeat can be regarded as a half-site of the operator.

Repressor protein binds to the operator | SECTION 3.10.10 © 2004. Virtual Text / www.ergito.com

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Figure 10.12 The lac operator has a symmetrical sequence. The sequence is numbered relative to the startpoint for transcription at +1. The pink arrows to left and right identify the two dyad repeats. The green blocks indicate the positions of identity.

We can use the same approaches to define the points that the repressor contacts in the operator that we used for analyzing the polymerase-promoter interaction (see Molecular Biology 3.9.14 RNA polymerase binds to one face of DNA). Deletions of material on either side define the end points of the region; constitutive point mutations identify individual base pairs that must be crucial. Experiments in which DNA bound to repressor is compared with unbound DNA for its susceptibility to methylation or UV crosslinking identify bases that are either protected or more susceptible when associated with the protein. Figure 10.13 shows that the region of DNA protected from nucleases by bound repressor lies within the region of symmetry, comprising the 26 bp region from -5 to +21. The area identified by constitutive mutations is even smaller. Within a central region extending over the 13 bp from +5 to +17, there are eight sites at which single base-pair substitutions cause constitutivity. This emphasizes the same point made by the promoter mutations summarized earlier in Figure 9.29. A small number of essential specific contacts within a larger region can be responsible for sequence-specific association of DNA with protein.


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Figure 10.13 Bases that contact the repressor can be identified by chemical crosslinking or by experiments to see whether modification prevents binding. They identify positions on both strands of DNA extending from +1 to +23. Constitutive mutations occur at 8 positions in the operator between +5 and +17.

The symmetry of the DNA sequence reflects the symmetry in the protein. Each of the identical subunits in a repressor tetramer has a DNA-binding site. Two of these sites contact the operator in such a way that each inverted repeat of the operator makes the same pattern of contacts with a repressor monomer. This is shown by symmetry in the contacts that repressor makes with the operator (the pattern between +1 and +6 is identical with that between +21 and +16) and by matching constitutive mutations in each inverted repeat. (However, the operator is not perfectly symmetrical; the left side binds more strongly than the right side to the repressor. A stronger operator would be created by a perfect inverted duplication of the left side.) Last updated on 7-29-2002


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References 509. Gilbert, W. and Muller-Hill, B. (1966). Isolation of the lac repressor. Proc. Natl. Acad. Sci. USA 56, 1891-1898. 510. Gilbert, W. and Muller-Hill, B. (1967). The lac operator is DNA. Proc. Natl. Acad. Sci. USA 58, 2415-2421.



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THE OPERON

3.10.11 Binding of inducer releases repressor from the operator Key Concepts

• Inducer binding causes a change in repressor conformation that reduces its affinity for DNA and releases it from the operator.

Various inducers cause characteristic reductions in the affinity of the repressor for the operator in vitro. These changes correlate with the effectiveness of the inducers in vivo. This suggests that induction results from a reduction in the attraction between operator and repressor. So when inducer enters the cell, it binds to free repressors and in effect prevents them from finding their operators. But consider a repressor tetramer that is already bound tightly to the operator. How does inducer cause this repressor to be released? Two models for repressor action are illustrated in Figure 10.14:

Figure 10.14 Does the inducer bind to free repressor to upset an equilibrium (left) or directly to repressor bound at the operator (right)? Binding of inducer releases repressor from the operator | SECTION 3.10.11 © 2004. Virtual Text / www.ergito.com

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• The equilibrium model (left) calls for repressor bound to DNA to be in rapid equilibrium with free repressor. Inducer would bind to the free form of repressor, and thus unbalance the equilibrium by preventing reassociation with DNA. • But the rate of dissociation of the repressor from the operator is much too slow to be compatible with this model (the half-life in vitro in the absence of inducer is >15 min). This means that instead the inducer must bind directly to repressor protein complexed with the operator. As indicated in the model on the right, inducer binding must produce a change in the repressor that makes it release the operator. Indeed, addition of IPTG causes an immediate destabilization of the repressor-operator complex in vitro. Binding of the repressor-IPTG complex to the operator can be studied by using greater concentrations of the protein in the methylation protection/enhancement assay. The large amount compensates for the low affinity of the repressor-IPTG complex for the operator. The complex makes exactly the same pattern of contacts with DNA as the free repressor. An analogous result is obtained with mutant repressors whose affinity for operator DNA is increased; they too make the same pattern of contacts. Overall, a range of repressor variants whose affinities for the operator span seven orders of magnitude all make the same contacts with DNA. Changes in the affinity of the repressor for DNA must therefore occur by influencing the general conformation of the protein in binding DNA, not by making or breaking one or a few individual bonds. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.10.11

Binding of inducer releases repressor from the operator | SECTION 3.10.11 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.12 The repressor monomer has several domains Key Terms The headpiece is the DNA-binding domain of the lac repressor. Key Concepts

• A single repressor subunit can be divided into the N-terminal DNA-binding domain, a hinge, and the core of the protein.

• The DNA-binding domain contains two short α-helical regions that bind the major groove of DNA.

• The inducer-binding site and the regions responsible for multimerization are located in the core.

The repressor has several domains. The DNA-binding domain occupies residues 1-59. It is known as the headpiece. It can be cleaved from the remainder of the monomer, which is known as the core, by trypsin. The crystal structure illustrated in Figure 10.15 offers a more detailed account of these regions (512; 513).

Figure 10.15 The structure of a monomer of Lac repressor identifies several independent domains. Photograph kindly provided by Mitchell Lewis.

The N-terminus of the monomer consists of two α-helices separated by a turn. This The repressor monomer has several domains | SECTION 3.10.12 © 2004. Virtual Text / www.ergito.com

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is a common DNA-binding motif, known as the HTH (helix-turn-helix); the two α-helices fit into the major groove of DNA, where they make contacts with specific bases (see Molecular Biology 3.12.12 Repressor uses a helix-turn-helix motif to bind DNA). This region is connected by a hinge to the main body of the protein. In the DNA-binding form of repressor, the hinge forms a small α-helix (as shown in the figure); but when the repressor is not bound to DNA, this region is disordered. The HTH and hinge together correspond to the headpiece. The bulk of the core consists of two regions with similar structures (core domains 1 and 2). Each has a six-stranded parallel β-sheet sandwiched between two α-helices on either side. The inducer binds in a cleft between the two regions . At the C-terminus, there is an α-helix that contains two leucine heptad repeats. This is the oligomerization domain. The oligomerization helices of four monomers associate to maintain the tetrameric structure. Last updated on 7-29-2002

The repressor monomer has several domains | SECTION 3.10.12 © 2004. Virtual Text / www.ergito.com

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References 512. Friedman, A. M., Fischmann, T. O., and Steitz, T. A. (1995). Crystal structure of lac repressor core tetramer and its implications for DNA looping. Science 268, 1721-1727. 513. Lewis, M. et al. (1996). Crystal structure of the lactose operon repressor and its complexes with DNA and inducer. Science 271, 1247-1254.


The repressor monomer has several domains | SECTION 3.10.12 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.13 Repressor is a tetramer made of two dimers Key Concepts

• Monomers form a dimer by making contacts between core domain 2 and between the oligomerization helices.

• Dimers form a tetramer by interactions between the oligomerization helices.

Figure 10.16 shows the structure of the tetrameric core (using a different modeling system from Figure 10.15). It consists in effect of two dimers. The body of the dimer contains a loose interface between the N-terminal regions of the core monomers, a cleft at which inducer binds, and a hydrophobic core (top). The C-terminal regions of each monomer protrude as parallel helices. (The headpiece would join on to the N-terminal regions at the top.) Together the dimers interact to form a tetramer (center) that is held together by a C-terminal bundle of four helices.

Repressor is a tetramer made of two dimers | SECTION 3.10.13 © 2004. Virtual Text / www.ergito.com

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Figure 10.16 The crystal structure of the core region of Lac repressor identifies the interactions between monomers in the tetramer. Each monomer is identified by a different color. Mutations are colored as: dimer interface - yellow; inducer-binding blue; oligomerization - white and purple. Photographs kindly provided by Alan Friedman.

Sites of mutations are shown by beads on the structure at the bottom. lacI s mutations make the repressor unresponsive to the inducer, so that the operon is uninducible. They map in two groups: gray shows those in the inducer-binding cleft, and yellow shows those that affect the dimer interface. The first group abolish the inducer binding site; the second group prevent the effects of inducer binding from being transmitted to the DNA-binding site. lacI –d mutations that affect oligomerization map in two groups. White shows mutations in core domain 2 that prevent dimer Repressor is a tetramer made of two dimers | SECTION 3.10.13 © 2004. Virtual Text / www.ergito.com

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formation. Purple shows those in the oligomerization helix that prevent tetramer formation from dimers. From these data we can derive the schematic of Figure 10.17, which shows how the monomers are organized into the tetramer. Two monomers form a dimer by means of contacts at core domain 2 and in the oligomerization helix. The dimer has two DNA-binding domains at one end of the structure, and the oligomerization helices at the other end. Two dimers then form a tetramer by interactions at the oligomerization interface.

Figure 10.17 The repressor tetramer consists of two dimers. Dimers are held together by contacts involving core domain 2 as well as by the oligomerization helix. The dimers are linked into the tetramer by the oligomerization interface.

Last updated on January 26, 2004 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.10.13

Repressor is a tetramer made of two dimers | SECTION 3.10.13 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.14 DNA-binding is regulated by an allosteric change in conformation Key Concepts

• The DNA-binding domain of a monomer inserts into the major groove of DNA. • Active repressor has a conformation in which the two DNA-binding domains of a dimer can insert into successive turns of the double helix.

• Inducer binding changes the conformation so that the two DNA-binding sites are not in the right geometry to make simultaneous contacts.

Early work suggested a model in which the headpiece is relatively independent of the core. It can bind to operator DNA by making the same pattern of contacts with a half-site as intact repressor. However, its affinity for DNA is many orders of magnitude less than that of intact repressor. The reason for the difference is that the dimeric form of intact repressor allows two headpieces to contact the operator simultaneously, each binding to one half-site. Figure 10.18 shows that the two DNA-binding domains in a dimeric unit contact DNA by inserting into successive turns of the major groove. This enormously increases affinity for the operator.

DNA-binding is regulated by an allosteric change in conformation | SECTION 3.10.14 © 2004. Virtual Text / www.ergito.com

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Figure 10.18 Inducer changes the structure of the core so that the headpieces of a repressor dimer are no longer in an orientation that permits binding to DNA.

Binding of inducer causes an immediate conformational change in the repressor protein. Binding of two molecules of inducer to the repressor tetramer is adequate to release repression. Binding of inducer changes the orientation of the headpieces relative to the core, with the result that the two headpieces in a dimer can no longer bind DNA simultaneously. This eliminates the advantage of the multimeric repressor, and reduces the affinity for the operator. Last updated on 7-29-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.10.14

DNA-binding is regulated by an allosteric change in conformation | SECTION 3.10.14 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.15 Mutant phenotypes correlate with the domain structure Key Concepts

• Different types of mutations occur in different domains of the repressor subunit.

Mutations in the Lac repressor identified the existence of different domains even before the structure was known (2568; 2569). We can now explain the nature of the mutations more fully by reference to the structure, as summarized inFigure 10.19 (for review see 2567).

Figure 10.19 The locations of three type of mutations in lactose repressor are mapped on the domain structure of the protein. Recessive lacI– mutants that cannot repress can map anywhere in the protein. Dominant negative lacI–d mutants that cannot repress map to the DNA-binding domain. Dominant lacIs mutants that cannot induce because they do not bind inducer map to core domain 1.

Recessive mutations of the lacI– type can occur anywhere in the bulk of the protein. Basically any mutation that inactivates the protein will have this phenotype. The more detailed mapping of mutations on to the crystal structure in Figure 10.16 identifies specific impairments for some of these mutations, for example, those that affect oligomerization. The special class of dominant-negative lacI-d mutations lie in the DNA-binding site Mutant phenotypes correlate with the domain structure | SECTION 3.10.15 © 2004. Virtual Text / www.ergito.com

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of the repressor subunit (see Molecular Biology 3.10.9 Multimeric proteins have special genetic properties). This explains their ability to prevent mixed tetramers from binding to the operator; a reduction in the number of binding sites reduces the specific affinity for the operator. The role of the N-terminal region in specifically binding DNA is shown also by its location as the site of occurrence of "tight binding" mutations. These increase the affinity of the repressor for the operator, sometimes so much that it cannot be released by inducer. They are rare. Uninducible lacIs mutations map in a region of the core domain 1 extending from the inducer-binding site to the hinge. One group lies in amino acids that contact the inducer, and these mutations function by preventing binding of inducer. The remaining mutations lie at sites that must be involved in transmitting the allosteric change in conformation to the hinge when inducer binds. Last updated on 7-29-2002

Mutant phenotypes correlate with the domain structure | SECTION 3.10.15 © 2004. Virtual Text / www.ergito.com

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Reviews 2567. Pace, H. C., Kercher, M. A., Lu, P., Markiewicz, P., Miller, J. H., Chang, G., and Lewis, M. (1997). Lac repressor genetic map in real space. Trends Biochem. Sci. 22, 334-339.

References 2568. Markiewicz, P., Kleina, L. G., Cruz, C., Ehret, S., and Miller, J. H. (1994). Genetic studies of the lac repressor. XIV. Analysis of 4000 altered E. coli lac repressors reveals essential and non-essential residues, as well as spacers which do not require a specific sequence. J. Mol. Biol. 240, 421-433. 2569. Suckow, J., Markiewicz, P., Kleina, L. G., Miller, J., Kisters-Woike, B., and Muller-Hill, B. (1996). Genetic studies of the Lac repressor. XV: 4000 single amino acid substitutions and analysis of the resulting phenotypes on the basis of the protein structure. J. Mol. Biol. 261, 509-523.


Mutant phenotypes correlate with the domain structure | SECTION 3.10.15 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.16 Repressor binds to three operators and interacts with RNA polymerase Key Concepts

• Each dimer in a repressor tetramer can bind an operator, so that the tetramer can bind two operators simultaneously.

• Full repression requires the repressor to bind to an additional operator downstream or upstream as well as to the operator at lacZ.

• Binding of repressor at the operator stimulates binding of RNA polymerase at the promoter.

The allosteric transition that results from binding of inducer occurs in the repressor dimer. So why is a tetramer required to establish full repression? Each dimer can bind an operator sequence. This enables the intact repressor to bind to two operator sites simultaneously. In fact, there are two further operator sites in the initial region of the lac operon. The original operator, O1, is located just at the start of the lacZ gene. It has the strongest affinity for repressor. Weaker operator sequences (sometimes called pseudo-operators) are located on either side; O2 is 410 bp downstream of the startpoint, and O3 is 83 bp upstream of it. Figure 10.20 shows what happens when a DNA-binding protein can bind simultaneously to two separated sites on DNA. The DNA between the two sites forms a loop from a base where the protein has bound the two sites. The length of the loop depends on the distance between the two binding sites. When Lac repressor binds simultaneously to O1 and to one of the other operators, it causes the DNA between them to form a rather short loop, significantly constraining the DNA structure A scale model for binding of tetrameric repressor to two operators is shown in Figure 10.21 (511).

Repressor binds to three operators and interacts with RNA polymerase | SECTION 3.10.16 © 2004. Virtual Text / www.ergito.com

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Figure 10.20 If both dimers in a repressor tetramer bind to DNA, the DNA between the two binding sites is held in a loop.

Figure 10.21 When a repressor tetramer binds to two operators, the stretch of DNA between them is forced into a tight loop. (The blue structure in the center of the looped DNA represents CAP, another regulator protein that binds in this region). Photograph kindly provided by Mitchell Lewis.

Binding at the additional operators affects the level of repression. Elimination of either the downstream operator (O2) or the upstream operator (O3) reduces the efficiency of repression by 2-4×. However, if both O2 and O3 are eliminated, repression is reduced 100×. This suggests that the ability of the repressor to bind to one of the two other operators as well as to O1 is important for establishing repression. We do not know how and why this simultaneous binding increases repression. We know most about the direct effects of binding of repressor to the operator (O1). It was originally thought that repressor binding would occlude RNA polymerase from binding to the promoter. However, we now know that the two proteins may be bound to DNA simultaneously, and the binding of repressor actually enhances the binding of RNA polymerase! But the bound enzyme is prevented from initiating transcription. Repressor binds to three operators and interacts with RNA polymerase | SECTION 3.10.16 © 2004. Virtual Text / www.ergito.com

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The equilibrium constant for RNA polymerase binding alone to the lac promoter is 1.9 × 107 M-1. The presence of repressor increases this constant by two orders of magnitude to 2.5 × 109 M-1. In terms of the range of values for the equilibrium constant KB given in Figure 9.20, repressor protein effectively converts the formation of closed complex by RNA polymerase at the lac promoter from a weak to a strong interaction. What does this mean for induction of the operon? The higher value for KB means that, when occupied by repressor, the promoter is 100 times more likely to be bound by an RNA polymerase. And by allowing RNA polymerase to be bound at the same time as repressor, it becomes possible for transcription to begin immediately upon induction, instead of waiting for an RNA polymerase to be captured. The repressor in effect causes RNA polymerase to be stored at the promoter. The complex of RNA polymerase·repressor·DNA is blocked at the closed stage. When inducer is added, the repressor is released, and the closed complex is converted to an open complex that initiates transcription. The overall effect of repressor has been to speed up the induction process. Does this model apply to other systems? The interaction between RNA polymerase, repressor, and the promoter/operator region is distinct in each system, because the operator does not always overlap with the same region of the promoter (see Figure 10.26). For example, in phage lambda, the operator lies in the upstream region of the promoter, and binding of repressor occludes the binding of RNA polymerase (see Molecular Biology 3.12 Phage strategies). So a bound repressor does not interact with RNA polymerase in the same way in all systems. Last updated on 7-29-2002


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References 511. Oehler, S. (1990). The three operators of the lac operon cooperate in repression. EMBO J. 9, 973-979.



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3.10.17 Repressor is always bound to DNA Key Concepts

• Proteins that have a high affinity for a specific DNA sequence also have a low affinity for other DNA sequences.

• Every base pair in the bacterial genome is the start of a low-affinity binding-site for repressor.

• The large number of low-affinity sites ensures that all repressor protein is bound to DNA.

• Repressor binds to the operator by moving from a low-affinity site rather than by equilibrating from solution.

Probably all proteins that have a high affinity for a specific sequence also possess a low affinity for any (random) DNA sequence. A large number of low-affinity sites will compete just as well for a repressor tetramer as a small number of high-affinity sites. There is only one high-affinity site in the E. coli genome: the operator. The remainder of the DNA provides low-affinity binding sites. Every base pair in the genome starts a new low-affinity site. (Just moving one base pair along the genome, out of phase with the operator itself, creates a low-affinity site!) So there are 4.2 × 106 low-affinity sites. The large number of low-affinity sites means that, even in the absence of a specific binding site, all or virtually all repressor is bound to DNA; none is free in solution. Figure 10.22 shows how the equation of describing the equilibrium between free repressor and DNA-bound repressor can be rearranged to give the proportion of free repressor.

Figure 10.22 Repressor binding to random sites is governed by an equilibrium equation. Repressor is always bound to DNA | SECTION 3.10.17 © 2004. Virtual Text / www.ergito.com

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Applying the parameters for the lac system, we find that: • The nonspecific equilibrium binding constant is KA = 2 × 106 M–1. • The concentration of nonspecific binding sites is 4 × 106 in a bacterial volume of 10–15 liter, which corresponds to [DNA] = 7 × 10–3 M (a very high concentration). Substituting these values gives: Free / Bound repressor = 10–4. So all but 0.01% of repressor is bound to (random) DNA. Since there are ~10 molecules of repressor per cell, this is tantamount to saying that there is no free repressor protein. This has an important implication for the interaction of repressor with the operator: it means that we are concerned with the partitioning of the repressor on DNA, in which the single high-affinity site of the operator competes with the large number of low-affinity sites. In this competition, the absolute values of the association constants for operator and random DNA are not important; what is important is the ratio of Ksp (the constant for binding a specific site) to Knsp (the constant for binding any random DNA sequence), that is, the specificity. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.10.17

Repressor is always bound to DNA | SECTION 3.10.17 © 2004. Virtual Text / www.ergito.com

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THE OPERON

3.10.18 The operator competes with low-affinity sites to bind repressor Key Concepts

• In the absence of inducer, the operator has an affinity for repressor that is 10 × that 7

of a low affinity site.

• The level of 10 repressor tetramers per cell ensures that the operator is bound by repressor 96% of the time.

• Induction reduces the affinity for the operator to 10 × that of low-affinity sites, so 4

that only 3% of operators are bound.

• Induction causes repressor to move from the operator to a low-affinity site by direct displacement.

• These parameters could be changed by a reduction in the effective concentration of DNA in vivo.

We can define the parameters that influence the ability of a regulator protein to saturate its target site by comparing the equilibrium equations for specific and nonspecific binding. As might be expected intuitively, the important parameters are: • The size of the genome dilutes the ability of a protein to bind specific target sites. • The specificity of the protein counters the effect of the mass of DNA. • The amount of protein that is required increases with the total amount of DNA in the genome and decreases with the specificity. • The amount of protein also must be in reasonable excess of the total number of specific target sites, so we expect regulators with many targets to be found in greater quantities than regulators with fewer targets. Figure 10.23 compares the equilibrium constants for lac repressor/operator binding with repressor/general DNA binding. From these constants, we can deduce how repressor is partitioned between the operator and the rest of DNA, and what happens to the repressor when inducer causes it to dissociate from the operator.

The operator competes with low-affinity sites to bind repressor | SECTION 3.10.18 © 2004. Virtual Text / www.ergito.com

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Figure 10.23 Lac repressor binds strongly and specifically to its operator, but is released by inducer. All equilibrium constants are in M-1.

Repressor binds ~107 times better to operator DNA than to any random DNA sequence of the same length. So the operator comprises a single high-affinity site that will compete for the repressor 107 better than any low-affinity (random) site. How does this ensure that the repressor can maintain effective control of the operon? Using the specificity, we can calculate the distribution between random sites and the operator, and can express this in terms of occupancy of the operator. If there are 10 molecules of lac repressor per cell with a specificity for the operator of 107, the operator will be bound by repressor 96% of the time. The role of specificity explains two features of the lac repressor-operator interaction: • When inducer binds to the repressor, the affinity for the operator is reduced by ~103-fold. The affinity for general DNA sequences remains unaltered. So the specificity is now only 104, which is insufficient to capture the repressor against competition from the excess of 4.2 × 106 low-affinity sites. Only 3% of operators would be bound under these conditions. • Mutations that reduce the affinity of the operator for the repressor by as little as 20-30× have sufficient effect to be constitutive. Within the genome, the mutant operators can be overwhelmed by the preponderance of random sites. The occupancy of the operator is reduced to ~50% if the repressor's specificity is reduced just 10× . The consequence of these affinities is that in an uninduced cell, one tetramer of repressor usually is bound to the operator. All or almost all of the remaining tetramers are bound at random to other regions of DNA, as illustrated in Figure 10.24. There are likely to be very few or no repressor tetramers free within the cell.


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Figure 10.24 Virtually all the repressor in the cell is bound to DNA.

The addition of inducer abolishes the ability of repressor to bind specifically at the operator. Those repressors bound at the operator are released, and bind to random (low-affinity) sites. So in an induced cell, the repressor tetramers are "stored" on random DNA sites. In a noninduced cell, a tetramer is bound at the operator, while the remaining repressor molecules are bound to nonspecific sites. The effect of induction is therefore to change the distribution of repressor on DNA, rather than to generate free repressor. When inducer is removed, repressor recovers its ability to bind specifically to the operator, and does so very rapidly. This must involve its movement from a nonspecific "storage" site on DNA. What mechanism is used for this rapid movement? The ability to bind to the operator very rapidly is not consistent with the time that would be required for multiple cycles of dissociation and reassociation with nonspecific sites on DNA. The discrepancy excludes random-hit mechanisms for finding the operator, suggesting that the repressor can move directly from a random site on DNA to the operator. This is the same issue that we encountered previously with the ability of RNA polymerase to find its promoters (see Figure 9.22 and The operator competes with low-affinity sites to bind repressor | SECTION 3.10.18 © 2004. Virtual Text / www.ergito.com

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Figure 9.23). The same solution is likely: movement could be accomplished by direct displacement from site to site (as indicated in Figure 10.24). A displacement reaction might be aided by the presence of more binding sites per tetramer (four) than are actually needed to contact DNA at any one time (two). The parameters involved in finding a high-affinity operator in the face of competition from many low-affinity sites pose a dilemma for repressor. Under conditions of repression, there must be high specificity for the operator. But under conditions of induction, this specificity must be relieved. Suppose, for example, that there were 1000 molecules of repressor per cell. Then only 0.04% of operators would be free under conditions of repression. But upon induction only 40% of operators would become free. We therefore see an inverse correlation between the ability to achieve complete repression and the ability to relieve repression effectively. We assume that the number of repressors synthesized in vivo has been subject to selective forces that balance these demands. The difference in expression of the lactose operon between its induced and repressed states in vivo is actually 103×. In other words, even when inducer is absent, there is a basal level of expression of ~0.1% of the induced level. This would be reduced if there were more repressor protein present, increased if there were less. So it could be impossible to establish tight repression if there were fewer repressors than the 10 found per cell; and it might become difficult to induce the operon if there were too many (514). It is possible to introduce the lac operator-repressor system into the mouse. When the lac operator is connected to a tyrosinase reporter gene, the enzyme is induced by the addition of IPTG (2240). This means that the repressor is finding its target in a genome 103 times larger than that of E. coli. Induction occurs at approximately the same concentration of IPTG as in bacteria. However, we do not know the concentration of Lac repressor and how effectively the target is induced. In order to extrapolate in vivo from the affinity of a DNA-protein interaction in vitro, we need to know the effective concentration of DNA in vivo. The "effective concentration" differs from the mass/volume because of several factors. The effective concentration is increased, for example, by molecular crowding, which occurs when polyvalent cations neutralize ~90% of the charges on DNA, and the nucleic acid collapses into condensed structures. The major force that decreases the effective concentration is the inaccessibility of DNA that results from occlusion or sequestration by DNA-binding proteins. One way to determine the effective concentration is to compare the rate of a reaction in vitro and in vivo that depends on DNA concentration. This has been done using intermolecular recombination between two DNA molecules. To provide a control, the same reaction is followed as an intramolecular recombination, that is, the two recombining sites are presented on the same DNA molecule. We assume that concentration is the same in vivo and in vitro for the intramolecular reaction, and therefore any difference in the ratio of intermolecular/ intramolecular recombination rates can be attributed to a change in the effective concentration in vivo. The results of such a comparison suggest that the effective concentration of DNA is reduced >10-fold in vivo (516).


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This could affect the rates of reactions that depend on DNA concentration, including DNA recombination, and protein-DNA binding. It emphasizes the problem encountered by all DNA-binding proteins in finding their targets with sufficient speed, and reinforces the conclusion that diffusion is not adequate (see Figure 9.22). Last updated on 12-20-2001


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References 514. Lin, S.-y. and Riggs, A. D. (1975). The general affinity of lac repressor for E. coli DNA: implications for gene regulation in prokaryotes and eukaryotes. Cell 4, 107-111. 516. Hildebrandt, E. R. et al. (1995). Comparison of recombination in vitro and in E. coli cells: measure of the effective concentration of DNA in vivo. Cell 81, 331-340. 2240. Cronin, C. A., Gluba, W., and Scrable, H. (2001). The lac operator-repressor system is functional in the mouse. Genes Dev. 15, 1506-1517.



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3.10.19 Repression can occur at multiple loci Key Terms Autogenous control describes the action of a gene product that either inhibits (negative autogenous control) or activates (positive autogenous control) expression of the gene coding for it. Key Concepts

• A repressor will act on all loci that have a copy of its target operator sequence.

The lac repressor acts only on the operator of the lacZYA cluster. However, some repressors control dispersed structural genes by binding at more than one operator. An example is the trp repressor, which controls three unlinked sets of genes: • An operator at the cluster of structural genes trpEDBCA controls coordinate synthesis of the enzymes that synthesize tryptophan from chorismic acid. • An operator at another locus controls the aroH gene, which codes for one of the three enzymes that catalyze the initial reaction in the common pathway of aromatic amino acid biosynthesis. • The trpR regulator gene is repressed by its own product, the trp repressor. So the repressor protein acts to reduce its own synthesis. This circuit is an example of autogenous control. Such circuits are quite common in regulatory genes, and may be either negative or positive (see Molecular Biology 3.11.11 r-protein synthesis is controlled by autogenous regulation and Molecular Biology 3.12.9 Repressor maintains an autogenous circuit). A related 21 bp operator sequence is present at each of the three loci at which the trp repressor acts. The conservation of sequence is indicated in Figure 10.25. Each operator contains appreciable (but not identical) dyad symmetry. The features conserved at all three operators include the important points of contact for trp repressor. This explains how one repressor protein acts on several loci: each locus has a copy of a specific DNA-binding sequence recognized by the repressor (just as each promoter shares consensus sequences with other promoters).

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Figure 10.25 The trp repressor recognizes operators at three loci. Conserved bases are shown in red. The location of the startpoint and mRNA varies, as indicated by the white arrows.

Figure 10.26 summarizes the variety of relationships between operators and promoters. A notable feature of the dispersed operators recognized by TrpR is their presence at different locations within the promoter in each locus. In trpR the operator lies between positions -12 and +9, while in the trp operon it occupies positions -23 to -3, but in the aroH locus it lies farther upstream, between -49 and -29. In other cases, the operator lies downstream from the promoter (as in lac), or apparently just upstream of the promoter (as in gal, where the nature of the repressive effect is not quite clear). The ability of the repressors to act at operators whose positions are different in each target promoter suggests that there could be differences in the exact mode of repression, the common feature being that RNA polymerase is prevented from initiating transcription at the promoter.

Figure 10.26 Operators may lie at various positions relative to the promoter.


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3.10.20 Summary Transcription is regulated by the interaction between trans-acting factors and cis-acting sites. A trans-acting factor is the product of a regulator gene. It is usually protein but can be RNA. Because it diffuses in the cell, it can act on any appropriate target gene. A cis-acting site in DNA (or RNA) is a sequence that functions by being recognized in situ. It has no coding function and can regulate only those sequences that are physically contiguous with it. Bacterial genes coding for proteins whose functions are related, such as successive enzymes in a pathway, may be organized in a cluster that is transcribed into a polycistronic mRNA from a single promoter. Control of this promoter regulates expression of the entire pathway. The unit of regulation, containing structural genes and cis-acting elements, is called the operon. Initiation of transcription is regulated by interactions that occur in the vicinity of the promoter. The ability of RNA polymerase to initiate at the promoter is prevented or activated by other proteins. Genes that are active unless they are turned off are said to be under negative control. Genes that are active only when specifically turned on are said to be under positive control. The type of control can be determined by the dominance relationships between wild type and mutants that are constitutive/derepressed (permanently on) or uninducible/super-repressed (permanently off). A repressor protein prevents RNA polymerase either from binding to the promoter or from activating transcription. The repressor binds to a target sequence, the operator, that usually is located around or upstream of the startpoint. Operator sequences are short and often are palindromic. The repressor is often a homomultimer whose symmetry reflects that of its target. The ability of the repressor protein to bind to its operator is regulated by a small molecule. An inducer prevents a repressor from binding; a corepressor activates it. Binding of the inducer or corepressor to its site produces a change in the structure of the DNA-binding site of the repressor. This allosteric reaction occurs in both free repressor proteins and directly in repressor proteins already bound to DNA. The lactose pathway operates by induction, when an inducer β-galactoside prevents the repressor from binding its operator; transcription and translation of the lacZ gene then produce β-galactosidase, the enzyme that metabolizes β-galactosides. The tryptophan pathway operates by repression; the corepressor (tryptophan) activates the repressor protein, so that it binds to the operator and prevents expression of the genes that code for the enzymes that biosynthesize tryptophan. A repressor can control multiple targets that have copies of an operator consensus sequence. A protein with a high affinity for a particular target sequence in DNA has a lower affinity for all DNA. The ratio defines the specificity of the protein. Because there are many more nonspecific sites (any DNA sequence) than specific target sites in a genome, a DNA-binding protein such as a repressor or RNA polymerase is "stored" on DNA; probably none or very little is free. The specificity for the target sequence must be great enough to counterbalance the excess of nonspecific sites over specific Summary | SECTION 3.10.20 © 2004. Virtual Text / www.ergito.com

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sites. The balance for bacterial proteins is adjusted so that the amount of protein and its specificity allow specific recognition of the target in "on" conditions, but allow almost complete release of the target in "off" conditions. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.10.20


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3.11.1 Introduction The basic concept of genetic regulation in bacteria is that the expression of a gene may be controlled by a regulator that interacts with a specific sequence or structure in DNA or mRNA at some stage prior to the synthesis of protein. This is an extremely flexible idea with many ramifications. The stage of expression that is controlled can involve transcription, when the target for regulation is DNA; or it can be at translation, when the target for regulation is RNA. When transcription is involved, the level of control can be at initiation or at termination. The regulator can be a protein or an RNA. "Controlled" can mean that the regulator turns off (represses) the target or that it turns on (activates) the target. Expression of many genes can be coordinately controlled by a single regulator gene on the principle that each target contains a copy of the sequence or structure that the regulator recognizes. Regulators may themselves be regulated, most typically in response to small molecules whose supply responds to environmental conditions. Regulators may be controlled by other regulators to make complex circuits. Let's compare the ways that different types of regulators work. Protein regulators work on the principle of allostery (see Molecular Biology Supplement 32.7 Allostery). The protein has two binding sites, one for a nucleic acid target, the other for a small molecule. Binding of the small molecule to its site changes the conformation in such a way as to alter the affinity of the other site for the nucleic acid. The way in which this happens is known in detail for the Lac repressor (see Molecular Biology 3.10.14 DNA-binding is regulated by an allosteric change in conformation). Protein regulators are often multimeric, with a symmetrical organization that allows two subunits to contact a palindromic target on DNA. This can generate cooperative binding effects that create a more sensitive response to regulation. RNA regulators use changes in secondary structure as the guiding principle. An RNA regulator recognizes its target by the familiar principle of complementary base pairing. Figure 11.1 shows that the regulator is usually a small RNA molecule with extensive secondary structure, but with a single-stranded region(s) that is complementary to a single-stranded region in its target. The formation of a double helical region between regulator and target can have two types of consequence:


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Figure 11.1 A regulator RNA is a small RNA with a single-stranded region that can pair with a single-stranded region in a target RNA.

• Formation of the double helical structure may itself be sufficient. In some cases, protein(s) can bind only to the single-stranded form of the target sequence, and are therefore prevented by acting by duplex formation. In other cases, the duplex region becomes a target for binding, for example, by nucleases that degrade the RNA and therefore prevent its expression. • Duplex formation may be important because it sequesters a region of the target RNA that would otherwise participate in some alternative secondary structure. Going beyond the interactions in which a protein or RNA regulates expression of a single gene, we find that bacteria have responses in which the expression of many genes is coordinated. The simplest form of regulating multiple genes occurs when they all have a copy of the same cis-acting regulatory elements. Genes that have a copy of the same operator sequence are coordinately repressed by a single repressor protein. Genes that have the same type of promoter are coordinately activated by the production of a sigma factor that causes RNA polymerase to use that promoter. Regulatory networks are created when one regulator is required for the production of another. This happens in situations in which an ordered temporal expression of genes is required, for example, during phage infection (see Molecular Biology 3.12.3 Lytic development is controlled by a cascade) or during the development of a new cell type, such as a spore (see Molecular Biology 3.9.18 Sigma factors may be organized into cascades ). These use one of the simplest relationships between regulators, in which one regulator is necessary for expression of the next regulator in a series, creating a cascade. Another type of relationship occurs when one regulator directly regulates the activity of another regulator. For example, a protein that represses or activates expression of a gene may itself by inhibited by an "anti-regulator" that responds to some other signal. Figure 11.2 shows an example. A series of such relationships can be Introduction | SECTION 3.11.1 © 2004. Virtual Text / www.ergito.com

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extended indefinitely. Some circuits are controlled by a series of regulators each of which antagonizes another. Such circuits allow the cell to control the target set of genes in response to multiple stimuli, since each stimulus can feed into the regulatory circuit at a different point.

Figure 11.2 A protein that antagonizes a repressor can turn a gene on.

A special type of circuit is created when a protein regulates expression of the gene that codes for it. This is called autogenous control. It allows a protein to regulate its own level of expression without reference to any other circuit. It is often used for regulating levels of proteins that are assembled into macromolecular complexes. Last updated on 11-15-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.11.1


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3.11.2 Distinguishing positive and negative control Key Terms An activator is a protein that stimulates the expression of a gene, typically by acting at a promoter to stimulate RNA polymerase. In eukaryotes, the sequence to which it binds in the promoter is called a response element. An inducible operon is expressed only in the presence of a specific small molecule (the inducer). A repressible operon is expressed unless the small molecule co-repressor is present. The derepressed state describes a gene that is turned on because a small molecule corepressor is absent. It has the same effect as the induced state that is produced by a small molecule inducer for a gene that is regulated by induction. In describing the effect of a mutation, derepressed and constitutive have the same meaning. Super-repressed is a mutant condition in which a repressible operon cannot be de-repressed, so it is always turned off. The level of response from a system in the absence of a stimulus is its basal level. (The basal level of transcription of a gene is the level that occurs in the absence of any specific activation.) Key Concepts

• Induction can be achieved by inactivating a repressor or activating an activator. • Repression can be achieved by activating a repressor or inactivating an activator.

Positive and negative control systems are defined by the response of the operon when no regulator protein is present. The characteristics of the two types of control system are mirror images. Genes under negative control are expressed unless they are switched off by a repressor protein (see Figure 10.2). Any action that interferes with gene expression can provide a negative control. Typically a repressor protein either binds to DNA to prevent RNA polymerase from initiating transcription, or binds to mRNA to prevent a ribosome from initiating translation. Negative control provides a fail-safe mechanism: if the regulator protein is inactivated, the system functions and so the cell is not deprived of these enzymes. It is easy to see how this might evolve. Originally a system functions constitutively, but then cells able to interfere specifically with its expression acquire a selective advantage by virtue of their increased efficiency. For genes under positive control, expression is possible only when an active regulator protein is present. The mechanism for controlling an individual operon is Distinguishing positive and negative control | SECTION 3.11.2 © 2004. Virtual Text / www.ergito.com

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an exact counterpart of negative control, but instead of interfering with initiation, the regulator protein is essential for it. It interacts with DNA and with RNA polymerase to assist the initiation event (see Figure 10.3). The use of sigma factors to regulate transcription formally is an example of positive control. A positive regulator protein that responds to a small molecule is usually called an activator. It is less obvious how positive control evolved, since the cell must have had the ability to express the regulated genes even before any control existed. Presumably some component of the control system must have changed its role. Perhaps originally it was used as a regular part of the apparatus for gene expression; then later it became restricted to act only in a particular system or systems. Operons are defined as inducible or repressible by the nature of their response to the small molecule that regulates their expression. Inducible operons function only in the presence of the small-molecule inducer. Repressible operons function only in the absence of the small-molecule corepressor (so called to distinguish it from the repressor protein). The terminology used for repressible systems describes the active state of the operon as derepressed; this has the same meaning as induced. The condition in which a (mutant) operon cannot be derepressed is sometimes called super-repressed; this is the exact counterpart of uninducible. Either positive or negative control could be used to achieve either induction or repression by utilizing appropriate interactions between the regulator protein and the small-molecule inducer or corepressor. Figure 11.3 summarizes four simple types of control circuit. Induction is achieved when an inducer inactivates a repressor protein or activates an activator protein. Repression is accomplished when a corepressor activates a repressor protein or inactivates an activator protein.

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Figure 11.3 Control circuits are versatile and can be designed to allow positive or negative control of induction or repression.

The trp operon is a repressible system. Tryptophan is the end product of the reactions catalyzed by a series of biosynthetic enzymes. Both the activity and the synthesis of the tryptophan enzymes are controlled by the level of tryptophan in the cell. Tryptophan functions as a corepressor that activates a repressor protein. This is the classic mechanism for repression, as seen in Figure 11.3 (lower left). In conditions when the supply of tryptophan is plentiful, the operon is repressed because the repressor protein·corepressor complex is bound at the operator. When tryptophan is in short supply, the corepressor is inactive, therefore has reduced specificity for the operator, and is stored elsewhere on DNA. Deprivation of repressor causes ~70-fold increase in the frequency of initiation events at the trp promoter. Even under repressing conditions, the structural genes Distinguishing positive and negative control | SECTION 3.11.2 © 2004. Virtual Text / www.ergito.com

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continue to be expressed at a low basal level (sometimes also called the repressed level). The efficiency of repression at the operator is much lower than in the lac operon (where the basal level is only ~1/1000 of the induced level). We have treated both induction and repression as phenomena that rely upon allosteric changes induced in regulator proteins by small molecules. Other types of interactions also can be used to control the activities of regulator proteins. One example is OxyR, a transcriptional activator of genes induced by hydrogen peroxide. The OxyR protein is directly activated by oxidation, so it provides a sensitive measure of oxidative stress. Another common type of signal is phosphorylation of a regulator protein. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.11.2

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3.11.3 Glucose repression controls use of carbon sources Key Terms Glucose repression (Catabolite repression) describes the decreased expression of many bacterial operons that results from addition of glucose. Inducer exclusion describes the inhibition of uptake of other carbon sources into the cell that is caused by uptake of glucose. Key Concepts

• E. coli uses glucose in preference to other carbon sources when it has a choice. • Glucose prevents uptake of alternative carbon sources from the medium. • Exclusion of the alternative carbon sources from the cell prevents expression of the operons coding for the enzymes that metabolize them.

Bacteria have distinct preferences among potential carbon sources when they are offered a choice. When glucose is available as an energy source, it is used in preference to other sugars. So when E. coli finds (for example) both glucose and lactose in the medium, it metabolizes the glucose and represses the use of lactose. The phenomenon of glucose repression follows from the ability of glucose to prevent the use of alternative carbon sources. It describes the general repression of transcription of the operons (such as lac, gal, and ara) coding for the enzymes required to metabolize the alternative carbon sources. The same effect is found in many bacteria, although different molecular mechanisms may be responsible in different classes of bacteria (for review see 2563). The basis for glucose repression in E. coli appears to be largely indirect. It involves two mechanisms: • Inducer exclusion describes the ability of glucose to prevent alternative carbon sources from being taken up from the medium. These include the inducers of the operons coding for the alternative metabolic systems. As a result of the exclusion of the small molecule inducers from the cell, the operons cannot be transcribed. • A general system for activating many operons is provided by the activator protein CRP, which is activated by the small molecule cyclic AMP (see Molecular Biology 3.11.5 CRP functions in different ways in different target operons). It was thought for many years that this system is inactivated by glucose, with the result that its target operons are not expressed under conditions of glucose repression. However, this view has been challenged recently, and the exact way in which it is controlled by glucose is not clear.

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Figure 11.4 outlines the interactions involved in inducer exclusion. The key molecular component in inducer exclusion is the PTS (phosphoenolpyruvate:glycose phosphotransferase system), a complex of proteins in the bacterial membrane, which simultaneously phosphorylates and transports sugars into the cell (for review see 2561). One of the proteins of this complex (IIAGlc, which is coded by the crr gene) becomes dephosphorylated as a result of glucose transport. It then binds to the Lac permease and prevents it from importing lactose into the cell.

Figure 11.4 Glucose is imported through the Pts system. This causes the Lac permease to be inactivated. The absence of lactose means that the lac repressor switches off the operon. When glucose is absent, lactose can be imported, and inactivates the lac repressor, so the operon is switched on.



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Reviews 2561. Meadow, N. D., Fox, D. K., and Roseman, S. (1990). The bacterial phosphoenolpyruvate: glycose phosphotransferase system. Annu. Rev. Biochem. 59, 497-542. 2563. Stalke, J. and Hillen, W. (2000). Regulation of carbon catabolism in Bacillus species. Annu. Rev. Microbiol. 54, 849-880.



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3.11.4 Cyclic AMP is an inducer that activates CRP to act at many operons Key Terms CRP activator (CAP activator) is a positive regulator protein activated by cyclic AMP. It is needed for RNA polymerase to initiate transcription of many operons of E. coli. Adenylate cyclase is an enzyme that uses ATP as a substrate to generate cyclic AMP, in which 5 ′ and 3 ′ positions of the sugar ring are connected via a phosphate group. Key Concepts

• CRP is an activator protein that binds to a target sequence at a promoter. • A dimer of CRP is activated by a single molecule of cyclic AMP.

So far we have dealt with the promoter as a DNA sequence that is competent to bind RNA polymerase, which then initiates transcription. But there are some promoters at which RNA polymerase cannot initiate transcription without assistance from an ancillary protein. Such proteins are positive regulators, because their presence is necessary to switch on the transcription unit. Typically the activator overcomes a deficiency in the promoter, for example, a poor consensus sequence at –35 or –10. One of the most widely acting activators is a protein called CRP activator that controls the activity of a large set of operons in E. coli. The protein is a positive control factor whose presence is necessary to initiate transcription at dependent promoters. CRP is active only in the presence of cyclic AMP, which behaves as the classic small-molecule inducer (see Figure 11.3; upper right). Cyclic AMP is synthesized by the enzyme adenylate cyclase. The reaction uses ATP as substrate and introduces a 3 ′ –5 ′ link via phosphodiester bonds, generating the structure drawn in Figure 11.5. Mutations in the gene coding for adenylate cyclase (cya–) do not respond to changes in glucose levels.

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Figure 11.5 Cyclic AMP has a single phosphate group connected to both the 3 ′ and 5 ′ positions of the sugar ring.

The level of cyclic AMP is inversely related to the level of glucose. The basis for this effect lies with the same component of the Pts system that is responsible for controlling lactose uptake. The phosphorylated form of protein IIAGlc stimulates adenylate cyclase. When glucose is imported, the dephosphorylation of IIAGlc leads to a fall in adenylate cyclase activity. Figure 11.6 shows that reducing the level of cyclic AMP renders the (wild-type) protein unable to bind to the control region, which in turn prevents RNA polymerase from initiating transcription. So the effect of glucose in reducing cyclic AMP levels is to deprive the relevant operons of a control factor necessary for their expression.

Figure 11.6 By reducing the level of cyclic AMP, glucose inhibits the transcription of operons that require CRP activity.


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3.11.5 CRP functions in different ways in different target operons Key Concepts

• CRP-binding sites lie at highly variable locations relative to the promoter. • CRP interacts with RNA polymerase, but the details of the interaction depend on the relative locations of the CRP-binding site and the promoter.

The CRP factor binds to DNA, and complexes of cyclic AMP·CRP·DNA can be isolated at each promoter at which it functions. The factor is a dimer of two identical subunits of 22.5 kD, which can be activated by a single molecule of cyclic AMP. A CRP monomer contains a DNA-binding region and a transcription-activating region (for review see 99). A CRP dimer binds to a site of ~22 bp at a responsive promoter. The binding sites include variations of the consensus sequence given in Figure 11.7. Mutations preventing CRP action usually are located within the well conserved pentamer , which appears to be the essential element in recognition. CRP binds most strongly to sites that contain two (inverted) versions of the pentamer, because this enables both subunits of the dimer to bind to the DNA. Many binding sites lack the second pentamer, however, and in these the second subunit must bind a different sequence (if it binds to DNA). The hierarchy of binding affinities for CRP helps to explain why different genes are activated by different levels of cyclic AMP in vivo.

Figure 11.7 The consensus sequence for CRP contains the well conserved pentamer TGTGA and (sometimes) an inversion of this sequence (TCANA).

The action of CRP has the curious feature that its binding sites lie at different locations relative to the startpoint in the various operons that it regulates. And the TGTGA pentamer may lie in either orientation. The three examples summarized in Figure 11.8 encompass the range of locations:

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Figure 11.8 The CRP protein can bind at different sites relative to RNA polymerase.

• The CRP-binding site is adjacent to the promoter, as in the lac operon, in which the region of DNA protected by CRP is centered on –61. It is possible that two dimers of CRP are bound. The binding pattern is consistent with the presence of CRP largely on one face of DNA, the same face that is bound by RNA polymerase. This location would place the two proteins just about in reach of each other. • Sometimes the CRP-binding site lies within the promoter, as in the gal locus, where the CRP-binding site is centered on –41. It is likely that only a single CRP dimer is bound, probably in quite intimate contact with RNA polymerase, since the CRP-binding site extends well into the region generally protected by the RNA polymerase. • In other operons, the CRP-binding site lies well upstream of the promoter. In the ara region, the binding site for a single CRP is the farthest from the startpoint, centered at –92. Dependence on CRP is related to the intrinsic efficiency of the promoter. No CRP-dependent promoter has a good –35 sequence and some also lack good –10 sequences. In fact, we might argue that effective control by CRP would be difficult if the promoter had effective –35 and –10 regions that interacted independently with RNA polymerase. There are in principle two ways in which CRP might activate transcription: it could interact directly with RNA polymerase; or it could act upon DNA to change its structure in some way that assists RNA polymerase to bind. In fact, CRP has effects upon both RNA polymerase and DNA (for review see 101). Binding sites for CRP at most promoters resemble either lac (centered at –61) or gal (centered at –41 bp). The basic difference between them is that in the first type (called class I) the CRP-binding site is entirely upstream of the promoter, whereas in the second type (called class II) the CRP-binding site overlaps the binding site for RNA polymerase. (The interactions at the ara promoter may be different.) CRP functions in different ways in different target operons | SECTION 3.11.5 © 2004. Virtual Text / www.ergito.com

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In both types of promoter, the CRP binding site is centered an integral number of turns of the double helix from the startpoint. This suggests that CRP is bound to the same face of DNA as RNA polymerase. However, the nature of the interaction between CRP and RNA polymerase is different at the two types of promoter. When the α subunit of RNA polymerase has a deletion in the C-terminal end, transcription appears normal except for the loss of ability to be activated by CRP. CRP has an "activating region", which consists of a small exposed loop of ~ 10 amino acids, that is required for activating both types of its promoters. The activating region is a small patch of amino acids that interacts directly with the α subunit of RNA polymerase to stimulate the enzyme (2565). At class I promoters, this interaction is sufficient. At class II promoters, a second interaction is also required, involving another region of CRP and the N-terminal region of the RNA polymerase α subunit (2564). Experiments using CRP dimers in which only one of the subunits has a functional transcription-activating region shows that, when CRP is bound at the lac promoter, only the activating region of the subunit nearer the startpoint is required, presumably because it touches RNA polymerase (2566). This offers an explanation for the lack of dependence on the orientation of the binding site: the dimeric structure of CRP ensures that one of the subunits is available to contact RNA polymerase, no matter which subunit binds to DNA and in which orientation. The effect upon RNA polymerase binding depends on the relative locations of the two proteins. At class I promoters, where CRP binds adjacent to the promoter, it increases the rate of initial binding to form a closed complex. At class II promoters, where CRP binds within the promoter, it increases the rate of transition from the closed to open complex. Last updated on 7-25-2002


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Reviews 99.

Botsford, J. L. and Harman, J. G. (1992). Cyclic AMP in prokaryotes. Microbiol. Rev. 56, 100-122.

101. Kolb, A. (1993). Transcriptional regulation by cAMP and its receptor protein. Annu. Rev. Biochem. 62, 749-795.

References 2564. Niu, W., Kim, Y., Tau, G., Heyduk, T., and Ebright, R. H. (1996). Transcription activation at class II CAP-dependent promoters: two interactions between CAP and RNA polymerase. Cell 87, 1123-1134. 2565. Zhou, Y., Merkel, T. J., and Ebright, R. H. (1994). Characterization of the activating region of E. coli catabolite gene activator protein (CAP). II. Role at Class I and class II CAP-dependent promoters. J. Mol. Biol. 243, 603-610. 2566. Zhou, Y., Busby, S., and Ebright, R. H. (1993). Identification of the functional subunit of a dimeric transcription activator protein by use of oriented heterodimers. Cell 73, 375-379.



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3.11.6 CRP bends DNA Key Concepts

• CRP introduces a 90° bend into DNA at its binding site.

The structure of the CRP-DNA complex is interesting: the DNA has a bend. Proteins may distort the double helical structure of DNA when they bind, and several regulator proteins induce a bend in the axis. Figure 11.9 illustrates a technique that can be used to measure the extent and location of a bend. A target sequence containing the site is cut with different restriction enzymes to generate a set of fragments all of the same length, but each containing the protein-binding site at a different location.

Figure 11.9 Gel electrophoresis can be used to analyze bending.

The fragments move at different speeds in an electrophoretic gel, depending on the position of the bend. (If there is no bend, all fragments move at the same rate.) The greatest impediment to motion, causing the lowest mobility, happens when the bend is in the center of the DNA fragment. The least impediment to motion, allowing the greatest mobility, happens when the bend is at one end. The results are analyzed by plotting mobility against the site of restriction cutting. The low point on the curve identifies the situation in which the restriction enzyme CRP bends DNA | SECTION 3.11.6 © 2004. Virtual Text / www.ergito.com

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has cut the sequence immediately adjacent to the site of bending. For the interaction of CRP with the lac promoter, this point lies at the center of dyad symmetry. The bend is quite severe, >90°, as illustrated in the model of Figure 11.10. There is therefore a dramatic change in the organization of the DNA double helix when CRP protein binds. The mechanism of bending is to introduce a sharp kink within the TGTGA consensus sequence. When there are inverted repeats of the consensus, the two kinks in each copy present in a palindrome cause the overall 90° bend. It is possible that the bend has some direct effect upon transcription, but it could be the case that it is needed simply to allow CRP to contact RNA polymerase at the promoter (518).

Figure 11.10 CRP bends DNA >90° around the center of symmetry.

Whatever the exact means by which CRP activates transcription at various promoters, it accomplishes the same general purpose: to turn off alternative metabolic pathways when they become unnecessary because the cell has an adequate supply of glucose. Again, this makes the point that coordinate control, of either negative or positive type, can extend over dispersed loci by repetition of binding sites for the regulator protein.

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References 518. Gaston, K. A. et al. (1990). Stringent spacing requirements for transcription activation by CRP. Cell 62, 733-743.


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3.11.7 The stringent response produces (p)ppGpp Key Terms Stringent response refers to the ability of a bacterium to shut down synthesis of tRNA and ribosomes in a poor-growth medium. An alarmone is a small molecule in bacteria that is produced as a result of stress and which acts to alter the state of gene expression. The unusual nucleotides ppGpp and pppGpp are examples. ppGpp is guanosine tetraphosphate. Diphosphate groups are attached to both the 5 ′ and 3 ′ positions. pppGpp is a guanosine pentaphosphate, with a triphosphate attached to the 5 ′ position and a diphosphate attached to the 3 ′ position. Key Concepts

• Poor growth conditions cause bacteria to produce the small molecule regulators ppGpp and pppGpp.

When bacteria find themselves in such poor growth conditions that they lack a sufficient supply of amino acids to sustain protein synthesis, they shut down a wide range of activities. This is called the stringent response. We can view it as a mechanism for surviving hard times: the bacterium husbands its resources by engaging in only the minimum of activities until nutrient conditions improve, when it reverses the response and again engages its full range of metabolic activities. The stringent response causes a massive (10-20×) reduction in the synthesis of rRNA and tRNA. This alone is sufficient to reduce the total amount of RNA synthesis to ~5-10% of its previous level. The synthesis of certain mRNAs is reduced, leading to an overall reduction of ~3× in mRNA synthesis. The rate of protein degradation is increased. Many metabolic adjustments occur, as seen in reduced synthesis of nucleotides, carbohydrates, lipids, etc. The stringent response causes the accumulation of two unusual nucleotides (sometimes called alarmones). ppGpp is guanosine tetraphosphate, with diphosphates attached to both 5 ′ and 3 ′ positions. pppGpp is guanosine pentaphosphate, with a 5 ′ triphosphate group and a 3 ′ diphosphate. These nucleotides are typical small-molecule effectors that function by binding to target proteins to alter their activities. Sometimes they are known collectively as (p)ppGpp (520; for review see 94). (p)ppGpp functions to regulate coordinately a large number of cellular activities. Its production is controlled in two ways. A drastic increase in (p)ppGpp is triggered by the stringent response. And there is also a general inverse correlation between (p)ppGpp levels and the bacterial growth rate, which is controlled by some unknown means. The stringent response produces (p)ppGpp | SECTION 3.11.7 © 2004. Virtual Text / www.ergito.com

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Reviews 94.

Cashel, M. and Rudd, K. E. (1987). The stringent response In E. coli and S. typhimurium. E. coli and S. typhimurium, 1410-1429.

References 520. Cashel, M. and Gallant, J. (1969). Two compounds implicated in the function of the RC gene of E. coli. Nature 221, 838-841.


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3.11.8 (p)ppGpp is produced by the ribosome Key Terms The idling reaction results in the production of pppGpp and ppGpp by ribosomes when an uncharged tRNA is present in the A site; this triggers the stringent response. Relaxed mutants of E. coli do not display the stringent response to starvation for amino acids (or other nutritional deprivation). The stringent factor is the protein RelA, which is associated with ribosomes. It synthesizes ppGpp and pppGpp when uncharged aminoacyl-tRNA enters the A site. Key Concepts

• The stringent factor RelA is a (p)ppGpp synthetase that is associated with ~5% of ribosomes.

• RelA is activated when the A site is occupied by an uncharged tRNA. • One (p)ppGpp is produced every time an uncharged tRNA enters the A site.

Deprivation of any one amino acid, or mutation to inactivate any aminoacyl-tRNA synthetase, is sufficient to initiate the stringent response. The trigger that sets the entire series of events in train is the presence of uncharged tRNA in the A site of the ribosome. Under normal conditions, of course, only aminoacyl-tRNA is placed in the A site by EF-Tu (see Molecular Biology 2.6.10 Elongation factor Tu loads aminoacyl-tRNA into the A site). But when there is no aminoacyl-tRNA available to respond to a particular codon, the uncharged tRNA becomes able to gain entry. Of course, this blocks any further progress by the ribosome; and it triggers an idling reaction. The components involved in producing (p)pGpp via the idling reaction have been identified through the existence of relaxed (rel) mutants. rel mutations abolish the stringent response, so that starvation for amino acids does not cause any reduction in stable RNA synthesis or alter any of the other reactions that are usually seen. The most common site of relaxed mutation lies in the gene relA, which codes for a protein called the stringent factor. This factor is associated with the ribosomes, although the amount is rather low – say, 10–6 M, it binds to gene 32 mRNA. At yet greater concentrations, it binds to other mRNA sequences, with a range of affinities.

Figure 11.20 Gene 32 protein binds to various substrates with different affinities, in the order single-stranded DNA, its own mRNA, and other mRNAs. Binding to its own mRNA prevents the level of p32 from rising >10-6 M.

These results imply that the level of p32 should be autoregulated to be 10 loci at the post-transcriptional level.

In bacteria, regulator RNAs are short molecules, collectively known as sRNAs; E. coli contains at least 17 different sRNAs (2945). Some of the sRNAs are general regulators that affect many target genes (for review see 3234). They function by base pairing with target RNAs (typically mRNAs) to control either their stability or function. An example of stability control is provided by the small antisense regulator RyhB, which regulates 6 mRNAs coding for proteins concerned with iron storage in E. coli. It base pairs with each of the target mRNAs to form double-stranded regions that are substrates for RNAase E. An interesting feature of the circuit is that the ribonuclease destroys the regulator RNA as well as the mRNA (4521). Oxidative stress provides an interesting example of a general control system in which RNA is the regulator. When exposed to reactive oxygen species, bacteria respond by inducing antioxidant defense genes. Hydrogen peroxide activates the transcription activator OxyR, which controls the expression of several inducible genes. One of these genes is oxyS, which codes for a small RNA. Figure 11.35 shows two salient features of the control of oxyS expression. In a wild-type bacterium under normal conditions, it is not expressed. The pair of gels on the left side of the figure shows that it is expressed at high levels in a mutant bacterium with a constitutively active oxyR gene. This identifies oxyS as a target for activation by oxyR. The pair of gels on the right side of the figure show that OxyS RNA is transcribed within 1 minute of exposure to hydrogen peroxide (2944).

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Figure 11.35 The gels on the left show that oxyS RNA is induced in an oxyR constitutive mutant. The gels on the right show that oxyS RNA is induced within 1 minute of adding hydrogen peroxide to a wild-type culture. Photograph kindly provided by Gisela Storz (see 2944).

The OxyS RNA is a short sequence (109 nucleotides) that does not code for protein. It is a trans-acting regulator that affects gene expression at post-transcriptional levels. It has >10 target loci; at some of them, it activates expression, at others it represses expression. Figure 11.36shows the mechanism of repression of one target, the FlhA mRNA. Three stem-loop structures protrude in the secondary structure of OxyR mRNA, and the loop close to the 3 ′ terminus is complementary to a sequence just preceding the initiation codon of FlhA mRNA. Base pairing between OxyS RNA and FlhA RNA prevents the ribosome from binding to the initiation codon, and therefore represses translation (2943). There is also a second pairing interaction that involves a sequence within the coding region of FlhA.


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Figure 11.36 oxyS RNA inhibits translation of flhA mRNA by base pairing with a sequence just upstream of the AUG initiation codon.

Another target for oxyS is rpoS, the gene coding for an alternative sigma factor (which activates a general stress response). By inhibiting production of the sigma factor, oxyS ensures that the specific response to oxidative stress does not trigger the response that is appropriate for other stress conditions. The rpoS gene is also regulated by two other sRNAs (DsrA and RprA), which activate it. These three sRNAs appear to be global regulators that coordinate responses to various environmental conditions. The actions of all three sRNAs are assisted by an RNA-binding protein called Hfq. The Hfq protein was originally identified as a bacterial host factor needed for replication of the RNA bacteriophage Q β. It is related to the Sm proteins of eukaryotes that bind to many of the snRNAs (small nuclear RNAs) that have regulatory roles in gene expression (see Molecular Biology 5.24.5 snRNAs are required for splicing) (2941; 2942). Mutations in its gene have many effects, identifying it as a pleiotropic protein. Hfq binds to many of the sRNAs of E. coli. It increases the effectiveness of OxyS RNA by enhancing its ability to bind to its target mRNAs. The effect of Hfq is probably mediated by causing a small change in the secondary structure of OxyS RNA that improves the exposure of the single-stranded sequences that pair with the target mRNAs. Last updated on January 7, 2004


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Reviews 3234. Gottesman, S. (2002). Stealth regulation: biological circuits with small RNA switches. Genes Dev. 16, 2829-2842.

References 2941. Zhang, A., Wassarman, K. M., Ortega, J., Steven, A. C., and Storz, G. (2002). The Sm-like Hfq protein increases OxyS RNA interaction with target mRNAs. Mol. Cell 9, 11-22. 2942. Moller, T., Franch, T., Hojrup, P., Keene, D. R., Bachinger, H. P., Brennan, R. G., and Valentin-Hansen, P. (2002). Hfq: a bacterial Sm-like protein that mediates RNA-RNA interaction. Mol. Cell 9, 23-30. 2943. Altuvia, S., Zhang, A., Argaman, L., Tiwari, A., and Storz, G. (1998). The E. coli OxyS regulatory RNA represses fhlA translation by blocking ribosome binding. EMBO J. 17, 6069-6075. 2944. Altuvia, S., Weinstein-Fischer, D., Zhang, A., Postow, L., and Storz, G. (1997). A small, stable RNA induced by oxidative stress: role as a pleiotropic regulator and antimutator. Cell 90, 43-53. 2945. Wassarman, K. M., Repoila, F., Rosenow, C., Storz, G., and Gottesman, S. (2001). Identification of novel small RNAs using comparative genomics and microarrays. Genes Dev. 15, 1637-1651. 4521. Massé, E., Escorcia, F. E., and Gottesman, S. (2003). Coupled degradation of a small regulatory RNA and its mRNA targets in Escherichia coli. Genes Dev. 17, 2374-2383.



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3.11.21 MicroRNAs are regulators in many eukaryotes Key Terms MicroRNAs are very short RNAs that may regulate gene expression. Key Concepts

• Animal and plant genomes code for many short (~22 base) RNA molecules, called microRNAs.

• MicroRNAs regulate gene expression by base pairing with complementary sequences in target mRNAs.

Very small RNAs are gene regulators in many eukaryotes. The first example was discovered in the nematode C. elegans as the result of the interaction between the regulator gene lin4 and its target gene, lin14. Figure 11.37 illustrates the behavior of this regulatory system. The lin14 target gene regulates larval development. Expression of lin14 is controlled by lin4, which codes for a small transcript of 22 nucleotides (2195; 2196). The lin4 transcripts are complementary to a 10 base sequence that is repeated 7 times in the 3 ′ nontranslated region of lin14. Expression of lin4 represses expression of lin14 post-transcriptionally, most likely because the base pairing reaction between the two RNAs leads to degradation of the mRNA. This system is especially interesting in implicating the 3 ′ end as a site for regulation.

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Figure 11.37 lin4 RNA regulates expression of lin14 by binding to the 3 ′ nontranslated region.

The lin4 RNA is an example of a microRNA (miRNA). There are ~80 genes in the C. elegans genome coding for microRNAs of 21-24 nucleotide length (2193; 2194). They have varying patterns of expression during development and are likely to be regulators of gene expression. Many of the microRNAs of C. elegans are contained in a large (15S) ribonucleoprotein particle (2510). Many of the C. elegans microRNAs have homologues in mammals, so the mechanism may be widespread. They are also found in plants. Of 16 microRNAs in Arabidopsis, 8 are completely conserved in rice, suggesting widespread conservation of this regulatory mechanism (3027). The mechanism of production of the microRNAs is also widely conserved. In the example of lin4, the gene is transcribed into a transcript that forms a double-stranded region that becomes a target for a nuclease called Dicer. This has an N-terminal helicase activity, enabling it to unwind the double-stranded region, and two nuclease domains that are related to the bacterial ribonuclease III. Related enzymes are found in flies, worms, and plants (3028; 3029; 3027). Cleavage of the initial transcript generates the active microRNA. Interfering with the enzyme activity blocks the production of microRNAs and causes developmental defects. Last updated on 10-16-2002 MicroRNAs are regulators in many eukaryotes | SECTION 3.11.21 © 2004. Virtual Text / www.ergito.com

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References 2193. Lau, N. C., Lim, l. e. E. P., Weinstein, E. G., and Bartel, d. a. V. P. (2001). An abundant class of tiny RNAs with probable regulatory roles in C. elegans. Science 294, 858-862. 2194. Lee, R. C. and Ambros, V. (2001). An extensive class of small RNAs in C. eleganss. Science 294, 862-864. 2195. Lee, R. C., Feinbaum, R. L., and Ambros, V. (1993). The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell 75, 843-854. 2196. Wightman, B., Ha, I., and Ruvkun, G. (1993). Posttranscriptional regulation of the heterochronic gene lin-14 by lin-4 mediates temporal pattern formation in C. elegans. Cell 75, 855-862. 2510. Mourelatos, Z., Dostie, J., Paushkin, S., Sharma, A., Charroux, B., Abel, L., Rappsilber, J., Mann, M., and Dreyfuss, G. (2002). miRNPs: a novel class of ribonucleoproteins containing numerous microRNAs. Genes Dev. 16, 720-728. 3027. Reinhart,B. J., Weinstein, E. G., Rhoades, M. W., Bartel, B., and Bartel, D. P. (2002). MicroRNAs in plants. Genes Dev. 16, 1616-1626. 3028. Bernstein, E., Caudy, A. A., Hammond, S. M., and Hannon, G. J. (2001). Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363-366. 3029. Ketting, R. F., Fischer, S. E., Bernstein, E., Sijen, T., Hannon, G. J., and Plasterk, R. H. (2001). Dicer functions in RNA interference and in synthesis of small RNA involved in developmental timing in C. elegans. Genes Dev. 15, 2654-2659.


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3.11.22 RNA interference is related to gene silencing Key Terms RNA interference (RNAi) describes the technique in which double-strand RNA is introduced into cells to eliminate or reduce the activity of a target gene. It is caused by using sequences complementary to the double-stranded RNA sequences to trigger degradation of the mRNA of the gene. RNA silencing describes the ability of a dsRNA to suppress expression of the corresponding gene systemically in a plant. Cosuppression describes the ability of a transgene (usually in plants) to inhibit expression of the corresponding endogenous gene. Key Concepts

• RNA interference triggers degradation of mRNAs complementary to either strand of a short dsRNA.

• dsRNA may cause silencing of host genes.

The regulation of mRNAs by microRNAs is mimicked by the phenomenon of RNA interference (RNAi). This was discovered when it was observed that antisense and sense RNAs can be equally effective in inhibiting gene expression (for review see 2077). The reason is that preparations of either type of (supposedly) single-stranded RNA are actually contaminated by small amounts of double-stranded RNA (1189). Work with an in vitro system shows that the dsRNA is degraded by ATP-dependent cleavage to give oligonucleotides of 21-23 bases. The short RNA is sometimes called siRNA (short interfering RNA). Figure 11.38 shows that the mechanism of cleavage involves making breaks relative to each 3 ′ end of a long dsRNA to generate siRNA fragments with short (2 base) protruding 3 ′ ends. The same enzyme (Dicer) that generates microRNAs is responsible for the cleavage.

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Figure 11.38 siRNA that mediates RNA interference is generated by cleaving dsRNA into smaller fragments. The cleavage reaction occurs 21-23 nucleotides from a 3 ′ end. The siRNA product has protruding bases on its 3 ′ ends.

RNAi occurs post-transcriptionally when an siRNA induces degradation of a complementary mRNA (1190; 1191). Figure 11.39 suggests that the siRNA may provide a template that directs a nuclease to degrade mRNAs that are complementary to one or both strands, perhaps by a process in which the mRNA pairs with the fragments (1192). It is likely that a helicase is required to assist the pairing reaction. The siRNA directs cleavage of the mRNA in the middle of the paired segment. These reactions occur within a ribonucleoprotein complex called RISC (RNA-induced silencing complex) (for review see 2511).

Figure 11.39 RNAi occurs when a dsRNA is cleaved into fragments that direct cleavage of the corresponding mRNA. RNA interference is related to gene silencing | SECTION 3.11.22 © 2004. Virtual Text / www.ergito.com

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RNAi has become a powerful technique for ablating the expression of a specific target gene in invertebrate cells, especially in C. elegans and D. melanogaster. However, the technique has been limited in mammalian cells, which have a more generalized response to dsRNA of shutting down protein synthesis and degrading mRNA. Figure 11.40 shows that this happens because of two reactions. The dsRNA activates the enzyme PKR, which inactivates the translation initiation factor eIF2a by phosphorylating it. And it activates 2 ′ 5 ′ oligoadenylate synthetase, whose product activates RNAase L, which degrades all mRNAs. However, it turns out that these reactions require dsRNA that is longer than 26 nucleotides. If shorter dsRNA (21-23 nucleotides) is introduced into mammalian cells, it triggers the specific degradation of complementary RNAs just as with the RNAi technique in worms and flies (1905). With this advance, it seems likely that RNAi will become the universal mechanism of choice for turning off the expression of a specific gene.

Figure 11.40 dsRNA inhibits protein synthesis and triggers degradation of all mRNA in mammalian cells as well as having sequence-specific effects.

As an example of the progress being made with the technique, it has been possible to use RNAi for a systematic analysis of gene expression in C. elegans. Loss of function phenotypes can be generated by feeding worms with bacteria expressing a dsRNA that is homologous to a target gene. By making a library of bacteria in which each bacterium expresses a dsRNA corresponding to a different gene, worms have been screened for the effects of knocking out most (86%) of the genes (3323). RNA interference is related to natural processes in which gene expression is silenced. Plants and fungi show RNA silencing (sometimes called post-transcriptional gene silencing) in which dsRNA inhibits expression of a gene (for review see 3242). The most common source of the RNA is a replicating virus. This mechanism may have evolved as a defense against viral infection. When a virus infects a plant cell, the formation of dsRNA triggers the suppression of expression from the plant genome (1396). RNA silencing has the further remarkable feature that it is not limited to the RNA interference is related to gene silencing | SECTION 3.11.22 © 2004. Virtual Text / www.ergito.com

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cell in which the viral infection occurs: it can spread throughout the plant systemically (1397). Presumably the propagation of the signal involves passage of RNA or fragments of RNA. It may require some of the same features that are involved in movement of the virus itself. It is possible that RNA silencing involves an amplification of the signal by an RNA-dependent RNA synthesis process in which a novel polymerase uses the siRNA as a primer to synthesize more RNA on a template of complementary RNA (for review see 2490). A related process is the phenomenon of cosuppression transgene causes the corresponding endogenous gene to largely characterized in plants (1193). The implication make both antisense and sense RNA copies, and this endogenous gene.

in which introduction of a be silenced. This has been is that the transgene must inhibits expression of the

Silencing takes place by RNA-RNA interactions. It is also possible that dsRNA may inhibit gene expression by interacting with the DNA. If a DNA copy of a viroid RNA sequence is inserted into a plant genome, it becomes methylated when the viroid RNA replicates (2149). This suggests that the RNA sequence could be inducing methylation of the DNA sequence. Similar targeting of methylation of DNA corresponding to sequences represented in dsRNA has been detected in plant cells (2150). Methylation of DNA is associated with repression of transcription, so this could be another means of silencing genes represented in dsRNA (see Molecular Biology 5.21.18 Gene expression is associated with demethylation). Nothing is known about the mechanism (for review see 2077; 2078). Last updated on 10-16-2002


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Reviews 2077. Sharp, P. A. (2001). RNA interference--2001. Genes Dev. 15, 485-490. 2078. Matzke, M., Matzke, A. J., and Kooter, J. M. (2001). RNA: guiding gene silencing. Science 293, 1080-1083. 2490. Ahlquist, P. (2002). RNA-Dependent RNA Polymerases, Viruses, and RNA Silencing. Science 296, 1270-1273. 2511. Schwartz, D. S. and Zamore, P. D. (2002). Why do miRNAs live in the miRNP? Genes Dev. 16, 1025-1031. 3242. Tijsterman, M., Ketting, R. F., and Plasterk, R. H. (2002). The genetics of RNA silencing. Annu. Rev. Genet. 36, 489-519.

References 1189. Fire, A.Xu, S.Montgomery, M. K.Kostas, S. A.Driver, and S. E.Mello, C. C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806-811. 1190. Montgomery, M. K., Xu, S., and Fire, A. (1998). RNA as a target of double-stranded RNA-mediated genetic interference in C. elegans. Proc. Natl. Acad. Sci. USA 95, 15502-15507. 1191. Ngo, H., Tschudi, C., Gull, K., and Ullu, E. (1998). Double-stranded RNA induces mRNA degradation in Trypanosoma brucei. Proc. Natl. Acad. Sci. USA 95, 14687-14692. 1192. Zamore, P. D., Tuschl, T., Sharp, P. A., and Bartel, D. P. (2000). RNAi: double-stranded RNA directs the ATP-dependent cleavage of mRNA at 21 to 23 nucleotide intervals. Cell 101, 25-33. 1193. Waterhouse, P. M., Graham, M. W., and Wang, M. B. (1998). Virus resistance and gene silencing in plants can be induced by simultaneous expression of sense and antisense RNA. Proc. Natl. Acad. Sci. USA 95, 13959-13964. 1396. Hamilton, A. J. and Baulcombe, D. C. (1999). A species of small antisense RNA in posttranscriptional gene silencing in plants. Science 286, 950-952. 1397. Voinnet, O., Pinto, Y. M., and Baulcombe, D. C. (1999). Suppression of gene silencing: a general strategy used by diverse DNA and RNA viruses of plants. Proc. Natl. Acad. Sci. USA 96, 14147-14152. 1905. Elbashir, S. M., Harborth, J., Lendeckel, W., Yalcin, A., Weber, K., and Tuschl, T. (2001). Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411, 494-498. 2149. Wassenegger, M., Heimes, S., Riedel, L., and Sanger, H. L. (1994). RNA-directed de novo methylation of genomic sequences in plants. Cell 76, 567-576. 2150. Mette, M. F., Aufsatz, W., van der Winden, J., Matzke, M. A., and Matzke, A. J. (2000). Transcriptional silencing and promoter methylation triggered by double-stranded RNA. EMBO J. 19, 5194-5201. 3323. Kamath, R. S., Fraser, A. G., Dong, Y., Poulin, G., Durbin, R., Gotta, M., Kanapin, A., Le Bot, N., Moreno, S., Sohrmann, M., Welchman, D. P., Zipperlen, P., and Ahringer, J. (2003). Systematic functional analysis of the C. elegans genome using RNAi. Nature 421, 231-237.



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3.11.23 Summary Gene expression can be regulated positively by factors that activate a gene or negatively by factors that repress a gene. The first and most common level of control is at the initiation of transcription, but termination of transcription may be also be controlled. Translation may be controlled by regulators that interact with mRNA. The regulatory products may be proteins, which often are controlled by allosteric interactions in response to the environment, or RNAs, which function by base pairing with the target RNA to change its secondary structure. Regulatory networks can be created by linking regulators so that the production or activity of one regulator is controlled by another. Bacteria respond to the supply of glucose by repressing the production of the enzyme systems that catabolize alternative carbon sources. Inducer exclusion is a major component of the response, and works by inhibiting the uptake of the other sugars into the bacterium, with the result that the operons coding for their enzyme systems fail to be turned on. Increase in glucose levels also may lead to a reduction in the level of the small nucleotide cyclic AMP, although this is now controversial. Some promoters cannot be recognized by RNA polymerase (or are recognized only poorly) unless a specific activator protein is present. Activator proteins also may be regulated by small molecules. The CRP activator becomes able to bind to target sequences in the presence of cyclic AMP. All promoters that respond to CRP have at least one copy of the target sequence. Binding of CRP to its target involves bending DNA. Direct contact between one subunit of CRP and RNA polymerase is required to activate transcription. A common means for controlling translation is for a regulator protein to bind to a site on the mRNA that overlaps the ribosome binding site at the initiation codon. This prevents ribosomes from initiating translation. RegA of T4 is a general regulator that functions on several target mRNAs at the level of translation. Most proteins that repress translation possess this capacity in addition to other functional roles; in particular, translation is controlled in some cases of autogenous regulation, when a gene product regulates expression of the operon containing its own gene. The level of protein synthesis itself provides an important coordinating signal. Deficiency in aminoacyl-tRNA causes an idling reaction on the ribosome, which leads to the synthesis of the unusual nucleotide ppGpp. This is an effector that inhibits initiation of transcription at certain promoters; it also has a general effect in inhibiting elongation on all templates. Attenuation is a mechanism that relies on regulation of termination to control transcription through bacterial operons. It is commonly used in operons that code for enzymes involved in biosynthesis of an amino acid. The polycistronic mRNA of the operon starts with a sequence that can form alternative secondary structures. One of the structures has a hairpin loop that provides an intrinsic terminator upstream of the structural genes; the alternative structure lacks the hairpin. Various types of interaction can be used to determine whether the hairpin forms. One is for a protein Summary | SECTION 3.11.23 © 2004. Virtual Text / www.ergito.com

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to bind to the mRNA to prevent formation of the alternative structure. In the trp operon of B. subtilis, the TRAP protein has this function; it is controlled by the anti-TRAP protein, whose production in turn is controlled by the level of uncharged aminoacyl-tRNATrp. In the trp operon of E. coli, the choice of which structure forms is controlled by the progress of translation through a short leader sequence that includes codons for the amino acid(s) that are the product of the system. In the presence of aminoacyl-tRNA bearing such amino acid(s), ribosomes translate the leader peptide, allowing a secondary structure to form that supports termination. In the absence of this aminoacyl-tRNA, the ribosome stalls, resulting in a new secondary structure in which the hairpin needed for termination cannot form. The supply of aminoacyl-tRNA therefore (inversely) controls amino acid biosynthesis. Small regulator RNAs are found in both bacteria and eukaryotes. E. coli has ~17 sRNA species. The oxyS sRNA controls about 10 target loci at the post-transcriptional level; some of them are repressed, and others are activated. Repression is caused when the sRNA binds to a target mRNA to form a duplex region that includes the ribosome-binding site. MicroRNAs are ~22 bases long and are produced in many eukaryotes by cleavage of a longer transcript. They function by base pairing with target mRNAs to form duplex regions that are susceptible to cleavage by endonucleases. The degradation of the mRNA prevents its expression. The technique of RNA interference is becoming the method of choice for inactivating eukaryotic genes. It uses the introduction of short dsRNA sequences with one strand complementary to the target RNA, and it works by inducing degradation of the targets. This may be related to a natural defense system in plants called RNA silencing. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.11.23


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PHAGE STRATEGIES

3.12.1 Introduction Key Terms Lytic infection of a bacterium by a phage ends in the destruction of the bacterium with release of progeny phage. Lysis describes the death of bacteria at the end of a phage infective cycle when they burst open to release the progeny of an infecting phage (because phage enzymes disrupt the bacterium's cytoplasmic membrane or cell wall). The same term also applies to eukaryotic cells; for example, when infected cells are attacked by the immune system. Prophage is a phage genome covalently integrated as a linear part of the bacterial chromosome. Lysogeny describes the ability of a phage to survive in a bacterium as a stable prophage component of the bacterial genome. Integration of viral or another DNA sequence describes its insertion into a host genome as a region covalently linked on either side to the host sequences. Induction of prophage describes its entry into the lytic (infective) cycle as a result of destruction of the lysogenic repressor, which leads to excision of free phage DNA from the bacterial chromosome. The excision of phage or episome or other sequence describes its release from the host chromosome as an autonomous DNA molecule. A plasmid is a circular, extrachromosomal DNA. It is autonomous and can replicate itself. An extrachromosomal genome in a bacterium is a self-replicating set of genes that is not part of the bacterial chromosome. In many cases, the genes are necessary for bacterial growth under certain environmental conditions. An episome is a plasmid able to integrate into bacterial DNA. Immunity in phages refers to the ability of a prophage to prevent another phage of the same type from infecting a cell. It results from the synthesis of phage repressor by the prophage genome. Immunity in plasmids describes the ability of a plasmid to prevent another of the same type from becoming established in a cell. It results usually from interference with the ability to replicate.

Some phages have only a single strategy for survival. On infecting a susceptible host, they subvert its functions to the purpose of producing a large number of progeny phage particles. As the result of this lytic infection, the host bacterium dies. In the typical lytic cycle, the phage DNA (or RNA) enters the host bacterium, its genes are transcribed in a set order, the phage genetic material is replicated, and the protein components of the phage particle are produced. Finally, the host bacterium is broken open (lysed) to release the assembled progeny particles by the process of lysis. Introduction | SECTION 3.12.1 © 2004. Virtual Text / www.ergito.com

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Other phages have a dual existence. They are able to perpetuate themselves via the same sort of lytic cycle in what amounts to an open strategy for producing as many copies of the phage as rapidly as possible. But they also have an alternative form of existence, in which the phage genome is present in the bacterium in a latent form known as prophage. This form of propagation is called lysogeny. In a lysogenic bacterium, the prophage is inserted into the bacterial genome, and is inherited in the same way as bacterial genes. The process by which it is converted from an independent phage genome into a prophage that is a linear part of the bacterial genome is described as integration. By virtue of its possession of a prophage, a lysogenic bacterium has immunity against infection by further phage particles of the same type. Immunity is established by a single integrated prophage, so usually a bacterial genome contains only one copy of a prophage of any particular type. Transitions occur between the lysogenic and lytic modes of existence. Figure 12.1 shows that when a phage produced by a lytic cycle enters a new bacterial host cell, it either repeats the lytic cycle or enters the lysogenic state. The outcome depends on the conditions of infection and the genotypes of phage and bacterium.


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Figure 12.1 Lytic development involves the reproduction of phage particles with destruction of the host bacterium, but lysogenic existence allows the phage genome to be carried as part of the bacterial genetic information.

A prophage is freed from the restrictions of lysogeny by the process called induction. First the phage DNA is released from the bacterial chromosome by excision; then the free DNA proceeds through the lytic pathway. The alternative forms in which these phages are propagated are determined by the regulation of transcription. Lysogeny is maintained by the interaction of a phage repressor with an operator. The lytic cycle requires a cascade of transcriptional controls. And the transition between the two life-styles is accomplished by the establishment of repression (lytic cycle to lysogeny) or by the relief of repression (induction of lysogen to lytic phage). Another type of existence within bacteria is represented by plasmids. These are autonomous units that exist in the cell as extrachromosomal genomes. Plasmids are self-replicating circular molecules of DNA that are maintained in the cell in a stable Introduction | SECTION 3.12.1 © 2004. Virtual Text / www.ergito.com

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and characteristic number of copies; that is, the number remains constant from generation to generation. Some plasmids also have alternative life-styles. They can exist either in the autonomous extrachromosomal state; or they can be inserted into the bacterial chromosome, and then are carried as part of it like any other sequence. Such units are properly called episomes (but the terms "plasmid" and "episome" are sometimes used loosely as though interchangeable). Like lysogenic phages, plasmids and episomes maintain a selfish possession of their bacterium and often make it impossible for another element of the same type to become established. This effect also is called immunity, although the basis for plasmid immunity is different from lysogenic immunity. (We discuss the control of plasmid perpetuation in Molecular Biology 4.13 The replicon.) Figure 12.2 summarizes the types of genetic units that can be propagated in bacteria as independent genomes. Lytic phages may have genomes of any type of nucleic acid; they transfer between cells by release of infective particles. Lysogenic phages have double-stranded DNA genomes, as do plasmids and episomes. Some plasmids and episomes transfer between cells by a conjugative process (involving direct contact between donor and recipient cells). A feature of the transfer process in both cases is that on occasion some bacterial host genes are transferred with the phage or plasmid DNA, so these events play a role in allowing exchange of genetic information between bacteria.

Figure 12.2 Several types of independent genetic units exist in bacteria. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.1


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PHAGE STRATEGIES

3.12.2 Lytic development is divided into two periods Key Terms Early infection is the part of the phage lytic cycle between entry and replication of the phage DNA. During this time, the phage synthesizes the enzymes needed to replicate its DNA. Late infection is the part of the phage lytic cycle from DNA replication to lysis of the cell. During this time, the DNA is replicated and structural components of the phage particle are synthesized. Key Concepts

• A phage infective cycle is divided into the early period (before replication) and the late period (after the onset of replication).

• A phage infection generates a pool of progeny phage genomes that replicate and recombine.

Phage genomes of necessity are small. As with all viruses, they are restricted by the need to package the nucleic acid within the protein coat. This limitation dictates many of the viral strategies for reproduction. Typically a virus takes over the apparatus of the host cell, which then replicates and expresses phage genes instead of the bacterial genes. Usually the phage includes genes whose function is to ensure preferential replication of phage DNA. These genes are concerned with the initiation of replication and may even include a new DNA polymerase. Changes are introduced in the capacity of the host cell to engage in transcription. They involve replacing the RNA polymerase or modifying its capacity for initiation or termination. The result is always the same: phage mRNAs are preferentially transcribed. So far as protein synthesis is concerned, usually the phage is content to use the host apparatus, redirecting its activities principally by replacing bacterial mRNA with phage mRNA. Lytic development is accomplished by a pathway in which the phage genes are expressed in a particular order. This ensures that the right amount of each component is present at the appropriate time. The cycle can be divided into the two general parts illustrated in Figure 12.3:

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Figure 12.3 Lytic development takes place by producing phage genomes and protein particles that are assembled into progeny phages.

• Early infection describes the period from entry of the DNA to the start of its replication. • Late infection defines the period from the start of replication to the final step of lysing the bacterial cell to release progeny phage particles. The early phase is devoted to the production of enzymes involved in the reproduction of DNA. These include the enzymes concerned with DNA synthesis, recombination, and sometimes modification. Their activities cause a pool of phage genomes to Lytic development is divided into two periods | SECTION 3.12.2 © 2004. Virtual Text / www.ergito.com

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accumulate. In this pool, genomes are continually replicating and recombining, so that the events of a single lytic cycle concern a population of phage genomes. During the late phase, the protein components of the phage particle are synthesized. Often many different proteins are needed to make up head and tail structures, so the largest part of the phage genome consists of late functions. In addition to the structural proteins, "assembly proteins" are needed to help construct the particle, although they are not themselves incorporated into it. By the time the structural components are assembling into heads and tails, replication of DNA has reached its maximum rate. The genomes then are inserted into the empty protein heads, tails are added, and the host cell is lysed to allow release of new viral particles. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.2

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PHAGE STRATEGIES

3.12.3 Lytic development is controlled by a cascade Key Terms A cascade is a sequence of events, each of which is stimulated by the previous one. In transcriptional regulation, as seen in sporulation and phage lytic development, it means that regulation is divided into stages, and at each stage, one of the genes that are expressed codes for a regulator needed to express the genes of the next stage. Early genes are transcribed before the replication of phage DNA. They code for regulators and other proteins needed for later stages of infection. Immediate early phage genes in phage lambda are equivalent to the early class of other phages. They are transcribed immediately upon infection by the host RNA polymerase. Delayed early genes in phage lambda are equivalent to the middle genes of other phages. They cannot be transcribed until regulator protein(s) coded by the immediate early genes have been synthesized. Middle genes are phage genes that are regulated by the proteins coded by early genes. Some proteins coded by middle genes catalyze replication of the phage DNA; others regulate the expression of a later set of genes. Late genes are transcribed when phage DNA is being replicated. They code for components of the phage particle. Key Concepts

• The early genes transcribed by host RNA polymerase following infection include or comprise regulators required for expression of the middle set of phage genes.

• The middle group of genes include regulators to transcribe the late genes. • This results in the ordered expression of groups of genes during phage infection.

The organization of the phage genetic map often reflects the sequence of lytic development. The concept of the operon is taken to somewhat of an extreme, in which the genes coding for proteins with related functions are clustered to allow their control with the maximum economy. This allows the pathway of lytic development to be controlled with a small number of regulatory switches. The lytic cycle is under positive control, so that each group of phage genes can be expressed only when an appropriate signal is given. Figure 12.4 shows that the regulatory genes function in a cascade, in which a gene expressed at one stage is necessary for synthesis of the genes that are expressed at the next stage.

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Figure 12.4 Phage lytic development proceeds by a regulatory cascade, in which a gene product at each stage is needed for expression of the genes at the next stage.

The first stage of gene expression necessarily relies on the transcription apparatus of the host cell. Usually only a few genes are expressed at this stage. Their promoters are indistinguishable from those of host genes. The name of this class of genes depends on the phage. In most cases, they are known as the early genes. In phage lambda, they are given the evocative description of immediate early. Irrespective of the name, they constitute only a preliminary, representing just the initial part of the early period. Sometimes they are exclusively occupied with the transition to the next period. At all events, one of these genes always codes for a protein that is necessary for transcription of the next class of genes. This second class of genes is known variously as the delayed early or middle gene group. Its expression typically starts as soon as the regulator protein coded by the early gene(s) is available. Depending on the nature of the control circuit, the initial set of early genes may or may not continue to be expressed at this stage. If control is at initiation, the two events are independent (see Figure 12.5), and early genes can be switched off when middle genes are transcribed. If control is at termination, the early genes must continue to be expressed (see Figure 12.6). Often the expression of Lytic development is controlled by a cascade | SECTION 3.12.3 © 2004. Virtual Text / www.ergito.com

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host genes is reduced. Together the two sets of early genes account for all necessary phage functions except those needed to assemble the particle coat itself and to lyse the cell. When the replication of phage DNA begins, it is time for the late genes to be expressed. Their transcription at this stage usually is arranged by embedding a further regulator gene within the previous (delayed early or middle) set of genes. This regulator may be another antitermination factor (as in lambda) or it may be another sigma factor (as in SPO1). A lytic infection often falls into three stages, as shown in Figure 12.4. The first stage consists of early genes transcribed by host RNA polymerase (sometimes the regulators are the only products at this stage). The second stage consists of genes transcribed under direction of the regulator produced in the first stage (most of these genes code for enzymes needed for replication of phage DNA). The final stage consists of genes for phage components, transcribed under direction of a regulator synthesized in the second stage. The use of these successive controls, in which each set of genes contains a regulator that is necessary for expression of the next set, creates a cascade in which groups of genes are turned on (and sometimes off) at particular times.The means used to construct each phage cascade are different, but the results are similar, as the following sections show. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.3

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PHAGE STRATEGIES

3.12.4 Two types of regulatory event control the lytic cascade Key Concepts

• Regulator proteins used in phage cascades may sponsor initiation at new (phage)

promoters or cause the host polymerase to read through transcription terminators.

At every stage of phage expression, one or more of the active genes is a regulator that is needed for the subsequent stage. The regulator may take the form of a new RNA polymerase, a sigma factor that redirects the specificity of the host RNA polymerase (see Molecular Biology 3.9.18 Sigma factors may be organized into cascades ), or an antitermination factor that allows it to read a new group of genes (see Molecular Biology 3.9.23 Antitermination is a regulatory event). The next two figures compare the use of switching at initiation or termination to control gene expression. One mechanism for recognizing new phage promoters is to replace the sigma factor of the host enzyme with another factor that redirects its specificity in initiation (see Figure 9.31). An alternative mechanism is to synthesize a new phage RNA polymerase. In either case, the critical feature that distinguishes the new set of genes is their possession of different promoters from those originally recognized by host RNA polymerase.Figure 12.5 shows that the two sets of transcripts are independent; as a consequence, early gene expression can cease after the new sigma factor or polymerase has been produced.

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Figure 12.5 Control at initiation utilizes independent transcription units, each with its own promoter and terminator, which produce independent mRNAs. The transcription units need not be located near one another.

Antitermination provides an alternative mechanism for phages to control the switch from early genes to the next stage of expression. The use of antitermination depends on a particular arrangement of genes. Figure 12.6 shows that the early genes lie adjacent to the genes that are to be expressed next, but are separated from them by terminator sites. If termination is prevented at these sites, the polymerase reads through into the genes on the other side. So in antitermination, the same promoters continue to be recognized by RNA polymerase. So the new genes are expressed only by extending the RNA chain to form molecules that contain the early gene sequences at the 5 ′ end and the new gene sequences at the 3 ′ end. Since the two types of sequence remain linked, early gene expression inevitably continues (for review see 102).


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Figure 12.6 Control at termination requires adjacent units, so that transcription can read from the first gene into the next gene. This produces a single mRNA that contains both sets of genes.

The regulator gene that controls the switch from immediate early to delayed early expression in phage lambda is identified by mutations in gene N that can transcribe only the immediate early genes; they proceed no further into the infective cycle (see Figure 9.53). The same effect is seen when gene 28 of phage SPO1 is mutated to prevent the production of σgp28 (see Figure 9.40). From the genetic point of view, the mechanisms of new initiation and antitermination are similar. Both are positive controls in which an early gene product must be made by the phage in order to express the next set of genes. By employing either sigma factors or antitermination proteins with different specificities, a cascade for gene expression can be constructed


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Reviews 102. Greenblatt, J., Nodwell, J. R., and Mason, S. W. (1993). Transcriptional antitermination. Nature 364, 401-406.



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PHAGE STRATEGIES

3.12.5 The T7 and T4 genomes show functional clustering Key Concepts

• Genes concerned with related functions are often clustered. • Phages T7 and T4 are examples of regulatory cascades in which phage infection is divided into three periods.

The genome of phage T7 has three classes of genes, each constituting a group of adjacent loci. As Figure 12.7 shows, the class I genes are the immediate early type, expressed by host RNA polymerase as soon as the phage DNA enters the cell. Among the products of these genes are a phage RNA polymerase and enzymes that interfere with host gene expression. The phage RNA polymerase is responsible for expressing the class II genes (concerned principally with DNA synthesis functions) and the class III genes (concerned with assembling the mature phage particle).

Figure 12.7 Phage T7 contains three classes of genes that are expressed sequentially. The genome is ~38 kb.

T4 has one of the larger phage genomes (165 kb), organized with extensive functional grouping of genes. Figure 12.8 presents the genetic map. Essential genes are numbered: a mutation in any one of these loci prevents successful completion of the lytic cycle. Nonessential genes are indicated by three-letter abbreviations. (They are defined as nonessential under the usual conditions of infection. We do not really understand the inclusion of many nonessential genes, but presumably they confer a selective advantage in some of T4's habitats. In smaller phage genomes, most or all The T7 and T4 genomes show functional clustering | SECTION 3.12.5 © 2004. Virtual Text / www.ergito.com

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of the genes are essential.)

Figure 12.8 The map of T4 is circular. There is extensive clustering of genes coding for components of the phage and processes such as DNA replication, but there is also dispersion of genes coding for a variety of enzymatic and other functions. Essential genes are indicated by numbers. Nonessential genes are identified by letters. Only some representative T4 genes are shown on the map.

There are three phases of gene expression. A summary of the functions of the genes expressed at each stage is given in Figure 12.9. The early genes are transcribed by host RNA polymerase. The middle genes are also transcribed by host RNA polymerase, but two phage-encoded products, MotA and AsiA, are also required. The middle promoters lack a consensus –30 sequence, and instead have a binding sequence for MotA. The phage protein is an activator that compensates for the deficiency in the promoter by assisting host RNA polymerase to bind. (This is similar to a mechanism employed by phage lambda, which is illustrated later in Figure 12.28.) The early and middle genes account for virtually all of the phage functions concerned with the synthesis of DNA, modifying cell structure, and transcribing and translating phage genes.

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Figure 12.9 The phage T4 lytic cascade falls into two parts: early functions are concerned with DNA synthesis and gene expression; late functions are concerned with particle assembly.

The two essential genes in the "transcription" category fulfill a regulatory function: their products are necessary for late gene expression. Phage T4 infection depends on a mechanical link between replication and late gene expression. Only actively replicating DNA can be used as template for late gene transcription. The connection is generated by introducing a new sigma factor and also by making other modifications in the host RNA polymerase so that it is active only with a template of replicating DNA. This link establishes a correlation between the synthesis of phage protein components and the number of genomes available for packaging. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.5

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PHAGE STRATEGIES

3.12.6 Lambda immediate early and delayed early genes are needed for both lysogeny and the lytic cycle Key Concepts

• Lambda has two immediate early genes, N and cro, which are transcribed by host RNA polymerase.

• N is required to express the delayed early genes. • Three of the delayed early genes are regulators. • Lysogeny requires the delayed early genes cII-cIII. • The lytic cycle requires the immediate early gene cro and the delayed early gene Q.

One of the most intricate cascade circuits is provided by phage lambda. Actually, the cascade for lytic development itself is straightforward, with two regulators controlling the successive stages of development. But the circuit for the lytic cycle is interlocked with the circuit for establishing lysogeny, as summarized in Figure 12.10 (for review see 100).

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Figure 12.10 The lambda lytic cascade is interlocked with the circuitry for lysogeny.

When lambda DNA enters a new host cell, the lytic and lysogenic pathways start off the same way. Both require expression of the immediate early and delayed early genes. But then they diverge: lytic development follows if the late genes are expressed; lysogeny ensues if synthesis of the repressor is established. Lambda has only two immediate early genes, transcribed independently by host RNA polymerase: • N codes for an antitermination factor whose action at the nut sites allows transcription to proceed into the delayed early genes (see Molecular Biology 3.9.24 Antitermination requires sites that are independent of the terminators). • cro has dual functions: it prevents synthesis of the repressor (a necessary action if the lytic cycle is to proceed); and it turns off expression of the immediate early genes (which are not needed later in the lytic cycle). The delayed early genes include two replication genes (needed for lytic infection), seven recombination genes (some involved in recombination during lytic infection, Lambda immediate early and delayed early genes are needed for both lysogeny and the lytic cycle | SECTION 3.12.6 © 2004. Virtual Text / www.ergito.com

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two necessary to integrate lambda DNA into the bacterial chromosome for lysogeny), and three regulators. The regulators have opposing functions: • The cII-cIII pair of regulators is needed to establish the synthesis of repressor. • The Q regulator is an antitermination factor that allows host RNA polymerase to transcribe the late genes. So the delayed early genes serve two masters: some are needed for the phage to enter lysogeny, the others are concerned with controlling the order of the lytic cycle.


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Reviews 100. Ptashne, M. (1992). A genetic switch. The Genetic Switch.



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3.12.7 The lytic cycle depends on antitermination Key Concepts

• pN is an antitermination factor that allows RNA polymerase to continue transcription past the ends of the two immediate early genes.

• pQ is the product of a delayed early gene and is an antiterminator that allows RNA polymerase to transcribe the late genes.

• Because lambda DNA circularizes after infection, the late genes form a single transcription unit.

To disentangle the two pathways, let's first consider just the lytic cycle. Figure 12.11 gives the map of lambda phage DNA. A group of genes concerned with regulation is surrounded by genes needed for recombination and replication. The genes coding for structural components of the phage are clustered. All of the genes necessary for the lytic cycle are expressed in polycistronic transcripts from three promoters.

Figure 12.11 The lambda map shows clustering of related functions. The genome is 48,514 bp.

Figure 12.12 shows that the two immediate early genes, N and cro, are transcribed by host RNA polymerase. N is transcribed toward the left, and cro toward the right. Each transcript is terminated at the end of the gene. pN is the regulator that allows transcription to continue into the delayed early genes. It is an antitermination factor that suppresses use of the terminators tL and tR (see Molecular Biology 3.9.25 Termination and anti-termination factors interact with RNA polymerase ). In the presence of pN, transcription continues to the left of N into the recombination genes, and to the right of cro into the replication genes.

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Figure 12.12 Phage lambda has two early transcription units; in the "leftward" unit, the "upper" strand is transcribed toward the left; in the "rightward" unit, the "lower" strand is transcribed toward the right. . Genes N and cro are the immediate early functions, and are separated from the delayed early genes by the terminators. Synthesis of N protein allows RNA polymerase to pass the terminators tL1 to the left and tR1 to the right.

The map in Figure 12.11 gives the organization of the lambda DNA as it exists in the phage particle. But shortly after infection, the ends of the DNA join to form a circle. Figure 12.13 shows the true state of lambda DNA during infection. The late genes are welded into a single group, containing the lysis genes S-R from the right end of the linear DNA, and the head and tail genes A-J from the left end.


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Figure 12.13 Lambda DNA circularizes during infection, so that the late gene cluster is intact in one transcription unit.

The late genes are expressed as a single transcription unit, starting from a promoter PR ′ that lies between Q and S. The late promoter is used constitutively. However, in the absence of the product of gene Q (which is the last gene in the rightward delayed early unit), late transcription terminates at a site tR3. The transcript resulting from this termination event is 194 bases long; it is known as 6S RNA. When pQ becomes available, it suppresses termination at tR3 and the 6S RNA is extended, with the result that the late genes are expressed. Late gene transcription does not seem to terminate at any specific point, but continues through all the late genes into the region beyond. A similar event happens with the leftward delayed early transcription, which continues past the recombination functions. Transcription in each direction is probably terminated before the The lytic cycle depends on antitermination | SECTION 3.12.7 © 2004. Virtual Text / www.ergito.com

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polymerases could crash into each other. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.7


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PHAGE STRATEGIES

3.12.8 Lysogeny is maintained by repressor protein Key Terms A plaque is an area of clearing in a bacterial lawn. It is created by a single phage particle that has undergone multiple rounds of lytic growth. A clear plaque is a type of plaque that contains only lysed bacterial cells. Key Concepts

• Mutants in the cI gene cannot maintain lysogeny. • cI codes for a repressor protein that acts at the OL and OR operators to block transcription of the immediate early genes.

• Because the immediate early genes trigger a regulatory cascade, their repression prevents the lytic cycle from proceeding.

Looking at the lambda lytic cascade, we see that the entire program is set in train by initiating transcription at the two promoters PL and PR for the immediate early genes N and cro. Because lambda uses antitermination to proceed to the next stage of (delayed early) expression, the same two promoters continue to be used throughout the early period. The expanded map of the regulatory region drawn in Figure 12.14 shows that the promoters PL and PR lie on either side of the cI gene. Associated with each promoter is an operator (OL, OR) at which repressor protein binds to prevent RNA polymerase from initiating transcription. The sequence of each operator overlaps with the promoter that it controls; so often these are described as the PL /OL and PR/OR control regions.

Figure 12.14 The lambda regulatory region contains a cluster of trans-acting functions and cis-acting elements.

Because of the sequential nature of the lytic cascade, the control regions provide a pressure point at which entry to the entire cycle can be controlled. By denying RNA Lysogeny is maintained by repressor protein | SECTION 3.12.8 © 2004. Virtual Text / www.ergito.com

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polymerase access to these promoters, a repressor protein prevents the phage genome from entering the lytic cycle. The repressor functions in the same way as repressors of bacterial operons: it binds to specific operators (525; 526; 527). The repressor protein is coded by the cI gene. Mutants in this gene cannot maintain lysogeny, but always enter the lytic cycle. Since the original isolation of the repressor protein (525; see Great Experiments 2.1 Isolation of repressor), its characterization has shown how it both maintains the lysogenic state and provides immunity for a lysogen against superinfection by new phage lambda genomes. When a bacterial culture is infected with a phage, the cells are lysed to generate regions that can be seen on a culture plate as small areas of clearing called plaques. With wild-type phages, the plaques are turbid or cloudy, because they contain some cells that have established lysogeny instead of being lysed. The effect of a cI mutation is to prevent lysogeny, so that the plaques contain only lysed cells. As a result, such an infection generates only clear plaques, and three genes (cI, cII, cIII) were named for their involvement in this phenotype. Figure 12.15 compares wild-type and mutant plaques.

Figure 12.15 Wild-type and virulent lambda mutants can be distinguished by their plaque types. Photograph kindly provided by Dale Kaiser.

The cI gene is transcribed from a promoter PRM that lies at its right end. (The subscript "RM" stands for repressor maintenance.) Transcription is terminated at the left end of the gene. The mRNA starts with the AUG initiation codon; because of the absence of the usual ribosome binding site, the mRNA is translated somewhat inefficiently, producing only a low level of repressor protein.

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References 525. Ptashne, M. (1967). Isolation of the λ phage repressor. Proc. Natl. Acad. Sci. USA 57, 306-313. 526. Ptashne, M. (1967). Specific binding of the λ phage repressor to DNA. Nature 214, 232-234. 527. Pirrotta, V., Chadwick, P., and Ptashne, M. (1970). Active form of two coliphage repressors. Nature 227, 41-44.


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PHAGE STRATEGIES

3.12.9 Repressor maintains an autogenous circuit Key Concepts

• Repressor binding at OL blocks transcription of gene N from PL. • Repressor binding at OR blocks transcription of cro but also is required for transcription of cI.

• Repressor binding to the operators therefore simultaneously blocks entry to the lytic cycle and promotes its own synthesis.

The repressor binds independently to the two operators. It has a single function at OL, but has dual functions at OR. These are illustrated in the upper part of Figure 12.16.

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Figure 12.16 Lysogeny is maintained by an autogenous circuit (upper). If this circuit is interrupted, the lytic cycle starts (lower).

At OL the repressor has the same sort of effect that we have already discussed for several other systems: it prevents RNA polymerase from initiating transcription at PL. This stops the expression of gene N. Since PL is used for all leftward early gene transcription, this action prevents expression of the entire leftward early transcription unit. So the lytic cycle is blocked before it can proceed beyond the early stages. At OR, repressor binding prevents the use of PR. So cro and the other rightward early genes cannot be expressed. (We see later why it is important to prevent the expression of cro when lysogeny is being maintained.) But the presence of repressor at OR also has another effect. The promoter for repressor synthesis, PRM, is adjacent to the rightward operator OR. It turns out that RNA polymerase can initiate efficiently at PRM only when repressor is bound at OR. The repressor behaves as a positive regulator protein that is necessary for transcription of the cI gene (see Molecular Biology 3.12.15 Repressor at OR2 Repressor maintains an autogenous circuit | SECTION 3.12.9 © 2004. Virtual Text / www.ergito.com

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interacts with RNA polymerase at PRM). Since the repressor is the product of cI, this interaction creates a positive autogenous circuit, in which the presence of repressor is necessary to support its own continued synthesis. The nature of this control circuit explains the biological features of lysogenic existence. Lysogeny is stable because the control circuit ensures that, so long as the level of repressor is adequate, there is continued expression of the cI gene. The result is that OL and OR remain occupied indefinitely. By repressing the entire lytic cascade, this action maintains the prophage in its inert form. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.9

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PHAGE STRATEGIES

3.12.10 The repressor and its operators define the immunity region Key Terms Immunity in phages refers to the ability of a prophage to prevent another phage of the same type from infecting a cell. It results from the synthesis of phage repressor by the prophage genome. Virulent phage mutants are unable to establish lysogeny. The immunity region is a segment of the phage genome that enables a prophage to inhibit additional phage of the same type from infecting the bacterium. This region has a gene that encodes for the repressor, as well as the sites to which the repressor binds. Key Concepts

• Several lambdoid phages have different immunity regions. • A lysogenic phage confers immunity to further infection by any other phage with the same immunity region.

The presence of repressor explains the phenomenon of immunity. If a second lambda phage DNA enters a lysogenic cell, repressor protein synthesized from the resident prophage genome will immediately bind to OL and OR in the new genome. This prevents the second phage from entering the lytic cycle. The operators were originally identified as the targets for repressor action by virulent mutations ( λvir). These mutations prevent the repressor from binding at OL or OR, with the result that the phage inevitably proceeds into the lytic pathway when it infects a new host bacterium. And λvir mutants can grow on lysogens because the virulent mutations in OL and OR allow the incoming phage to ignore the resident repressor and thus to enter the lytic cycle. Virulent mutations in phages are the equivalent of operator-constitutive mutations in bacterial operons. Prophage is induced to enter the lytic cycle when the lysogenic circuit is broken. This happens when the repressor is inactivated (see next section). The absence of repressor allows RNA polymerase to bind at PL and PR, starting the lytic cycle as shown in the lower part of Figure 12.16 (for review see 104). The autogenous nature of the repressor-maintenance circuit creates a sensitive response. Because the presence of repressor is necessary for its own synthesis, expression of the cI gene stops as soon as the existing repressor is destroyed. So no repressor is synthesized to replace the molecules that have been damaged. This enables the lytic cycle to start without interference from the circuit that maintains lysogeny.

The repressor and its operators define the immunity region | SECTION 3.12.10 © 2004. Virtual Text / www.ergito.com

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The region including the left and right operators, the cI gene, and the cro gene determines the immunity of the phage. Any phage that possesses this region has the same type of immunity, because it specifies both the repressor protein and the sites on which the repressor acts. Accordingly, this is called the immunity region (as marked in Figure 12.14). Each of the four lambdoid phages φ80, 21, 434, and λ has a unique immunity region. When we say that a lysogenic phage confers immunity to any other phage of the same type, we mean more precisely that the immunity is to any other phage that has the same immunity region (irrespective of differences in other regions).


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Reviews 104. Friedman, D. I. and Gottesman, M. (1982). Lytic mode of lambda development In Lambda. Lambda II, 21-51.



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PHAGE STRATEGIES

3.12.11 The DNA-binding form of repressor is a dimer Key Concepts

• A repressor monomer has two distinct domains. • The N-terminal domain contains the DNA-binding site. • The C-terminal domain dimerizes. • Binding to the operator requires the dimeric form so that two DNA-binding domains can contact the operator simultaneously.

• Cleavage of the repressor between the two domains reduces the affinity for the operator and induces a lytic cycle.

The repressor subunit is a polypeptide of 27 kD with the two distinct domains summarized in Figure 12.17.

Figure 12.17 The N-terminal and C-terminal regions of repressor form separate domains. The C-terminal domains associate to form dimers; the N-terminal domains bind DNA.

• The N-terminal domain, residues 1-92, provides the operator-binding site. • The C-terminal domain, residues 132-236, is responsible for dimerization. The two domains are joined by a connector of 40 residues. When repressor is digested by a protease, each domain is released as a separate fragment. Each domain can exercise its function independently of the other. The C-terminal fragment can form oligomers. The N-terminal fragment can bind the operators, although with a lower affinity than the intact repressor. So the information for The DNA-binding form of repressor is a dimer | SECTION 3.12.11 © 2004. Virtual Text / www.ergito.com

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specifically contacting DNA is contained within the N-terminal domain, but the efficiency of the process is enhanced by the attachment of the C-terminal domain (529). The dimeric structure of the repressor is crucial in maintaining lysogeny. The induction of a lysogenic prophage to enter the lytic cycle is caused by cleavage of the repressor subunit in the connector region, between residues 111 and 113. (This is a counterpart to the allosteric change in conformation that results when a small-molecule inducer inactivates the repressor of a bacterial operon, a capacity that the lysogenic repressor does not have.) Induction occurs under certain adverse conditions, such as exposure of lysogenic bacteria to UV irradiation, which leads to proteolytic inactivation of the repressor. In the intact state, dimerization of the C-terminal domains ensures that when the repressor binds to DNA its two N-terminal domains each contact DNA simultaneously. But cleavage releases the C-terminal domains from the N-terminal domains. As illustrated in Figure 12.18 this means that the N-terminal domains can no longer dimerize; this upsets the equilibrium between monomers and dimers, so that repressor dissociates from DNA, allowing lytic infection to start. (Another relevant parameter is the loss of cooperative effects between adjacent dimers.)

The DNA-binding form of repressor is a dimer | SECTION 3.12.11 © 2004. Virtual Text / www.ergito.com

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Figure 12.18 Repressor dimers bind to the operator. The affinity of the N-terminal domains for DNA is controlled by the dimerization of the C-terminal domains.

The balance between lysogeny and the lytic cycle depends on the concentration of repressor. Intact repressor is present in a lysogenic cell at a concentration sufficient to ensure that the operators are occupied. But if the repressor is cleaved, this concentration is inadequate, because of the lower affinity of the separate N-terminal domain for the operator. Too high a concentration of repressor would make it impossible to induce the lytic cycle in this way; too low a level, of course, would make it impossible to maintain lysogeny.


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References 529. Pabo, C. O. and Lewis, M. (1982). The operator-binding domain of λ repressor: structure and DNA recognition. Nature 298, 443-447.



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PHAGE STRATEGIES

3.12.12 Repressor uses a helix-turn-helix motif to bind DNA Key Terms The helix-turn-helix motif describes an arrangement of two α helices that form a site that binds to DNA, one fitting into the major groove of DNA and other lying across it. Key Concepts

• Each DNA-binding region in the repressor contacts a half-site in the DNA. • The DNA-binding site of repressor includes two short α-helical regions which fit into the successive turns of the major groove of DNA.

• A DNA-binding site is a (partially) palindromic sequence of 17 bp.

A repressor dimer is the unit that binds to DNA. It recognizes a sequence of 17 bp displaying partial symmetry about an axis through the central base pair. Figure 12.19 shows an example of a binding site. The sequence on each side of the central base pair is sometimes called a "half-site". Each individual N-terminal region contacts a half-site. Several DNA-binding proteins that regulate bacterial transcription share a similar mode of holding DNA, in which the active domain contains two short regions of α-helix that contact DNA. (Some transcription factors in eukaryotic cells use a similar motif; see Molecular Biology 5.22.14 Homeodomains bind related targets in DNA.)

Figure 12.19 The operator is a 17 bp sequence with an axis of symmetry through the central base pair. Each half site is marked in green. Base pairs that are identical in each operator half are in red.

The N-terminal domain of lambda repressor contains several stretches of α-helix, arranged as illustrated diagrammatically in Figure 12.20. Two of the helical regions are responsible for binding DNA. The helix-turn-helix model for contact is illustrated in Figure 12.21. Looking at a single monomer, α-helix-3 consists of 9 amino acids, lying at an angle to the preceding region of 7 amino acids that forms α-helix-2. In the dimer, the two apposed helix-3 regions lie 34 Å apart, enabling them to fit into successive major grooves of DNA. The helix-2 regions lie at an angle that would place them across the groove (530). The symmetrical binding of dimer to the site means that each N-terminal domain of the dimer contacts a similar set of Repressor uses a helix-turn-helix motif to bind DNA | SECTION 3.12.12 © 2004. Virtual Text / www.ergito.com

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bases in its half-site.

Figure 12.20 Lambda repressor''s N-terminal domain contains five stretches of α -helix; helices 2 and 3 are involved in binding DNA.

Figure 12.21 In the two-helix model for DNA binding, helix-3 of each monomer lies in the wide groove on the same face of DNA, and helix-2 lies across the groove.


Repressor uses a helix-turn-helix motif to bind DNA | SECTION 3.12.12 © 2004. Virtual Text / www.ergito.com

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References 530. Sauer, R. T. et al. (1982). Homology among DNA-binding proteins suggests use of a conserved super-secondary structure. Nature 298, 447-451.


Repressor uses a helix-turn-helix motif to bind DNA | SECTION 3.12.12 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.13 The recognition helix determines specificity for DNA Key Terms The recognition helix is the one of the two helices of the helix-turn-helix motif that makes contacts with DNA that are specific for particular bases. This determines the specificity of the DNA sequence that is bound. Key Concepts

• The amino acid sequence of the recognition helix makes contacts with particular bases in the operator sequence that it recognizes.

Related forms of the α-helical motifs employed in the helix-loop-helix of the lambda repressor are found in several DNA-binding proteins, including CRP, the lac repressor, and several other phage repressors. By comparing the abilities of these proteins to bind DNA, we can define the roles of each helix: • Contacts between helix-3 and DNA rely on hydrogen bonds between the amino acid side chains and the exposed positions of the base pairs. This helix is responsible for recognizing the specific target DNA sequence, and is therefore also known as the recognition helix. • Contacts from helix-2 to the DNA take the form of hydrogen bonds connecting with the phosphate backbone. These interactions are necessary for binding, but do not control the specificity of target recognition. In addition to these contacts, a large part of the overall energy of interaction with DNA is provided by ionic interactions with the phosphate backbone. What happens if we manipulate the coding sequence to construct a new protein by substituting the recognition helix in one repressor with the corresponding sequence from a closely related repressor? The specificity of the hybrid protein is that of its new recognition helix. The amino acid sequence of this short region determines the sequence specificities of the individual proteins, and is able to act in conjunction with the rest of the polypeptide chain (531). Figure 12.22 shows the details of the binding to DNA of two proteins that bind similar DNA sequences. Both lambda repressor and Cro protein have a similar organization of the helix-turn-helix motif, although their individual specificities for DNA are not identical:

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Figure 12.22 Two proteins that use the two-helix arrangement to contact DNA recognize lambda operators with affinities determined by the amino acid sequence of helix-3.

• Each protein uses similar interactions between hydrophobic amino acids to maintain the relationship between helix-2 and helix-3: repressor has an Ala-Val connection, while Cro has an Ala-Ile association. • Amino acids in helix-3 of the repressor make contacts with specific bases in the operator. Three amino acids in repressor recognize three bases in DNA; the amino acids at these positions and also at additional positions in Cro recognize five (or possibly six) bases in DNA. Two of the amino acids involved in specific recognition are identical in repressor and Cro (Gln and Ser at the N-terminal end of the helix), while the other contacts are different (Ala in repressor versus Lys and the additional Asn in Cro). Also, a Thr in helix-2 of Cro directly contacts DNA. The interactions shown in the figure represent binding to the DNA sequence that each protein recognizes most tightly. The sequences shown at the bottom of the figure with the contact points in color differ at 3 of the 9 base pairs. The use of overlapping, but not identical contacts between amino acids and bases shows how related recognition helices confer recognition of related DNA sequences. This enables repressor and Cro to recognize the same set of sequences, but with different relative affinities for particular members of the group. The bases contacted by helix-3 of repressor or Cro lie on one face of DNA, as can be seen from the positions indicated on the helical diagram in Figure 12.22. However, repressor makes an additional contact with the other face of DNA. Removing the last six N-terminal amino acids (which protrude from helix-1) eliminates some of the contacts. This observation provides the basis for the idea that the bulk of the N-terminal domain contacts one face of DNA, while the last six N-terminal amino The recognition helix determines specificity for DNA | SECTION 3.12.13 © 2004. Virtual Text / www.ergito.com

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acids form an "arm" extending around the back. Figure 12.23 shows the view from the back. Lysine residues in the arm make contacts with G residues in the major groove, and also with the phosphate backbone. The interaction between the arm and DNA contributes heavily to DNA binding; the affinity of the armless repressor for DNA is reduced by ~1000-fold (532).

Figure 12.23 A view from the back shows that the bulk of the repressor contacts one face of DNA, but its N-terminal arms reach around to the other face.

Bases that are not contacted directly by repressor protein may have an important effect on binding. The related phage 434 repressor binds DNA via a helix-turn-helix motif, and the crystal structure shows that helix-3 is positioned at each half-site so that it contacts the 5 outermost base pairs but not the inner 2. However, operators with A·T base pairs at the inner positions bind 434 repressor more strongly than operators with G·C base pairs. The reason is that 434 repressor binding slightly twists DNA at the center of the operator, widening the angle between the two half-sites of DNA by ~3°. This is probably needed to allow each monomer of the repressor dimer to make optimal contacts with DNA. A·T base pairs allow this twist more readily than G·C pairs, thus affecting the affinity of the operator for repressor. Last updated on 2-28-2002


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References 531. Wharton, R. L., Brown, E. L., and Ptashne, M. (1984). Substituting an α -helix switches the sequence specific DNA interactions of a repressor. Cell 38, 361-369. 532. Brennan, R. G. et al. (1990). Protein-DNA conformational changes in the crystal structure of a λCro-operator complex. Proc. Natl. Acad. Sci. USA 87, 8165-8169.



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3.12.14 Repressor dimers bind cooperatively to the operator Key Concepts

• Repressor binding to one operator increases the affinity for binding a second repressor dimer to the adjacent operator.

• The affinity is 10× greater for OL1 and OR1 than other operators, so they are bound first.

• Cooperativity allows repressor to bind the O1/O2 sites at lower concentrations.

Each operator contains three repressor-binding sites. As can be seen from Figure 12.24, no two of the six individual repressor-binding sites are identical, but they all conform with a consensus sequence. The binding sites within each operator are separated by spacers of 3-7 bp that are rich in A·T base pairs. The sites at each operator are numbered so that OR consists of the series of binding sites OR1-OR2-OR3, while OL consists of the series OL1-OL2-OL3. In each case, site 1 lies closest to the startpoint for transcription in the promoter, and sites 2 and 3 lie farther upstream.

Figure 12.24 Each operator contains three repressor-binding sites, and overlaps with the promoter at which RNA polymerase binds. The orientation of OL has been reversed from usual to facilitate comparison with OR.

Faced with the triplication of binding sites at each operator, how does repressor Repressor dimers bind cooperatively to the operator | SECTION 3.12.14 © 2004. Virtual Text / www.ergito.com

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decide where to start binding? At each operator, site 1 has a greater affinity (roughly tenfold) than the other sites for the repressor. So the repressor always binds first to OL1 and OR1. Lambda repressor binds to subsequent sites within each operator in a cooperative manner. The presence of a dimer at site 1 greatly increases the affinity with which a second dimer can bind to site 2. When both sites 1 and 2 are occupied, this interaction does not extend farther, to site 3. At the concentrations of repressor usually found in a lysogen, both sites 1 and 2 are filled at each operator, but site 3 is not occupied (528). If site 1 is inactive (because of mutation), then repressor binds cooperatively to sites 2 and 3. That is, binding at site 2 assists another dimer to bind at site 3. This interaction occurs directly between repressor dimers and not via conformational change in DNA. The C-terminal domain is responsible for the cooperative interaction between dimers as well as for the dimer formation between subunits. Figure 12.25 shows that it involves both subunits of each dimer, that is, each subunit contacts its counterpart in the other dimer, forming a tetrameric structure (1199).

Figure 12.25 When two lambda repressor dimers bind cooperatively, each of the subunits of one dimer contacts a subunit in the other dimer.

A result of cooperative binding is to increase the effective affinity of repressor for the operator at physiological concentrations. This enables a lower concentration of repressor to achieve occupancy of the operator. This is an important consideration in a system in which release of repression has irreversible consequences. In an operon coding for metabolic enzymes, after all, failure of repression will merely allow unnecessary synthesis of enzymes. But failure to repress lambda prophage will lead to induction of phage and lysis of the cell. From the sequences shown in Figure 12.24, we see that OL1 and OR1 lie more or less in the center of the RNA polymerase binding sites of PL and PR, respectively. Occupancy of OL1-OL2 and OR1-OR2 thus physically blocks access of RNA polymerase to the corresponding promoters.

Repressor dimers bind cooperatively to the operator | SECTION 3.12.14 © 2004. Virtual Text / www.ergito.com

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References 528. Johnson, A. D., Meyer, B. J., and Ptashne, M. (1979). Interactions between DNA-bound repressors govern regulation by the phage λrepressor. Proc. Natl. Acad. Sci. USA 76, 5061-5065. 1199. Bell, C. E., Frescura, P., Hochschild, A., and Lewis, M. (2000). Crystal structure of the lambda repressor C-terminal domain provides a model for cooperative operator binding. Cell 101, 801-811.


Repressor dimers bind cooperatively to the operator | SECTION 3.12.14 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.15 Repressor at OR2 interacts with RNA polymerase at PRM Key Concepts

• The DNA-binding region of repressor at OR2 contacts RNA polymerase and stabilizes its binding to PRM.

• This is the basis for the autogenous control of repressor maintenance.

A different relationship is shown between OR and the promoter PRM for transcription of cI. The RNA polymerase binding site is adjacent to OR2. This explains how repressor autogenously regulates its own synthesis. When two dimers are bound at OR1-OR2, the dimer at OR2 interacts with RNA polymerase (see Figure 12.16 in Molecular Biology 3.12.9 Repressor maintains an autogenous circuit). This effect resides in the amino terminal domain of repressor. Mutations that abolish positive control map in the cI gene. One interesting class of mutants remain able to bind the operator to repress transcription, but cannot stimulate RNA polymerase to transcribe from PRM. They map within a small group of amino acids, located on the outside of helix-2 or in the turn between helix-2 and helix-3. The mutations reduce the negative charge of the region; conversely, mutations that increase the negative charge enhance the activation of RNA polymerase. This suggests that the group of amino acids constitutes an "acidic patch" that functions by an electrostatic interaction with a basic region on RNA polymerase. The location of these "positive control mutations" in the repressor is indicated on Figure 12.26. They lie at a site on repressor that is close to a phosphate group on DNA that is also close to RNA polymerase. So the group of amino acids on repressor that is involved in positive control is in a position to contact the polymerase. The interaction between repressor and polymerase is needed for the polymerase to make the transition from a closed complex to an open complex (see also Figure 12.29). The important principle is that protein-protein interactions can release energy that is used to help to initiate transcription.

Repressor at OR2 interacts with RNA polymerase at PRM | SECTION 3.12.15 © 2004. Virtual Text / www.ergito.com

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Figure 12.26 Positive control mutations identify a small region at helix-2 that interacts directly with RNA polymerase.

What happens if a repressor dimer binds to OR3? This site overlaps with the RNA polymerase binding site at PRM. So if the repressor concentration becomes great enough to cause occupancy of OR3, the transcription of cI is prevented. This leads in due course to a reduction in repressor concentration; OR3 then becomes empty, and the autogenous loop can start up again because OR2 remains occupied. This mechanism could prevent the concentration of repressor from becoming too great, although it would require repressor concentration in lysogens to reach unusually high levels. In the formal sense, the repressor is an autogenous regulator of its own expression that functions positively at low concentrations and negatively at high concentrations. Virulent mutations occur in sites 1 and 2 of both OL and OR. The mutations vary in their degree of virulence, according to the extent to which they reduce the affinity of the binding site for repressor, and also depending on the relationship of the affected site to the promoter. Consistent with the conclusion that OR3 and OL3 usually are not occupied, virulent mutations are not found in these sites. Last updated on 10-17-2000 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.15

Repressor at OR2 interacts with RNA polymerase at PRM | SECTION 3.12.15 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.16 The cII and cIII genes are needed to establish lysogeny Key Concepts

• The delayed early gene products cII and cIII are necessary for RNA polymerase to initiate transcription at the promoter PRE.

• cII acts direct at the promoter and cIII protects cII from degradation. • Transcription from PRE leads to synthesis of repressor and also blocks the transcription of cro.

The control circuit for maintaining lysogeny presents a paradox. The presence of repressor protein is necessary for its own synthesis. This explains how the lysogenic condition is perpetuated. But how is the synthesis of repressor established in the first place? When a lambda DNA enters a new host cell, RNA polymerase cannot transcribe cI, because there is no repressor present to aid its binding at PRM. But this same absence of repressor means that PR and PL are available. So the first event when lambda DNA infects a bacterium is for genes N and cro to be transcribed. Then pN allows transcription to be extended farther. This allows cIII (and other genes) to be transcribed on the left, while cII (and other genes) are transcribed on the right (see Figure 12.14). The cII and cIII genes share with cI the property that mutations in them cause clear plaques. But there is a difference. The cI mutants can neither establish nor maintain lysogeny. The cII or cIII mutants have some difficulty in establishing lysogeny, but once established, they are able to maintain it by the cI autogenous circuit. This implicates the cII and cIII genes as positive regulators whose products are needed for an alternative system for repressor synthesis. The system is needed only to initiate the expression of cI in order to circumvent the inability of the autogenous circuit to engage in de novo synthesis. They are not needed for continued expression. The cII protein acts directly on gene expression. Between the cro and cII genes is another promoter, called PRE. (The subscript "RE" stands for repressor establishment.) This promoter can be recognized by RNA polymerase only in the presence of cII, whose action is illustrated in Figure 12.27.

The cII and cIII genes are needed to establish lysogeny | SECTION 3.12.16 © 2004. Virtual Text / www.ergito.com

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Figure 12.27 Repressor synthesis is established by the action of cII and RNA polymerase at PRE to initiate transcription that extends from the antisense strand of cro through the cI gene.

The cII protein is extremely unstable in vivo, because it is degraded as the result of the activity of a host protein called HflA. The role of cIII is to protect cII against this degradation. Transcription from PRE promotes lysogeny in two ways. Its direct effect is that cI is translated into repressor protein. An indirect effect is that transcription proceeds through the cro gene in the "wrong" direction. So the 5 ′ part of the RNA corresponds to an antisense transcript of cro; in fact, it hybridizes to authentic cro mRNA, inhibiting its translation. This is important because cro expression is needed to enter the lytic cycle (see Molecular Biology 3.12.19 The cro repressor is needed for lytic infection). The cI coding region on the PRE transcript is very efficiently translated, in contrast with the weak translation of the PRM transcript. In fact, repressor is synthesized ~7-8 times more effectively via expression from PRE than from PRM. This reflects the fact that the PRE transcript has an efficient ribosome-binding site, whereas the PRM transcript has no ribosome-binding site and actually starts with the AUG initiation codon. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.16

The cII and cIII genes are needed to establish lysogeny | SECTION 3.12.16 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.17 A poor promoter requires cII protein Key Concepts

• PRE has atypical sequences at –10 and –35. • RNA polymerase binds the promoter only in the presence of cII. • cII binds to sequences close to the –35 region.

The PRE promoter has a poor fit with the consensus at –10 and lacks a consensus sequence at –35. This deficiency explains its dependence on cII. The promoter cannot be transcribed by RNA polymerase alone in vitro, but can be transcribed when cII is added. The regulator binds to a region extending from about –25 to –45. When RNA polymerase is added, an additional region is protected, extending from –12 to +13. As summarized in Figure 12.28, the two proteins bind to overlapping sites.

Figure 12.28 RNA polymerase binds to PRE only in the presence of cII, which contacts the region around -35.

The importance of the –35 and –10 regions for promoter function, in spite of their lack of resemblance with the consensus, is indicated by the existence of cy mutations. These have effects similar to those of cII and cIII mutations in preventing the establishment of lysogeny; but they are cis-acting instead of trans-acting. They fall A poor promoter requires cII protein | SECTION 3.12.17 © 2004. Virtual Text / www.ergito.com

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into two groups, cyL and cyR, localized at the consensus operator positions of –10 and –35. The cyL mutations are located around –10, and probably prevent RNA polymerase from recognizing the promoter. The cyR mutations are located around –35, and fall into two types, affecting either RNA polymerase or cII binding. Mutations in the center of the region do not affect cII binding; presumably they prevent RNA polymerase binding. On either side of this region, mutations in short tetrameric repeats, TTGC, prevent cII from binding. Each base in the tetramer is 10 bp (one helical turn) separated from its homologue in the other tetramer, so that when cII recognizes the two tetramers, it lies on one face of the double helix. Positive control of a promoter implies that an accessory protein has increased the efficiency with which RNA polymerase initiates transcription. Figure 12.29 reports that either or both stages of the interaction between promoter and polymerase can be the target for regulation. Initial binding to form a closed complex or its conversion into an open complex can be enhanced.

Figure 12.29 Positive regulation can influence RNA polymerase at either stage of initiating transcription. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.17

A poor promoter requires cII protein | SECTION 3.12.17 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.18 Lysogeny requires several events Key Concepts

• cII/cIII cause repressor synthesis to be established and also trigger inhibition of late gene transcription.

• Establishment of repressor turns off immediate and delayed early gene expression. • Repressor turns on the maintenance circuit for its own synthesis. • Lambda DNA is integrated into the bacterial genome at the final stage in establishing lysogeny.

Now we can see how lysogeny is established during an infection. Figure 12.30 recapitulates the early stages and shows what happens as the result of expression of cIII and cII. The presence of cII allows PRE to be used for transcription extending through cI. Repressor protein is synthesized in high amounts from this transcript. Immediately it binds to OL and OR.

Lysogeny requires several events | SECTION 3.12.18 © 2004. Virtual Text / www.ergito.com

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Figure 12.30 A cascade is needed to establish lysogeny, but then this circuit is switched off and replaced by the autogenous repressor-maintenance circuit.

By directly inhibiting any further transcription from PL and PR, repressor binding turns off the expression of all phage genes. This halts the synthesis of cII and cIII, which are unstable; they decay rapidly, with the result that PRE can no longer be used. So the synthesis of repressor via the establishment circuit is brought to a halt. But repressor now is present at OR. It switches on the maintenance circuit for Lysogeny requires several events | SECTION 3.12.18 © 2004. Virtual Text / www.ergito.com

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expression from PRM. Repressor continues to be synthesized, although at the lower level typical of PRM function. So the establishment circuit starts off repressor synthesis at a high level; then repressor turns off all other functions, while at the same time turning on the maintenance circuit, which functions at the low level adequate to sustain lysogeny. We shall not now deal in detail with the other functions needed to establish lysogeny, but we can just briefly remark that the infecting lambda DNA must be inserted into the bacterial genome (see Molecular Biology 4.15.16 Specialized recombination involves specific sites). The insertion requires the product of gene int, which is expressed from its own promoter PI, at which cII also is necessary. The sequence of PI shows homology with PRE in the cII binding site (although not in the –10 region). The functions necessary for establishing the lysogenic control circuit are therefore under the same control as the function needed to integrate the phage DNA into the bacterial genome. So the establishment of lysogeny is under a control that ensures all the necessary events occur with the same timing. Emphasizing the tricky quality of lambda's intricate cascade, we now know that cII promotes lysogeny in another, indirect manner. It sponsors transcription from a promoter called Panti-Q, which is located within the Q gene. This transcript is an antisense version of the Q region, and it hybridizes with Q mRNA to prevent translation of Q protein, whose synthesis is essential for lytic development. So the same mechanisms that directly promote lysogeny by causing transcription of the cI repressor gene also indirectly help lysogeny by inhibiting the expression of cro (see above) and Q, the regulator genes needed for the antagonistic lytic pathway. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.18

Lysogeny requires several events | SECTION 3.12.18 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.19 The cro repressor is needed for lytic infection Key Concepts

• Cro binds to the same operators as repressor but with different affinities. • When Cro binds to OR3, it prevents RNA polymerase from binding to PRM, and blocks maintenance of repressor.

• When Cro binds to other operators at OR or OL, it prevents RNA polymerase from expressing immediate early genes, which (indirectly) blocks repressor establishment.

Lambda has the alternatives of entering lysogeny or starting a lytic infection. Lysogeny is initiated by establishing an autogenous maintenance circuit that inhibits the entire lytic cascade through applying pressure at two points. The program for establishing lysogeny proceeds through some of the same events that are required for the lytic cascade (expression of delayed early genes via expression of N is needed). We now face a problem. How does the phage enter the lytic cycle? The key influence on the lytic cycle is the role of gene cro, which codes for another repressor. Cro is responsible for preventing the synthesis of the repressor protein; this action shuts off the possibility of establishing lysogeny. cro mutants usually establish lysogeny rather than entering the lytic pathway, because they lack the ability to switch events away from the expression of repressor. Cro forms a small dimer (the subunit is 9 kD) that acts within the immunity region. It has two effects: • It prevents the synthesis of repressor via the maintenance circuit; that is, it prevents transcription via PRM. • It also inhibits the expression of early genes from both PL and PR. This means that, when a phage enters the lytic pathway, Cro has responsibility both for preventing the synthesis of repressor and (subsequently) for turning down the expression of the early genes. Cro achieves its function by binding to the same operators as (cI) repressor protein. Cro includes a region with the same general structure as the repressor; a helix-2 is offset at an angle from recognition helix-3. (The remainder of the structure is different, demonstrating that the helix-turn-helix motif can operate within various contexts.) Like repressor, Cro binds symmetrically at the operators.

The cro repressor is needed for lytic infection | SECTION 3.12.19 © 2004. Virtual Text / www.ergito.com

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The sequences of Cro and repressor in the helix-turn-helix region are related, explaining their ability to contact the same DNA sequences (see Figure 12.22). Cro makes similar contacts to those made by repressor, but binds to only one face of DNA; it lacks the N-terminal arms by which repressor reaches around to the other side. How can two proteins have the same sites of action, yet have such opposite effects? The answer lies in the different affinities that each protein has for the individual binding sites within the operators. Let us just consider OR, where more is known, and where Cro exerts both its effects. The series of events is illustrated in Figure 12.31. (Note that the first two stages are identical to those of the lysogenic circuit shown in Figure 12.30.)


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Figure 12.31 The lytic cascade requires Cro protein, which directly prevents repressor maintenance via PRM, as well as turning off delayed early gene expression, indirectly preventing repressor establishment.

The affinity of Cro for OR3 is greater than its affinity for OR2 or OR1. So it binds first to OR3. This inhibits RNA polymerase from binding to PRM. So Cro's first action is to prevent the maintenance circuit for lysogeny from coming into play. Then Cro binds to OR2 or OR1. Its affinity for these sites is similar, and there is no cooperative effect. Its presence at either site is sufficient to prevent RNA polymerase The cro repressor is needed for lytic infection | SECTION 3.12.19 © 2004. Virtual Text / www.ergito.com

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from using PR. This in turn stops the production of the early functions (including Cro itself). Because cII is unstable, any use of PRE is brought to a halt. So the two actions of Cro together block all production of repressor. So far as the lytic cycle is concerned, Cro turns down (although it does not completely eliminate) the expression of the early genes. Its incomplete effect is explained by its affinity for OR1 and OR2, which is about eight times lower than that of repressor. This effect of Cro does not occur until the early genes have become more or less superfluous, because pQ is present; by this time, the phage has started late gene expression, and is concentrating on the production of progeny phage particles. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.19


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PHAGE STRATEGIES

3.12.20 What determines the balance between lysogeny and the lytic cycle? Key Concepts

• The delayed early stage when both Cro and repressor are being expressed is common to lysogeny and the lytic cycle.

• The critical event is whether cII causes sufficient synthesis of repressor to overcome the action of Cro.

The programs for the lysogenic and lytic pathways are so intimately related that it is impossible to predict the fate of an individual phage genome when it enters a new host bacterium. Will the antagonism between repressor and Cro be resolved by establishing the autogenous maintenance circuit shown in Figure 12.30, or by turning off repressor synthesis and entering the late stage of development shown in Figure 12.31? The same pathway is followed in both cases right up to the brink of decision. Both involve the expression of the immediate early genes and extension into the delayed early genes. The difference between them comes down to the question of whether repressor or Cro will obtain occupancy of the two operators. The early phase during which the decision is taken is limited in duration in either case. No matter which pathway the phage follows, expression of all early genes will be prevented as PL and PR are repressed; and, as a consequence of the disappearance of cII and cIII, production of repressor via PRE will cease. The critical question comes down to whether the cessation of transcription from PRE is followed by activation of PRM and the establishment of lysogeny, or whether PRM fails to become active and the pQ regulator commits the phage to lytic development. Figure 12.32 shows the critical stage, at which both repressor and Cro are being synthesized.

What determines the balance between lysogeny and the lytic cycle? | SECTION 3.12.20 © 2004. Virtual Text / www.ergito.com

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Figure 12.32 The critical stage in deciding between lysogeny and lysis is when delayed early genes are being expressed. If cII causes sufficient synthesis of repressor, lysogeny will result because repressor occupies the operators. Otherwise Cro occupies the operators, resulting in a lytic cycle.

The initial event in establishing lysogeny is the binding of repressor at OL1 and OR1. Binding at the first sites is rapidly succeeded by cooperative binding of further repressor dimers at OL2 and OR2. This shuts off the synthesis of Cro and starts up the synthesis of repressor via PRM. The initial event in entering the lytic cycle is the binding of Cro at OR3. This stops the lysogenic-maintenance circuit from starting up at PRM. Then Cro must bind to OR1 or OR2, and to OL1 or OL2, to turn down early gene expression. By halting production of cII and cIII, this action leads to the cessation of repressor synthesis via What determines the balance between lysogeny and the lytic cycle? | SECTION 3.12.20 © 2004. Virtual Text / www.ergito.com

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PRE. The shutoff of repressor establishment occurs when the unstable cII and cIII proteins decay. The critical influence over the switch between lysogeny and lysis is cII. If cII is active, synthesis of repressor via the establishment promoter is effective; and, as a result, repressor gains occupancy of the operators. If cII is not active, repressor establishment fails, and Cro binds to the operators. The level of cII protein under any particular set of circumstances determines the outcome of an infection. Mutations that increase the stability of cII increase the frequency of lysogenization. Such mutations occur in cII itself or in other genes. The cause of cII's instability is its susceptibility to degradation by host proteases. Its level in the cell is influenced by cIII as well as by host functions. The effect of the lambda protein cIII is secondary: it helps to protect cII against degradation. Although the presence of cIII does not guarantee the survival of cII, in the absence of cIII, cII is virtually always inactivated. Host gene products act on this pathway. Mutations in the host genes hflA and hflB increase lysogeny – hfl stands for high frequency lysogenization. The mutations stabilize cII because they inactivate host protease(s) that degrade it. The influence of the host cell on the level of cII provides a route for the bacterium to interfere with the decision-taking process. For example, host proteases that degrade cII are activated by growth on rich medium, so lambda tends to lyse cells that are growing well, but is more likely to enter lysogeny on cells that are starving (and which lack components necessary for efficient lytic growth). This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.20

What determines the balance between lysogeny and the lytic cycle? | SECTION 3.12.20 © 2004. Virtual Text / www.ergito.com

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PHAGE STRATEGIES

3.12.21 Summary Phages have a lytic life cycle, in which infection of a host bacterium is followed by production of a large number of phage particles, lysis of the cell, and release of the viruses. Some phages also can exist in lysogenic form, in which the phage genome is integrated into the bacterial chromosome and is inherited in this inert, latent form like any other bacterial gene. Lytic infection falls typically into three phases. In the first phase a small number of phage genes are transcribed by the host RNA polymerase. One or more of these genes is a regulator that controls expression of the group of genes expressed in the second phase. The pattern is repeated in the second phase, when one or more genes is a regulator needed for expression of the genes of the third phase. Genes of the first two phases code for enzymes needed to reproduce phage DNA; genes of the final phase code for structural components of the phage particle. It is common for the very early genes to be turned off during the later phases. In phage lambda, the genes are organized into groups whose expression is controlled by individual regulatory events. The immediate early gene N codes for an antiterminator that allows transcription of the leftward and rightward groups of delayed early genes from the early promoters PR and P L. The delayed early gene Q has a similar antitermination function that allows transcription of all late genes from the promoter PR ′ . The lytic cycle is repressed, and the lysogenic state maintained, by expression of the cI gene, whose product is a repressor protein that acts at the operators OR and OL to prevent use of the promoters PR and PL, respectively. A lysogenic phage genome expresses only the cI gene, from its promoter PRM. Transcription from this promoter involves positive autogenous regulation, in which repressor bound at OR activates RNA polymerase at PRM. Each operator consists of three binding sites for repressor. Each site is palindromic, consisting of symmetrical half-sites. Repressor functions as a dimer. Each half binding site is contacted by a repressor monomer. The N-terminal domain of repressor contains a helix-turn-helix motif that contacts DNA. Helix-3 is the recognition helix, responsible for making specific contacts with base pairs in the operator. Helix-2 is involved in positioning helix-3; it is also involved in contacting RNA polymerase at PRM. The C-terminal domain is required for dimerization. Induction is caused by cleavage between the N- and C-terminal domains, which prevents the DNA-binding regions from functioning in dimeric form, thereby reducing their affinity for DNA and making it impossible to maintain lysogeny. Repressor-operator binding is cooperative, so that once one dimer has bound to the first site, a second dimer binds more readily to the adjacent site. The helix-turn-helix motif is used by other DNA-binding proteins, including lambda Cro, which binds to the same operators, but has a different affinity for the individual operator sites, determined by the sequence of helix-3. Cro binds individually to operator sites, starting with OR3, in a noncooperative manner. It is needed for progression through the lytic cycle. Its binding to OR3 first prevents synthesis of repressor from PRM; then its binding to OR2 and OR1 prevents continued expression Summary | SECTION 3.12.21 © 2004. Virtual Text / www.ergito.com

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of early genes, an effect also seen in its binding to OL1 and OL2. Establishment of repressor synthesis requires use of the promoter PRE, which is activated by the product of the cII gene. The product of cIII is required to stabilize the cII product against degradation. By turning off cII and cIII expression, Cro acts to prevent lysogeny. By turning off all transcription except that of its own gene, repressor acts to prevent the lytic cycle. The choice between lysis and lysogeny depends on whether repressor or Cro gains occupancy of the operators in a particular infection. The stability of cII protein in the infected cell is a primary determinant of the outcome. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.3.12.21


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THE REPLICON

4.13.1 Introduction Key Terms The replicon is a unit of the genome in which DNA is replicated. Each replicon contains an origin for initiation of replication. The origin is a sequence of DNA at which replication is initiated. A terminus is a segment of DNA at which replication ends. Single copy replication describes a control system in which there is only one copy of a replicon per unit bacterium. The bacterial chromosome and some plasmids have this type of regulation. A plasmid is said to be under multicopy control when the control system allows the plasmid to exist in more than one copy per individual bacterial cell.

Whether a cell has only one chromosome (as in prokaryotes) or has many chromosomes (as in eukaryotes), the entire genome must be replicated precisely once for every cell division. How is the act of replication linked to the cell cycle? Two general principles are used to compare the state of replication with the condition of the cell cycle: • Initiation of DNA replication commits the cell (prokaryotic or eukaryotic) to a further division. From this standpoint, the number of descendants that a cell generates is determined by a series of decisions on whether or not to initiate DNA replication. Replication is controlled at the stage of initiation. Once replication has started, it continues until the entire genome has been duplicated. • If replication proceeds, the consequent division cannot be permitted to occur until the replication event has been completed. Indeed, the completion of replication may provide a trigger for cell division. Then the duplicate genomes are segregated one to each daughter cell. The unit of segregation is the chromosome. In prokaryotes, the initiation of replication is a single event involving a unique site on the bacterial chromosome, and the process of division is accomplished by the development of a septum that grows from the cell wall and divides the cell into two. In eukaryotic cells, initiation of replication is identified by the start of S phase, a protracted period during which DNA synthesis occurs, and which involves many individual initiation events. The act of division is accomplished by the reorganization of the cell at mitosis. In this chapter, we are concerned with the regulation of DNA replication. How is a cycle of replication initiated? What controls its progress and how is its termination signaled? In Molecular Biology 6.29 Cell cycle and growth regulation, we discuss the regulatory processes in eukaryotic cells that control entry into S phase and into mitosis, and also the "checkpoints" that postpone these actions until the appropriate conditions have been fulfilled. Introduction | SECTION 4.13.1 © 2004. Virtual Text / www.ergito.com

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The unit of DNA in which an individual act of replication occurs is called the replicon. Each replicon "fires" once and only once in each cell cycle. The replicon is defined by its possession of the control elements needed for replication. It has an origin at which replication is initiated. It may also have a terminus at which replication stops (535). Any sequence attached to an origin – or, more precisely, not separated from an origin by a terminus – is replicated as part of that replicon. The origin is a cis-acting site, able to affect only that molecule of DNA on which it resides. (The original formulation of the replicon [in prokaryotes] viewed it as a unit possessing both the origin and the gene coding for the regulator protein. Now, however, "replicon" is usually applied to eukaryotic chromosomes to describe a unit of replication that contains an origin; trans-acting regulator protein(s) may be coded elsewhere.) A genome in a prokaryotic cell constitutes a single replicon; so the units of replication and segregation coincide. Initiation at a single origin sponsors replication of the entire genome, once for every cell division. Each haploid bacterium has a single chromosome, so this type of replication control is called single copy. Bacteria may contain additional genetic information in the form of plasmids. A plasmid is an autonomous circular DNA genome that constitutes a separate replicon (see Figure 12.2). A plasmid replicon may show single copy control, which means that it replicates once every time the bacterial chromosome replicates. Or it may be under multicopy control, when it is present in a greater number of copies than the bacterial chromosome. Each phage or virus DNA also constitutes a replicon, able to initiate many times during an infectious cycle. Perhaps a better way to view the prokaryotic replicon, therefore, is to reverse the definition: any DNA molecule that contains an origin can be replicated autonomously in the cell. A major difference in the organization of bacterial and eukaryotic genomes is seen in their replication. Each eukaryotic chromosome contains a large number of replicons. So the unit of segregation includes many units of replication. This adds another dimension to the problem of control. All the replicons on a chromosome must be fired during one cell cycle, although they are not active simultaneously, but are activated over a fairly protracted period. Yet each of these replicons must be activated no more than once in each cell cycle. Some signal must distinguish replicated from nonreplicated replicons, so that replicons do not fire a second time. And because many replicons are activated independently, another signal must exist to indicate when the entire process of replicating all replicons has been completed. We have begun to collect information about the construction of individual replicons, but we still have little information about the relationship between replicons. We do not know whether the pattern of replication is the same in every cell cycle. Are all origins always used or are some origins sometimes silent? Do origins always fire in the same order? If there are different classes of origins, what distinguishes them? In contrast with nuclear chromosomes, which have a single-copy type of control, the Introduction | SECTION 4.13.1 © 2004. Virtual Text / www.ergito.com

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DNA of mitochondria and chloroplasts may be regulated more like plasmids that exist in multiple copies per bacterium. There are multiple copies of each organelle DNA per cell, and the control of organelle DNA replication must be related to the cell cycle. In all these systems, the key question is to define the sequences that function as origins and to determine how they are recognized by the appropriate proteins of the apparatus for replication. We start by considering the basic construction of replicons and the various forms that they take; following the consideration of the origin, we turn to the question of how replication of the genome is coordinated with bacterial division, and what is responsible for segregating the genomes to daughter bacteria.


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References 535. Jacob, F., Brenner, S., and Cuzin, F. (1963). On the regulation of DNA replication in bacteria. Cold Spring Harbor Symp. Quant. Biol. 28, 329-348.



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THE REPLICON

4.13.2 Replicons can be linear or circular Key Terms A replication eye is a region in which DNA has been replicated within a longer, unreplicated region. A replication fork (Growing point) is the point at which strands of parental duplex DNA are separated so that replication can proceed. A complex of proteins including DNA polymerase is found at the fork. Unidirectional replication refers to the movement of a single replication fork from a given origin. Bidirectional replication describes a system in which an origin generates two replication forks that proceed away from the origin in opposite directions. Key Concepts

• A replicated region appears as an eye within nonreplicated DNA. • A replication fork is initiated at the origin and then moves sequentially along DNA. • Replication is unidirectional when a single replication fork is created at an origin. • Replication is bidirectional when an origin creates two replication forks that move in opposite directions.

A molecule of DNA engaged in replication has two types of regions. Figure 13.1 shows that when replicating DNA is viewed by electron microscopy, the replicated region appears as a replication eye within the nonreplicated DNA. The nonreplicated region consists of the parental duplex; this opens into the replicated region where the two daughter duplexes have formed.

Replicons can be linear or circular | SECTION 4.13.2 © 2004. Virtual Text / www.ergito.com

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Figure 13.1 Replicated DNA is seen as a replication eye flanked by nonreplicated DNA.

The point at which replication is occurring is called the replication fork (sometimes also known as the growing point). A replication fork moves sequentially along the DNA, from its starting point at the origin. The origin may be used to start either unidirectional replication or bidirectional replication. The type of event is determined by whether one or two replication forks set out from the origin. In unidirectional replication, one replication fork leaves the origin and proceeds along the DNA. In bidirectional replication, two replication forks are formed; they proceed away from the origin in opposite directions. The appearance of a replication eye does not distinguish between unidirectional and bidirectional replication. As depicted in Figure 13.2, the eye can represent either of two structures. If generated by unidirectional replication, the eye represents one fixed origin and one moving replication fork. If generated by bidirectional replication, the eye represents a pair of replication forks. In either case, the progress of replication expands the eye until ultimately it encompasses the whole replicon.


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Figure 13.2 Replicons may be unidirectional or bidirectional, depending on whether one or two replication forks are formed at the origin.

When a replicon is circular, the presence of an eye forms the θ-structure drawn in Figure 13.3. The successive stages of replication of the circular DNA of polyoma virus are visualized by electron microscopy in Figure 13.4.

Figure 13.3 A replication eye forms a theta structure in circular DNA.


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Figure 13.4 The replication eye becomes larger as the replication forks proceed along the replicon. Note that the "eye" becomes larger than the nonreplicated segment. The two sides of the eye can be defined because they are both the same length. Photograph kindly provided by Bernard Hirt. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.13.2


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THE REPLICON

4.13.3 Origins can be mapped by autoradiography and electrophoresis Key Concepts

• Replication fork movement can be detected by autoradiography using radioactive pulses.

• Replication forks create Y-shaped structures that change the electrophoretic migration of DNA fragments.

Whether a replicating eye has one or two replication forks can be determined in two ways. The choice of method depends on whether the DNA is a defined molecule or an unidentified region of a cellular genome. With a defined linear molecule, we can use electron microscopy to measure the distance of each end of the eye from the end of the DNA. Then the positions of the ends of the eyes can be compared in molecules that have eyes of different sizes. If replication is unidirectional, only one of the ends will move; the other is the fixed origin. If replication is bidirectional, both will move; the origin is the point midway between them. With undefined regions of large genomes, two successive pulses of radioactivity can be used to label the movement of the replication forks. If one pulse has a more intense label than the other, they can be distinguished by the relative intensities of labeling. These can be visualized by autoradiography. Figure 13.5 shows that unidirectional replication causes one type of label to be followed by the other at one end of the eye. Bidirectional replication produces a (symmetrical) pattern at both ends of the eye. This is the pattern usually observed in replicons of eukaryotic chromosomes (537).

Origins can be mapped by autoradiography and electrophoresis | SECTION 4.13.3 © 2004. Virtual Text / www.ergito.com

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Figure 13.5 Different densities of radioactive labeling can be used to distinguish unidirectional and bidirectional replication.

A more recent method for mapping origins with greater resolution takes advantage of the effects that changes in shape have upon electrophoretic migration of DNA. Figure 13.6 illustrates the two dimensional mapping technique, in which restriction fragments of replicating DNA are electrophoresed in a first dimension that separates by mass, and a second dimension where movement is determined more by shape. Different types of replicating molecules follow characteristic paths, measured by their deviation from the line that would be followed by a linear molecule of DNA that doubled in size.


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Figure 13.6 The position of the origin and the number of replicating forks determine the shape of a replicating restriction fragment, which can be followed by its electrophoretic path (solid line). The dashed line shows the path for a linear DNA.

A simple Y-structure, in which one fork moves along a linear fragment, follows a continuous path. An inflection point occurs when all three branches are the same length, and the structure therefore deviates most extensively from linear DNA. Analogous considerations determine the paths of double Y-structures or bubbles. An asymmetric bubble follows a discontinuous path, with a break at the point at which the bubble is converted to a Y-structure as one fork runs off the end. Taken together, the various techniques for characterizing replicating DNA show that origins are most often used to initiate bidirectional replication. From this level of resolution, we must now proceed to the molecular level, to identify the cis-acting sequences that comprise the origin, and the trans-acting factors that recognize it.


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References 537. Huberman, J. and Riggs, A. D. (1968). On the mechanism of DNA replication in mammalian chromosomes. J. Mol. Biol. 32, 327-341.



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THE REPLICON

4.13.4 The bacterial genome is a single circular replicon Key Concepts

• Bacterial replicons are usually circles that are replicated bidirectionally from a single origin.

• The origin of E. coli,oriC, is 245 bp in length. • The two replication forks usually meet halfway round the circle, but there are ter sites that cause termination if they go too far.

To be properly inherited, a bacterial replicon should support several functions: • Initiating a replication cycle. • Controlling the frequency of initiation events. • Segregating replicated chromosomes to daughter cells. The first two functions both are properties of the origin. Segregation could be an independent function, but in prokaryotic systems it is usually determined by sequences in the vicinity of the origin. Origins in eukaryotes do not function in segregation, but are concerned only with replication. As a general principle, the DNA constituting an origin can be isolated by its ability to support replication of any DNA sequence to which it is joined. When DNA from the origin is cloned into a molecule that lacks an origin, this will create a plasmid capable of autonomous replication only if the DNA from the origin contains all the sequences needed to identify itself as an authentic origin for replication. Origins now have been identified in bacteria, yeast, chloroplasts, and mitochondria, although not in higher eukaryotes. A general feature is that the overall sequence composition is A·T-rich. We assume this is related to the need to melt the DNA duplex to initiate replication (539). The genome of E. coli is replicated bidirectionally from a single origin, identified as the genetic locus oriC. The addition of oriC to any piece of DNA creates an artificial plasmid that can replicate in E. coli. By reducing the size of the cloned fragment of oriC, the region required to initiate replication has been equated with a fragment of 245 bp. (We discuss the properties of oriC and its interaction with the replication apparatus in more detail in Molecular Biology 4.14.15 Creating the replication forks at an origin .) Prokaryotic replicons are usually circular, so that the DNA forms a closed circle The bacterial genome is a single circular replicon | SECTION 4.13.4 © 2004. Virtual Text / www.ergito.com

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with no free ends. Circular structures include the bacterial chromosome itself, all plasmids, and many bacteriophages. They are also common in chloroplast and mitochondrial DNAs. Replication of a circular molecule avoids the problem of how to replicate the ends of a linear molecule, but poses the problem of how to terminate replication (534). The bacterial chromosome is replicated bidirectionally as a single unit from oriC. Two replication forks initiate at oriC and move around the genome (at approximately the same speed) to a meeting point. Termination occurs in a discrete region. One interesting question is what ensures that the DNA is replicated right across the region where the forks meet. Following the termination of DNA replication itself, enzymes that manipulate higher-order structure of DNA are required for the two daughter chromosomes to be physically separated (540). Sequences that cause termination are called ter sites. A ter site contains a short (~23 bp) sequence that causes termination in vitro. The termination sequences function in only one orientation. The ter site is recognized by a protein (called Tus in E. coli and RTP in B. subtilis) that recognizes the consensus sequence and prevents the replication fork from proceeding (see Molecular Biology 4.14.17 The primosome is needed to restart replication). However, deletion of the ter sites does not prevent normal replication cycles from occurring (2223), although it does affect segregation of the daughter chromosomes (see Molecular Biology 4.13.19 Chromosomal segregation may require site-specific recombination). Termination in E. coli and B. subtilis has the interesting features reported in Figure 13.7. We know that the replication forks usually meet and halt replication at a point midway round the chromosome from the origin. But two termination regions (terE,D,A and terC,B in E. coli, and terI, terII and also some other sites in B. subtilis) have been identified, located ~100 kb on either side of this meeting point. Each contains multiple terminators. Each terminus is specific for one direction of fork movement, and they are arranged in such a way that each fork would have to pass the other in order to reach the terminus to which it is susceptible. This arrangement creates a "replication fork trap". If for some reason one fork is delayed, so that the forks fail to meet at the usual central position, the more rapid fork will be trapped at the ter region to wait for the arrival of the slow fork.

The bacterial genome is a single circular replicon | SECTION 4.13.4 © 2004. Virtual Text / www.ergito.com

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Figure 13.7 Replication termini in E. coli are located beyond the point at which the replication forks actually meet.

What happens when a replication fork encounters a protein bound to DNA? We assume that repressors (for example) are displaced and then reattach. A particularly interesting question is what happens when a replication fork encounters an RNA polymerase engaged in transcription. A replication fork moves >10× faster than RNA polymerase. If they are proceeding in the same direction, either the replication fork must displace the polymerase or it must slow down as it waits for the RNA polymerase to reach its terminator. It appears that a DNA polymerase moving in the same direction as an RNA polymerase can "bypass" it without disrupting transcription, but we do not understand how this happens (533; for review see 108). A conflict arises when the replication fork meets an RNA polymerase traveling in the opposite direction, that is, toward it. Can it displace the RNA polymerase? Or do both replication and transcription come to a halt? An indication that these encounters cannot easily be resolved is provided by the organization of the E. coli chromosome. Almost all active transcription units are oriented so that they are expressed in the same direction as the replication fork that passes them. The exceptions all comprise small transcription units that are infrequently expressed. The difficulty of generating inversions containing highly expressed genes argues that head-on encounters between a replication fork and a series of transcribing RNA polymerases may be lethal. Last updated on 12-13-2001


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Reviews 108. Brewer, B. J. (1988). When polymerases collide: replication and transcriptional organization of the E. coli chromosome. Cell 53, 679-686.

References 533. Liu, B., Wong, M. L., and Alberts, B. (1994). A transcribing RNA polymerase molecule survives DNA replication without aborting its growing RNA chain. Proc. Natl. Acad. Sci. USA 91, 10660-10664. 534. Cairns, J. (1963). The bacterial chromosome and its manner of replication as seen by autoradiography. J. Mol. Biol. 6, 208-213. 539. Zyskind, J. W. and Smith, D. W. (1980). Nucleotide sequence of the S. typhimurium origin of DNA replication. Proc. Natl. Acad. Sci. USA 77, 2460-2464. 540. Steck, T. R. and Drlica, K. (1984). Bacterial chromosome segregation: evidence for DNA gyrase involvement in decatenation. Cell 36, 1081-1088. 2223. Iismaa, T. P. and Wake, R. G. (1987). The normal replication terminus of the B. subtilis chromosome, terC, is dispensable for vegetative growth and sporulation. J. Mol. Biol. 195, 299-310.



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THE REPLICON

4.13.5 Each eukaryotic chromosome contains many replicons Key Terms S phase is the restricted part of the eukaryotic cell cycle during which synthesis of DNA occurs. Key Concepts

• Eukaryotic replicons are 40-100 kb in length. • A chromosome is divided into many replicons. • Individual replicons are activated at characteristic times during S phase. • Regional activation patterns suggest that replicons near one another are activated at the same time.

In eukaryotic cells, the replication of DNA is confined to part of the cell cycle. S phase usually lasts a few hours in a higher eukaryotic cell. Replication of the large amount of DNA contained in a eukaryotic chromosome is accomplished by dividing it into many individual replicons. Only some of these replicons are engaged in replication at any point in S phase. Presumably each replicon is activated at a specific time during S phase, although the evidence on this issue is not decisive (for review see 114). The start of S phase is signaled by the activation of the first replicons. Over the next few hours, initiation events occur at other replicons in an ordered manner. Much of our knowledge about the properties of the individual replicons is derived from autoradiographic studies, generally using the types of protocols illustrated in Figure 13.5 and Figure 13.6. Chromosomal replicons usually display bidirectional replication. How large is the average replicon, and how many are there in the genome? A difficulty in characterizing the individual unit is that adjacent replicons may fuse to give large replicated eyes, as illustrated in Figure 13.8. The approach usually used to distinguish individual replicons from fused eyes is to rely on stretches of DNA in which several replicons can be seen to be active, presumably captured at a stage when all have initiated around the same time, but before the forks of adjacent units have met.

Each eukaryotic chromosome contains many replicons | SECTION 4.13.5 © 2004. Virtual Text / www.ergito.com

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Figure 13.8 Measuring the size of the replicon requires a stretch of DNA in which adjacent replicons are active.

In groups of active replicons, the average size of the unit is measured by the distance between the origins (that is, between the midpoints of adjacent replicons). The rate at which the replication fork moves can be estimated from the maximum distance that the autoradiographic tracks travel during a given time. Individual replicons in eukaryotic genomes are relatively small, typically ~40 kb in yeast or fly, ~100 kb in animals cells. However, they can vary >10-fold in length within a genome. The rate of replication is ~2000 bp/min, which is much slower than the 50,000 bp/min of bacterial replication fork movement. From the speed of replication, it is evident that a mammalian genome could be replicated in ~1 hour if all replicons functioned simultaneously. But S phase actually lasts for >6 hours in a typical somatic cell, which implies that no more than 15% of the replicons are likely to be active at any given moment. There are some exceptional cases, such as the early embryonic divisions of Drosophila embryos, where the duration of S phase is compressed by the simultaneous functioning of a large number of replicons (558). How are origins selected for initiation at different times during S phase? In S. cerevisiae, the default appears to be for origins to replicate early, but cis-acting sequences can cause origins linked to them to replicate at late times. Available evidence suggests that chromosomal replicons do not have termini at which the replication forks cease movement and (presumably) dissociate from the Each eukaryotic chromosome contains many replicons | SECTION 4.13.5 © 2004. Virtual Text / www.ergito.com

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DNA. It seems more likely that a replication fork continues from its origin until it meets a fork proceeding toward it from the adjacent replicon. We have already mentioned the potential topological problem of joining the newly synthesized DNA at the junction of the replication forks. The propensity of replicons located in the same vicinity to be active at the same time could be explained by "regional" controls, in which groups of replicons are initiated more or less coordinately, as opposed to a mechanism in which individual replicons are activated one by one in dispersed areas of the genome. Two structural features suggest the possibility of large-scale organization. Quite large regions of the chromosome can be characterized as "early replicating" or "late replicating," implying that there is little interspersion of replicons that fire at early or late times. And visualization of replicating forks by labeling with DNA precursors identifies 100-300 "foci" instead of uniform staining; each focus shown in Figure 13.9 probably contains >300 replication forks. The foci could represent fixed structures through which replicating DNA must move.

Figure 13.9 Replication forks are organized into foci in the nucleus. Cells were labeled with BrdU. The leftmost panel was stained with propidium iodide to identify bulk DNA. The right panel was stained using an antibody to BrdU to identify replicating DNA. Photographs kindly provided by A. D. Mills and Ron Laskey.


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Reviews 114. Fangman, W. L. and Brewer, B. J. (1991). Activation of replication origins within yeast chromosomes. Annu. Rev. Cell Biol. 7, 375-402.

References 558. Blumenthal, A. B., Kriegstein, H. J., and Hogness, D. S. (1974). The units of DNA replication inD. melanogaster chromosomes. Cold Spring Harbor Symp. Quant. Biol. 38, 205-223.



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4.13.6 Replication origins can be isolated in yeast Key Terms ARS (autonomous replication sequence) is an origin for replication in yeast. The common feature among different ARS sequences is a conserved 11 bp sequence called the A-domain. The A domain is the conserved 11 bp sequence of A-T base pairs in the yeast ARS element that comprises the replication origin. Key Concepts

• Origins in S. cerevisiae are short A·T-rich sequences that have an essential 11 bp sequence.

• The ORC is a complex of 6 proteins that binds to an ARS.

Any segment of DNA that has an origin should be able to replicate. So although plasmids are rare in eukaryotes, it may be possible to construct them by suitable manipulation in vitro. This has been accomplished in yeast, although not in higher eukaryotes. S. cerevisiae mutants can be "transformed" to the wild phenotype by addition of DNA that carries a wild-type copy of the gene. The discovery of yeast origins resulted from the observation that some yeast DNA fragments (when circularized) are able to transform defective cells very efficiently. These fragments can survive in the cell in the unintegrated (autonomous) state, that is, as self-replicating plasmids. A high-frequency transforming fragment possesses a sequence that confers the ability to replicate efficiently in yeast. This segment is called an ARS (for autonomously replicating sequence). ARS elements are derived from origins of replication. Where ARS elements have been systematically mapped over extended chromosomal regions, it seems that only some of them are actually used to initiate replication. The others are silent, or possibly used only occasionally. If it is true that some origins have varying probabilities of being used, it follows that there can be no fixed termini between replicons. In this case, a given region of a chromosome could be replicated from different origins in different cell cycles. An ARS element consists of an A·T-rich region that contains discrete sites in which mutations affect origin function. Base composition rather than sequence may be important in the rest of the region. Figure 13.10 shows a systematic mutational analysis along the length of an origin. Origin function is abolished completely by mutations in a 14 bp "core" region, called the A domain, that contains an 11 bp consensus sequence consisting of A·T base pairs. This consensus sequence (sometimes called the ACS for ARS Consensus Sequence) is the only homology Replication origins can be isolated in yeast | SECTION 4.13.6 © 2004. Virtual Text / www.ergito.com

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between known ARS elements (541).

Figure 13.10 An ARS extends for ~50 bp and includes a consensus sequence (A) and additional elements (B1-B3).

Mutations in three adjacent elements, numbered B1-B3, reduce origin function. An origin can function effectively with any 2 of the B elements, so long as a functional A element is present. (Imperfect copies of the core consensus, typically conforming at 9/11 positions, are found close to, or overlapping with, each B element, but they do not appear to be necessary for origin function.) The ORC (origin recognition complex) is a complex of 6 proteins with a mass of ~400 kD (for review see 2222). ORC binds to the A and B1 elements on the A·T-rich strand, and is associated with ARS elements throughout the cell cycle. This means that initiation depends on changes in its condition rather than de novo association with an origin (see Molecular Biology 4.14.21 Licensing factor consists of MCM proteins). By counting the number of sites to which ORC binds, we can estimate that there are about 400 origins of replication in the yeast genome (2247). This means that the average length of a replicon is ~35,000 bp. Counterparts to ORC are found in higher eukaryotic cells (2199). ORC was first found in S. cerevisiae (where it is called scORC), but similar complexes have now been characterized in S. pombe (spORC), Drosophila (DmORC) and Xenopus (XlORC). All of the ORC complexes bind to DNA. Replication origins can be isolated in yeast | SECTION 4.13.6 © 2004. Virtual Text / www.ergito.com

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Although none of the binding sites have been characterized in the same detail as in S. cerevisiae, in several cases they are at locations associated with the initiation of replication. It seems clear that ORC is an initiation complex whose binding identifies an origin of replication (for review see 3089). However, details of the interaction are clear only in S. cerevisiae; it is possible that additional components are required to recognize the origin in the other cases. ARS elements satisfy the classic definition of an origin as a cis-acting sequence that causes DNA replication to initiate. Are similar elements to be found in higher eukaryotes? The conservation of the ORC suggests that origins are likely to take the same sort of form in other eukaryotes, but in spite of this, there is little conservation of sequence among putative origins in different organisms (for review see 4186). Difficulties in finding consensus origin sequences cells suggest the possibility that origins may be more complex (or determined by features other than discrete cis-acting sequences). There are suggestions that some animal cell replicons may have complex patterns of initiation: in some cases, many small replication bubbles are found in one region, posing the question of whether there are alternative or multiple starts to replication, and whether there is a small discrete origin (for review see 116). A reconciliation between this phenomenon and the use of ORCs is suggested by the discovery that environmental effects can influence the use of origins 4185). At one location where multiple bubbles are found, there is a primary origin that is used predominantly when the nucleotide supply is high. But when the nucleotide supply is limiting, many secondary origins are also used, giving rise to a pattern of multiple bubbles. One possible molecular explanation is that ORCs dissociate from the primary origin and initiate elsewhere in the vicinity if the supply of nucleotides is insufficient for the initiation reaction to occur quickly. At all events, it now seems likely that we will be able in due course to characterize discrete sequences that function as origins of replication in higher eukaryotes. Last updated on 9-26-2003

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Reviews 116. DePamphlis, M. L. (1993). Eukaryotic DNA replication: anatomy of an origin. Annu. Rev. Biochem. 62, 29-63. 2222. Kelly, T. J. and Brown, G. W. (2000). Regulation of chromosome replication. Annu. Rev. Biochem. 69, 829-880. 3089. Bell, S. P. and Dutta, A. (2002). DNA replication in eukaryotic cells. Annu. Rev. Biochem. 71, 333-374. 4186. Gilbert, D. M. (2001). Making sense of eukaryotic DNA replication origins. Science 294, 96-100.

References 541. Marahrens, Y. and Stillman, B. (1992). A yeast chromosomal origin of DNA replication defined by multiple functional elements. Science 255, 817-823. 2199. Chesnokov, I., Remus, D., and Botchan, M. (2001). Functional analysis of mutant and wild-type Drosophila origin recognition complex. Proc. Natl. Acad. Sci. USA 98, 11997-12002. 2247. Wyrick, J. J., Aparicio, J. G., Chen, T., Barnett, J. D., Jennings, E. G., Young, R. A., Bell, S. P., and Aparicio, O. M. (2001). Genome-Wide Distribution of ORC and MCM Proteins in S. cerevisiae: High-Resolution Mapping of Replication Origins. Science 294, 2357-2360. 4185. Anglana, M., Apiou, F., Bensimon, A., and Debatisse, M. (2003). Dynamics of DNA replication in mammalian somatic cells: nucleotide pool modulates origin choice and interorigin spacing. Cell 114, 385-394.


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4.13.7 D loops maintain mitochondrial origins Key Terms A D loop is a region within mitochondrial DNA in which a short stretch of RNA is paired with one strand of DNA, displacing the original partner DNA strand in this region. The same term is used also to describe the displacement of a region of one strand of duplex DNA by a complementary single-stranded invader. Key Concepts

• Mitochondria use different origin sequences to initiate replication of each DNA strand.

• Replication of the H-strand is initiated in a D-loop. • Replication of the L-strand is initiated when its origin is exposed by the movement of the first replication fork.

The origins of replicons in both prokaryotic and eukaryotic chromosomes are static structures: they comprise sequences of DNA that are recognized in duplex form and used to initiate replication at the appropriate time. Initiation requires separating the DNA strands and commencing bidirectional DNA synthesis. A different type of arrangement is found in mitochondria. Replication starts at a specific origin in the circular duplex DNA. But initially only one of the two parental strands (the H strand in mammalian mitochondrial DNA) is used as a template for synthesis of a new strand. Synthesis proceeds for only a short distance, displacing the original partner (L) strand, which remains single-stranded, as illustrated in Figure 13.11. The condition of this region gives rise to its name as the displacement or D loop (for review see 105; 113).

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Figure 13.11 The D loop maintains an opening in mammalian mitochondrial DNA, which has separate origins for the replication of each strand.

DNA polymerases cannot initiate synthesis, but require a priming 3 ′ end (see Molecular Biology 4.14.8 Priming is required to start DNA synthesis). Replication at the H strand origin is initiated when RNA polymerase transcribes a primer. 3 ′ ends are generated in the primer by an endonuclease that cleaves the DNA-RNA hybrid at several discrete sites. The endonuclease is specific for the triple structure of DNA-RNA hybrid plus the displaced DNA single strand. The 3 ′ end is then extended into DNA by the DNA polymerase. D loops maintain mitochondrial origins | SECTION 4.13.7 © 2004. Virtual Text / www.ergito.com

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A single D loop is found as an opening of 500-600 bases in mammalian mitochondria. The short strand that maintains the D loop is unstable and turns over; it is frequently degraded and resynthesized to maintain the opening of the duplex at this site. Some mitochondrial DNAs possess several D loops, reflecting the use of multiple origins. The same mechanism is employed in chloroplast DNA, where (in higher plants) there are two D loops. To replicate mammalian mitochondrial DNA, the short strand in the D loop is extended. The displaced region of the original L strand becomes longer, expanding the D loop. This expansion continues until it reaches a point about two-thirds of the way around the circle. Replication of this region exposes an origin in the displaced L strand. Synthesis of an H strand initiates at this site, which is used by a special primase that synthesizes a short RNA. The RNA is then extended by DNA polymerase, proceeding around the displaced single-stranded L template in the opposite direction from L-strand synthesis. Because of the lag in its start, H-strand synthesis has proceeded only a third of the way around the circle when L-strand synthesis finishes. This releases one completed duplex circle and one gapped circle, which remains partially single-stranded until synthesis of the H strand is completed. Finally, the new strands are sealed to become covalently intact (for review see 122). The existence of D loops exposes a general principle. An origin can be a sequence of DNA that serves to initiate DNA synthesis using one strand as template. The opening of the duplex does not necessarily lead to the initiation of replication on the other strand. In the case of mitochondrial DNA replication, the origins for replicating the complementary strands lie at different locations. Origins that sponsor replication of only one strand are also found in the rolling circle mode of replication (see Molecular Biology 4.13.10 Rolling circles produce multimers of a replicon ).


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Reviews 105. Clayton, D. (1982). Replication of animal mitochondrial DNA. Cell 28, 693-705. 113. Clayton, D., A. (1991). Replication and transcription of vertebrate mitochondrial DNA. Annu. Rev. Cell Biol. 7, 453-478. 122. Shadel, G. S. and Clayton, D. A. (1997). Mitochondrial DNA maintenance in vertebrates. Annu. Rev. Biochem. 66, 409-435.



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4.13.8 The ends of linear DNA are a problem for replication Key Concepts

• Special arrangements must be made to replicate the DNA strand with a 5 ′ end.

None of the replicons that we have considered so far have a linear end: either they are circular (as in the E. coli or mitochondrial genomes) or they are part of longer segregation units (as in eukaryotic chromosomes). But linear replicons occur, in some cases as single extrachromosomal units, and of course at the ends of eukaryotic chromosomes. The ability of all known nucleic acid polymerases, DNA or RNA, to proceed only in the 5 ′ – 3 ′ direction poses a problem for synthesizing DNA at the end of a linear replicon. Consider the two parental strands depicted in Figure 13.12. The lower strand presents no problem: it can act as template to synthesize a daughter strand that runs right up to the end, where presumably the polymerase falls off. But to synthesize a complement at the end of the upper strand, synthesis must start right at the very last base (or else this strand would become shorter in successive cycles of replication).

Figure 13.12 Replication could run off the 3 ′ end of a newly synthesized linear strand, but could it initiate at a 5 ′ end?

We do not know whether initiation right at the end of a linear DNA is feasible. We usually think of a polymerase as binding at a site surrounding the position at which a base is to be incorporated. So a special mechanism must be employed for replication at the ends of linear replicons. Several types of solution may be imagined to accommodate the need to copy a terminus: • The problem may be circumvented by converting a linear replicon into a circular or multimeric molecule. Phages such as T4 or lambda use such mechanisms (see The ends of linear DNA are a problem for replication | SECTION 4.13.8 © 2004. Virtual Text / www.ergito.com

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Molecular Biology 4.13.10 Rolling circles produce multimers of a replicon ). • The DNA may form an unusual structure – for example, by creating a hairpin at the terminus, so that there is no free end. Formation of a crosslink is involved in replication of the linear mitochondrial DNA of Paramecium. • Instead of being precisely determined, the end may be variable. Eukaryotic chromosomes may adopt this solution, in which the number of copies of a short repeating unit at the end of the DNA changes (see Molecular Biology 5.19.18 Telomeres are synthesized by a ribonucleoprotein enzyme). A mechanism to add or remove units makes it unnecessary to replicate right up to the very end. • A protein may intervene to make initiation possible at the actual terminus. Several linear viral nucleic acids have proteins that are covalently linked to the 5 ′ terminal base. The best characterized examples are adenovirus DNA, phage φ29 DNA, and poliovirus RNA. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.13.8

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4.13.9 Terminal proteins enable initiation at the ends of viral DNAs Key Terms Strand displacement is a mode of replication of some viruses in which a new DNA strand grows by displacing the previous (homologous) strand of the duplex. A terminal protein allows replication of a linear phage genome to start at the very end. The protein attaches to the 5 ′ end of the genome through a covalent bond, is associated with a DNA polymerase, and contains a cytosine residue that serves as a primer. Key Concepts

• A terminal protein binds to the 5 ′ end of DNA and provides a cytidine nucleotide with a 3 ′ –OH end that primes replication.

An example of initiation at a linear end is provided by adenovirus and φ29 DNAs, which actually replicate from both ends, using the mechanism of strand displacement illustrated in Figure 13.13. The same events can occur independently at either end. Synthesis of a new strand starts at one end, displacing the homologous strand that was previously paired in the duplex. When the replication fork reaches the other end of the molecule, the displaced strand is released as a free single strand. It is then replicated independently; this requires the formation of a duplex origin by base pairing between some short complementary sequences at the ends of the molecule.

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Figure 13.13 Adenovirus DNA replication is initiated separately at the two ends of the molecule and proceeds by strand displacement.

In several viruses that use such mechanisms, a protein is found covalently attached to each 5 ′ end. In the case of adenovirus, a terminal protein is linked to the mature viral DNA via a phosphodiester bond to serine, as indicated in Figure 13.14.


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Figure 13.14 The 5 ′ terminal phosphate at each end of adenovirus DNA is covalently linked to serine in the 55 kD Ad-binding protein.

How does the attachment of the protein overcome the initiation problem? The terminal protein has a dual role: it carries a cytidine nucleotide that provides the primer; and it is associated with DNA polymerase. In fact, linkage of terminal protein to a nucleotide is undertaken by DNA polymerase in the presence of adenovirus DNA. This suggests the model illustrated in Figure 13.15. The complex of polymerase and terminal protein, bearing the priming C nucleotide, binds to the end of the adenovirus DNA. The free 3 ′ –OH end of the C nucleotide is used to prime the elongation reaction by the DNA polymerase. This generates a new strand whose 5 ′ end is covalently linked to the initiating C nucleotide. (The reaction actually involves displacement of protein from DNA rather than binding de novo. The 5 ′ end of adenovirus DNA is bound to the terminal protein that was used in the previous replication cycle. The old terminal protein is displaced by the new terminal protein for each new replication cycle.)


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Figure 13.15 Adenovirus terminal protein binds to the 5 ′ end of DNA and provides a C-OH end to prime synthesis of a new DNA strand.

Terminal protein binds to the region located between 9 and 18 bp from the end of the DNA. The adjacent region, between positions 17 and 48, is essential for the binding of a host protein, nuclear factor I, which is also required for the initiation reaction. The initiation complex may therefore form between positions 9 and 48, a fixed distance from the actual end of the DNA. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.13.9


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4.13.10 Rolling circles produce multimers of a replicon Key Terms The rolling circle is a mode of replication in which a replication fork proceeds around a circular template for an indefinite number of revolutions; the DNA strand newly synthesized in each revolution displaces the strand synthesized in the previous revolution, giving a tail containing a linear series of sequences complementary to the circular template strand. Key Concepts

• A rolling circle generates single-stranded multimers of the original sequence.

The structures generated by replication depend on the relationship between the template and the replication fork. The critical features are whether the template is circular or linear, and whether the replication fork is engaged in synthesizing both strands of DNA or only one. Replication of only one strand is used to generate copies of some circular molecules. A nick opens one strand, and then the free 3 ′ –OH end generated by the nick is extended by the DNA polymerase. The newly synthesized strand displaces the original parental strand. The ensuing events are depicted in Figure 13.16.

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Figure 13.16 The rolling circle generates a multimeric single-stranded tail.

This type of structure is called a rolling circle, because the growing point can be envisaged as rolling around the circular template strand. It could in principle continue to do so indefinitely. As it moves, the replication fork extends the outer strand and displaces the previous partner (549). An example is shown in the electron micrograph of Figure 13.17.


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Figure 13.17 A rolling circle appears as a circular molecule with a linear tail by electron microscopy. Photograph kindly provided by David Dressler.

Because the newly synthesized material is covalently linked to the original material, the displaced strand has the original unit genome at its 5 ′ end. The original unit is followed by any number of unit genomes, synthesized by continuing revolutions of the template. Each revolution displaces the material synthesized in the previous cycle. The rolling circle is put to several uses in vivo. Some pathways that are used to replicate DNA are depicted in Figure 13.18.


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Figure 13.18 The fate of the displaced tail determines the types of products generated by rolling circles. Cleavage at unit length generates monomers, which can be converted to duplex and circular forms. Cleavage of multimers generates a series of tandemly repeated copies of the original unit. Note that the conversion to double-stranded form could occur earlier, before the tail is cleaved from the rolling circle.

Cleavage of a unit length tail generates a copy of the original circular replicon in linear form. The linear form may be maintained as a single strand or may be converted into a duplex by synthesis of the complementary strand (which is identical in sequence to the template strand of the original rolling circle). The rolling circle provides a means for amplifying the original (unit) replicon. This mechanism is used to generate amplified rDNA in the Xenopus oocyte. The genes for rRNA are organized as a large number of contiguous repeats in the genome. A single repeating unit from the genome is converted into a rolling circle. The displaced tail, containing many units, is converted into duplex DNA; later it is cleaved from the circle so that the two ends can be joined together to generate a large circle of amplified rDNA. The amplified material therefore consists of a large number of identical repeating units.


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References 549. Gilbert, W. and Dressler, D. (1968). DNA replication: the rolling circle model. Cold Spring Harbor Symp. Quant. Biol. 33, 473-484.



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4.13.11 Rolling circles are used to replicate phage genomes Key Terms A relaxase is an enzyme that cuts one strand of DNA, and binds to the free 5 ′ end. Key Concepts

• The φX A protein is a cis-acting relaxase that generates single-stranded circles from the tail produced by rolling circle replication.

Replication by rolling circles is common among bacteriophages. Unit genomes can be cleaved from the displaced tail, generating monomers that can be packaged into phage particles or used for further replication cycles. A more detailed view of a phage replication cycle that is centered on the rolling circle is given in Figure 13.19.

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Figure 13.19 φ X174 RF DNA is a template for synthesizing single-stranded viral circles. The A protein remains attached to the same genome through indefinite revolutions, each time nicking the origin on the viral (+) strand and transferring to the new 5 ′ end. At the same time, the released viral strand is circularized. This is a static version of an interactive figure; see http://www.ergito.com/main.jsp?bcs=MBIO.4.13.11 to view properly.

Phage φX174 consists of a single-stranded circular DNA, known as the plus (+) strand. A complementary strand, called the minus (–) strand, is synthesized. This action generates the duplex circle shown at the top of the figure, which is then replicated by a rolling circle mechanism. The duplex circle is converted to a covalently closed form, which becomes supercoiled. A protein coded by the phage genome, the A protein, nicks the (+) strand of the duplex DNA at a specific site that defines the origin for replication. Rolling circles are used to replicate phage genomes | SECTION 4.13.11 © 2004. Virtual Text / www.ergito.com

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Molecular Biology After nicking the origin, the A protein remains connected to the 5 ′ end that it generates, while the 3 ′ end is extended by DNA polymerase. The structure of the DNA plays an important role in this reaction, for the DNA can be nicked only when it is negatively supercoiled (wound about its axis in space in the opposite sense from the handedness of the double helix; see Molecular Biology 4.15.12 Supercoiling affects the structure of DNA). The A protein is able to bind to a single-stranded decamer fragment of DNA that surrounds the site of the nick. This suggests that the supercoiling is needed to assist the formation of a single-stranded region that provides the A protein with its binding site. (An enzymatic activity in which a protein cleaves duplex DNA and binds to a released 5 ′ end is sometimes called a relaxase.) The nick generates a 3 ′ –OH end and a 5 ′ –phosphate end (covalently attached to the A protein), both of which have roles to play in φX174 replication. Using the rolling circle, the 3 ′ –OH end of the nick is extended into a new chain. The chain is elongated around the circular (–) strand template, until it reaches the starting point and displaces the origin. Now the A protein functions again. It remains connected with the rolling circle as well as to the 5 ′ end of the displaced tail, and it is therefore in the vicinity as the growing point returns past the origin. So the same A protein is available again to recognize the origin and nick it, now attaching to the end generated by the new nick. The cycle can be repeated indefinitely. Following this nicking event, the displaced single (+) strand is freed as a circle. The A protein is involved in the circularization. In fact, the joining of the 3 ′ and 5 ′ ends of the (+) strand product is accomplished by the A protein as part of the reaction by which it is released at the end of one cycle of replication, and starts another cycle. The A protein has an unusual property that may be connected with these activities. It is cis-acting in vivo. (This behavior is not reproduced in vitro, as can be seen from its activity on any DNA template in a cell-free system.) The implication is that in vivo the A protein synthesized by a particular genome can attach only to the DNA of that genome. We do not know how this is accomplished. However, its activity in vitro shows how it remains associated with the same parental (–) strand template. The A protein has two active sites; this may allow it to cleave the "new" origin while still retaining the "old" origin; then it ligates the displaced strand into a circle. The displaced (+) strand may follow either of two fates after circularization. During the replication phase of viral infection, it may be used as a template to synthesize the complementary (–) strand. The duplex circle may then be used as a rolling circle to generate more progeny. During phage morphogenesis, the displaced (+) strand is packaged into the phage virion. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.13.11

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4.13.12 The F plasmid is transferred by conjugation between bacteria Key Terms Conjugation is a process in which two cells come in contact and exchange genetic material. In bacteria, DNA is transferred from a donor to a recipient cell. In protozoa, DNA passes from each cell to the other. The F plasmid is an episome that can be free or integrated in E. coli, and which in either form can sponsor conjugation. The transfer region is a segment on the F plasmid that is required for bacterial conjugation. A pilus (pili) is a surface appendage on a bacterium that allows the bacterium to attach to other bacterial cells. It appears like a short, thin, flexible rod. During conjugation, pili are used to transfer DNA from one bacterium to another. Pilin is the subunit that is polymerized into the pilus in bacteria. Key Concepts

• A free F factor is a replicon that is maintained at the level of one plasmid per bacterial chromosome.

• An F factor can integrate into the bacterial chromosome, in which case its own replication system is suppressed.

• The F factor codes for specific pili that form on the surface of the bacterium. • An F-pilus enables an F-positive bacterium to contact an F-negative bacterium and to initiate conjugation.

Another example of a connection between replication and the propagation of a genetic unit is provided by bacterial conjugation, in which a plasmid genome or host chromosome is transferred from one bacterium to another. Conjugation is mediated by the F plasmid, which is the classic example of an episome, an element that may exist as a free circular plasmid, or that may become integrated into the bacterial chromosome as a linear sequence (like a lysogenic bacteriophage). The F plasmid is a large circular DNA, ~100 kb in length. The F factor can integrate at several sites in the E. coli chromosome, often by a recombination event involving certain sequences (called IS sequences; see Molecular Biology 4.16.5 Transposons cause rearrangement of DNA) that are present on both the host chromosome and F plasmid. In its free (plasmid) form, the F plasmid utilizes its own replication origin (oriV) and control system, and is maintained at a level of one copy per bacterial chromosome. When it is integrated into the bacterial chromosome, this system is suppressed, and F DNA is replicated as a part of the The F plasmid is transferred by conjugation between bacteria | SECTION 4.13.12 © 2004. Virtual Text / www.ergito.com

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chromosome. The presence of the F plasmid, whether free or integrated, has important consequences for the host bacterium. Bacteria that are F-positive are able to conjugate (or mate) with bacteria that are F-negative. Conjugation involves a contact between donor (F-positive) and recipient (F-negative) bacteria; contact is followed by transfer of the F factor. If the F factor exists as a free plasmid in the donor bacterium, it is transferred as a plasmid, and the infective process converts the F-negative recipient into an F-positive state. If the F factor is present in an integrated form in the donor, the transfer process may also cause some or all of the bacterial chromosome to be transferred. Many plasmids have conjugation systems that operate in a generally similar manner, but the F factor was the first to be discovered, and remains the paradigm for this type of genetic transfer (550). A large (~33 kb) region of the F plasmid, called the transfer region, is required for conjugation. It contains ~40 genes that are required for the transmission of DNA; their organization is summarized in Figure 13.20. The genes are named as tra and trb loci. Most of them are expressed coordinately as part of a single 32 kb transcription unit (the traY-I unit). traM and traJ are expressed separately. traJ is a regulator that turns on both traM and traY-I. On the opposite strand, finP is a regulator that codes for a small antisense RNA that turns off traJ. Its activity requires expression of another gene, finO. Only four of the tra genes in the major transcription unit are concerned directly with the transfer of DNA; most are concerned with the properties of the bacterial cell surface and with maintaining contacts between mating bacteria.

Figure 13.20 The tra region of the F plasmid contains the genes needed for bacterial conjugation.

F-positive bacteria possess surface appendages called pili (singular pilus) that are coded by the F factor. The gene traA codes for the single subunit protein, pilin, that is polymerized into the pilus. At least 12 tra genes are required for the modification and assembly of pilin into the pilus. The F-pili are hair-like structures, 2-3 µm long, that protrude from the bacterial surface. A typical F-positive cell has 2-3 pili. The pilin subunits are polymerized into a hollow cylinder, ~8 nm in diameter, with a 2 nm axial hole. Mating is initiated when the tip of the F-pilus contacts the surface of the recipient cell. Figure 13.21 shows an example of E. coli cells beginning to mate. A donor cell does not contact other cells carrying the F factor, because the genes traS and traT The F plasmid is transferred by conjugation between bacteria | SECTION 4.13.12 © 2004. Virtual Text / www.ergito.com

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code for "surface exclusion" proteins that make the cell a poor recipient in such contacts. This effectively restricts donor cells to mating with F-negative cells. (And the presence of F-pili has secondary consequences; they provide the sites to which RNA phages and some single-stranded DNA phages attach, so F-positive bacteria are susceptible to infection by these phages, whereas F-negative bacteria are resistant.)

Figure 13.21 Mating bacteria are initially connected when donor F pili contact the recipient bacterium. Photograph kindly provided by Ron Skurray.

The initial contact between donor and recipient cells is easily broken, but other tra genes act to stabilize the association, bringing the mating cells closer together. The F pili are essential for initiating pairing, but retract or disassemble as part of the process by which the mating cells are brought into close contact. There must be a channel through which DNA is transferred, but the pilus itself does not appear to provide it. TraD is an inner membrane protein in F+ bacteria that is necessary for transport of DNA and it may provide or be part of the channel.

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References 550. Ihler, G. and Rupp, W. D. (1969). Strand-specific transfer of donor DNA during conjugation in E. coli. Proc. Natl. Acad. Sci. USA 63, 138-143.


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THE REPLICON

4.13.13 Conjugation transfers single-stranded DNA Key Terms An Hfr cell is a bacterium that has an integrated F plasmid within its chromosome. Hfr stands for high frequency recombination, referring to the fact that chromosomal genes are transferred from an Hfr cell to an F# cell much more frequently than from an F+ cell. Key Concepts

• Transfer of an F factor is initiated when rolling circle replication begins at oriT. • The free 5 ′ end initiates transfer into the recipient bacterium. • The transferred DNA is converted into double-stranded form in the recipient bacterium.

• When an F factor is free, conjugation "infects" the recipient bacterium with a copy of the F factor.

• When an F factor is integrated, conjugation causes transfer of the bacterial

chromosome until the process is interrupted by (random) breakage of the contact between donor and recipient bacteria.

Transfer of the F factor is initiated at a site called oriT, the origin of transfer, which is located at one end of the transfer region. The transfer process may be initiated when TraM recognizes that a mating pair has formed. Then TraY binds near oriT and causes TraI to bind. TraI is a relaxase, like φX174 A protein. TraI nicks oriT at a unique site (called nic), and then forms a covalent link to the 5 ′ end that has been generated. TraI also catalyzes the unwinding of ~200 bp of DNA (this is a helicase activity; see Molecular Biology 4.14.7 The φX model system shows how single-stranded DNA is generated for replication). Figure 13.22 shows that the freed 5 ′ end leads the way into the recipient bacterium. A complement for the transferred single strand is synthesized in the recipient bacterium, which as a result is converted to the F-positive state (for review see 138).

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Figure 13.22 Transfer of DNA occurs when the F factor is nicked at oriT and a single strand is led by the 5 ′ end into the recipient. Only one unit length is transferred. Complementary strands are synthesized to the single strand remaining in the donor and to the strand transferred into the recipient.

A complementary strand must be synthesized in the donor bacterium to replace the strand that has been transferred. If this happens concomitantly with the transfer process, the state of the F plasmid will resemble the rolling circle of Figure 13.16 (and will not generate the extensive single-stranded regions shown in Figure 13.22). Conjugating DNA usually appears like a rolling circle, but replication as such is not necessary to provide the driving energy, and single-strand transfer is independent of DNA synthesis. Only a single unit length of the F factor is transferred to the recipient bacterium. This implies that some (unidentified) feature terminates the process after one revolution, after which the covalent integrity of the F plasmid is restored (for review see 106; 111). Conjugation transfers single-stranded DNA | SECTION 4.13.13 © 2004. Virtual Text / www.ergito.com

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When an integrated F plasmid initiates conjugation, the orientation of transfer is directed away from the transfer region, into the bacterial chromosome. Figure 13.23 shows that, following a short leading sequence of F DNA, bacterial DNA is transferred. The process continues until it is interrupted by the breaking of contacts between the mating bacteria. It takes ~100 minutes to transfer the entire bacterial chromosome, and under standard conditions, contact is often broken before the completion of transfer (for review see 120).

Figure 13.23 Transfer of chromosomal DNA occurs when an integrated F factor is nicked at oriT. Transfer of DNA starts with a short sequence of F DNA and continues until prevented by loss of contact between the bacteria.

Donor DNA that enters a recipient bacterium is converted to double-stranded form, and may recombine with the recipient chromosome. (Note that two recombination events are required to insert the donor DNA.) So conjugation affords a means to exchange genetic material between bacteria (a contrast with their usual asexual growth). A strain of E. coli with an integrated F factor supports such recombination at relatively high frequencies (compared to strains that lack integrated F factors); Conjugation transfers single-stranded DNA | SECTION 4.13.13 © 2004. Virtual Text / www.ergito.com

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such strains are described as Hfr (for high frequency recombination). Each position of integration for the F factor gives rise to a different Hfr strain, with a characteristic pattern of transferring bacterial markers to a recipient chromosome. Contact between conjugating bacteria is usually broken before transfer of DNA is complete. As a result, the probability that a region of the bacterial chromosome will be transferred depends upon its distance from oriT. Bacterial genes located close to the site of F integration (in the direction of transfer) enter recipient bacteria first, and are therefore found at greater frequencies than those located farther away that enter later. This gives rise to a gradient of transfer frequencies around the chromosome, declining from the position of F integration. Marker positions on the donor chromosome can be assayed in terms of the time at which transfer occurs, and this gave rise to the standard description of the E. coli chromosome as a map divided into 100 minutes. The map refers to transfer times from a particular Hfr strain; the starting point for the gradient of transfer is different for each Hfr strain, being determined by the site where the F factor has integrated into the bacterial genome.


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Reviews 106. Ippen-Ihler, K. A. and Minkley, E. G. (1986). The conjugation system of F, the fertility factor of E. coli. Annu. Rev. Genet. 20, 593-624. 111. Willetts, N. and Skurray, R. (1987). Structure and function of the F factor and mechanism of conjugation. E. coli and S. typhimurium, 1110-1131. 120. Lanka, E. and Wilkins, B. M. (1995). DNA processing reactions in bacterial conjugation. Annu. Rev. Biochem. 64, 141-169. 138. Frost, L. S., Ippen-Ihler, K., and Skurray, R. A. (1994). Analysis of the sequence and gene products of the transfer region of the F sex factor. Microbiol. Rev. 58, 162-210.



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THE REPLICON

4.13.14 Replication is connected to the cell cycle Key Terms The doubling time is the period (usually measured in minutes) that it takes for a bacterial cell to reproduce. A multiforked chromosome (in bacterium) has more than one replication fork, because a second initiation has occurred before the first cycle of replication has been completed. The unit cell describes the state of an E. coli bacterium generated by a new division. It is 1.7 µm long and has a single replication origin. Key Concepts

• The doubling time of E. coli can vary over a 10× range, depending on growth conditions.

• It requires 40 minutes to replicate the bacterial chromosome (at normal temperature).

• Completion of a replication cycle triggers a bacterial division 20 minutes later. • If the doubling time is 60 minutes.) A cycle of chromosome replication must be initiated a fixed time before a cell division, C + D = 60 minutes. For bacteria dividing more frequently than every 60 minutes, a cycle of replication must be initiated before the end of the preceding division cycle. Consider the example of cells dividing every 35 minutes. The cycle of replication connected with a division must have been initiated 25 minutes before the preceding division. This situation is illustrated in Figure 13.24, which shows the chromosomal complement of a bacterial cell at 5-minute intervals throughout the cycle.

Figure 13.24 The fixed interval of 60 minutes between initiation of replication and cell division produces multiforked chromosomes in rapidly growing cells. Note that only the replication forks moving in one direction are shown; actually the chromosome is replicated symmetrically by two sets of forks moving in opposite directions on circular chromosomes.

At division (35/0 minutes), the cell receives a partially replicated chromosome. The Replication is connected to the cell cycle | SECTION 4.13.14 © 2004. Virtual Text / www.ergito.com

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replication fork continues to advance. At 10 minutes, when this "old" replication fork has not yet reached the terminus, initiation occurs at both origins on the partially replicated chromosome. The start of these "new" replication forks creates a multiforked chromosome. At 15 minutes – that is, at 20 minutes before the next division – the old replication fork reaches the terminus. Its arrival allows the two daughter chromosomes to separate; each of them has already been partially replicated by the new replication forks (which now are the only replication forks). These forks continue to advance. At the point of division, the two partially replicated chromosomes segregate. This recreates the point at which we started. The single replication fork becomes "old," it terminates at 15 minutes, and 20 minutes later there is a division. We see that the initiation event occurs 1 25/35 cell cycles before the division event with which it is associated. The general principle of the link between initiation and the cell cycle is that, as cells grow more rapidly (the cycle is shorter), the initiation event occurs an increasing number of cycles before the related division. There are correspondingly more chromosomes in the individual bacterium. This relationship can be viewed as the cell's response to its inability to reduce the periods of C and D to keep pace with the shorter cycle. How does the cell know when to initiate the replication cycle? The initiation event occurs at a constant ratio of cell mass to the number of chromosome origins. Cells growing more rapidly are larger and possess a greater number of origins. The growth of the bacterium can be described in terms of the unit cell, an entity 1.7 µm long. A bacterium contains one origin per unit cell; a rapidly growing cell with two origins will be 1.7-3.4 µm long. In terms of Figure 13.24, it is at the point 10 minutes after division that the cell mass has increased sufficiently to support an initiation event at both available origins (543; 544). How is cell mass titrated? An initiator protein could be synthesized continuously throughout the cell cycle; accumulation of a critical amount would trigger initiation. This explains why protein synthesis is needed for the initiation event. An alternative possibility is that an inhibitor protein might be synthesized at a fixed point, and diluted below an effective level by the increase in cell volume (for review see 117).

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Reviews 117. Donachie, W. D. (1993). The cell cycle of E. coli. Annu. Rev. Immunol. 47, 199-230.

References 543. Donachie, W. D. and Begg, K. J. (1970). Growth of the bacterial cell. Nature 227, 1220-1224. 544. Donachie, W. D., Begg, K. J., and Vicente, M. (1976). Cell length, cell growth and cell division. Nature 264, 328-333.


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THE REPLICON

4.13.15 The septum divides a bacterium into progeny each containing a chromosome Key Terms A septum is the structure that forms in the center of a dividing bacterium, providing the site at which the daughter bacteria will separate. The same term is used to describe the cell wall that forms between plant cells at the end of mitosis. A periseptal annulus is an ring-like area where inner and outer membrane appear fused. Formed around the circumference of the bacterium, the periseptal annulus determines the location of the septum. Key Concepts

• Septum formation is initiated at the annulus, which is a ring around the cell where the structure of the envelope is altered.

• New annuli are initiated at 50% of the distance from the septum to each end of the bacterium.

• When the bacterium divides, each daughter has an annulus at the mid center position.

• Septation starts when the cell reaches a fixed length. • The septum consists of the same peptidoglycans that comprise the bacterial envelope.

Chromosome segregation in bacteria is especially interesting because the DNA itself is involved in the mechanism for partition. (This contrasts with eukaryotic cells, in which segregation is achieved by the complex apparatus of mitosis.) The bacterial apparatus is quite accurate, however; anucleate cells form 1 plasmid copy per bacterial chromosome origin. • Homologous recombination between circular plasmids generates dimers and higher multimers.

• Plasmids have site-specific recombination systems that undertake intramolecular recombination to regenerate monomers.

• Partition systems ensure that duplicate plasmids are segregated to different daughter cells produced by a division.

The type of system that a plasmid uses to ensure that it is distributed to both daughter cells at division depends upon its type of replication system. Each type of plasmid is maintained in its bacterial host at a characteristic copy number: • Single-copy control systems resemble that of the bacterial chromosome and result in one replication per cell division. A single-copy plasmid effectively maintains parity with the bacterial chromosome. • Multicopy control systems allow multiple initiation events per cell cycle, with the result that there are several copies of the plasmid per bacterium. Multicopy plasmids exist in a characteristic number (typically 10-20) per bacterial chromosome. Copy number is primarily a consequence of the type of replication control mechanism. The system responsible for initiating replication determines how many origins can be present in the bacterium. Since each plasmid consists of a single replicon, the number of origins is the same as the number of plasmid molecules. Single-copy plasmids have a system for replication control whose consequences are similar to that governing the bacterial chromosome. A single origin can be replicated once; then the daughter origins are segregated to the different daughter cells. Single-copy plasmids have a partitioning system | SECTION 4.13.21 © 2004. Virtual Text / www.ergito.com

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Multicopy plasmids have a replication system that allows a pool of origins to exist. If the number is great enough (in practice >10 per bacterium), an active segregation system becomes unnecessary, because even a statistical distribution of plasmids to daughter cells will result in the loss of plasmids at frequencies 10 copies are required to ensure that each daughter gains at least one copy (see Molecular Biology 4.13.21 Single-copy plasmids have a partitioning system). When there are mtDNAs with allelic variations (either because of inheritance from different parents or because of mutation), the stochastic distribution may generate cells that have only one of the alleles. Replication of mtDNA may be stochastic because there is no control over which particular copies are replicated, so that in any cycle some mtDNA molecules may replicate more times than others. The total number of copies of the genome may be controlled by titrating mass in a way similar to bacteria (see Molecular Biology 4.13.14 Replication is connected to the cell cycle ). A mitochondrion divides by developing a ring around the organelle that constricts to pinch it into two halves. The mechanism is similar in principle to that involved in bacterial division. The apparatus that is used in plant cell mitochondria is similar to bacteria and uses a homologue of the bacterial protein FtsZ (see Molecular Biology 4.13.17 FtsZ is necessary for septum formation). The molecular apparatus is different in animal cell mitochondria, and uses the protein dynamin that is involved in formation of membranous vesicles (see Molecular Biology 6.27.5 Different types of coated vesicles exist in each pathway). An individual organelle may have more than one copy of its genome. We do not know whether there is a partitioning mechanism for segregating mtDNA molecules within the mitochondrion, or whether they are simply inherited by daughter mitochondria according to which half of the mitochondrion they happen to lie in. Figure 13.44 shows that the combination of replication and segregation mechanisms can result in a stochastic assignment of DNA to each of the copies, that is, so that the distribution of mitochondrial genomes to daughter mitochondria does not depend on their parental origins. How do mitochondria replicate and segregate? | SECTION 4.13.24 © 2004. Virtual Text / www.ergito.com

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Figure 13.44 Mitochondrial DNA replicates by increasing the number of genomes in proportion to mitochondrial mass, but without ensuring that each genome replicates the same number of times. This can lead to changes in the representation of alleles in the daughter mitochondria.

The assignment of mitochondria to daughter cells at mitosis also appears to be random. Indeed, it was the observation of somatic variation in plants that first suggested the existence of genes that could be lost from one of the daughter cells because they were not inherited according to Mendel's laws (see Figure 3.37). In some situations a mitochondrion has both paternal and maternal alleles. This has two requirements: that both parents provide alleles to the zygote (which of course is not the case when there is maternal inheritance; see Molecular Biology 1.3.19 Organelles have DNA); and that the parental alleles are found in the same mitochondrion. For this to happen, parental mitochondria must have fused. The size of the individual mitochondrion may not be precisely defined. Indeed, there is a continuing question as to whether an individual mitochondrion represents a unique and discrete copy of the organelle or whether it is in a dynamic flux in which it can fuse with other mitochondria. We know that mitochondria can fuse in yeast, because recombination between mtDNAs can occur after two haploid yeast strains How do mitochondria replicate and segregate? | SECTION 4.13.24 © 2004. Virtual Text / www.ergito.com

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have mated to produce a diploid strain. This implies that the two mtDNAs must have been exposed to one another in the same mitochondrial compartment. Attempts have been made to test for the occurrence of similar events in animal cells by looking for complementation between alleles after two cells have been fused, but the results are not clear. Last updated on 2-8-2002

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Reviews 2288. Birky, C. W. (2001). The inheritance of genes in mitochondria and chloroplasts: laws, mechanisms, and models. Annu. Rev. Genet. 35, 125-148.


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THE REPLICON

4.13.25 Summary The entire chromosome is replicated once for every cell division cycle. Initiation of replication commits the cell to a cycle of division; completion of replication may provide a trigger for the actual division process. The bacterial chromosome consists of a single replicon, but a eukaryotic chromosome is divided into many replicons that function over the protracted period of S phase. The problem of replicating the ends of a linear replicon is solved in a variety of ways, most often by converting the replicon to a circular form. Some viruses have special proteins that recognize ends. Eukaryotic chromosomes encounter the problem at their terminal replicons. Eukaryotic replication is (at least) an order of magnitude slower than bacterial replication. Origins sponsor bidirectional replication, and are probably used in a fixed order during S phase. The only eukaryotic origins identified at the sequence level are those of S. cerevisiae, which have a core consensus sequence consisting of 11 base pairs, mostly A·T. The minimal E. coli origin consists of ~245 bp and initiates bidirectional replication. Any DNA molecule with this sequence can replicate in E. coli. Two replication forks leave the origin and move around the chromosome, apparently until they meet, although ter sequences that would cause the forks to terminate after meeting have been identified. Transcription units are organized so that transcription usually proceeds in the same direction as replication. The rolling circle is an alternative form of replication for circular DNA molecules in which an origin is nicked to provide a priming end. One strand of DNA is synthesized from this end, displacing the original partner strand, which is extruded as a tail. Multiple genomes can be produced by continuing revolutions of the circle. Rolling circles are used to replicate some phages. The A protein that nicks the φX174 origin has the unusual property of cis-action. It acts only on the DNA from which it was synthesized. It remains attached to the displaced strand until an entire strand has been synthesized, and then nicks the origin again, releasing the displaced strand and starting another cycle of replication. Rolling circles also are involved in bacterial conjugation, when an F plasmid is transferred from a donor to a recipient cell, following the initiation of contact between the cells by means of the F-pili. A free F plasmid infects new cells by this means; an integrated F factor creates an Hfr strain that may transfer chromosomal DNA. In the case of conjugation, replication is used to synthesize complements to the single strand remaining in the donor and to the single strand transferred to the recipient, but does not provide the motive power. A fixed time of 40 minutes is required to replicate the E. coli chromosome and a further 20 minutes is required before the cell can divide. When cells divide more rapidly than every 60 minutes, a replication cycle is initiated before the end of the preceding division cycle. This generates multiforked chromosomes. The initiation Summary | SECTION 4.13.25 © 2004. Virtual Text / www.ergito.com

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event depends on titration of cell mass, probably by accumulating an initiator protein. Initiation may occur at the cell membrane, since the origin is associated with the membrane for a short period after initiation. The septum that divides the cell grows at a location defined by the pre-existing periseptal annulus; a locus of three genes (minCDE) codes for products that regulate whether the midcell periseptal annulus or the polar sites derived from previous annuli are used for septum formation. Absence of septum formation generates multinucleated filaments; excess of septum formation generates anucleate minicells. Many transmembrane proteins interact to form the septum. ZipA is located in the inner bacterial membrane and binds to FtsZ, which is a tubulin-like protein that can polymerize into a filamentous structure called a Z-ring. FtsA is a cytosolic protein that binds to FtsZ. Several other fts products, all transmembrane proteins, join the Z-ring in an ordered process that generates a septal ring. The last proteins to bind are the SEDS protein FtsW and the transpeptidase ftsI (PBP3), which together function to produce the peptidoglycans of the septum. Chloroplasts use a related division mechanism that has an FtsZ-like protein, but mitochondria use a different process in which the membrane is constricted by a dynamin-like protein. Plasmids and bacteria have site-specific recombination systems that regenerate pairs of monomers by resolving dimers created by general recombination. The Xer system acts on a target sequence located in the terminus region of the chromosome. The system is active only in the presence of the FtsK protein of the septum, which may ensure that it acts only when a dimer needs to be resolved. Partitioning involves the interaction of the ParB protein with the parS target site to build a structure that includes the IHF protein. This partition complex ensures that replica chromosomes segregate into different daughter cells. The mechanism of segregation may involve movement of DNA, possibly by the action of MukB in condensing chromosomes into masses at different locations as they emerge from replication. Plasmids have a variety of systems that ensure or assist partition, and an individual plasmid may carry systems of several types. The copy number of a plasmid describes whether it is present at the same level as the bacterial chromosome (one per unit cell) or in greater numbers. Plasmid incompatibility can be a consequence of the mechanisms involved in either replication or partition (for single-copy plasmids). Two plasmids that share the same control system for replication are incompatible because the number of replication events ensures that there is only one plasmid for each bacterial genome. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.13.25


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DNA REPLICATION

4.14.1 Introduction Key Terms The replisome is the multiprotein structure that assembles at the bacterial replicating fork to undertake synthesis of DNA. It contains DNA polymerase and other enzymes. A dna mutant of bacteria is temperature-sensitive; it cannot synthesize DNA at 42°C, but can do so at 37°C. A quick-stop mutant is a type of DNA replication temperature-sensitive mutant (dna )in E. coli that immediately stops DNA replication when the temperature is increased to 42°C. A slow-stop mutant is a type of DNA replication temperature-sensitive mutant in E. coli that can finish a round of replication at the unpermissive temperature, but cannot start another. In vitro complementation is a functional assay used to identify components of a process. The reaction is reconstructed using extracts from a mutant cell. Fractions from wild-type cells are then tested for restoration of activity.

Replication of duplex DNA is a complex endeavor involving a conglomerate of enzyme activities. Different activities are involved in the stages of initiation, elongation, and termination. • Initiation involves recognition of an origin by a complex of proteins. Before DNA synthesis begins, the parental strands must be separated and (transiently) stabilized in the single-stranded state. Then synthesis of daughter strands can be initiated at the replication fork. • Elongation is undertaken by another complex of proteins. The replisome exists only as a protein complex associated with the particular structure that DNA takes at the replication fork. It does not exist as an independent unit (for example, analogous to the ribosome). As the replisome moves along DNA, the parental strands unwind and daughter strands are synthesized. • At the end of the replicon, joining and/or termination reactions are necessary. Following termination, the duplicate chromosomes must be separated from one another, which requires manipulation of higher-order DNA structure. Inability to replicate DNA is fatal for a growing cell. Mutants in replication must therefore be obtained as conditional lethals. These are able to accomplish replication under permissive conditions (provided by the normal temperature of incubation), but they are defective under nonpermissive conditions (provided by the higher temperature of 42°C). A comprehensive series of such temperature-sensitive mutants in E. coli identifies a set of loci called the dna genes. The dna mutants distinguish two stages of replication by their behavior when the temperature is raised (545): Introduction | SECTION 4.14.1 © 2004. Virtual Text / www.ergito.com

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• The major class of quick-stop mutants cease replication immediately on a temperature rise. They are defective in the components of the replication apparatus, typically in the enzymes needed for elongation (but also include defects in the supply of essential precursors). • The smaller class of slow-stop mutants complete the current round of replication, but cannot start another. They are defective in the events involved in initiating a cycle of replication at the origin. An important assay used to identify the components of the replication apparatus is called in vitro complementation. An in vitro system for replication is prepared from a dna mutant and operated under conditions in which the mutant gene product is inactive. Extracts from wild-type cells are tested for their ability to restore activity. The protein coded by the dna locus can be purified by identifying the active component in the extract. Each component of the bacterial replication apparatus is now available for study in vitro as a biochemically pure product, and is implicated in vivo by mutations in its gene. Eukaryotic replication systems are highly purified, and usually have components analogous to the bacterial proteins, but have not necessarily reached the stage of identification of every single component.


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References 545. Hirota, Y., Ryter, A., and Jacob, F. (1968). Thermosensitive mutants of E. coli affected In the processes of DNA synthesis and cellular division. Cold Spring Harbor Symp. Quant. Biol. 33, 677-693.



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DNA REPLICATION

4.14.2 DNA polymerases are the enzymes that make DNA Key Terms Replication of duplex DNA takes place by synthesis of two new strands that are complementary to the parental strands. The parental duplex is replaced by two identical daughter duplexes, each of which has one parental strand and one newly synthesized strand. Replication is called semiconservative because the conserved units are the single strands of the parental duplex. Repair of damaged DNA can take place by repair synthesis, when a strand that has been damaged is excised and replaced by the synthesis of a new stretch. It can also take place by recombination reactions, when the duplex region containing the damaged is replaced by an undamaged region from another copy of the genome. A DNA polymerase is an enzyme that synthesizes a daughter strand(s) of DNA (under direction from a DNA template). Any particular enzyme may be involved in repair or replication (or both). A DNA replicase is a DNA-synthesizing enzyme required specifically for replication. Key Concepts

• DNA is synthesized in both semiconservative replication and repair reactions. • A bacterium or eukaryotic cell has several different DNA polymerase enzymes. • One bacterial DNA polymerase undertakes semiconservative replication; the others are involved in repair reactions.

• Eukaryotic nuclei, mitochondria, and chloroplasts each have a single unique DNA polymerase required for replication, and other DNA polymerases involved in ancillary or repair activities.

There are two basic types of DNA synthesis. Figure 14.1 shows the result of semiconservative replication. The two strands of the parental duplex are separated, and each serves as a template for synthesis of a new strand. The parental duplex is replaced with two daughter duplexes, each of which has one parental strand and one newly synthesized strand.

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Figure 14.1 Semiconservative replication synthesizes two new strands of DNA.

Figure 14.2 shows the consequences of a repair reaction. One strand of DNA has been damaged. It is excised and new material is synthesized to replace it. (Repair synthesis is not the only way to replace damaged DNA; the reactions involved in replacement of damaged sequences are discussed in Molecular Biology 4.15 Recombination and repair.)

Figure 14.2 Repair synthesis replaces a short stretch of one strand of DNA containing a damaged base.

An enzyme that can synthesize a new DNA strand on a template strand is called a DNA polymerase. Both prokaryotic and eukaryotic cells contain multiple DNA polymerase activities. Only some of these enzymes actually undertake replication; sometimes they are called DNA replicases. The others are involved in subsidiary roles in replication and/or participate in repair synthesis. All prokaryotic and eukaryotic DNA polymerases share the same fundamental type of synthetic activity. Each can extend a DNA chain by adding nucleotides one at a time to a 3 ′ –OH end, as illustrated diagrammatically in Figure 14.3. The choice of the nucleotide to add to the chain is dictated by base pairing with the template strand.


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Figure 14.3 DNA synthesis occurs by adding nucleotides to the 3 ′ -OH end of the growing chain, so that the new chain grows in the 5 ′ → 3 ′ direction. The precursor for DNA synthesis is a nucleoside triphosphate, which loses the terminal two phosphate groups in the reaction.

Some DNA polymerases function as independent enzymes, but others (most notably the replicases) are incorporated into large protein assemblies. The DNA-synthesizing subunit is only one of several functions of the replicase, which typically contains many other activities concerned with unwinding DNA, initiating new strand synthesis, and so on. Figure 14.4 summarizes the DNA polymerases that have been characterized in E. coli. DNA polymerase III, a multisubunit protein, is the replicase responsible for de novo synthesis of new strands of DNA. DNA polymerase I (coded by polA) is involved in the repair of damaged DNA and, in a subsidiary role, in semiconservative replication. DNA polymerase II is required to restart a replication fork when its progress is blocked by damage in DNA. DNA polymerases IV and V are involved in allowing replication to bypass certain types of damage.


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Figure 14.4 Only one DNA polymerase is the replicase. The others participate in repair of damaged DNA, restarting stalled replication forks, or bypassing damage in DNA.

When extracts of E. coli are assayed for their ability to synthesize DNA, the predominant enzyme activity is DNA polymerase I. Its activity is so great that it makes it impossible to detect the activities of the enzymes actually responsible for DNA replication! To develop in vitro systems in which replication can be followed, extracts are therefore prepared from polA mutant cells. Some phages code for DNA polymerases. They include T4, T5, T7, and SPO1. The enzymes all possess 5 ′ –3 ′ synthetic activities and 3 ′ –5 ′ exonuclease proofreading activities (see Molecular Biology 4.14.3 DNA polymerases have various nuclease activities). In each case, a mutation in the gene that codes for a single phage polypeptide prevents phage development. Each phage polymerase polypeptide associates with other proteins, of either phage or host origin, to make the intact enzyme. Several classes of eukaryotic DNA polymerases have been identified. DNA polymerases δ and ε are required for nuclear replication; DNA polymerase α is concerned with "priming" (initiating) replication. Other DNA polymerases are involved in repairing damaged nuclear DNA ( β and also ε) or with mitochondrial DNA replication ( γ). Last updated on January 8, 2004 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.14.2


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4.14.3 DNA polymerases have various nuclease activities Key Terms Nick translation describes the ability of E. coli DNA polymerase I to use a nick as a starting point from which one strand of a duplex DNA can be degraded and replaced by resynthesis of new material; is used to introduce radioactively labeled nucleotides into DNA in vitro. Key Concepts

• DNA polymerase I has a unique 5 ′ –3 ′ exonuclease activity that can be combined with DNA synthesis to perform nick translation.

Replicases often have nuclease activities as well as the ability to synthesize DNA. A 3 ′ –5 ′ exonuclease activity is typically used to excise bases that have been added to DNA incorrectly. This provides a "proofreading" error-control system (see Molecular Biology 4.14.4 DNA polymerases control the fidelity of replication). The first DNA-synthesizing enzyme to be characterized was DNA polymerase I, which is a single polypeptide of 103 kD. The chain can be cleaved into two parts by proteolytic treatment. The C-terminal two-thirds of the protein contains the polymerase active site, while the N-terminal third contains the proofreading exonuclease. The larger cleavage product (68 kD) is called the Klenow fragment. It is used in synthetic reactions in vitro. It contains the polymerase and the 3 ′ –5 ′ exonuclease activities. The active sites are ~30 Å apart in the protein, indicating that there is spatial separation between adding a base and removing one. The small fragment (35 kD) possesses a 5 ′ –3 ′ exonucleolytic activity, which excises small groups of nucleotides, up to ~10 bases at a time. This activity is coordinated with the synthetic/proofreading activity. It provides DNA polymerase I with a unique ability to start replication in vitro at a nick in DNA. (No other DNA polymerase has this ability.) At a point where a phosphodiester bond has been broken in a double-stranded DNA, the enzyme extends the 3 ′ –OH end. As the new segment of DNA is synthesized, it displaces the existing homologous strand in the duplex. This process of nick translation is illustrated in Figure 14.5. The displaced strand is degraded by the 5 ′ –3 ′ exonucleolytic activity of the enzyme. The properties of the DNA are unaltered, except that a segment of one strand has been replaced with newly synthesized material, and the position of the nick has been moved along the duplex. This is of great practical use; nick translation has been a major technique for introducing radioactively labeled nucleotides into DNA in vitro.

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Figure 14.5 Nick translation replaces part of a pre-existing strand of duplex DNA with newly synthesized material.

The 5 ′ –3 ′ synthetic/3 ′ –5 ′ exonucleolytic action is probably used in vivo mostly for filling in short single-stranded regions in double-stranded DNA. These regions arise during replication, and also when bases that have been damaged are removed from DNA (see Molecular Biology 4.15 Recombination and repair). . This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.14.3

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4.14.4 DNA polymerases control the fidelity of replication Key Terms Processivity describes the ability of an enzyme to perform multiple catalytic cycles with a single template instead of dissociating after each cycle. Proofreading refers to any mechanism for correcting errors in protein or nucleic acid synthesis that involves scrutiny of individual units after they have been added to the chain. Key Concepts

• DNA polymerases often have a 3 ′ –5 ′ exonuclease activity that is used to excise incorrectly paired bases.

• The fidelity of replication is improved by proofreading by a factor of ~100.

The fidelity of replication poses the same sort of problem we have encountered already in considering (for example) the accuracy of translation. It relies on the specificity of base pairing. Yet when we consider the interactions involved in base pairing, we would expect errors to occur with a frequency of ~10-3 per base pair replicated. The actual rate in bacteria seems to be ~10-8-10-10. This corresponds to ~1 error per genome per 1000 bacterial replication cycles, or ~10-6 per gene per generation. We can divide the errors that DNA polymerase makes during replication into two classes. • Frameshifts occur when an extra nucleotide is inserted or omitted. Fidelity with regard to frameshifts is affected by the processivity of the enzyme: the tendency to remain on a single template rather than to dissociate and reassociate. This is particularly important for the replication of a homopolymeric stretch, for example, a long sequence of dTn:dAn, in which "replication slippage" can change the length of the homopolymeric run. As a general rule, increased processivity reduces the likelihood of such events. In multimeric DNA polymerases, processivity is usually increased by a particular subunit that is not needed for catalytic activity per se. • Substitutions occur when the wrong (improperly paired) nucleotide is incorporated. The error level is determined by the efficiency of proofreading, in which the enzyme scrutinizes the newly formed base pair and removes the nucleotide if it is mispaired. All of the bacterial enzymes possess a 3 ′ –5 ′ exonucleolytic activity that proceeds in the reverse direction from DNA synthesis. This provides the proofreading function DNA polymerases control the fidelity of replication | SECTION 4.14.4 © 2004. Virtual Text / www.ergito.com

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illustrated diagrammatically in Figure 14.6. In the chain elongation step, a precursor nucleotide enters the position at the end of the growing chain. A bond is formed. The enzyme moves one base pair farther, ready for the next precursor nucleotide to enter. If a mistake has been made, however, the enzyme uses the exonucleolytic activity to excise the last base that was added.

Figure 14.6 Bacterial DNA polymerases scrutinize the base pair at the end of the growing chain and excise the nucleotide added in the case of a misfit.

Different DNA polymerases handle the relationship between the polymerizing and proofreading activities in different ways. In some cases, the activities are part of the same protein subunit, but in others they are contained in different subunits. Each DNA polymerase has a characteristic error rate that is reduced by its proofreading activity. Proofreading typically decreases the error rate in replication from ~10-5 to ~10-7 per base pair replicated. Systems that recognize errors and correct them following replication then eliminate some of the errors, bringing the overall rate to 103-fold. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.14.4

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4.14.5 DNA polymerases have a common structure Key Concepts

• Many DNA polymerases have a large cleft composed of three domains that resemble a hand.

• DNA lies across the "palm" in a groove created by the "fingers" and "thumb".

Figure 14.7 shows that all DNA polymerases share some common structural features (3091; 3092). The enzyme structure can be divided into several independent domains, which are described by analogy with a human right hand. DNA binds in a large cleft composed of three domains. The "palm" domain has important conserved sequence motifs that provide the catalytic active site. The "fingers" are involved in positioning the template correctly at the active site. The "thumb" binds the DNA as it exits the enzyme, and is important in processivity. The most important conserved regions of each of these three domains converge to form a continuous surface at the catalytic site. The exonuclease activity resides in an independent domain with its own catalytic site. The N-terminal domain extends into the nuclease domain. DNA polymerases fall into five families based on sequence homologies; the palm is well conserved among them, but the thumb and fingers provide analogous secondary structure elements from different sequences.

Figure 14.7 The common organization of DNA polymerases has a palm that contains the catalytic site, fingers that position the template, a thumb that binds DNA and is important in processivity, an exonuclease domain with its own active site, and an N-terminal domain.

The catalytic reaction in a DNA polymerase occurs at an active site in which a nucleotide triphosphate pairs with an (unpaired) single strand of DNA. The DNA lies DNA polymerases have a common structure | SECTION 4.14.5 © 2004. Virtual Text / www.ergito.com

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across the palm in a groove that is created by the thumb and fingers. Figure 14.8 shows the crystal structure of the T7 enzyme complexed with DNA (in the form of a primer annealed to a template strand) and an incoming nucleotide that is about to be added to the primer. The DNA is in the classic B-form duplex up to the last 2 base pairs at the 3 ′ end of the primer, which are in the more open A-form. A sharp turn in the DNA exposes the template base to the incoming nucleotide. The 3 ′ end of the primer (to which bases are added) is anchored by the fingers and palm. The DNA is held in position by contacts that are made principally with the phosphodiester backbone (thus enabling the polymerase to function with DNA of any sequence).

Figure 14.8 The crystal structure of phage T7 DNA polymerase shows that the template strand takes a sharp turn in order to be exposed to the incoming nucleotide. Photograph kindly provided by Charles Richardson and Tom Ellenberger (see 3748).

In structures of DNA polymerases of this family complexed only with DNA (that is, lacking the incoming nucleotide), the orientation of the fingers and thumb relative to the palm is more open, with the O helix (O, O1, O2; see Figure 14.8) rotated away from the palm. This suggests that an inward rotation of the O helix occurs to grasp the incoming nucleotide and create the active catalytic site. When a nucleotide binds, the fingers domain rotates 60° toward the palm, with the tops of the fingers moving by 30 Å. The thumb domain also rotates toward the palm by 8°. These changes are cyclical: they are reversed when the nucleotide is incorporated into the DNA chain, which then translocates through the enzyme to recreate an empty site. The exonuclease activity is responsible for removing mispaired bases. But the catalytic site of the exonuclease domain is distant from the active site of the catalytic domain. The enzyme alternates between polymerizing and editing modes, as determined by a competition between the two active sites for the 3 ′ primer end of the DNA (3093). Amino acids in the active site contact the incoming base in such a way that the enzyme structure is affected by a mismatched base. When a mismatched base pair occupies the catalytic site, the fingers cannot rotate toward the palm to bind the incoming nucleotide. This leaves the 3 ′ end free to bind to the active site in the exonuclease domain, which is accomplished by a rotation of the DNA in the enzyme structure (3094). DNA polymerases have a common structure | SECTION 4.14.5 © 2004. Virtual Text / www.ergito.com

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Reviews 3091. Joyce, C. M. and Steitz, T. A. (1994). Function and structure relationships in DNA polymerases. Annu. Rev. Biochem. 63, 777-822. 3092. Hubscher, U., Maga, G., and Spadari, S. (2002). Eukaryotic DNA polymerases. Annu. Rev. Biochem. 71, 133-163. 3093. Johnson, K. A. (1993). Conformational coupling in DNA polymerase fidelity. Annu. Rev. Biochem. 62, 685-713.

References 3094. Shamoo, Y. and Steitz, T. A. (1999). Building a replisome from interacting pieces: sliding clamp complexed to a peptide from DNA polymerase and a polymerase editing complex. Cell 99, 155-166.


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4.14.6 DNA synthesis is semidiscontinuous Key Terms The leading strand of DNA is synthesized continuously in the 5 ′ -3 ′ direction. The lagging strand of DNA must grow overall in the 3 ′ -5 ′ direction and is synthesized discontinuously in the form of short fragments (5 ′ -3 ′ ) that are later connected covalently. Okazaki fragments are the short stretches of 1000-2000 bases produced during discontinuous replication; they are later joined into a covalently intact strand. Semidiscontinuous replication is mode in which one new strand is synthesized continuously while the other is synthesized discontinuously. Key Concepts

• The DNA replicase advances continuously when it synthesizes the leading strand (5 ′ –3 ′ ), but synthesizes the lagging strand by making short fragments that are subsequently joined together.

The antiparallel structure of the two strands of duplex DNA poses a problem for replication. As the replication fork advances, daughter strands must be synthesized on both of the exposed parental single strands. The fork moves in the direction from 5 ′ –3 ′ on one strand, and in the direction from 3 ′ –5 ′ on the other strand. Yet nucleic acids are synthesized only from a 5 ′ end toward a 3 ′ end. The problem is solved by synthesizing the strand that grows overall from 3 ′ –5 ′ in a series of short fragments, each actually synthesized in the "backwards" direction, that is, with the customary 5 ′ –3 ′ polarity. Consider the region immediately behind the replication fork, as illustrated in Figure 14.9. We describe events in terms of the different properties of each of the newly synthesized strands:

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Figure 14.9 The leading strand is synthesized continuously while the lagging strand is synthesized discontinuously.

• On the leading strand DNA synthesis can proceed continuously in the 5 ′ to 3 ′ direction as the parental duplex is unwound. • On the lagging strand a stretch of single-stranded parental DNA must be exposed, and then a segment is synthesized in the reverse direction (relative to fork movement). A series of these fragments are synthesized, each 5 ′ –3 ′ ; then they are joined together to create an intact lagging strand. Discontinuous replication can be followed by the fate of a very brief label of radioactivity. The label enters newly synthesized DNA in the form of short fragments, sedimenting in the range of 7-11S, corresponding to ~1000-2000 bases in length. These Okazaki fragments are found in replicating DNA in both prokaryotes and eukaryotes. After longer periods of incubation, the label enters larger segments of DNA. The transition results from covalent linkages between Okazaki fragments. (The lagging strand must be synthesized in the form of Okazaki fragments. For a long time it was unclear whether the leading strand is synthesized in the same way or is synthesized continuously. All newly synthesized DNA is found as short fragments in E. coli. Superficially, this suggests that both strands are synthesized discontinuously. However, it turns out that not all of the fragment population represents bona fide Okazaki fragments; some are pseudofragments, generated by breakage in a DNA strand that actually was synthesized as a continuous chain. The source of this breakage is the incorporation of some uracil into DNA in place of thymine. When the uracil is removed by a repair system, the leading strand has breaks until a thymine is inserted.) So the lagging strand is synthesized discontinuously and the leading strand is synthesized continuously. This is called semidiscontinuous replication. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.14.6

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4.14.7 The φX model system shows how single-stranded DNA is generated for replication Key Terms A helicase is an enzyme that uses energy provided by ATP hydrolysis to separate the strands of a nucleic acid duplex. The single-strand binding protein (SSB) attaches to single-stranded DNA, thereby preventing the DNA from forming a duplex. Key Concepts

• Replication requires a helicase to separate the strands of DNA using energy provided by hydrolysis of ATP.

• A single-strand binding protein is required to maintain the separated strands. • The combination of helicase, SSB, and A protein separates a φX174 duplex into a single-stranded circle and a single-stranded linear strand.

As the replication fork advances, it unwinds the duplex DNA. One of the template strands is rapidly converted to duplex DNA as the leading daughter strand is synthesized. The other remains single-stranded until a sufficient length has been exposed to initiate synthesis of an Okazaki fragment of the lagging strand in the backward direction. The generation and maintenance of single-stranded DNA is therefore a crucial aspect of replication. Two types of function are needed to convert double-stranded DNA to the single-stranded state: • A helicase is an enzyme that separates the strands of DNA, usually using the hydrolysis of ATP to provide the necessary energy. • A single-strand binding protein (SSB) binds to the single-stranded DNA, preventing it from reforming the duplex state. The SSB binds as a monomer, but typically in a cooperative manner in which the binding of additional monomers to the existing complex is enhanced. Helicases separate the strands of a duplex nucleic acid in a variety of situations, ranging from strand separation at the growing point of a replication fork to catalyzing migration of Holliday (recombination) junctions along DNA. There are 12 different helicases in E. coli. A helicase is generally multimeric. A common form of helicase is a hexamer (1195). This typically translocates along DNA by using its multimeric structure to provide multiple DNA-binding sites. Figure 14.10 shows a generalized schematic model for the action of a hexameric helicase. It is likely to have one conformation that binds to duplex DNA and another that binds to single-stranded DNA. Alternation between them drives the motor that The φX model system shows how single-stranded DNA is generated for replication | SECTION 4.14.7 © 2004. Virtual Text / www.ergito.com

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melts the duplex, and requires ATP hydrolysis – typically 1 ATP is hydrolyzed for each base pair that is unwound (1194). A helicase usually initiates unwinding at a single-stranded region adjacent to a duplex, and may function with a particular polarity, preferring single-stranded DNA with a 3 ′ end (3 ′ –5 ′ helicase) or with a 5 ′ end (5 ′ –3 ′ helicase).

Figure 14.10 A hexameric helicase moves along one strand of DNA. It probably changes conformation when it binds to the duplex, uses ATP hydrolysis to separate the strands, and then returns to the conformation it has when bound only to a single strand.

The conversion of φX174 double-stranded DNA into individual single strands illustrates the features of the strand separation process. Figure 14.11 shows that a single strand is peeled off the circular strand, resembling the rolling circle described previously in Figure 13.16. The reaction can occur in the absence of DNA synthesis when the appropriate 3 proteins are provided in vitro.

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Figure 14.11 φ X174 DNA can be separated into single strands by the combined effects of 3 functions: nicking with A protein, unwinding by Rep, and single-strand stabilization by SSB.

The phage A protein nicks the viral (+) strand at the origin of replication. In the presence of 2 host proteins, Rep and SSB, and ATP, the nicked DNA unwinds. The Rep protein provides a helicase that separates the strands; the SSB traps them in single-stranded form. The E. coli SSB is a tetramer of 74 kD that binds cooperatively to single-stranded DNA. The significance of the cooperative mode of binding is that the binding of one protein molecule makes it much easier for another to bind. So once the binding reaction has started on a particular DNA molecule, it is rapidly extended until all of the single-stranded DNA is covered with the SSB protein. Note that this protein is not a DNA-unwinding protein; its function is to stabilize DNA that is already in the single-stranded condition. Under normal circumstances in vivo, the unwinding, coating, and replication The φX model system shows how single-stranded DNA is generated for replication | SECTION 4.14.7 © 2004. Virtual Text / www.ergito.com

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reactions proceed in tandem. The SSB binds to DNA as the replication fork advances, keeping the two parental strands separate so that they are in the appropriate condition to act as templates. SSB is needed in stoichiometric amounts at the replication fork. It is required for more than one stage of replication; ssb mutants have a quick-stop phenotype, and are defective in repair and recombination as well as in replication. (Some phages use different SSB proteins, notably T4; this shows that there may be specific interactions between components of the replication apparatus and the SSB; see Molecular Biology 4.14.14 Phage T4 provides its own replication apparatus).


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References 1194. Dillingham, M. S., Wigley, D. B., and Webb, M. R. (2000). Demonstration of unidirectional single-stranded DNA translocation by PcrA helicase: measurement of step size and translocation speed. Biochemistry 39, 205-212. 1195. Singleton, M. R., Sawaya, M. R., Ellenberger, T., and Wigley, D. B. (2000). Crystal structure of T7 gene 4 ring helicase indicates a mechanism for sequential hydrolysis of nucleotides. Cell 101, 589-600.



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4.14.8 Priming is required to start DNA synthesis Key Terms A primer is a short sequence (often of RNA) that is paired with one strand of DNA and provides a free 3 ′ -OH end at which a DNA polymerase starts synthesis of a deoxyribonucleotide chain. The primase is a type of RNA polymerase that synthesizes short segments of RNA that will be used as primers for DNA replication. Key Concepts

• All DNA polymerases require a 3 ′ –OH priming end to initiate DNA synthesis. • The priming end can be provided by an RNA primer, a nick in DNA, or a priming protein.

• For DNA replication, a special RNA polymerase called a primase synthesizes an RNA chain that provides the priming end.

• E. coli has two types of priming reaction, which occur at the bacterial origin (oriC) and the φX174 origin.

• Priming of replication on double-stranded DNA always requires a replicase, SSB, and primase.

• DnaB is the helicase that unwinds DNA for replication in E. coli.

A common feature of all DNA polymerases is that they cannot initiate synthesis of a chain of DNA de novo. Figure 14.12 shows the features required for initiation. Synthesis of the new strand can only start from a pre-existing 3 ′ –OH end; and the template strand must be converted to a single-stranded condition.

Figure 14.12 A DNA polymerase requires a 3 ′ -OH end to initiate replication.

The 3 ′ –OH end is called a primer. The primer can take various forms. Types of priming reaction are summarized in Figure 14.13: Priming is required to start DNA synthesis | SECTION 4.14.8 © 2004. Virtual Text / www.ergito.com

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Figure 14.13 There are several methods for providing the free 3 ′ -OH end that DNA polymerases require to initiate DNA synthesis.

• A sequence of RNA is synthesized on the template, so that the free 3 ′ –OH end of the RNA chain is extended by the DNA polymerase. This is commonly used in replication of cellular DNA, and by some viruses (see Figure 13.40 in Molecular Biology 4.13.23 The ColE1 compatibility system is controlled by an RNA regulator).

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• A preformed RNA pairs with the template, allowing its 3 ′ –OH end to be used to prime DNA synthesis. This mechanism is used by retroviruses to prime reverse transcription of RNA (see Figure 17.6 in Molecular Biology 4.17.4 Viral DNA is generated by reverse transcription). • A primer terminus is generated within duplex DNA. The most common mechanism is the introduction of a nick, as used to initiate rolling circle replication (see Figure 13.16). In this case, the pre-existing strand is displaced by new synthesis. (Note the difference from nick translation shown in Figure 14.5, in which DNA polymerase I simultaneously synthesizes and degrades DNA from a nick.) • A protein primes the reaction directly by presenting a nucleotide to the DNA polymerase. This reaction is used by certain viruses (see Figure 13.15 in Molecular Biology 4.13.8 The ends of linear DNA are a problem for replication ). Priming activity is required to provide 3 ′ –OH ends to start off the DNA chains on both the leading and lagging strands. The leading strand requires only one such initiation event, which occurs at the origin. But there must be a series of initiation events on the lagging strand, since each Okazaki fragment requires its own start de novo. Each Okazaki fragment starts with a primer sequence of RNA, ~10 bases long, that provides the 3 ′ –OH end for extension by DNA polymerase. A primase is required to catalyze the actual priming reaction. This is provided by a special RNA polymerase activity, the product of the dnaG gene. The enzyme is a single polypeptide of 60 kD (much smaller than RNA polymerase). The primase is an RNA polymerase that is used only under specific circumstances, that is, to synthesize short stretches of RNA that are used as primers for DNA synthesis. DnaG primase associates transiently with the replication complex, and typically synthesizes an 11-12 base primer. Primers start with the sequence pppAG, opposite the sequence 3 ′ –GTC-5 ′ in the template. (Some systems use alternatives to the DnaG primase. In the examples of the two phages M13 and G4, which were used for early work on replication, an interesting difference emerged. G4 priming uses DnaG, but M13 priming uses bacterial RNA polymerase. These phages have another unusual feature, which is that the site of priming is indicated by a region of secondary structure.) There are two types of priming reaction in E. coli. • The oriC system, named for the bacterial origin, basically involves the association of the DnaG primase with the protein complex at the replication fork. • The φX system, named for phage φX174, requires an initiation complex consisting of additional components, called the primosome (see Molecular Biology 4.14.17 The primosome is needed to restart replication). Sometimes replicons are referred to as being of the φX or oriC type. Priming is required to start DNA synthesis | SECTION 4.14.8 © 2004. Virtual Text / www.ergito.com

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The types of activities involved in the initiation reaction are summarized in Figure 14.14. Although other replicons in E. coli may have alternatives for some of these particular proteins, the same general types of activity are required in every case. A helicase is required to generate single strands, a single-strand binding protein is required to maintain the single-stranded state, and the primase synthesizes the RNA primer.

Figure 14.14 Initiation requires several enzymatic activities, including helicases, single-strand binding proteins, and synthesis of the primer.

DnaB is the central component in both φX and oriC replicons. It provides the 5 ′ –3 ′ helicase activity that unwinds DNA. Energy for the reaction is provided by cleavage of ATP. Basically DnaB is the active component of the growing point. In oriC replicons, DnaB is initially loaded at the origin as part of a large complex (see Molecular Biology 4.14.15 Creating the replication forks at an origin ). It forms the growing point at which the DNA strands are separated as the replication fork advances. It is part of the DNA polymerase complex and interacts with the DnaG primase to initiate synthesis of each Okazaki fragment on the lagging strand. Priming is required to start DNA synthesis | SECTION 4.14.8 © 2004. Virtual Text / www.ergito.com

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4.14.9 Coordinating synthesis of the lagging and leading strands Key Concepts

• Different enzyme units are required to synthesize the leading and lagging strands. • In E. coli both these units contain the same catalytic subunit (DnaE). • In other organisms, different catalytic subunits may be required for each strand.

Each new DNA strand is synthesized by an individual catalytic unit. Figure 14.15 shows that the behavior of these two units is different because the new DNA strands are growing in opposite directions. One enzyme unit is moving with the unwinding point and synthesizing the leading strand continuously. The other unit is moving "backwards," relative to the DNA, along the exposed single strand. Only short segments of template are exposed at any one time. When synthesis of one Okazaki fragment is completed, synthesis of the next Okazaki fragment is required to start at a new location approximately in the vicinity of the growing point for the leading strand. This requires a translocation relative to the DNA of the enzyme unit that is synthesizing the lagging strand.

Figure 14.15 Leading and lagging strand polymerases move apart. This is a static version of an interactive figure; see http://www.ergito.com/main.jsp?bcs=MBIO.4.14.9 to view properly.

The term "enzyme unit" avoids the issue of whether the DNA polymerase that synthesizes the leading strand is the same type of enzyme as the DNA polymerase that synthesizes the lagging strand. In the case that we know best, E. coli, there is only a single type of DNA polymerase catalytic subunit used in replication, the DnaE protein. The active replicase is a dimer, and each half of the dimer contains DnaE as the catalytic subunit, supported by other proteins (which differ between the leading and lagging strands). The use of a single type of catalytic subunit, however, may be atypical. In the bacterium B. subtilis, there are two different catalytic subunits (2184). PolC is the homologue to E. coli's DnaE, and is responsible for synthesizing the leading strand. A related protein, DnaEBS, is the catalytic subunit that synthesizes the lagging strand. Eukaryotic DNA polymerases have the same general structure, with different enzyme Coordinating synthesis of the lagging and leading strands | SECTION 4.14.9 © 2004. Virtual Text / www.ergito.com

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units synthesizing the leading and lagging strands, but it is not clear whether the same or different types of catalytic subunits are used (see Molecular Biology 4.14.13 Separate eukaryotic DNA polymerases undertake initiation and elongation). A major problem of the semidiscontinuous mode of replication follows from the use of different enzyme units to synthesize each new DNA strand: how is synthesis of the lagging strand coordinated with synthesis of the leading strand? As the replisome moves along DNA, unwinding the parental strands, one enzyme unit elongates the leading strand. Periodically the primosome activity initiates an Okazaki fragment on the lagging strand, and the other enzyme unit must then move in the reverse direction to synthesize DNA. Figure 14.16 proposes two types of model for what happens to this enzyme unit when it completes synthesis of an Okazaki fragment. The same complex may be reutilized for synthesis of successive Okazaki fragments. Or the complex might dissociate from the template, so that a new complex must be assembled to elongate the next Okazaki fragment. We see in the Molecular Biology 4.14.11 The clamp controls association of core enzyme with DNA that the first model applies.

Figure 14.16 The upper model for the action of lagging strand polymerase is that when an enzyme unit completes one Okazaki fragment, it moves to a new position to synthesize the next fragment. The lower model is that the lagging strand polymerase dissociates when it completes an Okazaki fragment, and a new enzyme unit associates with DNA to synthesize the next Okazaki fragment.


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References 2184. Dervyn, E., Suski, C., Daniel, R., Bruand, C., Chapuis, J., Errington, J., Janniere, L., and Ehrlich, S. D. (2001). Two essential DNA polymerases at the bacterial replication fork. Science 294, 1716-1719.


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4.14.10 DNA polymerase holoenzyme has 3 subcomplexes Key Terms The clamp loader is a 5 subunit protein complex which is responsible for loading the β clamp on to DNA at the replication fork. Key Concepts

• The E. coli replicase DNA polymerase III is a 900 kD complex with a dimeric structure.

• Each monomeric unit has a catalytic core, a dimerization subunit, and a processivity component.

• A clamp loader places the processivity subunits on DNA, and they form a circular clamp around the nucleic acid.

• One catalytic core is associated with each template strand.

We can now relate the subunit structure of E. coli DNA polymerase III to the activities required for DNA synthesis and propose a model for its action. The holoenzyme is a complex of 900 kD that contains 10 proteins organized into four types of subcomplex: • There are two copies of the catalytic core. Each catalytic core contains the α subunit (the DNA polymerase activity), ε subunit (3 ′ –5 ′ proofreading exonuclease), and θ subunit (stimulates exonuclease). • There are two copies of the dimerizing subunit, τ, which link the two catalytic cores together (2185). • There are two copies of the clamp, which is responsible for holding catalytic cores on to their template strands. Each clamp consists of a homodimer of β subunits that binds around the DNA and ensures processivity. • The γ complex is a group of 5 proteins, the Clamp loader, that places the clamp on DNA (2185). A model for the assembly of DNA polymerase III is shown in Figure 14.17. The holoenzyme assembles on DNA in three stages:

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Figure 14.17 DNA polymerase III holoenzyme assembles in stages, generating an enzyme complex that synthesizes the DNA of both new strands.

• First the clamp loader uses hydrolysis of ATP to bind β subunits to a template-primer complex. • Binding to DNA changes the conformation of the site on β that binds to the clamp loader, and as a result it now has a high affinity for the core polymerase. This enables core polymerase to bind, and this is the means by which the core polymerase is brought to DNA. • A τ dimer binds to the core polymerase, and provides a dimerization function that binds a second core polymerase (associated with another β clamp). The holoenzyme is asymmetric, because it has only 1 clamp loader. The clamp loader is responsible for adding a pair of β dimers to each parental strand of DNA. Each of the core complexes of the holoenzyme synthesizes one of the new strands of DNA. Because the clamp loader is also needed for unloading the β complex from DNA, the two cores have different abilities to dissociate from DNA. This corresponds to the need to synthesize a continuous leading strand (where polymerase DNA polymerase holoenzyme has 3 subcomplexes | SECTION 4.14.10 © 2004. Virtual Text / www.ergito.com

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remains associated with the template) and a discontinuous lagging strand (where polymerase repetitively dissociates and reassociates). The clamp loader is associated with the core polymerase that synthesizes the lagging strand, and plays a key role in the ability to synthesize individual Okazaki fragments (2186). Last updated on 5-14-2002


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References 2185. Studwell-Vaughan, P. S. and O'Donnell, M. (1991). Constitution of the twin polymerase of DNA polymerase III holoenzyme. J. Biol. Chem. 266, 19833-19841. 2186. Stukenberg, P. T., Studwell-Vaughan, P. S., and O'Donnell, M. (1991). Mechanism of the sliding beta-clamp of DNA polymerase III holoenzyme. J. Biol. Chem. 266, 11328-11334.



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4.14.11 The clamp controls association of core enzyme with DNA Key Concepts

• The core on the leading strand is processive because its clamp keeps it on the DNA.

• The clamp associated with the core on the lagging strand dissociates at the end of each Okazaki fragment and reassembles for the next fragment.

• The helicase DnaB is responsible for interacting with the primase DnaG to initiate each Okazaki fragment.

The β dimer makes the holoenzyme highly processive. β is strongly bound to DNA, but can slide along a duplex molecule. The crystal structure of β shows that it forms a ring-shaped dimer (2187). The model in Figure 14.18 shows the β-ring in relationship to a DNA double helix. The ring has an external diameter of 80 Å and an internal cavity of 35 Å, almost twice the diameter of the DNA double helix (20 Å). The space between the protein ring and the DNA is filled by water. Each of the β subunits has three globular domains with similar organization (although their sequences are different). As a result, the dimer has 6-fold symmetry, reflected in 12 α-helices that line the inside of the ring.

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Figure 14.18 The β subunit of DNA polymerase III holoenzyme consists of a head to tail dimer (the two subunits are shown in red and orange) that forms a ring completely surrounding a DNA duplex (shown in the center). Photograph kindly provided by John Kuriyan.

The dimer surrounds the duplex, providing the "sliding clamp" that allows the holoenzyme to slide along DNA. The structure explains the high processivity – there is no way for the enzyme to fall off! The α-helices on the inside have some positive charges that may interact with the DNA via the intermediate water molecules. Because the protein clamp does not directly contact the DNA, it may be able to "ice-skate" along the DNA, making and breaking contacts via the water molecules. How does the clamp get on to the DNA? Because the clamp is a circle of subunits surrounding DNA, its assembly or removal requires the use of an energy-dependent process by the clamp loader. The γ clamp loader is a pentameric circular structure that binds an open form of the β ring preparatory to loading it on to DNA (2188). In effect, the ring is opened at one of the interfaces between the two β subunits by the δ subunit of the clamp loader. The clamp loader uses hydrolysis of ATP to provide the energy to open the ring of the clamp and insert DNA into its central cavity. The relationship between the β clamp and the γ clamp loader is a paradigm for similar systems used by DNA replicases ranging from bacteriophages to animal cells. The clamp is a heteromer (sometimes a dimer, sometimes a trimer) that forms a ring around DNA with a set of 12 α-helices forming 6-fold symmetry for the structure as The clamp controls association of core enzyme with DNA | SECTION 4.14.11 © 2004. Virtual Text / www.ergito.com

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a whole. The clamp loader has some subunits that hydrolyze ATP to provide energy for the reaction (for review see 3441). The basic principle that is established by the dimeric polymerase model is that, while one polymerase subunit synthesizes the leading strand continuously, the other cyclically initiates and terminates the Okazaki fragments of the lagging strand within a large single-stranded loop formed by its template strand. Figure 14.19 draws a generic model for the operation of such a replicase. The replication fork is created by a helicase, typically forming a hexameric ring, that translocates in the 5 ′ –3 ′ direction on the template for the lagging strand. The helicase is connected to two DNA polymerase catalytic subunits, each of which is associated with a sliding clamp.

Figure 14.19 The helicase creating the replication fork is connected to two DNA polymerase catalytic subunits, each of which is held on to DNA by a sliding clamp. The polymerase that synthesizes the leading strand moves continuously. The polymerase that synthesizes the lagging strand dissociates at the end of an Okazaki fragment and then reassociates with a primer in the single-stranded template loop to synthesize the next fragment.

We can describe this model for DNA polymerase III in terms of the individual components of the enzyme complex, as illustrated in Figure 14.20. A catalytic core is associated with each template strand of DNA. The holoenzyme moves continuously along the template for the leading strand; the template for the lagging strand is "pulled through," creating a loop in the DNA. DnaB creates the unwinding point, and translocates along the DNA in the "forward" direction (for review see 2279)


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Figure 14.20 Each catalytic core of Pol III synthesizes a daughter strand. DnaB is responsible for forward movement at the replication fork.

DnaB contacts the τ subunit(s) of the clamp loader. This establishes a direct connection between the helicase-primase complex and the catalytic cores. This link has two effects. One is to increase the speed of DNA synthesis by increasing the rate of movement by DNA polymerase core by 10×. The second is to prevent the leading strand polymerase from falling off, that is, to increase its processivity. Synthesis of the leading strand creates a loop of single-stranded DNA that provides the template for lagging strand synthesis, and this loop becomes larger as the unwinding point advances. After initiation of an Okazaki fragment, the lagging strand core complex pulls the single-stranded template through the β clamp while synthesizing the new strand. The single-stranded template must extend for the length of at least one Okazaki fragment before the lagging polymerase completes one fragment and is ready to begin the next. What happens to the loop when the Okazaki fragment is completed? Figure 14.21 suggests that the core complex dissociates when it completes synthesis of each fragment, releasing the loop. The core complex then associates with a β clamp to The clamp controls association of core enzyme with DNA | SECTION 4.14.11 © 2004. Virtual Text / www.ergito.com

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Figure 14.21 Core polymerase and the β clamp dissociate at completion of Okazaki fragment synthesis and reassociate at the beginning.

What is responsible for recognizing the sites for initiating synthesis of Okazaki fragments? In oriC replicons, the connection between priming and the replication fork is provided by the dual properties of DnaB: it is the helicase that propels the replication fork; and it interacts with the DnaG primase at an appropriate site. Following primer synthesis, the primase is released. The length of the priming RNA is limited to 8-14 bases. Apparently DNA polymerase III is responsible for displacing the primase. Last updated on 11-20-2002


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Reviews 2279. Benkovic, S. J., Valentine, A. M., and Salinas, F. (2001). Replisome-mediated DNA replication. Annu. Rev. Biochem. 70, 181-208. 3441. Davey, M. J., Jeruzalmi, D., Kuriyan, J., and O'Donnell, M. (2002). Motors and switches: AAA+ machines within the replisome. Nat. Rev. Mol. Cell Biol. 3, 826-835.

References 2187. Kong, X. P., Onrust, R., O'Donnell, M., and Kuriyan, J. (1992). Three-dimensional structure of the beta subunit ofE. coli DNA polymerase III holoenzyme: a sliding DNA clamp. Cell 69, 425-437. 2188. Jeruzalmi, D., O'Donnell, M., and Kuriyan, J. (2001). Crystal structure of the processivity clamp loader gamma (gamma) complex of E. coli DNA polymerase III. Cell 106, 429-441.



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4.14.12 Okazaki fragments are linked by ligase Key Terms DNA ligase makes a bond between an adjacent 3 ′ -OH and 5 ′ -phosphate end where there is a nick in one strand of duplex DNA. Key Concepts

• Each Okazaki fragment starts with a primer and stops before the next fragment. • DNA polymerase I removes the primer and replaces it with DNA in an action that resembles nick translation.

• DNA ligase makes the bond that connects the 3 ′ end of one Okazaki fragment to the 5 ′ beginning of the next fragment.

We can now expand our view of the actions involved in joining Okazaki fragments, as illustrated in Figure 14.22. The complete order of events is uncertain, but must involve synthesis of RNA primer, its extension with DNA, removal of the RNA primer, its replacement by a stretch of DNA, and the covalent linking of adjacent Okazaki fragments.

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Figure 14.22 Synthesis of Okazaki fragments requires priming, extension, removal of RNA, gap filling, and nick ligation.

The figure suggests that synthesis of an Okazaki fragment terminates just before the start of the RNA primer of the preceding fragment. When the primer is removed, there will be a gap. The gap is filled by DNA polymerase I; polA mutants fail to join their Okazaki fragments properly. The 5 ′ –3 ′ exonuclease activity removes the RNA primer while simultaneously replacing it with a DNA sequence extended from the 3 ′ –OH end of the next Okazaki fragment. This is equivalent to nick translation, except that the new DNA replaces a stretch of RNA rather than a segment of DNA. In mammalian systems (where the DNA polymerase does not have a 5 ′ –3 ′ exonuclease activity), Okazaki fragments are removed by a two-step process. First RNAase HI (an enzyme that is specific for a DNA-RNA hybrid substrate) makes an endonucleolytic cleavage; then a 5 ′ –3 ′ exonuclease called FEN1 removes the RNA. Once the RNA has been removed and replaced, the adjacent Okazaki fragments must be linked together. The 3 ′ –OH end of one fragment is adjacent to the 5 ′ –phosphate end of the previous fragment. The responsibility for sealing this nick lies with the enzyme DNA ligase. Ligases are present in both prokaryotes and eukaryotes. Unconnected fragments persist in lig– mutants, because they fail to join Okazaki fragments together. The E. coli and T4 ligases share the property of sealing nicks that have 3 ′ –OH and 5 Okazaki fragments are linked by ligase | SECTION 4.14.12 © 2004. Virtual Text / www.ergito.com

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Figure 14.23 DNA ligase seals nicks between adjacent nucleotides by employing an enzyme-AMP intermediate. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.14.12

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4.14.13 Separate eukaryotic DNA polymerases undertake initiation and elongation Key Concepts

• A replication fork has 1 complex of DNA polymerase α/primase and 2 complexes of DNA polymerase δ and/or ε.

• The DNA polymerase α/primase complex initiates the synthesis of both DNA strands.

• DNA polymerase δ elongates the leading strand and a second DNA polymerase δ or DNA polymerase elongates the lagging strand

Eukaryotic cells have a large number of DNA polymerases (for review see 3092). They can be broadly divided into those required for semiconservative replication and those involved in synthesizing material to repair damaged DNA. Nuclear DNA replication requires DNA polymerases α, δ, and ε, and mitochondrial replication requires DNA polymerase γ. All the other enzymes are concerned with synthesizing stretches of new DNA to replace damaged material. Figure 14.24 shows that all of the nuclear replicases are large heterotetrameric enzymes. In each case, one of the subunits has the responsibility for catalysis, and the others are concerned with ancillary functions, such as priming, processivity, or proofreading. These enzymes all replicate DNA with high fidelity, as does the slightly less complex mitochondrial enzyme. The repair polymerases have much simpler structures, often consisting of a single monomeric subunit (although it may function in the context of a complex of other repair enzymes). Of the enzymes involved in repair, only DNA polymerase β has a fidelity approaching the replicases: all of the others have much greater error rates (for review see 3097).

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Figure 14.24 Eukaryotic cells have many DNA polymerases. The replicative enzymes operate with high fidelity. Except for the β enzyme, the repair enzymes all have low fidelity. Replicative enzymes have large structures, with separate subunits for different activities. Repair enzymes have much simpler structures.

Each of the three nuclear DNA replicases has a different function: • DNA polymerase α initiates the synthesis of new strands. • DNA polymerase δ elongates the leading strand. • DNA polymerase ε may be involved in lagging strand synthesis, but also has other roles. DNA polymerase α is unusual because it has the ability to initiate a new strand. It is used to initiate both the leading and lagging strands. The enzyme exists as a complex consisting of a 180 kD catalytic subunit, associated with the B subunit that appears necessary for assembly, and two smaller proteins that provide a primase activity. Reflecting its dual capacity to prime and extend chains, it is sometimes called pol α/primase. The pol α/primase enzyme binds to the initiation complex at the origin and synthesizes a short strand consisting of ~10 bases of RNA followed by 20-30 bases of DNA (sometimes called iDNA). Then it is replaced by an enzyme that will extend the chain. On the leading strand, this is DNA polymerase δ. This event is called the pol switch. It involves interactions among several components of the initiation complex. Separate eukaryotic DNA polymerases undertake initiation and elongation | SECTION 4.14.13 © 2004. Virtual Text / www.ergito.com

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Molecular Biology DNA polymerase δ is a highly processive enzyme that continuously synthesizes the leading strand. Its processivity results from its interaction with two other proteins, RF-C and PCNA. The roles of RF-C and PCNA are analogous to the E. coli γ clamp loader and β processivity unit (see Molecular Biology 4.14.11 The clamp controls association of core enzyme with DNA). RF-C is a clamp loader that catalyzes the loading of PCNA on to DNA. It binds to the 3 ′ end of the iDNA and uses ATP-hydrolysis to open the ring of PCNA so that it can encircle the DNA (3095). The processivity of DNA polymerase δ is maintained by PCNA, which tethers DNA polymerase δ to the template. (PCNA is called proliferating cell nuclear antigen for historical reasons.) The crystal structure of PCNA closely resembles the E. coli β subunit: a trimer forms a ring that surrounds the DNA. Although the sequence and subunit organization are different from the dimeric β clamp, the function is likely to be similar. We are less certain about events on the lagging strand. One possibility is that DNA polymerase δ also elongates the lagging strand. It has the capability to dimerize, which suggests a model analogous to the behavior of E. coli replicase (see Molecular Biology 4.14.10 DNA polymerase holoenzyme has 3 subcomplexes) (3096). However, there are some indications that DNA polymerase ε may elongate the lagging strand (2189; 2208), although it also has been identified with other roles. A general model suggests that a replication fork contains 1 complex of DNA polymerase α/primase and two other DNA polymerase complexes. One is DNA polymerase δ and the other is either a second DNA polymerase δ or may possibly be a DNA polymerase ε. The two complexes of DNA polymerase δ/ ε behave in the same way as the two complexes of DNA polymerase III in the E. coli replisome: one synthesizes the leading strand, and the other synthesizes Okazaki fragments on the lagging strand. The exonuclease MF1 removes the RNA primers of Okazaki fragments. The enzyme DNA ligase I is specifically required to seal the nicks between the completed Okazaki fragments. Last updated on 11-20-2002


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Reviews 3092. Hubscher, U., Maga, G., and Spadari, S. (2002). Eukaryotic DNA polymerases. Annu. Rev. Biochem. 71, 133-163. 3097. Goodman, M. F. (2002). Error-prone repair DNA polymerases in prokaryotes and eukaryotes. Annu. Rev. Biochem. 71, 17-50.

References 2189. Karthikeyan, R., Vonarx, E. J., Straffon, A. F., Simon, M., Faye, G., and Kunz, B. A. (2000). Evidence from mutational specificity studies that yeast DNA polymerases delta and epsilon replicate different DNA strands at an intracellular replication fork. J. Mol. Biol. 299, 405-419. 2208. Waga, S., Masuda, T., Takisawa, H., and Sugino, A. (2001). DNA polymerase epsilon is required for coordinated and efficient chromosomal DNA replication in Xenopus egg extracts. Proc. Natl. Acad. Sci. USA 98, 4978-4983. 3095. Shiomi, Y., Usukura, J., Masamura, Y., Takeyasu, K., Nakayama, Y., Obuse, C., Yoshikawa, H., and Tsurimoto, T. (2000). ATP-dependent structural change of the eukaryotic clamp-loader protein, replication factor C. Proc. Natl. Acad. Sci. USA 97, 14127-14132. 3096. Zuo, S., Bermudez, V., Zhang, G., Kelman, Z., and Hurwitz, J. (2000). Structure and activity associated with multiple forms of S. pombe DNA polymerase delta. J. Biol. Chem. 275, 5153-5162.



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4.14.14 Phage T4 provides its own replication apparatus Key Concepts

• Phage T4 provides its own replication apparatus, which consists of DNA

polymerase, the gene 32 SSB, a helicase, a primase, and accessory proteins that increase speed and processivity.

When phage T4 takes over an E. coli cell, it provides several functions of its own that either replace or augment the host functions. The phage places little reliance on expression of host functions. The degradation of host DNA is important in releasing nucleotides that are reused in the synthesis of phage DNA. (The phage DNA differs in base composition from cellular DNA in using hydroxymethylcytosine instead of the customary cytosine.) The phage-coded functions concerned with DNA synthesis in the infected cell can be identified by mutations that impede the production of mature phages. Essential phage functions are identified by conditional lethal mutations, which fall into three phenotypic classes: • Those in which there is no DNA synthesis at all identify genes whose products either are components of the replication apparatus or are involved in the provision of precursors (especially the hydroxymethylcytosine). • Those in which the onset of DNA synthesis is delayed are concerned with the initiation of replication. • Those in which DNA synthesis starts but then is arrested include regulatory functions, the DNA ligase, and some of the enzymes concerned with host DNA degradation. • There are also nonessential genes concerned with replication; for example, including those involved in glucosylating the hydroxymethylcytosine in the DNA. Synthesis of T4 DNA is catalyzed by a multienzyme aggregate assembled from the products of a small group of essential genes. The gene 32 protein (gp32) is a highly cooperative single-strand binding protein, needed in stoichiometric amounts. It was the first example of its type to be characterized. The geometry of the T4 replication fork may specifically require the phage-coded protein, since the E. coli SSB cannot substitute. The gp32 forms a complex with the T4 DNA polymerase; this interaction could be important in constructing the replication fork. Phage T4 provides its own replication apparatus | SECTION 4.14.14 © 2004. Virtual Text / www.ergito.com

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The T4 system uses an RNA priming event that is similar to that of its host. With single-stranded T4 DNA as template, the gene 41 and 61 products act together to synthesize short primers. Their behavior is analogous to that of DnaB and DnaG in E. coli. The gene 41 protein is the counterpart to DnaB. It is a hexameric helicase that uses hydrolysis of GTP to provide the energy to unwind DNA (3100). The p41/p61 complex moves processively in the 5 ′ –3 ′ direction in lagging strand synthesis, periodically initiating Okazaki fragments. Another protein, the product of gene 59, loads the p41/p61 complex on to DNA; it is required to displace the p32 protein in order to allow the helicase to assemble on DNA (3099). The gene 61 protein is needed in much smaller amounts than most of the T4 replication proteins. There are as few as 10 copies of gp61 per cell. (This impeded its characterization. It is required in such small amounts that originally it was missed as a necessary component, because enough was present as a contaminant of the gp32 preparation!) Gene 61 protein has the primase activity, analogous to DnaG of E. coli. The primase recognizes the template sequence 3 ′ –TTG-5 ′ and synthesizes pentaribonucleotide primers that have the general sequence pppApCpNpNpNp. If the complete replication apparatus is present, these primers are extended into DNA chains. The gene 43 DNA polymerase has the usual 5 ′ –3 ′ synthetic activity, associated with a 3 ′ –5 ′ exonuclease proofreading activity. It catalyzes DNA synthesis and removes the primers. When T4 DNA polymerase uses a single-stranded DNA as template, its rate of progress is uneven. The enzyme moves rapidly through single-stranded regions, but proceeds much more slowly through regions that have a base-paired intrastrand secondary structure. The accessory proteins assist the DNA polymerase in passing these roadblocks, and maintaining its speed. The remaining three proteins are referred to as "polymerase accessory proteins". They increase the affinity of the DNA polymerase for the DNA, and also its processivity and speed. The gene 45 product is a trimer that acts as a sliding clamp. The structure of the trimer is similar to that of the E. coli β dimer, forming a circle around DNA that holds the DNA polymerase subunit more tightly on the template. The products of genes 44 and 62 form a tight complex, which has ATPase activity. They are the equivalent of the γ δ clamp loader complex, and their role is to load p45 on to DNA. Four molecules of ATP are hydrolyzed in loading the p45 clamp and the p43 DNA polymerase on to DNA. The overall structure of the replisome is similar to that of E. coli. It consists of two coupled holoenzyme complexes, one synthesizing the leading strand and the other synthesizing the lagging strand. In this case, the dimerization involves a direct interaction between the p43 DNA polymerase subunits, and p32 plays a role in coordinating the actions of the two DNA polymerase units (3101). We have dealt with DNA replication so far solely in terms of the progression of the replication fork. The need for other functions is shown by the DNA-delay and DNA-arrest mutants. The four genes of the DNA-delay mutants include 39, 52, and 60, which code for the three subunits of T4 topoisomerase II, an activity needed for removing supercoils in the template (see Molecular Biology 4.15.13 Topoisomerases relax or introduce supercoils in DNA). The essential role of this enzyme suggests Phage T4 provides its own replication apparatus | SECTION 4.14.14 © 2004. Virtual Text / www.ergito.com

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that T4 DNA does not remain in a linear form, but becomes topologically constrained during some stage of replication. The topoisomerase could be needed to allow rotation of DNA ahead of the replication fork. Comparison of the T4 apparatus with the E. coli apparatus suggests that DNA replication poses a set of problems that are solved in analogous ways in different systems. We may now compare the enzymatic and structural activities found at the replication fork in E. coli, T4, and HeLa (human) cells. Figure 14.25 summarizes the functions and assigns them to individual proteins. We can interpret the known properties of replication complex proteins in terms of similar functions, involving the unwinding, priming, catalytic, and sealing reactions. The components of each system interact in restricted ways, as shown by the fact that phage T4 requires its own helicase, primase, clamp, etc., and the bacterial proteins cannot substitute for their phage counterparts.

Figure 14.25 Similar functions are required at all replication forks.

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References 3099. Ishmael, F. T., Alley, S. C., and Benkovic, S. J. (2002). Assembly of the bacteriophage T4 helicase: architecture and stoichiometry of the gp41-gp59 complex. J. Biol. Chem. 277, 20555-20562. 3100. Schrock, R. D. and Alberts, B. (1996). Processivity of the gene 41 DNA helicase at the bacteriophage T4 DNA replication fork. J. Biol. Chem. 271, 16678-16682. 3101. Salinas, F., and Benkovic, S. J. (2000). Characterization of bacteriophage T4-coordinated leading- and lagging-strand synthesis on a minicircle substrate. Proc. Natl. Acad. Sci. USA 97, 7196-7201.


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4.14.15 Creating the replication forks at an origin Key Concepts

• Initiation at oriC requires the sequential assembly of a large protein complex. • DnaA binds to short repeated sequences and forms an oligomeric complex that melts DNA.

• 6 DnaC monomers bind each hexamer of DnaB and this complex binds to the origin.

• A hexamer of DnaB forms the replication fork. Gyrase and SSB are also required.

Starting a cycle of replication of duplex DNA requires several successive activities: • The two strands of DNA must suffer their initial separation. This is in effect a melting reaction over a short region. • An unwinding point begins to move along the DNA; this marks the generation of the replication fork, which continues to move during elongation. • The first nucleotides of the new chain must be synthesized into the primer. This action is required once for the leading strand, but is repeated at the start of each Okazaki fragment on the lagging strand. Some events that are required for initiation therefore occur uniquely at the origin; others recur with the initiation of each Okazaki fragment during the elongation phase. Plasmids carrying the E. coli oriC sequence have been used to develop a cell-free system for replication. Initiation of replication at oriC in vitro starts with formation of a complex that requires six proteins: DnaA, DnaB, DnaC, HU, Gyrase, and SSB. Of the six proteins involved in prepriming, DnaA draws our attention as the only one uniquely involved in initiation vis-À-vis elongation. DnaB/DnaC provides the "engine" of initiation at the origin. The first stage in complex formation is binding to oriC by DnaA protein (3102). The reaction involves action at two types of sequences: 9 bp and 13 bp repeats. Together the 9 bp and 13 bp repeats define the limits of the 245 bp minimal origin, as indicated in Figure 14.26. An origin is activated by the sequence of events summarized in Figure 14.27, in which binding of DnaA is succeeded by association with the other proteins.

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Figure 14.26 The minimal origin is defined by the distance between the outside members of the 13-mer and 9-mer repeats.

Figure 14.27 Prepriming involves formation of a complex by sequential association of proteins, leading to the separation of DNA strands.

The four 9 bp consensus sequences on the right side of oriC provide the initial binding sites for DnaA. It binds cooperatively to form a central core around which oriC DNA is wrapped. Then DnaA acts at three A-T-rich 13 bp tandem repeats located in the left side of oriC. In the presence of ATP, DnaA melts the DNA strands at each of these sites to form an open complex (3103; 3104). All three 13 bp repeats must be opened for the reaction to proceed to the next stage. Altogether, 2-4 monomers of DnaA bind at the origin, and they recruit 2 Creating the replication forks at an origin | SECTION 4.14.15 © 2004. Virtual Text / www.ergito.com

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"prepriming" complexes of DnaB-DnaC to bind, so that there is one for each of the two (bidirectional) replication forks. Each DnaB-DnaC complex consists of 6 DnaC monomers bound to a hexamer of DnaB. Each DnaB·DnaC complex transfers a hexamer of DnaB to an opposite strand of DNA. DnaC hydrolyzes ATP in order to release DnaB (3106). The prepriming complex generates a protein aggregate of 480 kD, corresponding to a sphere of radius 6 nm (3107). The formation of a complex at oriC is detectable in the form of the large protein blob visualized in Figure 14.28. When replication begins, a replication bubble becomes visible next to the blob.

Figure 14.28 The complex at oriC can be detected by electron microscopy. Both complexes were visualized with antibodies against DnaB protein. Photographs kindly provided by Barbara Funnell.

The region of strand separation in the open complex is large enough for both DnaB hexamers to bind, initiating the two replication forks. As DnaB binds, it displaces DnaA from the 13 bp repeats, and extends the length of the open region. Then it uses its helicase activity to extend the region of unwinding. Each DnaB activates a DnaG primase, in one case to initiate the leading strand, and in the other to initiate the first Okazaki fragment of the lagging strand. Two further proteins are required to support the unwinding reaction. Gyrase provides a swivel that allows one strand to rotate around the other (a reaction discussed in more detail in Molecular Biology 4.15.15 Gyrase functions by coil inversion); without this reaction, unwinding would generate torsional strain in the DNA. The protein SSB stabilizes the single-stranded DNA as it is formed. The length of duplex Creating the replication forks at an origin | SECTION 4.14.15 © 2004. Virtual Text / www.ergito.com

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DNA that usually is unwound to initiate replication is probably 50 types of transposon, with a total of several hundred individual elements. Transposable elements can promote rearrangements of the genome, directly or indirectly: • The transposition event itself may cause deletions or inversions or lead to the movement of a host sequence to a new location. • Transposons serve as substrates for cellular recombination systems by functioning as "portable regions of homology"; two copies of a transposon at different locations (even on different chromosomes) may provide sites for reciprocal recombination. Such exchanges result in deletions, insertions, inversions, or translocations. The intermittent activities of a transposon seem to provide a somewhat nebulous target for natural selection. This concern has prompted suggestions that (at least some) transposable elements confer neither advantage nor disadvantage on the phenotype, but could constitute "selfish DNA," concerned only with their own propagation. Indeed, in considering transposition as an event that is distinct from other cellular recombination systems, we tacitly accept the view that the transposon is an independent entity that resides in the genome. Such a relationship of the transposon to the genome would resemble that of a parasite with its host. Presumably the propagation of an element by transposition is balanced by the harm done if a transposition event inactivates a necessary gene, or if the number of transposons becomes a burden on cellular systems. Yet we must remember that any transposition event conferring a selective advantage – for example, a genetic rearrangement – will lead to preferential survival of the genome carrying the active transposon (for review see 146).


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Reviews 146. Campbell, A. (1981). Evolutionary significance of accessory DNA elements in bacteria. Annu. Rev. Immunol. 35, 55-83. 164. Finnegan, D. J. (1985). Transposable elements in eukaryotes. Int. Rev. Cytol. 93, 281-326.



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4.16.2 Insertion sequences are simple transposition modules Key Terms An insertion sequence (IS) is a small bacterial transposon that carries only the genes needed for its own transposition. Inverted terminal repeats are the short related or identical sequences present in reverse orientation at the ends of some transposons. Direct repeats are identical (or closely related) sequences present in two or more copies in the same orientation in the same molecule of DNA; they are not necessarily adjacent. A transposase is the enzyme activity involved in insertion of transposon at a new site. Key Concepts

• An insertion sequence is a transposon that codes for the enzyme(s) needed for transposition flanked by short inverted terminal repeats.

• The target site at which a transposon is inserted is duplicated during the insertion process to form two repeats in direct orientation at the ends of the transposon.

• The length of the direct repeat is 5-9 bp and is characteristic for any particular transposon.

Transposable elements were first identified at the molecular level in the form of spontaneous insertions in bacterial operons. Such an insertion prevents transcription and/or translation of the gene in which it is inserted. Many different types of transposable elements have now been characterized (for review see 143; 144; 145; 149). The simplest transposons are called insertion sequences (reflecting the way in which they were detected). Each type is given the prefix IS, followed by a number that identifies the type. (The original classes were numbered IS1-4; later classes have numbers reflecting the history of their isolation, but not corresponding to the total number of elements so far isolated!) The IS elements are normal constituents of bacterial chromosomes and plasmids. A standard strain of E. coli is likely to contain several (10-15 kb away from the original insertion. This is called "target immunity." It is demonstrated in an in vitro reaction containing donor (Mu-containing) and target (Mu-deficient) plasmids, MuA and MuB proteins, E. coli HU protein, and Mg2+ and ATP. The presence of MuB and ATP restricts transposition exclusively to the target plasmid. The reason is that when MuB binds to the MuA-Mu DNA complex, MuA causes MuB to hydrolyze ATP, after which MuB is released. However, MuB binds (nonspecifically) to the target DNA, where it stimulates the recombination activity of MuA when a transposition complex forms. In effect, the prior presence of MuA "clears" MuB from the donor, thus giving a preference for transposition to the target. The product of these reactions is a strand transfer complex in which the transposon is connected to the target site through one strand at each end. The next step of the reaction differs and determines the type of transposition. We see in the next two sections how the common structure can be a substrate for replication (leading to replicative transposition) or used directly for breakage and reunion (leading to nonreplicative transposition).

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Reviews 157. Pato, M. L. (1989). Bacteriophage Mu. Mobile DNA, 23-52. 160. Mizuuchi, K. (1992). Transpositional recombination: mechanistic insights from studies of Mu and other elements. Annu. Rev. Biochem. 61, 1011-1051.

References 571. Aldaz, H., Schuster, E., and Baker, T. A. (1996). The interwoven architecture of the Mu transposase couples DNA synthesis to catalysis. Cell 85, 257-269. 572. Savilahti, H. and Mizuuchi, K. (1996). Mu transpositional recombination: donor DNA cleavage and strand transfer in trans by the Mu transpose. Cell 85, 271-280.


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4.16.7 Replicative transposition proceeds through a cointegrate Key Terms A cointegrate structure is produced by fusion of two replicons, one originally possessing a transposon, the other lacking it; the cointegrate has copies of the transposon present at both junctions of the replicons, oriented as direct repeats. Resolution occurs by a homologous recombination reaction between the two copies of the transposon in a cointegrate. The reaction generates the donor and target replicons, each with a copy of the transposon. Resolvase is the enzyme activity involved in site-specific recombination between two transposons present as direct repeats in a cointegrate structure. Key Concepts

• Replication of a strand transfer complex generates a cointegrate, which is a fusion of the donor and target replicons.

• The cointegrate has two copies of the transposon, which lie between the original replicons.

• Recombination between the transposon copies regenerates the original replicons, but the recipient has gained a copy of the transposon.

• The recombination reaction is catalyzed by a resolvase coded by the transposon.

The basic structures involved in replicative transposition are illustrated in Figure 16.13:

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Figure 16.13 Transposition may fuse a donor and recipient replicon into a cointegrate. Resolution releases two replicons, each containing a copy of the transposon.

• The 3 ′ ends of the strand transfer complex are used as primers for replication. This generates a structure called a cointegrate, which represents a fusion of the two original molecules. The cointegrate has two copies of the transposon, one at each junction between the original replicons, oriented as direct repeats. The crossover is formed by the transposase, as described in the previous section. Its conversion into the cointegrate requires host replication functions. • A homologous recombination between the two copies of the transposon releases two individual replicons, each of which has a copy of the transposon. One of the replicons is the original donor replicon. The other is a target replicon that has gained a transposon flanked by short direct repeats of the host target sequence. The recombination reaction is called resolution; the enzyme activity responsible is called the resolvase. The reactions involved in generating a cointegrate have been defined in detail for phage Mu, and are illustrated in Figure 16.14. The process starts with the formation of the strand transfer complex (sometimes also called a crossover complex). The donor and target strands are ligated so that each end of the transposon sequence is joined to one of the protruding single strands generated at the target site. The strand transfer complex generates a crossover-shaped structure held together at the duplex transposon. The fate of the crossover structure determines the mode of transposition.


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Figure 16.14 Mu transposition generates a crossover structure, which is converted by replication into a cointegrate.

The principle of replicative transposition is that replication through the transposon duplicates it, creating copies at both the target and donor sites. The product is a cointegrate. The crossover structure contains a single-stranded region at each of the staggered ends. These regions are pseudoreplication forks that provide a template for DNA synthesis. (Use of the ends as primers for replication implies that the strand breakage Replicative transposition proceeds through a cointegrate | SECTION 4.16.7 © 2004. Virtual Text / www.ergito.com

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4.16.8 Nonreplicative transposition proceeds by breakage and reunion Key Concepts

• Nonreplicative transposition results if a crossover structure is nicked on the unbroken pair of donor strands, and the target strands on either side of the transposon are ligated.

• Two pathways for nonreplicative transposition differ according to whether the first pair of transposon strands are joined to the target before the second pair are cut (Tn5), or whether all four strands are cut before joining to the target (Tn10).

The crossover structure can also be used in nonreplicative transposition. The principle of nonreplicative transposition by this mechanism is that a breakage and reunion reaction allows the target to be reconstructed with the insertion of the transposon; the donor remains broken. No cointegrate is formed. Figure 16.15 shows the cleavage events that generate nonreplicative transposition of phage Mu. Once the unbroken donor strands have been nicked, the target strands on either side of the transposon can be ligated. The single-stranded regions generated by the staggered cuts must be filled in by repair synthesis. The product of this reaction is a target replicon in which the transposon has been inserted between repeats of the sequence created by the original single-strand nicks. The donor replicon has a double-strand break across the site where the transposon was originally located.

Figure 16.15 Nonreplicative transposition results when a crossover structure is released by nicking. This inserts the transposon into the target DNA, flanked by the direct repeats of the target, and the donor is left with a double-strand break.

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Nonreplicative transposition can also occur by an alternative pathway in which nicks are made in target DNA, but a double-strand break is made on either side of the transposon, releasing it entirely from flanking donor sequences (as envisaged in Figure 16.7). This "cut and paste" pathway is used by Tn10, as illustrated in Figure 16.16 (564).

Figure 16.16 Both strands of Tn10 are cleaved sequentially, and then the transposon is joined to the nicked target site.

A neat experiment to prove that Tn10 transposes nonreplicatively made use of an artificially constructed heteroduplex of Tn10 that contained single base mismatches. If transposition involves replication, the transposon at the new site will contain information from only one of the parent Tn10 strands. But if transposition takes place by physical movement of the existing transposon, the mismatches will be conserved at the new site, which proved to be the case (567). The basic difference in Figure 16.16 from the model of Figure 16.15 is that both strands of Tn10 are cleaved before any connection is made to the target site. The first step in the reaction is recognition of the transposon ends by the transposase, forming a proteinaceous structure within which the reaction occurs. At each end of the transposon, the strands are cleaved in a specific order – first the transferred strand (the one to be connected to the target site) is cleaved, then the other strand (this is the same order as in the Mu transposition of Figure 16.14 and Figure 16.15). Nonreplicative transposition proceeds by breakage and reunion | SECTION 4.16.8 © 2004. Virtual Text / www.ergito.com

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Tn5 also transposes by nonreplicative transposition, and Figure 16.17 shows the interesting cleavage reaction that separates the transposon from the flanking sequences (1084). First one DNA strand is nicked. The 3 ′ –OH end that is released then attacks the other strand of DNA. This releases the flanking sequence and joins the two strands of the transposon in a hairpin. Then an activated water molecule attacks the hairpin to generate free ends for each strand of the transposon.

Figure 16.17 Cleavage of Tn5 from flanking DNA involves nicking, interstrand reaction, and hairpin cleavage.

Then the cleaved donor DNA is released, and the transposon is joined to the nicked ends at the target site. The transposon and the target site remain constrained in the proteinaceous structure created by the transposase (and other proteins). The double-strand cleavage at each end of the transposon precludes any replicative-type transposition and forces the reaction to proceed by nonreplicative transposition, thus giving the same outcome as in Figure 16.14, but with the individual cleavage and joining steps occurring in a different order. The Tn5 and Tn10 transposases both function as dimers. Each subunit in the dimer has an active site that successively catalyzes the double-strand breakage of the two strands at one end of the transposon and then catalyzes staggered cleavage of the target site (570). Figure 16.18 illustrates the structure of the Tn5 transposase bound to the cleaved transposon (1085). Each end of the transposon is located in the active site of one subunit. One end of the subunit also contacts the other end of the transposon. This controls the geometry of the transposition reaction. Each of the active sites will cleave one strand of the target DNA. It is the geometry of the complex that determines the distance between these sites on the two target strands (9 base pairs in the case of Tn5).


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Figure 16.18 Each subunit of the Tn5 transposase has one end of the transposon located in its active site and also makes contact at a different site with the other end of the transposon.



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References 564. Haniford, D. B., Benjamin, H. W., and Kleckner, N. (1991). Kinetic and structural analysis of a cleaved donor intermediate and a strand transfer intermediate in Tn10 transposition. Cell 64, 171-179. 567. Bender, J. and Kleckner, N. (1986). Genetic evidence that Tn10 transposes by a nonreplicative mechanism. Cell 45, 801-815. 570. Bolland, S. and Kleckner, N. (1996). The three chemical steps of Tn10/IS10 transposition involve repeated utilization of a single active site. Cell 84, 223-233. 1084. Kennedy, A. K., Guhathakurta, A., Kleckner, N., and Haniford, D. B. (1998). Tn10 transposition via a DNA hairpin intermediate. Cell 95, 125-134. 1085. Davies, D. R., Goryshin, I. Y., Reznikoff, W. S., Rayment, I., Davies, D. R., Goryshin, I. Y., Reznikoff, W. S., and Rayment, I. (2000). Three-dimensional structure of the Tn5 synaptic complex transposition intermediate. Science 289, 77-85.



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4.16.9 TnA transposition requires transposase and resolvase Key Concepts

• Replicative transposition of TnA requires a transposase to form the cointegrate structure and a resolvase to release the two replicons.

• The action of the resolvase resembles lambda Int protein and belongs to the general family of topoisomerase-like site-specific recombination reactions, which pass through an intermediate in which the protein is covalently bound to the DNA.

Replicative transposition is the only mode of mobility of the TnA family, which consists of large (~5 kb) transposons. They are not composites relying on IS-type transposition modules, but comprise independent units carrying genes for transposition as well as for features such as drug resistance. The TnA family includes several related transposons, of which Tn3 and Tn1000 (formerly called γ δ) are the best characterized. They have the usual terminal feature of closely related inverted repeats, generally ~38 bp in length. Cis-acting deletions in either repeat prevent transposition of an element. A 5 bp direct repeat is generated at the target site. They carry resistance markers such as ampr. The two stages of TnA-mediated transposition are accomplished by the transposase and the resolvase, whose genes, tnpA and tnpR, are identified by recessive mutations. The transposition stage involves the ends of the element, as it does in IS-type elements. Resolution requires a specific internal site. This feature is unique to the TnA family (for review see 155). Mutants in tnpA cannot transpose. The gene product is a transposase that binds to a sequence of ~25 bp located within the 38 bp of the inverted terminal repeat. A binding site for the E. coli protein IHF exists adjacent to the transposase binding site; and transposase and IHF bind cooperatively. The transposase recognizes the ends of the element and also makes the staggered 5 bp breaks in target DNA where the transposon is to be inserted. IHF is a DNA-binding protein that is often involved in assembling large structures in E. coli; its role in the transposition reaction may not be essential. The tnpR gene product has dual functions. It acts as a repressor of gene expression and it provides the resolvase function. Mutations in tnpR increase the transposition frequency. The reason is that TnpR represses the transcription of both tnpA and its own gene. So inactivation of TnpR protein allows increased synthesis of TnpA, which results in an increased frequency of transposition. This implies that the amount of the TnpA transposase must be a limiting factor in transposition.

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The tnpA and tnpR genes are expressed divergently from an A·T-rich intercistronic control region, indicated in the map of Tn3 given in Figure 16.19. Both effects of TnpR are mediated by its binding in this region.

Figure 16.19 Transposons of the TnA family have inverted terminal repeats, an internal res site, and three known genes.

In its capacity as the resolvase, TnpR is involved in recombination between the direct repeats of Tn3 in a cointegrate structure. A cointegrate can in principle be resolved by a homologous recombination between any corresponding pair of points in the two copies of the transposon. But the Tn3 resolution reaction occurs only at a specific site (569). The site of resolution is called res. It is identified by cis-acting deletions that block completion of transposition, causing the accumulation of cointegrates. In the absence of res, the resolution reaction can be substituted by RecA-mediated general recombination, but this is much less efficient. The sites bound by the TnpR resolvase are summarized in the lower part of Figure 16.19. Binding occurs independently at each of three sites, each 30-40 bp long. The three binding sites share a sequence homology that defines a consensus sequence with dyad symmetry (565). Site I includes the region genetically defined as the res site; in its absence, the resolution reaction does not proceed at all. However, resolution also involves binding at sites II and III, since the reaction proceeds only poorly if either of these sites is deleted. Site I overlaps with the startpoint for tnpA transcription. Site II overlaps with the startpoint for tnpR transcription; an operator mutation maps just at the left end of the site. Do the sites interact? One possibility is that binding at all three sites is required to hold the DNA in an appropriate topology. Binding at a single set of sites may repress tnpA and tnpR transcription without introducing any change in the DNA. An in vitro resolution assay uses a cointegrate-like DNA molecule as substrate. The TnA transposition requires transposase and resolvase | SECTION 4.16.9 © 2004. Virtual Text / www.ergito.com

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substrate must be supercoiled; its resolution produces two catenated circles, each containing one res site. The reaction requires large amounts of the TnpR resolvase; no host factors are needed. Resolution occurs in a large nucleoprotein structure. Resolvase binds to each res site, and then the bound sites are brought together to form a structure ~10 nm in diameter. Changes in supercoiling occur during the reaction, and DNA is bent at the res sites by the binding of transposase. Resolution occurs by breaking and rejoining bonds without input of energy. The products identify an intermediate stage in cointegrate resolution; they consist of resolvase covalently attached to both 5 ′ ends of double-stranded cuts made at the res site. The cleavage occurs symmetrically at a short palindromic region to generate two base extensions. Expanding the view of the crossover region located in site I, we can describe the cutting reaction as:

The reaction resembles the action of lambda Int at the att sites (see Molecular Biology 4.15.17 Site-specific recombination involves breakage and reunion). Indeed, 15 of the 20 bp of the res site are identical to the bases at corresponding positions in att. This suggests that the site-specific recombination of lambda and resolution of TnA have evolved from a common type of recombination reaction; and indeed, we see in Molecular Biology 5.25.9 The RAG proteins catalyze breakage and reunion that recombination involving immunoglobulin genes has the same basis. The common feature in all these reactions is the transfer of the broken end to the catalytic protein as an intermediate stage before it is rejoined to another broken end (see Molecular Biology 4.15.18 Site-specific recombination resembles topoisomerase activity). The reactions themselves are analogous in terms of manipulation of DNA, although resolution occurs only between intramolecular sites, whereas the recombination between att sites is intermolecular and directional (as seen by the differences in attB and attP sites). However, the mechanism of protein action is different in each case. Resolvase functions in a manner in which four subunits bind to the recombining res sites. Each subunit makes a single-strand cleavage. Then a reorganization of the subunits relative to one another physically moves the DNA strands, placing them in a recombined conformation. This allows the nicks to be sealed, along with the release of resolvase.


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Reviews 155. Sherratt, D. (1989). Tn3 and related transposable elements: site-specific recombination and transposition. Mobile DNA, 163-185.

References 565. Grindley, N. D. et al. (1982). Transposon-mediated site-specific recombination: identification of three binding sites for resolvase at the res sites of γ δ and Tn3. Cell 30, 19-27. 569. Droge, P. et al. (1990). The two functional domains of gamma delta resolvase act on the same recombination site: implications for the mechanism of strand exchange. Proc. Natl. Acad. Sci. USA 87, 5336-5340.



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4.16.10 Transposition of Tn10 has multiple controls Key Concepts

• Multicopy inhibition reduces the rate of transposition of any one copy of a

transposon when other copies of the same transposon are introduced into the genome.

• Multiple mechanisms affect the rate of transposition.

Control of the frequency of transposition is important for the cell. A transposon must be able to maintain a certain minimum frequency of movement in order to survive; but too great a frequency could be damaging to the host cell. Every transposon appears to have mechanisms that control its frequency of transposition. A variety of mechanisms have been characterized for Tn10 (for review see 154; 159). Tn10 is a composite transposon in which the element IS10R provides the active module. The organization of IS10R is summarized in Figure 16.20. Two promoters are found close to the outside boundary. The promoter PIN is responsible for transcription of IS10R. The promoter POUT causes transcription to proceed toward the adjacent flanking DNA. Transcription usually terminates within the transposon, but occasionally continues into the host DNA; sometimes this readthrough transcription is responsible for activating adjacent bacterial genes.

Figure 16.20 Two promoters in opposite orientation lie near the outside boundary of IS10R. The strong promoter POUT sponsors transcription toward the flanking host DNA. The weaker promoter PIN causes transcription of an RNA that extends the length of IS10R and is translated into the transposase.

The phenomenon of "multicopy inhibition" reveals that expression of the IS10R transposase gene is regulated. Transposition of a Tn10 element on the bacterial chromosome is reduced when additional copies of IS10R are introduced via a multicopy plasmid. The inhibition requires the POUT promoter, and is exercised at the Transposition of Tn10 has multiple controls | SECTION 4.16.10 © 2004. Virtual Text / www.ergito.com

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Molecular Biology level of translation. The basis for the effect lies with the overlap in the 5 ′ terminal regions of the transcripts from PIN and POUT. OUT RNA is a transcript of 69 bases. It is present at >100× the level of IN RNA for two reasons: POUT is a much stronger promoter than PIN; and OUT RNA is more stable than IN RNA. OUT RNA functions as an antisense RNA (see Molecular Biology 3.11.19 Small RNA molecules can regulate translation). The level of OUT RNA has no effect in a single-copy situation, but has a significant effect when >5 copies are present. There are usually ~5 copies of OUT RNA per copy of IS10 (which corresponds to ~150 copies of OUT RNA in a typical multicopy situation). OUT RNA base pairs with IN RNA; and the excess of OUT RNA ensures that IN RNA is bound rapidly, before a ribosome can attach. So the paired IN RNA cannot be translated. The quantity of transposase protein is often a critical feature. Tn10, whose transposase is synthesized at the low level of 0.15 molecules per cell per generation, displays several interesting mechanisms. Figure 16.21 summarizes the various effects that influence transposition frequency.

Figure 16.21 Several mechanisms restrain the frequency of Tn10 transposition, by affecting either the synthesis or function of transposase protein. Transposition of an individual transposon is restricted by methylation to occur only after replication. In multicopy situations, cis-preference restricts the choice of target, and OUT/IN RNA pairing inhibits synthesis of transposase.

A continuous reading frame on one strand of IS10R codes for the transposase. The level of the transposase limits the rate of transposition. Mutants in this gene can be complemented in trans by another, wild-type IS10 element, but only with some difficulty. This reflects a strong preference of the transposase for cis-action; the enzyme functions efficiently only with the DNA template from which it was transcribed and translated. Cis-preference is a common feature of transposases coded by IS elements. (Other proteins that display cis-preference include the A protein involved in φX174 replication; see Molecular Biology 4.13.11 Rolling circles are used to replicate phage genomes.) Does cis-preference reflect an ability of the transposase to recognize more efficiently Transposition of Tn10 has multiple controls | SECTION 4.16.10 © 2004. Virtual Text / www.ergito.com

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those DNA target sequences that lie nearer to the site where the enzyme is synthesized? One possible explanation is that the transposase binds to DNA so tightly after (or even during) protein synthesis that it has a very low probability of diffusing elsewhere. Another possibility is that the enzyme may be unstable when it is not bound to DNA, so that protein molecules failing to bind quickly (and therefore nearby) never have a chance to become active. Together the results of cis-preference and multicopy inhibition ensure that an increase in the number of copies of Tn10 in a bacterial genome does not cause an increased frequency of transposition that could damage the genome. The effects of methylation provide the most important system of regulation for an individual element. They reduce the frequency of transposition and (more importantly) couple transposition to passage of the replication fork. The ability of IS10 to transpose is related to the replication cycle by the transposon's response to the state of methylation at two sites. One site is within the inverted repeat at the end of IS10R, where the transposase binds. The other site is in the promoter PIN, from which the transposase gene is transcribed. Both of these sites are methylated by the dam system described in Molecular Biology 4.14.18 Does methylation at the origin regulate initiation? The Dam methylase modifies the adenine in the sequence GATC on a newly synthesized strand generated by replication. The frequency of Tn10 transposition is increased 1000-fold in dam– strains in which the two target sites lack methyl groups. Passage of a replication fork over these sites generates hemimethylated sequences; this activates the transposon by a combination of transcribing the transposase gene more frequently from PIN and enhancing binding of transposase to the end of IS10R. In a wild-type bacterium, the sites remain hemimethylated for a short period after replication (566). Why should it be desirable for transposition to occur soon after replication? The nonreplicative mechanism of Tn10 transposition places the donor DNA at risk of being destroyed (see Figure 16.7). The cell's chances of survival may be increased if replication has just occurred to generate a second copy of the donor sequence. The mechanism is effective because only 1 of the 2 newly replicated copies gives rise to a transposition event (determined by which strand of the transposon is unmethylated at the dam sites). Since a transposon selects its target site at random, there is a reasonable probability that it may land in an active operon. Will transcription from the outside continue through the transposon and thus activate the transposase, whose overproduction may in turn lead to high (perhaps lethal) levels of transposition? Tn10 protects itself against such events by two mechanisms. Transcription across the IS10R terminus decreases its activity, presumably by inhibiting its ability to bind transposase. And the mRNA that extends from upstream of the promoter is poorly translated, because it has a secondary structure in which the initiation codon is inaccessible.

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Reviews 154. Kleckner, N. (1989). Transposon Tn10. Mobile DNA, 227-268. 159. Kleckner, N. (1990). Regulation of transposition in bacteria. Annu. Rev. Cell Biol. 6, 297-327.

References 566. Roberts, D. et al. (1985). IS10 transposition is regulated by DNA adenine methylation. Cell 43, 117-130.


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4.16.11 Controlling elements in maize cause breakage and rearrangements Key Terms Controlling elements of maize are transposable units originally identified solely by their genetic properties. They may be autonomous (able to transpose independently) or nonautonomous (able to transpose only in the presence of an autonomous element). A sector is a patch of cells made up of a single altered cell and its progeny. Variegation of phenotype is produced by a change in genotype during somatic development. An acentric fragment of a chromosome (generated by breakage) lacks a centromere and is lost at cell division. A dicentric chromosome is the product of fusing two chromosome fragments, each of which has a centromere. It is unstable and may be broken when the two centromeres are pulled to opposite poles in mitosis. The breakage-fusion-bridge cycle is a type of chromosomal behavior in which a broken chromatid fuses to its sister, forming a "bridge". When the centromeres separate at mitosis, the chromosome breaks again (not necessarily at the bridge), thereby restarting the cycle. Key Concepts

• Transposition in maize was discovered because of the effects of the chromosome breaks generated by transposition of "controlling elements".

• The break generates one chromosome that has a centromere and a broken end and one acentric fragment.

• The acentric fragment is lost during mitosis, and this can be detected by the disappearance of dominant alleles in a heterozygote.

• Fusion between the broken ends of the chromosome generates dicentric chromosomes, which undergo further cycles of breakage and fusion.

• The fusion-breakage-bridge cycle is responsible for the occurrence of somatic variegation.

One of the most visible consequences of the existence and mobility of transposons occurs during plant development, when somatic variation occurs. This is due to changes in the location or behavior of controlling elements (the name that transposons were given in maize before their molecular nature was discovered. For an account of the discovery see Great Experiments 2.6 The discovery of transposition).

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Two features of maize have helped to follow transposition events. Controlling elements often insert near genes that have visible but nonlethal effects on the phenotype. And because maize displays clonal development, the occurrence and timing of a transposition event can be visualized as depicted diagrammatically in Figure 16.22.

Figure 16.22 Clonal analysis identifies a group of cells descended from a single ancestor in which a transpositionmediated event altered the phenotype. Timing of the event during development is indicated by the number of cells; tissue specificity of the event may be indicated by the location of the cells.

The nature of the event does not matter: it may be a point mutation, insertion, excision, or chromosome break. What is important is that it occurs in a heterozygote to alter the expression of one allele. Then the descendants of a cell that has suffered the event display a new phenotype, while the descendants of cells not affected by the event continue to display the original phenotype. Mitotic descendants of a given cell remain in the same location and give rise to a sector of tissue. A change in phenotype during somatic development is called variegation; it is revealed by a sector of the new phenotype residing within the tissue of the original phenotype. The size of the sector depends on the number of divisions in the lineage giving rise to it; so the size of the area of the new phenotype is determined by the timing of the change in genotype. The earlier its occurrence in the Controlling elements in maize cause breakage and rearrangements | SECTION 4.16.11 © 2004. Virtual Text / www.ergito.com

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cell lineage, the greater the number of descendants and thus the size of patch in the mature tissue. This is seen most vividly in the variation in kernel color, when patches of one color appear within another color. Insertion of a controlling element may affect the activity of adjacent genes. Deletions, duplications, inversions, and translocations all occur at the sites where controlling elements are present. Chromosome breakage is a common consequence of the presence of some elements. A unique feature of the maize system is that the activities of the controlling elements are regulated during development. The elements transpose and promote genetic rearrangements at characteristic times and frequencies during plant development. The characteristic behavior of controlling elements in maize is typified by the Ds element, which was originally identified by its ability to provide a site for chromosome breakage. The consequences are illustrated in Figure 16.23. Consider a heterozygote in which Ds lies on one homologue between the centromere and a series of dominant markers. The other homologue lacks Ds and has recessive markers (C, bz, wx). Breakage at Ds generates an acentric fragment carrying the dominant markers. Because of its lack of a centromere, this fragment is lost at mitosis. So the descendant cells have only the recessive markers carried by the intact chromosome. This gives the type of situation whose results are depicted in Figure 16.22.

Figure 16.23 A break at a controlling element causes loss of an acentric fragment; if the fragment carries the dominant markers of a heterozygote, its loss changes the phenotype. The effects of the dominant markers, CI, Bz, Wx, can be visualized by the color of the cells or by appropriate staining.

Figure 16.24 shows that breakage at Ds leads to the formation of two unusual chromosomes. These are generated by joining the broken ends of the products of replication. One is a U-shaped acentric fragment consisting of the joined sister chromatids for the region distal to Ds (on the left as drawn in the figure). The other is a U-shaped dicentric chromosome comprising the sister chromatids proximal to Ds Controlling elements in maize cause breakage and rearrangements | SECTION 4.16.11 © 2004. Virtual Text / www.ergito.com

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(on its right in the figure). The latter structure leads to the classic breakage-fusion-bridge cycle illustrated in the figure.

Figure 16.24 Ds provides a site to initiate the chromatid fusion-bridge-breakage cycle. The products can be followed by clonal analysis.

Follow the fate of the dicentric chromosome when it attempts to segregate on the mitotic spindle. Each of its two centromeres pulls toward an opposite pole. The tension breaks the chromosome at a random site between the centromeres. In the example of the figure, breakage occurs between loci A and B, with the result that one daughter chromosome has a duplication of A, while the other has a deletion. If A is a dominant marker, the cells with the duplication will retain a phenotype, but cells Controlling elements in maize cause breakage and rearrangements | SECTION 4.16.11 © 2004. Virtual Text / www.ergito.com

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with the deletion will display the recessive a phenotype. The breakage-fusion-bridge cycle continues through further cell generations, allowing genetic changes to continue in the descendants. For example, consider the deletion chromosome that has lost A. In the next cycle, a break occurs between B and C, so that the descendants are divided into those with a duplication of B and those with a deletion. Successive losses of dominant markers are revealed by subsectors within sectors. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.4.16.11

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TRANSPOSONS

4.16.12 Controlling elements form families of transposons Key Terms An autonomous controlling element in maize is an active transposon with the ability to transpose (compare with nonautonomous controlling element). A nonautonomous controlling element is a transposon in maize that encodes a non-functional transposase; it can transpose only in the presence of a trans-acting autonomous member of the same family. Key Concepts

• Each family of transposons in maize has both autonomous and nonautonomous controlling elements.

• Autonomous controlling elements code for proteins that enable them to transpose. • Nonautonomous controlling elements have mutations that eliminate their capacity to catalyze transposition, but they can transpose when an autonomous element provides the necessary proteins.

• Autonomous controlling elements have changes of phase, when their properties alter as a result of changes in the state of methylation.

The maize genome contains several families of controlling elements. The numbers, types, and locations of the elements are characteristic for each individual maize strain. They may occupy a significant part of the genome (for review see 1182). The members of each family are divided into two classes: • Autonomous controlling elements have the ability to excise and transpose. Because of the continuing activity of an autonomous element, its insertion at any locus creates an unstable or "mutable" allele. Loss of the autonomous element itself, or of its ability to transpose, converts a mutable allele to a stable allele. • Nonautonomous controlling elements are stable; they do not transpose or suffer other spontaneous changes in condition. They become unstable only when an autonomous member of the same family is present elsewhere in the genome. When complemented in trans by an autonomous element, a nonautonomous element displays the usual range of activities associated with autonomous elements, including the ability to transpose to new sites. Nonautonomous elements are derived from autonomous elements by loss of trans-acting functions needed for transposition. Families of controlling elements are defined by the interactions between autonomous and nonautonomous elements. A family consists of a single type of autonomous element accompanied by many varieties of nonautonomous elements. A Controlling elements form families of transposons | SECTION 4.16.12 © 2004. Virtual Text / www.ergito.com

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nonautonomous element is placed in a family by its ability to be activated in trans by the autonomous elements. The major families of controlling elements in maize are summarized in Figure 16.25 (for review see 152; 158).

Figure 16.25 Each controlling element family has both autonomous and nonautonomous members. Autonomous elements are capable of transposition. Nonautonomous elements are deficient in transposition. Pairs of autonomous and nonautonomous elements can be classified in >4 families.

Characterized at the molecular level, the maize transposons share the usual form of organization – inverted repeats at the ends and short direct repeats in the adjacent target DNA – but otherwise vary in size and coding capacity. All families of transposons share the same type of relationship between the autonomous and nonautonomous elements. The autonomous elements have open reading frames between the terminal repeats, whereas the nonautonomous elements do not code for functional proteins. Sometimes the internal sequences are related to those of autonomous elements; sometimes they have diverged completely. The Mutator transposon is one of the simplest elements. The autonomous element MuDR codes for the genes mudrA (which codes for the MURA transposase) and mudrB (which codes for a nonessential accessory protein). The ends of the elements are marked by 200 bp inverted repeats. Nonautonomous elements – basically any unit that has the inverted repeats, which may not have any internal sequence relationship to MuDR – are also mobilized by MURA (1860). There are typically several members (~10) of each transposon family in a plant genome. By analyzing autonomous and nonautonomous elements of the Ac/Ds family, we have molecular information about many individual examples of these elements. Figure 16.26 summarizes their structures.

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Figure 16.26 The Ac element has two open reading frames; Ds elements have internal deletions.

Most of the length of the autonomous Ac element is occupied by a single gene consisting of 5 exons. The product is the transposase. The element itself ends in inverted repeats of 11 bp; and a target sequence of 8 bp is duplicated at the site of insertion. Ds elements vary in both length and sequence, but are related to Ac. They end in the same 11 bp inverted repeats. They are shorter than Ac, and the length of deletion varies. At one extreme, the element Ds9 has a deletion of only 194 bp. In a more extensive deletion, the Ds6 element retains a length of only 2 kb, representing 1 kb from each end of Ac. A complex double Ds element has one Ds6 sequence inserted in reverse orientation into another. Nonautonomous elements lack internal sequences, but possess the terminal inverted repeats (and possibly other sequence features). Nonautonomous elements are derived from autonomous elements by deletions (or other changes) that inactivate the trans-acting transposase, but leave intact the sites (including the termini) on which the transposase acts. Their structures range from minor (but inactivating) mutations of Ac to sequences that have major deletions or rearrangements. At another extreme, the Ds1 family members comprise short sequences whose only relationship to Ac lies in the possession of terminal inverted repeats. Elements of this class need not be directly derived from Ac, but could be derived by any event that generates the inverted repeats. Their existence suggests that the transposase recognizes only the terminal inverted repeats, or possibly the terminal repeats in conjunction with some short internal sequence. Transposition of Ac/Ds occurs by a nonreplicative mechanism, and is accompanied by its disappearance from the donor location. Clonal analysis suggests that transposition of Ac/Ds almost always occurs soon after the donor element has been replicated. These features resemble transposition of the bacterial element Tn10 (see Molecular Biology 4.16.10 Transposition of Tn10 has multiple controls ). The cause Controlling elements form families of transposons | SECTION 4.16.12 © 2004. Virtual Text / www.ergito.com

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is the same: transposition does not occur when the DNA of the transposon is methylated on both strands (the typical state before methylation), and is activated when the DNA is hemimethylated (the typical state immediately after replication) (3143). The recipient site is frequently on the same chromosome as the donor site, and often quite close to it. Replication generates two copies of a potential Ac/Ds donor, but usually only one copy actually transposes. What happens to the donor site? The rearrangements that are found at sites from which controlling elements have been lost could be explained in terms of the consequences of a chromosome break, as illustrated previously in Figure 16.23. Autonomous and nonautonomous elements are subject to a variety of changes in their condition. Some of these changes are genetic, others are epigenetic. The major change is (of course) the conversion of an autonomous element into a nonautonomous element, but further changes may occur in the nonautonomous element. Cis-acting defects may render a nonautonomous element impervious to autonomous elements. So a nonautonomous element may become permanently stable because it can no longer be activated to transpose. Autonomous elements are subject to "changes of phase," heritable but relatively unstable alterations in their properties. These take the form of a reversible inactivation in which the element cycles between an active and inactive condition during plant development. Phase changes in both the Ac and Mu types of autonomous element result from changes in the methylation of DNA. Comparisons of the susceptibilities of active and inactive elements to restriction enzymes suggest that the inactive form of the element is methylated in the target sequence . There are several target sites in each element, and we do not know which sites control the effect. In the case of MuDR, demethylation of the terminal repeats increases transposase expression, suggesting that the effect may mediated through control of the promoter for the transposase gene (1862). We should like to know what controls the methylation and demethylation of the elements. The effect of methylation is common generally among transposons in plants. The best demonstration of the effect of methylation on activity comes from observations made with the Arabidopsis mutant ddm1, which causes a loss of methylation in heterochromatin. Among the targets that lose methyl groups is a family of transposons related to MuDR. Direct analysis of genome sequences shows that the demethylation causes transposition events to occur (1861). Methylation is probably the major mechanism that is used to prevent transposons from damaging the genome by transposing too frequently. There may be self-regulating controls of transposition, analogous to the immunity effects displayed by bacterial transposons. An increase in the number of Ac elements in the genome decreases the frequency of transposition. The Ac element may code for a repressor of transposition; the activity could be carried by the same protein that Controlling elements form families of transposons | SECTION 4.16.12 © 2004. Virtual Text / www.ergito.com

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provides transposase function. Last updated on 5-29-2001


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Reviews 152. Fedoroff, N. (1989). Maize transposable elements. Mobile DNA, 375-412. 158. Gierl, A., Saedler, H., and Peterson, P.A. (1989). Maize transposable elements. Annu. Rev. Genet. 23, 71-85. 1182. Fedoroff, N. (2000). Transposons and genome evolution in plants. Proc. Natl. Acad. Sci. USA 97, 7002-7007.

References 1860. Benito, M. I. and Walbot, V. (1997). Characterization of the maize Mutator transposable element MURA transposase as a DNA-binding protein. Mol. Cell Biol. 17, 5165-5175. 1861. Singer, T., Yordan, C., and Martienssen, R. A. (2001). Robertson's Mutator transposons in A. thaliana are regulated by the chromatin-remodeling gene Decrease in DNA Methylation (DDM1). Genes Dev. 15, 591-602. 1862. Chandler, V. L. and Walbot, V. (1986). DNA modification of a maize transposable element correlates with loss of activity. Proc. Natl. Acad. Sci. USA 83, 1767-1771. 3143. Ros, F. and Kunze, R. (2001). Regulation of activator/dissociation transposition by replication and DNA methylation. Genetics 157, 1723-1733.



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TRANSPOSONS

4.16.13 Spm elements influence gene expression Key Concepts

• Spm elements affect gene expression at their sites of insertion, when the TnpA protein binds to its target sites at the ends of the transposon.

• Spm elements are inactivated by methylation.

The Spm and En autonomous elements are virtually identical; they differ at 500 kD), comparable in size to RNA polymerase itself, and containing 6 subunits. TFIIIA is a member of an interesting class of proteins containing a nucleic acid-binding motif called a zinc finger (see Molecular Biology 5.22.9 A zinc finger motif is a DNA-binding domain). The positioning factor, TFIIIIB, consists of three subunits. It includes the same protein, TBP, that is present in the core-binding factor for pol I promoters, and also in the corresponding transcription factor (TFIID) for RNA polymerase II (1677). It also contains Brf, which is related to the factor TFIIB that is used by RNA polymerase II. The third subunit is called B ′ ′ ; it is dispensable if the DNA duplex is partially melted, which suggests that its function is to initiate the transcription bubble (945). The role of B ′ ′ may be comparable to the role played by sigma factor in bacterial RNA polymerase (see Molecular Biology 3.9.16 Substitution of sigma factors may control initiation ). TFIIIB is the commitment factor for pol III promoters | SECTION 5.21.6 © 2004. Virtual Text / www.ergito.com

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The upstream region has a conventional role in the third class of polymerase III promoters. In the example shown in Figure 21.7, there are three upstream elements. These elements are also found in promoters for snRNA genes that are transcribed by RNA polymerase II. (Genes for some snRNAs are transcribed by RNA polymerase II, while others are transcribed by RNA polymerase III.) The upstream elements function in a similar manner in promoters for both polymerases II and III. Initiation at an upstream promoter for RNA polymerase III can occur on a short region that immediately precedes the startpoint and contains only the TATA element. However, efficiency of transcription is much increased by the presence of the PSE and OCT elements. The factors that bind at these elements interact cooperatively. (The PSE element may be essential at promoters used by RNA polymerase II, whereas it is stimulatory in promoters used by RNA polymerase III; its name stands for proximal sequence element.) The TATA element confers specificity for the type of polymerase (II or III) that is recognized by an snRNA promoter. It is bound by a factor that includes the TBP, which actually recognizes the sequence in DNA. The TBP is associated with other proteins, which are specific for the type of promoter. The function of TBP and its associated proteins is to position the RNA polymerase correctly at the startpoint. We discuss this in more detail for RNA polymerase II (see Molecular Biology 5.21.8 TBP is a universal factor). The factors work in the same way for both types of promoters for RNA polymerase III. The factors bind at the promoter before RNA polymerase itself can bind. They form a preinitiation complex that directs binding of the RNA polymerase. RNA polymerase III does not itself recognizes the promoter sequence, but binds adjacent to factors that are themselves bound just upstream of the startpoint. For the type 1 and type 2 internal promoters, the assembly factors ensure that TFIIIB (which includes TBP) is bound just upstream of the startpoint, to provide the positioning information. For the upstream promoters, TFIIIB binds directly to the region including the TATA box. So irrespective of the location of the promoter sequences, factor(s) are bound close to the startpoint in order to direct binding of RNA polymerase III (for review see 220; 3231). Last updated on 1-6-2003

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Reviews 220. Geiduschek, E. P. and Tocchini-Valentini, G. P. (1988). Transcription by RNA polymerase III. Annu. Rev. Biochem. 57, 873-914. 3231. Schramm, L. and Hernandez, N. (2002). Recruitment of RNA polymerase III to its target promoters. Genes Dev. 16, 2593-2620.

References 643. Kassavatis, G. A., Braun, B. R., Nguyen, L. H., and Geiduschek, E. P. (1990). S. cerevisiae TFIIIB is the transcription initiation factor proper of RNA polymerase III, while TFIIIA and TFIIIC are assembly factors. Cell 60, 235-245. 945. Kassavetis, G. A., Letts, G. A., and Geiduschek, E. P. (1999). A minimal RNA polymerase III transcription system. EMBO J. 18, 5042-5051. 1677. Kassavetis, G. A., Joazeiro, C. A., Pisano, M., Geiduschek, E. P., Colbert, T., Hahn, S., and Blanco, J. A. (1992). The role of the TATA-binding protein in the assembly and function of the multisubunit yeast RNA polymerase III transcription factor, TFIIIB. Cell 71, 1055-1064.


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PROMOTERS AND ENHANCERS

5.21.7 The startpoint for RNA polymerase II Key Terms TAFs (TFIIX) are the subunits of TFIID that assist TBP in binding to DNA. They also provide points of contact for other components of the transcription apparatus. The core promoter of RNA polymerase I is the region immediately surrounding the startpoint. It is necessary and sufficient to initiate transcription, but only at a low level. A core promoter is the shortest sequence at which an RNA polymerase can initiate transcription (typically at much lower level than that displayed by a promoter containing additional elements). For RNA polymerase II it is the minimal sequence at which the basal transcription apparatus can assemble, and includes two sequence elements, the InR and TATA box. It is typically ~40 bp long. The Inr is the sequence of a pol II promoter between –3 and +5 and has the general sequence Py2CAPy5. It is the simplest possible pol II promoter. TATA box is a conserved A·T-rich septamer found about 25 bp before the startpoint of each eukaryotic RNA polymerase II transcription unit; may be involved in positioning the enzyme for correct initiation. A TATA-less promoter does not have a TATA box in the sequence upstream of its startpoint. Key Concepts

• RNA polymerase II requires general transcription factors (called TFIIX) to initiate transcription.

• RNA polymerase II promoters have a short conserved sequence Py2CAPy5 (the initiator InR) at the startpoint.

• The TATA box is a common component of RNA polymerase II promoters and consists of an A·T-rich octamer located ~25 bp upstream of the startpoint

• The DPE is a common component of RNA polymerase II promoters that do not contain a TATA box.

• A core promoter for RNA polymerase II includes the InR and either a TATA box or a DPE.

The basic organization of the apparatus for transcribing protein-coding genes was revealed by the discovery that purified RNA polymerase II can catalyze synthesis of mRNA, but cannot initiate transcription unless an additional extract is added (2406). The purification of this extract led to the definition of the general transcription factors – a group of proteins that are needed for initiation by RNA polymerase II at all promoters (for review see 2407). RNA polymerase II in conjunction with these factors constitutes the basal transcription apparatus that is needed to transcribe any promoter. The general factors are described as TFIIX, where "X" is a letter that The startpoint for RNA polymerase II | SECTION 5.21.7 © 2004. Virtual Text / www.ergito.com

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identifies the individual factor. The subunits of RNA polymerase II and the general transcription factors are conserved among eukaryotes. Our starting point for considering promoter organization is to define the core promoter as the shortest sequence at which RNA polymerase II can initiate transcription. A core promoter can in principle be expressed in any cell. It comprises the minimum sequence that enables the general transcription factors to assemble at the startpoint. They are involved in the mechanics of binding to DNA and enable RNA polymerase II to initiate transcription. A core promoter functions at only a low efficiency. Other proteins, called activators, are required for a proper level of function (see Molecular Biology 5.21.13 Short sequence elements bind activators). The activators are not described systematically, but have casual names reflecting their histories of identification. We may expect any sequence components involved in the binding of RNA polymerase and general transcription factors to be conserved at most or all promoters. As with bacterial promoters, when promoters for RNA polymerase II are compared, homologies in the regions near the startpoint are restricted to rather short sequences. These elements correspond with the sequences implicated in promoter function by mutation. Figure 21.10 shows the construction of a typical pol II core promoter (for review see 3225, 4527).

Figure 21.10 The minimal pol II promoter has a TATA box ~25 bp upstream of the InR. The TATA box has the consensus sequence of TATAA. The Inr has pyrimidines (Y) surrounding the CA at the startpoint. The sequence shows the coding strand.

At the startpoint, there is no extensive homology of sequence, but there is a tendency for the first base of mRNA to be A, flanked on either side by pyrimidines. (This description is also valid for the CAT start sequence of bacterial promoters.) This region is called the initiator (Inr), and may be described in the general form Py2CAPy5 (3228; 3229). The Inr is contained between positions –3 and +5. Many promoters have a sequence called the TATA box, usually located ~25 bp upstream of the startpoint. It constitutes the only upstream promoter element that has a relatively fixed location with respect to the startpoint. The core sequence is TATAA, usually followed by three more A·T base pairs (see 3227). The TATA box tends to be surrounded by G·C-rich sequences, which could be a factor in its function. It is almost identical with the –10 sequence found in bacterial promoters; in fact, it could pass for one except for the difference in its location at –25 instead of –10. The startpoint for RNA polymerase II | SECTION 5.21.7 © 2004. Virtual Text / www.ergito.com

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Single base substitutions in the TATA box act as strong down mutations. Some mutations reverse the orientation of an A·T pair, so base composition alone is not sufficient for its function. So the TATA box comprises an element whose behavior is analogous to our concept of the bacterial promoter: a short, well-defined sequence just upstream of the startpoint, which is necessary for transcription. Promoters that do not contain a TATA element are called TATA-less promoters. Surveys of promoter sequences suggest that 50% or more of promoters may be TATA-less. When a promoter does not contain a TATA box, it usually contains another element, the DPE (downstream promoter element) which is located at +28 +32 (3230). A core promoter can consist either of a TATA box plus InR or of an InR plus DPE. Last updated on January 12, 2004

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Reviews 2407. Woychik, N. A. and Hampsey, M. (2002). The RNA polymerase II machinery: structure illuminates function. Cell 108, 453-463. 3225. Butler, J. E. and Kadonaga, J. T. (2002). The RNA polymerase II core promoter: a key component in the regulation of gene expression. Genes Dev. 16, 2583-2592. 3229. Smale, S. T., Jain, A., Kaufmann, J., Emami, K. H., Lo, K., and Garraway, I. P. (1998). The initiator element: a paradigm for core promoter heterogeneity within metazoan protein-coding genes. Cold Spring Harb Symp Quant Biol 63, 21-31. 4527. Smale, S. T. and Kadonaga, J. T. (2003). The RNA polymerase II core promoter. Annu. Rev. Biochem. 72, 449-479.

References 2406. Weil, P. A., Luse, D. S., Segall, J., and Roeder, R. G. (1979). Selective and accurate initiation of transcription at the Ad2 major late promoter in a soluble system dependent on purified RNA polymerase II and DNA. Cell 18, 469-484. 3227. Singer, V. L., Wobbe, C. R., and Struhl, K. (1990). A wide variety of DNA sequences can functionally replace a yeast TATA element for transcriptional activation. Genes Dev. 4, 636-645. 3228. Smale, S. T. and Baltimore, D. (1989). The "initiator" as a transcription control element. Cell 57, 103-113. 3230. Burke, T. W. and Kadonaga, J. T. (1996). Drosophila TFIID binds to a conserved downstream basal promoter element that is present in many TATA-box-deficient promoters. Genes Dev. 10, 711-724.


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5.21.8 TBP is a universal factor Key Terms TFIID is the transcription factor that binds to the TATA sequence upstream of the startpoint of promoters for RNA polymerase II. It consists of TBP (TATA binding protein) and the TAF subunits that bind to TBP. The TATA-binding protein (TBP) is the subunit of transcription factor TFIID that binds to DNA. TAFs (TFIIX) are the subunits of TFIID that assist TBP in binding to DNA. They also provide points of contact for other components of the transcription apparatus. Key Concepts

• TBP is a component of the positioning factor that is required for each type of RNA polymerase to bind its promoter.

• The factor for RNA polymerase II is TFIID, which consists of TBP and 11 TAFs, with a total mass ~800 kD.

The first step in complex formation at a promoter containing a TATA box is binding of the factor TFIID to a region that extends upstream from the TATA sequence. TFIID contains two types of component. Recognition of the TATA box is conferred by the TATA-binding protein (TBP), a small protein of ~30 kD. The other subunits are called TAFs (for TBP-associated factors). Some TAFs are stoichiometric with TBP; others are present in lesser amounts. TFIIDs containing different TAFs could recognize different promoters. Some (substoichiometric) TAFs are tissue-specific. The total mass of TFIID typically is ~800 kD, containing TBP and 11 TAFs, varying in mass from 30-250 kD. The TAFs in TFIID are named in the form TAFII00, where "00" gives the molecular mass of the subunit. Positioning factors that consist of TBP associated with a set of TAFs are responsible for identifying all classes of promoters. TFIIIB (for pol III promoters) and SL1 (for pol I promoters) may both be viewed as consisting of TBP associated with a particular group of proteins that substitute for the TAFs that are found in TFIID (for review see 1709). TBP is the key component, and is incorporated at each type of promoter by a different mechanism (for review see 2394). In the case of promoters for RNA polymerase II, the key feature in positioning is the fixed distance of the TATA box from the startpoint. Figure 21.11 shows that the positioning factor recognizes the promoter in a different way in each case. At promoters for RNA polymerase III, TFIIIB binds adjacent to TFIIIC. At promoters for RNA polymerase I, SL1 binds in conjunction with UBF. TFIID is solely responsible for recognizing promoters for RNA polymerase II. At a promoter that has a TATA element, TBP binds specifically to DNA, but at other promoters it may be incorporated by association with other proteins that bind to DNA. Whatever its means of entry into the initiation complex, it has the common TBP is a universal factor | SECTION 5.21.8 © 2004. Virtual Text / www.ergito.com

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purpose of interaction with the RNA polymerase.

Figure 21.11 RNA polymerases are positioned at all promoters by a factor that contains TBP.

TFIID is ubiquitous, but not unique. All multicellular eukaryotes also express an alternative complex, which has TLF (TBP like factor) instead of TBP (1708). A TLF is typically ~60% similar to TBP. It probably initiates complex formation by the usual set of TFII factors. However, TLF does not bind to the TATA box, and we do not yet know how it works. Drosophila also has a third factor, TRF1, which behaves in the same way as TBP and binds its own set of TAFs, to form a complex that functions as an alternative to TFIID at a specific set of promoters (1707). Last updated on 4-30-2001 TBP is a universal factor | SECTION 5.21.8 © 2004. Virtual Text / www.ergito.com

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Reviews 1708. Berk, A. J. (2000). TBP-like factors come into focus. Cell 103, 5-8. 1709. Lee, T. I. and Young, R. A. (1998). Regulation of gene expression by TBP-associated proteins. Genes Dev. 12, 1398-1408. 2394. Hernandez, N. (1993). TBP, a universal eukaryotic transcription factor? Genes Dev. 7, 1291-1308.

References 1707. Crowley, T. E., Hoey, T., Liu, J. K., Jan, Y. N., Jan, L. Y., and Tjian, R. (1993). A new factor related to TATA-binding protein has highly restricted expression patterns in Drosophila. Nature 361, 557-561.


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5.21.9 TBP binds DNA in an unusual way Key Concepts

• TBP binds to the TATA box in the minor groove of DNA. • It forms a saddle around the DNA and bends it by ~80°. • Some of the TAFs resemble histones and may form a structure resembling a histone octamer.

TBP has the unusual property of binding to DNA in the minor groove. (Virtually all known DNA-binding proteins bind in the major groove.) The crystal structure of TBP suggests a detailed model for its binding to DNA. Figure 21.12 shows that it surrounds one face of DNA, forming a "saddle" around the double helix. In effect, the inner surface of TBP binds to DNA, and the larger outer surface is available to extend contacts to other proteins. The DNA-binding site consists of a C-terminal domain that is conserved between species, while the variable N-terminal tail is exposed to interact with other proteins (647; 648; 649). It is a measure of the conservation of mechanism in transcriptional initiation that the DNA-binding sequence of TBP is 80% conserved between yeast and Man.

Figure 21.12 A view in cross-section shows that TBP surrounds DNA from the side of the narrow groove. TBP consists of two related (40% identical) conserved domains, which are shown in light and dark blue. The N-terminal region varies extensively and is shown in green. The two strands of the DNA double helix are in light and dark grey. Photograph kindly provided by Stephen Burley.

Binding of TBP may be inconsistent with the presence of nucleosomes. Because nucleosomes form preferentially by placing A·T-rich sequences with the minor grooves facing inward, they could prevent binding of TBP. This may explain why the presence of nucleosomes prevents initiation of transcription. TBP binds to the minor groove and bends the DNA by ~80°, as illustrated in Figure TBP binds DNA in an unusual way | SECTION 5.21.9 © 2004. Virtual Text / www.ergito.com

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21.13. The TATA box bends towards the major groove, widening the minor groove. The distortion is restricted to the 8 bp of the TATA box; at each end of the sequence, the minor groove has its usual width of ~5 Å, but at the center of the sequence the minor groove is >9 Å. This is a deformation of the structure, but does not actually separate the strands of DNA, because base pairing is maintained. The extent of the bend can vary with the exact sequence of the TATA box, and is correlated with the efficiency of the promoter (4532).

Figure 21.13 The cocrystal structure of TBP with DNA from -40 to the startpoint shows a bend at the TATA box that widens the narrow groove where TBP binds. Photograph provided by Stephen Burley.

This structure has several functional implications. By changing the spatial organization of DNA on either side of the TATA box, it allows the transcription factors and RNA polymerase to form a closer association than would be possible on linear DNA. The bending at the TATA box corresponds to unwinding of about 1/3 of a turn of DNA, and is compensated by a positive writhe. The presence of TBP in the minor groove, combined with other proteins binding in the major groove, creates a high density of protein-DNA contacts in this region. Binding of purified TBP to DNA in vitro protects ~1 turn of the double helix at the TATA box, typically extending from –37 to –25; but binding of the TFIID complex in the initiation reaction regularly protects the region from –45 to –10, and also extends farther upstream beyond the startpoint. TBP is the only general transcription factor that makes sequence-specific contacts with DNA. Within TFIID as a free protein complex, the factor TAFII230 binds to TBP, where it occupies the concave DNA-binding surface. In fact, the structure of the binding site, which lies in the N-terminal domain of TAFII230, mimics the surface of the minor groove in DNA. This molecular mimicry allows TAFII230 to control the ability of TBP to bind to DNA; the N-terminal domain of TAFII230 must be displaced from the DNA-binding surface of TBP in order for TFIID to bind to DNA (654). Some TAFs resemble histones; in particular TAFII42 and TAFII62 appear to be (distant) homologues of histones H3 and H4, and they form a heterodimer using the same motif (the histone fold) that histones use for the interaction. (Histones H3 and H4 form the kernel of the histone octamer – the basic complex that binds DNA in TBP binds DNA in an unusual way | SECTION 5.21.9 © 2004. Virtual Text / www.ergito.com

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eukaryotic chromatin; see Molecular Biology 5.20.8 Organization of the histone octamer.) Together with other TAFs, TAFII42 and TAFII62 may form the basis for a structure resembling a histone octamer; such a structure may be responsible for the nonsequence-specific interactions of TFIID with DNA. Histone folds are also used in pairwise interactions between other TAFIIs. Some of the TAFIIs may be found in other complexes as well as in TFIID. In particular, the histone-like TAFIIs are found also in protein complexes that modify the structure of chromatin prior to transcription (see Molecular Biology 5.23.7 Acetylases are associated with activators) (651; 653; 657; 695; for review see 225; 1709; 2395). Last updated on January 19, 2004

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Reviews 225. Burley, S. K. and Roeder, R. G. (1996). Biochemistry and structural biology of TFIID. Annu. Rev. Biochem. 65, 769-799. 1709. Lee, T. I. and Young, R. A. (1998). Regulation of gene expression by TBP-associated proteins. Genes Dev. 12, 1398-1408. 2395. Orphanides, G., Lagrange, T., and Reinberg, D. (1996). The general transcription factors of RNA polymerase II. Genes Dev. 10, 2657-2683.

References 647. Nikolov, D. B. et al. (1992). Crystal structure of TFIID TATA-box binding protein. Nature 360, 40-46. 648. Kim, Y. et al. (1993). Crystal structure of a yeast TBP/TATA box complex. Nature 365, 512-520. 649. Kim, J. L., Nikolov, D. B., and Burley, S. K. (1993). Cocrystal structure of TBP recognizing the minor groove of a TATA element. Nature 365, 520-527. 651. Martinez, E. et al. (1994). TATA-binding protein-associated factors in TFIID function through the initiator to direct basal transcription from a TATA-less class II promoter. EMBO J. 13, 3115-3126. 653. Verrijzer, C. P. et al. (1995). Binding of TAFs to core elements directs promoter selectivity by RNA polymerase II. Cell 81, 1115-1125. 654. Liu, D. et al. (1998). Solution structure of a TBP-TAFII230 complex: protein mimicry of the minor groove surface of the TATA box unwound by TBP. Cell 94, 573-583. 657. Horikoshi, M. et al. (1988). Transcription factor ATD interacts with a TATA factor to facilitate establishment of a preinitiation complex. Cell 54, 1033-1042. 695. Ogryzko, V. V. et al. (1998). Histone-like TAFs within the PCAF histone acetylase complex. Cell 94, 35-44. 4532. Wu, J., Parkhurst, K. M., Powell, R. M., Brenowitz, M., and Parkhurst, L. J. (2001). DNA bends in TATA-binding protein-TATA complexes in solution are DNA sequence-dependent. J. Biol. Chem. 276, 14614-14622.


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5.21.10 The basal apparatus assembles at the promoter Key Concepts

• Binding of TFIID to the TATA box is the first step in initiation. • Other transcription factors bind to the complex in a defined order, extending the length of the protected region on DNA.

• When RNA polymerase II binds to the complex, it initiates transcription.

Initiation requires the transcription factors to act in a defined order to build a complex that is joined by RNA polymerase. The series of events was initially defined by following the increasing size of the protein complex associated with DNA. Now we can define the events in more detail in terms of the interactions revealed by the crystal structures of the various factors, and of RNA polymerase bound to DNA. Footprinting of the DNA regions protected by each complex suggests the model summarized in Figure 21.15. As each TFII factor joins the complex, an increasing length of DNA is covered. RNA polymerase is incorporated at a late stage (644; for review see 223; 226).

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Figure 21.15 An initiation complex assembles at promoters for RNA polymerase II by an ordered sequence of association with transcription factors.

Commitment to a promoter is initiated when TFIID binds the TATA box. (TFIID also recognizes the InR sequence at the startpoint.) When TFIIA joins the complex, TFIID becomes able to protect a region extending farther upstream. TFIIA may activate TBP by relieving the repression that is caused by the TAFII230. Addition of TFIIB gives partial protection of the region of the template strand in the vicinity of the startpoint, from –10 to +10. This suggests that TFIIB is bound downstream of the TATA box, perhaps loosely associated with DNA and asymmetrically oriented with regard to the two DNA strands. The crystal structure shown in Figure 21.16 extends this model. TFIIB binds adjacent to TBP, extending contacts along one face of DNA. It makes contacts in the minor groove downstream of the TATA box, and contacts the major groove upstream of the TATA box, in a region called the BRE (2408). In archaea, the homologue of TFIIB actually makes sequence-specific contacts with the promoter in the BRE region (652). TFIIB may provide the surface that is in turn recognized by RNA polymerase, so that it is responsible for the directionality of the binding of the enzyme.


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Figure 21.16 Two views of the ternary complex of TFIIB-TBP-DNA show that TFIIB binds along the bent face of DNA. The two strands of DNA are green and yellow, TBP is blue, and TFIIB is red and purple. Photograph kindly provided by Stephen Burley.

The crystal structure of TFIIB with RNA polymerase shows that three domains of the factor interact with the enzyme(4832). As illustrated schematically in Figure 21.14, an N-terminal zinc ribbon from TFIIB contacts the enzyme near the site where RNA exits; it is possible that this interferes with the exit of RNA and influences the switch from abortive initiation to promoter escape. An elongated "finger" of TFIIB is inserted into the polymerase active center. The C-terminal domain interacts with the RNA polymerase and with TFIID to orient the DNA. It also determines the path of the DNA where it contacts the factors TFIIE, TFIIF, and TFIIH, which may align them in the basal factor complex.


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Figure 21.14 TFIIB binds to DNA and contacts RNA polymerase near the RNA exit site and at the active center, and orients it on DNA. Compare with 1170, which shows the polymerase structure engaged in transcription.

The factor TFIIF is a heterotetramer consisting of two types of subunit. The larger subunit (RAP74) has an ATP-dependent DNA helicase activity that could be involved in melting the DNA at initiation. The smaller subunit (RAP38) has some homology to the regions of bacterial sigma factor that contact the core polymerase; it binds tightly to RNA polymerase II. TFIIF may bring RNA polymerase II to the assembling transcription complex and provide the means by which it binds. The complex of TBP and TAFs may interact with the CTD tail of RNA polymerase, and interaction with TFIIB may also be important when TFIIF/polymerase joins the complex. Polymerase binding extends the sites that are protected downstream to +15 on the template strand and +20 on the nontemplate strand. The enzyme extends the full length of the complex, since additional protection is seen at the upstream boundary. What happens at TATA-less promoters? The same general transcription factors, including TFIID, are needed. The Inr provides the positioning element; TFIID binds to it via an ability of one or more of the TAFs to recognize the Inr directly. Other TAFs in TFIID also recognize the DPE element downstream from the startpoint (3230). The function of TBP at these promoters is more like that at promoters for RNA polymerase I and at internal promoters for RNA polymerase III. Assembly of the RNA polymerase II initiation complex provides an interesting contrast with prokaryotic transcription. Bacterial RNA polymerase is essentially a coherent aggregate with intrinsic ability to bind DNA; the sigma factor, needed for initiation but not for elongation, becomes part of the enzyme before DNA is bound, although it is later released. But RNA polymerase II can bind to the promoter only after separate transcription factors have bound. The factors play a role analogous to that of bacterial sigma factor – to allow the basic polymerase to recognize DNA The basal apparatus assembles at the promoter | SECTION 5.21.10 © 2004. Virtual Text / www.ergito.com

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specifically at promoter sequences – but have evolved more independence. Indeed, the factors are primarily responsible for the specificity of promoter recognition. Only some of the factors participate in protein-DNA contacts (and only TBP makes sequence-specific contacts); thus protein-protein interactions are important in the assembly of the complex. When a TATA box is present, it determines the location of the startpoint. Its deletion causes the site of initiation to become erratic, although any overall reduction in transcription is relatively small. Indeed, some TATA-less promoters lack unique startpoints; initiation occurs instead at any one of a cluster of startpoints. The TATA box aligns the RNA polymerase (via the interaction with TFIID and other factors) so that it initiates at the proper site. This explains why its location is fixed with respect to the startpoint. Binding of TBP to TATA is the predominant feature in recognition of the promoter, but two large TAFs (TAFII250 and TAFII150) also contact DNA in the vicinity of the startpoint and influence the efficiency of the reaction. Although assembly can take place just at the core promoter in vitro, this reaction is not sufficient for transcription in vivo, where interactions with activators that recognize the more upstream elements are required. The activators interact with the basal apparatus at various stages during its assembly (see Molecular Biology 5.22.5 Activators interact with the basal apparatus). Last updated on March 9, 2004


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Reviews 223. Zawel, L. and Reinberg, D. (1993). Initiation of transcription by RNA polymerase II: a multi-step process. Prog. Nucleic Acid Res. Mol. Biol. 44, 67-108. 226. Nikolov, D. B. and Burley, S. K. (1997). RNA polymerase II transcription initiation: a structural view. Proc. Natl. Acad. Sci. USA 94, 15-22.

References 644. Buratowski, S., Hahn, S., Guarente, L., and Sharp, P. A. (1989). Five intermediate complexes in transcription initiation by RNA polymerase II. Cell 56, 549-561. 652. Nikolov, D. B. et al. (1995). Crystal structure of a TFIIB-TBP-TATA-element ternary complex. Nature 377, 119-128. 2408. Littlefield, O., Korkhin, Y., and Sigler, P. B. (1999). The structural basis for the oriented assembly of a TBP/TFB/promoter complex. Proc. Natl. Acad. Sci. USA 96, 13668-13673. 3230. Burke, T. W. and Kadonaga, J. T. (1996). Drosophila TFIID binds to a conserved downstream basal promoter element that is present in many TATA-box-deficient promoters. Genes Dev. 10, 711-724. 4832. Bushnell, D. A., Westover, K. D., Davis, R. E., and Kornberg, R. D. (2004). Structural basis of transcription: an RNA polymerase II-TFIIB cocrystal at 4.5 Angstroms. Science 303, 983-988.



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5.21.11 Initiation is followed by promoter clearance Key Concepts

• TFIIE and TFIIH are required to melt DNA to allow polymerase movement. • Phosphorylation of the CTD may be required for elongation to begin. • Further phosphorylation of the CTD is required at some promoters to end abortive initiation.

• The CTD may coordinate processing of RNA with transcription.

Most of the transcription factors are required solely to bind RNA polymerase to the promoter, but some act at a later stage (for review see 2407). Binding of TFIIE causes the boundary of the region protected downstream to be extended by another turn of the double helix, to +30. Two further factors, TFIIH and TFIIJ, join the complex after TFIIE. They do not change the pattern of binding to DNA. TFIIH is the only general transcription factor that has independent enzymatic activities. Its several activities include an ATPase, helicases of both polarities, and a kinase activity that can phosphorylate the CTD tail of RNA polymerase II. TFIIH is an exceptional factor that may play a role also in elongation. Its interaction with DNA downstream of the startpoint is required for RNA polymerase to escape from the promoter (2207). TFIIH is also involved in repair of damage to DNA (see Molecular Biology 5.21.12 A connection between transcription and repair) (650). The initiation reaction, as defined by formation of the first phosphodiester bond, occurs once RNA polymerase has bound. Figure 21.17 proposes a model in which phosphorylation of the tail is needed to release RNA polymerase II from the transcription factors so that it can make the transition to the elongating form. Most of the transcription factors are released from the promoter at this stage.

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Figure 21.17 Phosphorylation of the CTD by the kinase activity of TFIIH may be needed to release RNA polymerase to start transcription.

On a linear template, ATP hydrolysis, TFIIE, and the helicase activity of TFIIH (provided by the XPB subunit) are required for polymerase movement. This requirement is bypassed with a supercoiled template. This suggests that TFIIE and TFIIH are required to melt DNA to allow polymerase movement to begin (946). The helicase activity of the XPB subunit of TFIIH is responsible for the actual melting of DNA (2409; 2410). RNA polymerase II stutters at some genes when it starts transcription. (The result is not dissimilar to the abortive initiation of bacterial RNA polymerase discussed in Molecular Biology 3.9.11 Sigma factor controls binding to DNA, although the mechanism is different.) At many genes, RNA polymerase II terminates after a short distance. The short RNA product is degraded rapidly. To extend elongation into the gene, a kinase called P-TEFb is required (for review see 948). This kinase is a member of the cdk family that controls the cell cycle (see Molecular Biology 6.29 Cell cycle and growth regulation). P-TEFb acts on the CTD, to Initiation is followed by promoter clearance | SECTION 5.21.11 © 2004. Virtual Text / www.ergito.com

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phosphorylate it further. We do not yet understand why this effect is required at some promoters but not others or how it is regulated. The CTD may also be involved, directly or indirectly, in processing RNA after it has been synthesized by RNA polymerase II (for review see 2421, 4528). Figure 21.18 summarizes processing reactions in which the CTD may be involved. The capping enzyme (guanylyl transferase), which adds the G residue to the 5 ′ end of newly synthesized mRNA, binds to the phosphorylated CTD: this may be important in enabling it to modify the 5 ′ end as soon as it is synthesized. A set of proteins called SCAFs bind to the CTD, and they may in turn bind to splicing factors. This may be a means of coordinating transcription and splicing. Some components of the cleavage/polyadenylation apparatus also bind to the CTD. Oddly enough, they do so at the time of initiation, so that RNA polymerase is all ready for the 3 ′ end processing reactions as soon as it sets out! All of this suggests that the CTD may be a general focus for connecting other processes with transcription (2239; for review see 2007, 4181). In the cases of capping and splicing, the CTD functions indirectly to promote formation of the protein complexes that undertake the reactions. In the case of 3 ′ end generation, it may participate directly in the reaction.


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Figure 21.18 The CTD is important in recruiting enzymes that modify RNA.

The general process of initiation is similar to that catalyzed by bacterial RNA polymerase. Binding of RNA polymerase generates a closed complex, which is converted at a later stage to an open complex in which the DNA strands have been separated. In the bacterial reaction, formation of the open complex completes the necessary structural change to DNA; a difference in the eukaryotic reaction is that further unwinding of the template is needed after this stage. Last updated on 9-24-2003


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Reviews 948. Price, D. H. (2000). P-TEFb, a cyclin dependent kinase controlling elongation by RNA polymerase II. Mol. Cell Biol. 20, 2629-2634. 2007. Hirose, Y. and Manley, J. L. (2000). RNA polymerase II and the integration of nuclear events. Genes Dev. 14, 1415-1429. 2407. Woychik, N. A. and Hampsey, M. (2002). The RNA polymerase II machinery: structure illuminates function. Cell 108, 453-463. 2421. Proudfoot, N. J., Furger, A., and Dye, M. J. (2002). Integrating mRNA processing with transcription. Cell 108, 501-512. 4181. Calvo, O. and Manley, J. L. (2003). Strange bedfellows: polyadenylation factors at the promoter. Genes Dev. 17, 1321-1327. 4528. Shilatifard, A., Conaway, R. C., and Conaway, J. W. (2003). The RNA polymerase II elongation complex. Annu. Rev. Biochem. 72, 693-715.

References 650. Goodrich, J. A. and Tjian, R. (1994). Transcription factors IIE and IIH and ATP hydrolysis direct promoter clearance by RNA polymerase II. Cell 77, 145-156. 946. Holstege, F. C., van der Vliet, P. C., and Timmers, H. T. (1996). Opening of an RNA polymerase II promoter occurs in two distinct steps and requires the basal transcription factors IIE and IIH. EMBO J. 15, 1666-1677. 2207. Spangler, L., Wang, X., Conaway, J. W., Conaway, R. C., and Dvir, A. (2001). TFIIH action in transcription initiation and promoter escape requires distinct regions of downstream promoter DNA. Proc. Natl. Acad. Sci. USA 98, 5544-5549. 2239. Fong, N. and Bentley, D. L. (2001). Capping, splicing, and 3 ′ processing are independently stimulated by RNA polymerase II: different functions for different segments of the CTD. Genes Dev. 15, 1783-1795. 2409. Douziech, M., Coin, F., Chipoulet, J. M., Arai, Y., Ohkuma, Y., Egly, J. M., and Coulombe, B. (2000). Mechanism of promoter melting by the xeroderma pigmentosum complementation group B helicase of transcription factor IIH revealed by protein-DNA photo-cross-linking. Mol. Cell Biol. 20, 8168-8177. 2410. Kim, T. K., Ebright, R. H., and Reinberg, D. (2000). Mechanism of ATP-dependent promoter melting by transcription factor IIH. Science 288, 1418-1422.



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5.21.12 A connection between transcription and repair Key Concepts

• Transcribed genes are preferentially repaired when DNA damage occurs. • TFIIH provides the link to a complex of repair enzymes. • Mutations in the XPD component of TFIIH cause three types of human diseases

In both bacteria and eukaryotes, there is a direct link from RNA polymerase to the activation of repair. The basic phenomenon was first observed because transcribed genes are preferentially repaired. Then it was discovered that it is only the template strand of DNA that is the target – the nontemplate strand is repaired at the same rate as bulk DNA. In bacteria, the repair activity is provided by the uvr excision-repair system (see Molecular Biology 4.15.21 Excision repair systems in E. coli). Preferential repair is abolished by mutations in the gene mfd, whose product provides the link from RNA polymerase to the Uvr enzymes (for review see 224). Figure 21.19 shows a model for the link between transcription and repair. When RNA polymerase encounters DNA damage in the template strand, it stalls because it cannot use the damaged sequences as a template to direct complementary base pairing. This explains the specificity of the effect for the template strand (damage in the nontemplate strand does not impede progress of the RNA polymerase).

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Figure 21.19 Mfd recognizes a stalled RNA polymerase and directs DNA repair to the damaged template strand.

The Mfd protein has two roles. First, it displaces the ternary complex of RNA polymerase from DNA. Second, it causes the UvrABC enzyme to bind to the damaged DNA. This leads to repair of DNA by the excision-repair mechanism (see Figure 15.40). After the DNA has been repaired, the next RNA polymerase to traverse the gene is able to produce a normal transcript (661). A similar mechanism, although relying on different components, is used in eukaryotes. The template strand of a transcribed gene is preferentially repaired following UV-induced damage. The general transcription factor TFIIH is involved. TFIIH is found in alternative forms, which consist of a core associated with other subunits. A connection between transcription and repair | SECTION 5.21.12 © 2004. Virtual Text / www.ergito.com

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TFIIH has a common function in both initiating transcription and repairing damage. The same helicase subunit (XPD) creates the initial transcription bubble and melts DNA at a damaged site. Its other functions differ between transcription and repair, as provided by the appropriate form of the complex. Figure 21.20 shows that the basic factor involved in transcription consists of a core (of 5 subunits) associated with other subunits that have a kinase activity; this complex also includes a repair subunit. The kinase catalytic subunit that phosphorylates the CTD of RNA polymerase belongs to a group of kinases that are involved in cell cycle control (see Molecular Biology 6.29 Cell cycle and growth regulation). It is possible that this connection influences transcription in response to the stage of the cell cycle.

Figure 21.20 The TFIIH core may associate with a kinase at initiation and associate with a repair complex when damaged DNA is encountered.

The alternative complex consists of the core associated with a large group of proteins that are coded by repair genes. (The basic model for repair is shown in Figure 15.53.) The repair proteins include a subunit (XPC) that recognizes damaged DNA, which provides the coupling function that enables a template strand to be preferentially repaired when RNA polymerase becomes stalled at damaged DNA. Other proteins associated with the complex include endonucleases (XPG, XPF, ERCC1). Homologous proteins are found in the complexes in yeast (where they are often identified by rad mutations that are defective in repair) and in Man (where they are identified by mutations that cause diseases resulting from deficiencies in A connection between transcription and repair | SECTION 5.21.12 © 2004. Virtual Text / www.ergito.com

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repairing damaged DNA) (662; 663). (Subunits with the name XP are coded by genes in which mutations cause the disease xeroderma pigmentosum (see Molecular Biology 4.15.28 Eukaryotic cells have conserved repair systems). The kinase complex and the repair complex can associate and dissociate reversibly from the core TFIIH. This suggests a model in which the first form of TFIIH is required for initiation, but may be replaced by the other form (perhaps in response to encountering DNA damage). TFIIH dissociates from RNA polymerase at an early stage of elongation (after transcription of ~50 bp); its reassociation at a site of damaged DNA may require additional coupling components. The repair function may require modification or degradation of RNA polymerase. The large subunit of RNA polymerase is degraded when the enzyme stalls at sites of UV damage. We do not yet understand the connection between the transcription/repair apparatus as such and the degradation of RNA polymerase. It is possible that removal of the polymerase is necessary once it has become stalled (664). This degradation of RNA polymerase is deficient in cells from patients with Cockayne's syndrome (a repair disorder). Cockayne's syndrome is caused by mutations in either of two genes (CSA and CSB), both of whose products appear to be part of or bound to TFIIH. Cockayne's syndrome is also occasionally caused by mutations in XPD. Another disease that can be caused by mutations in XPD is trichothiodystrophy, which has little in common with XP or Cockayne's (it involves mental retardation and is marked by changes in the structure of hair). All of this marks XPD as a pleiotropic protein, in which different mutations can affect different functions. In fact, XPD is required for the stability of the TFIIH complex during transcription, but the helicase activity as such is not needed. Mutations that prevent XPD from stabilizing the complex cause trichothiodystrophy. The helicase activity is required for the repair function. Mutations that affect the helicase activity cause the repair deficiency that results in XP or Cockayne's syndrome (for review see 1641). Last updated on 4-30-2001


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Reviews 224. Selby, C. P. and Sancar, A. (1994). Mechanisms of transcription-repair coupling and mutation frequency decline. Microbiol. Rev. 58, 317-329. 1641. Lehmann, A. R. (2001). The xeroderma pigmentosum group D (XPD) gene: one gene, two functions, three diseases. Genes Dev. 15, 15-23.

References 661. Selby, C. P. and Sancar, A. (1993). Molecular mechanism of transcription-repair coupling. Science 260, 53-58. 662. Schaeffer, L. et al. (1993). DNA repair helicase: a component of BTF2 (TFIIH) basic transcription factor. Science 260, 58-63. 663. Svejstrup, J. Q. et al. (1995). Different forms of TFIIH for transcription and DNA repair: holo-TFIIH and a nucleotide excision repairosome. Cell 80, 21-28. 664. Bregman, D. et al. (1996). UV-induced ubiquitination of RNA polymerase II: a novel modification deficient in Cockayne syndrome cells. Proc. Natl. Acad. Sci. USA 93, 11586-11590.



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5.21.13 Short sequence elements bind activators Key Terms An activator is a protein that stimulates the expression of a gene, typically by acting at a promoter to stimulate RNA polymerase. In eukaryotes, the sequence to which it binds in the promoter is called a response element. A CAAT box is part of a conserved sequence located upstream of the startpoints of eukaryotic transcription units; it is recognized by a large group of transcription factors. The GC box is a common pol II promoter element consisting of the sequence GGGCGG. Key Concepts

• Short conserved sequence elements are dispersed in the region preceding the startpoint.

• The upstream elements increase the frequency of initiation. • The factors that bind to them to stimulate transcription are called activators.

A promoter for RNA polymerase II consists of two types of region. The startpoint itself is identified by the Inr and/or by the TATA box close by. In conjunction with the general transcription factors, RNA polymerase II forms an initiation complex surrounding the startpoint, as we have just described. The efficiency and specificity with which a promoter is recognized, however, depend upon short sequences, farther upstream, which are recognized by a different group of factors, usually called activators. Usually the target sequences are ~100 bp upstream of the startpoint, but sometimes they are more distant. Binding of activators at these sites may influence the formation of the initiation complex at (probably) any one of several stages. An analysis of a typical promoter is summarized in Figure 21.21. Individual base substitutions were introduced at almost every position in the 100 bp upstream of the β-globin startpoint. The striking result is that most mutations do not affect the ability of the promoter to initiate transcription. Down mutations occur in three locations, corresponding to three short discrete elements. The two upstream elements have a greater effect on the level of transcription than the element closest to the startpoint. Up mutations occur in only one of the elements. We conclude that the three short sequences centered at –30, –75, and –90 constitute the promoter. Each of them corresponds to the consensus sequence for a common type of promoter element.

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Figure 21.21 Saturation mutagenesis of the upstream region of the β -globin promoter identifies three short regions (centered at -30, -75, and -90) that are needed to initiate transcription. These correspond to the TATA, CAAT, and GC boxes.

The TATA box (centered at –30) is the least effective component of the promoter as measured by the reduction in transcription that is caused by mutations. But although initiation is not prevented when a TATA box is mutated, the startpoint varies from its usual precise location. This confirms the role of the TATA box as a crucial positioning component of the core promoter. The basal elements and the elements upstream of them have different types of functions. The basal elements (the TATA box and Inr) primarily determine the location of the startpoint, but can sponsor initiation only at a rather low level. They identify the location at which the general transcription factors assemble to form the basal complex. The sequence elements farther upstream influence the frequency of initiation, most likely by acting directly on the general transcription factors to enhance the efficiency of assembly into an initiation complex (see Molecular Biology 5.22.5 Activators interact with the basal apparatus). The sequence at –75 is the CAAT box. Named for its consensus sequence, it was one of the first common elements to be described. It is often located close to –80, but it can function at distances that vary considerably from the startpoint. It functions in either orientation. Susceptibility to mutations suggests that the CAAT box plays a strong role in determining the efficiency of the promoter, but does not influence its specificity. The GC box at –90 contains the sequence GGGCGG. Often multiple copies are present in the promoter, and they occur in either orientation. It too is a relatively common promoter component. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.21.13

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5.21.14 Promoter construction is flexible but context can be important Key Concepts

• No individual upstream element is essential for promoter function, although one or more elements must be present for efficient initiation.

• Some elements are recognized by multiple factors, and the factor that is used at any particular promoter may be determined by the context of the other factors that are bound.

Promoters are organized on a principle of "mix and match." A variety of elements can contribute to promoter function, but none is essential for all promoters. Some examples are summarized in Figure 21.22. Four types of elements are found altogether in these promoters: TATA, GC boxes, CAAT boxes, and the octamer (an 8 bp element). The elements found in any individual promoter differ in number, location, and orientation. No element is common to all of the promoters. Although the promoter conveys directional information (transcription proceeds only in the downstream direction), the GC and CAAT boxes seem to be able to function in either orientation. This implies that the elements function solely as DNA-binding sites to bring transcription factors into the vicinity of the startpoint; the structure of a factor must be flexible enough to allow it to make protein-protein contacts with the basal apparatus irrespective of the way in which its DNA-binding domain is oriented and its exact distance from the startpoint.

Figure 21.22 Promoters contain different combinations of TATA boxes, CAAT boxes, GC boxes, and other elements.

Activators that are more or less ubiquitous are assumed to be available to any promoter that has a copy of the element that they recognize. Common elements Promoter construction is flexible but context can be important | SECTION 5.21.14 © 2004. Virtual Text / www.ergito.com

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recognized by ubiquitous activators include the CAAT box, GC box, and the octamer. All promoters probably require one or more of these elements in order to function efficiently. An activator typically has a consensus sequence of 40 proteins. Is it feasible for this apparatus to assemble step by step at the promoter? Some activators, coactivators, and basal factors may assemble stepwise at the promoter, but then may be joined by a very large complex consisting of RNA polymerase preassembled with further activators and coactivators, as illustrated in Figure 22.7 (for review see 1710).

Activators interact with the basal apparatus | SECTION 5.22.5 © 2004. Virtual Text / www.ergito.com

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Figure 22.7 RNA polymerase exists as a holoenzyme containing many activators.

Several forms of RNA polymerase have been found in which the enzyme is associated with various transcription factors. The most prominent "holoenzyme complex" in yeast (defined as being capable of initiating transcription without additional components) consists of RNA polymerase associated with a 20-subunit complex called mediator (1713; for review see 1711). The mediator includes products of several genes in which mutations block transcription, including some SRB loci (so named because many of their genes were originally identified as suppressors of mutations in RNA polymerase B.) The name was suggested by its ability to mediate the effects of activators. Mediator is necessary for transcription of most yeast genes. Homologous complexes are required for the transcription of most higher eukaryotic genes (2411). Mediator undergoes a conformational change when it interacts with the CTD domain of RNA polymerase (2412). It can transmit either activating or repressing effects from upstream components to the RNA polymerase. It is probably released when a polymerase starts elongation. Some transcription factors influence transcription directly by interacting with RNA polymerase or the basal apparatus, but others work by manipulating structure of chromatin (see Molecular Biology 5.23.3 Chromatin remodeling is an active process). Last updated on 4-22-2002


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Reviews 218. Maniatis, T., Goodbourn, S., and Fischer, J. A. (1987). Regulation of inducible and tissue-specific gene expression. Science 236, 1237-1245. 221. Mitchell, P., and Tjian, R. (1989). Transcriptional regulation in mammalian cells by sequence-specific DNA-binding proteins. Science 245, 371-378. 1710. Lemon, B. and Tjian, R. (2000). Orchestrated response: a symphony of transcription factors for gene control. Genes Dev. 14, 2551-2569. 1711. Myers, L. C. and Kornberg, R. D. (2000). Mediator of transcriptional regulation. Annu. Rev. Biochem. 69, 729-749.

References 645. Pugh, B. F. and Tjian, R. (1990). Mechanism of transcriptional activation by Sp1: evidence for coactivators. Cell 61, 1187-1197. 646. Dynlacht, B. D., Hoey, T., and Tjian, R. (1991). Isolation of coactivators associated with the TATA-binding protein that mediate transcriptional activation. Cell 66, 563-576. 658. Ma, J. and Ptashne, M. (1987). A new class of yeast transcriptional activators. Cell 51, 113-119. 659. Chen, J.-L. et al. (1994). Assembly of recombinant TFIID reveals differential coactivator requirements for distinct transcriptional activators. Cell 79, 93-105. 1713. 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. 2411. Asturias, F. J., Jiang, Y. W., Myers, L. C., Gustafsson, C. M., and Kornberg, R. D. (1999). Conserved structures of mediator and RNA polymerase II holoenzyme. Science 283, 985-987. 2412. Dotson, M. R., Yuan, C. X., Roeder, R. G., Myers, L. C., Gustafsson, C. M., Jiang, Y. W., Li, Y., Kornberg, R. D., and Asturias, F. J. (2000). Structural organization of yeast and mammalian mediator complexes. Proc. Natl. Acad. Sci. USA 97, 14307-14310.



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ACTIVATING TRANSCRIPTION

5.22.6 Some promoter-binding proteins are repressors Key Concepts

• Repression is usually achieved by affecting chromatin structure, but there are repressors that act by binding to specific promoters.

Repression of transcription in eukaryotes is generally accomplished at the level of influencing chromatin structure; regulator proteins that function like trans-acting bacterial repressors to block transcription are relatively rare, but some examples are known. One case is the global repressor NC2/Dr1/DRAP1, a heterodimer that binds to TBP to prevent it from interacting with other components of the basal apparatus (1741; 1742; 1743). The importance of this interaction is suggested by the lethality of null mutations in the genes that code for the repressor in yeast. Repressors that work in this way have an active role in inhibiting basal apparatus function. In a more specific case, the CAAT sequence is a target for regulation. Two copies of this element are found in the promoter of a gene for histone H2B (see Figure 21.22) that is expressed only during spermatogenesis in a sea urchin. CAAT-binding factors can be extracted from testis tissue and also from embryonic tissues, but only the former can bind to the CAAT box. In the embryonic tissues, another protein, called the CAAT-displacement protein (CDP), binds to the CAAT boxes, preventing the activator from recognizing them. Figure 22.10 illustrates the consequences for gene expression. In testis, the promoter is bound by transcription factors at the TATA box, CAAT boxes, and octamer sequences. In embryonic tissue, the exclusion of the CAAT-binding factor from the promoter prevents a transcription complex from being assembled. The analogy with the effect of a bacterial repressor in preventing RNA polymerase from initiating at the promoter is obvious. These results also make the point that the function of a protein in binding to a known promoter element cannot be assumed: it may be an activator, a repressor, or even irrelevant to gene transcription.

Some promoter-binding proteins are repressors | SECTION 5.22.6 © 2004. Virtual Text / www.ergito.com

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Figure 22.10 A transcription complex involves recognition of several elements in the sea urchin H2B promoter in testis. Binding of the CAAT displacement factor in embryo prevents the CAAT-binding factor from binding, so an active complex cannot form.


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References 1741. Inostroza, J. A., Mermelstein, F. H., Ha, I., Lane, W. S., and Reinberg, D. (1992). Dr1, a TATA-binding protein-associated phosphoprotein and inhibitor of class II gene transcription. Cell 70, 477-489. 1742. Goppelt, A., Stelzer, G., Lottspeich, F., and Meisterernst, M. (1996). A mechanism for repression of class II gene transcription through specific binding of NC2 to TBP-promoter complexes via heterodimeric histone fold domains. EMBO J. 15, 3105-3116. 1743. Kim, T. K., Kim, T. K., Zhao, Y., Ge, H., Bernstein, R., and Roeder, R. G. (1995). TATA-binding protein residues implicated in a functional interplay between negative cofactor NC2 (Dr1) and general factors TFIIA and TFIIB. J. Biol. Chem. 270, 10976-10981.



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5.22.7 Response elements are recognized by activators Key Terms A response element is a sequence in a eukaryotic promoter or enhancer that is recognized by a specific transcription factor. The heat shock response element (HSE) is a sequence in a promoter or enhancer that is used to activate a gene by an activator induced by heat shock. The glucocorticoid response element (GRE) is a sequence in a promoter or enhancer that is recognized by the glucocorticoid receptor ,which is activated by glucocorticoid steroids. The serum response element (SRE) is a sequence in a promoter or enhancer that is activated by transcription factor(s) induced by treatment with serum. This activates genes that stimulate cell growth. Heat shock genes are a set of loci that are activated in response to an increase in temperature (and other abuses to the cell). They occur in all organisms. They usually include chaperones that act on denatured proteins. Key Concepts

• Response elements may be located in promoters or enhancers. • Each response element is recognized by a specific activator. • A promoter may have many response elements, which may activate transcription independently or in certain combinations.

The principle that emerges from characterizing groups of genes under common control is that they share a promoter (or enhancer) element that is recognized by an activator. An element that causes a gene to respond to such a factor is called a response element; examples are the HSE (heat shock response element), GRE (glucocorticoid response element), SRE (serum response element). Response elements contain short consensus sequences; copies of the response elements found in different genes are closely related, but not necessarily identical. The region bound by the factor extends for a short distance on either side of the consensus sequence. In promoters, the elements are not present at fixed distances from the startpoint, but are usually 30 steroids, the two major groups being the glucocorticoids and mineralocorticoids. Steroids provide the reproductive hormones (androgen male sex hormones and estrogen female sex hormones). Vitamin D is required for bone development. Other hormones, with unrelated structures and physiological purposes, function at the molecular level in a similar way to the steroid hormones. Thyroid hormones, based on iodinated forms of tyrosine, control basal metabolic rate in animals. Steroid and thyroid hormones also may be important in metamorphosis (ecdysteroids in insects, and thyroid hormones in frogs).

Steroid receptors are activators | SECTION 5.22.10 © 2004. Virtual Text / www.ergito.com

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Retinoic acid (vitamin A) is a morphogen responsible for development of the anterior-posterior axis in the developing chick limb bud. Its metabolite, 9-cis retinoic acid, is found in tissues that are major sites for storage and metabolism of vitamin A. We may account for these various actions in terms of pathways for regulating gene expression. These diverse compounds share a common mode of action: each is a small molecule that binds to a specific receptor that activates gene transcription. ("Receptor" may be a misnomer: the protein is a receptor for steroid or thyroid hormone in the same sense that lac repressor is a receptor for a β-galactoside: it is not a receptor in the sense of comprising a membrane-bound protein that is exposed to the cell surface.) Receptors for the diverse groups of steroid hormones, thyroid hormones, and retinoic acid represent a new "superfamily" of gene regulators, the ligand-responsive activators. All the receptors have independent domains for DNA-binding and hormone binding, in the same relative locations. Their general organization is summarized in Figure 22.16 (for review see 231).

Figure 22.16 Receptors for many steroid and thyroid hormones have a similar organization, with an individual N-terminal region, conserved DNA-binding region, and a C-terminal hormone-binding region.

The central part of the protein is the DNA-binding domain. These regions are closely related for the various steroid receptors (from the most closely related pair with 94% sequence identity to the least well related pair at 42% identity). The act of binding DNA cannot be disconnected from the ability to activate transcription, because mutations in this domain affect both activities. The N-terminal regions of the receptors show the least conservation of sequence. They include other regions that are needed to activate transcription. Steroid receptors are activators | SECTION 5.22.10 © 2004. Virtual Text / www.ergito.com

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The C-terminal domains bind the hormones. Those in the steroid receptor family show identities ranging from 30-57%, reflecting specificity for individual hormones. Their relationships with the other receptors are minimal, reflecting specificity for a variety of compounds – thyroid hormones, vitamin D, retinoic acid, etc. This domain also has the motifs responsible for dimerization and a region involved in transcriptional activation (for review see 1436). Some ligands have multiple receptors that are closely related, such as the 3 retinoic acid receptors (RAR α , β, γ) and the three receptors for 9-cis-retinoic acid (RXR α , β, γ). Last updated on 2-12-2001


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Reviews 231. Evans, R. M. (1988). The steroid and thyroid hormone receptor superfamily. Science 240, 889-895. 1436. Mangelsdorf, D. J. and Evans, R. (1995). The RXR heterodimers and orphan receptors. Cell 83, 841-850.



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5.22.11 Steroid receptors have zinc fingers Key Concepts

• The DNA binding domain of a steroid receptor is a type of zinc finger that has Cys but not His residues.

• Glucocorticoid and estrogen receptors each have two zinc fingers, the first of which determines the DNA target sequence.

• Steroid receptors bind to DNA as dimers.

Steroid receptors (and some other proteins) have another type of zinc finger that is different from Cys2/His2 fingers. The structure is based on a sequence with the zinc-binding consensus: Cys-X2-Cys-X13-Cys-X2-Cys These are called Cys2/Cys2 fingers. Proteins with Cys2/Cys2 fingers often have nonrepetitive fingers, in contrast with the tandem repetition of the Cys2/His2 type. Binding sites in DNA (where known) are short and palindromic. The glucocorticoid and estrogen receptors each have two fingers, each with a zinc atom at the center of a tetrahedron of cysteines. The two fingers form α-helices that fold together to form a large globular domain. The aromatic sides of the α-helices form a hydrophobic center together with a β-sheet that connects the two helices. One side of the N-terminal helix makes contacts in the major groove of DNA. Two glucocorticoid receptors dimerize upon binding to DNA, and each engages a successive turn of the major groove. This fits with the palindromic nature of the response element (see Molecular Biology 5.22.13 Steroid receptors recognize response elements by a combinatorial code). Each finger controls one important property of the receptor. Figure 22.18 identifies the relevant amino acids. Those on the right side of the first finger determine the sequence of the target in DNA; those on the left side of the second finger control the spacing between the target sites recognized by each subunit in the dimer (see Molecular Biology 5.22.13 Steroid receptors recognize response elements by a combinatorial code).

Steroid receptors have zinc fingers | SECTION 5.22.11 © 2004. Virtual Text / www.ergito.com

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Figure 22.18 The first finger of a steroid receptor controls which DNA sequence is bound (positions shown in red); the second finger controls spacing between the sequences (positions shown in blue).

Direct evidence that the first finger binds DNA was obtained by a "specificity swap" experiment. The finger of the estrogen receptor was deleted and replaced by the sequence of the glucocorticoid receptor. The new protein recognized the GRE sequence (the usual target of the glucocorticoid receptor) instead of the ERE (the usual target of the estrogen receptor). This region therefore establishes the specificity with which DNA is recognized. The differences between the sequences of the glucocorticoid receptor and estrogen receptor fingers lie mostly at the base of the finger. The substitution at two positions shown in Figure 22.17 allows the glucocorticoid receptor to bind at an ERE instead of a GRE (1768; for review see 240).

Figure 22.17 Discrimination between GRE and ERE target sequences is determined by two amino acids at the base of the first zinc finger in the receptor.


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Reviews 240. Tsai, M J, and O'Malley, B W (1994). Molecular mechanisms of action of steroid/thyroid receptor superfamily members. Annu. Rev. Biochem. 63, 451-486.

References 1768. Umesono, K. and Evans, R. M. (1989). Determinants of target gene specificity for steroid/thyroid hormone receptors. Cell 57, 1139-1146.



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5.22.12 Binding to the response element is activated by ligand-binding Key Concepts

• Binding of ligand to the C-terminal domain increases the affinity of the DNA-binding domain for its specific target site in DNA.

We know most about the interaction of glucocorticoids with their receptor, whose action is illustrated in Figure 22.19. A steroid hormone can pass through the cell membrane to enter the cell by simple diffusion. Within the cell, a glucocorticoid binds the glucocorticoid receptor. (Work on the glucocorticoid receptor has relied on the synthetic steroid hormone, dexamethasone.) The localization of free receptors is not entirely clear; they may be in equilibrium between the nucleus and cytoplasm. But when hormone binds to the receptor, the protein is converted into an activated form that has an increased affinity for DNA, so the hormone-receptor complex is always localized in the nucleus.

Figure 22.19 Glucocorticoids regulate gene transcription by causing their receptor to bind to an enhancer whose action is needed for promoter function.

The activated receptor recognizes a specific consensus sequence that identifies the GRE, the glucocorticoid response element. The GRE is typically located in an enhancer that may be several kb upstream or downstream of the promoter. When the steroid-receptor complex binds to the enhancer, the nearby promoter is activated, and transcription initiates there. Enhancer activation provides the general mechanism by which steroids regulate a wide set of target genes. Binding to the response element is activated by ligand-binding | SECTION 5.22.12 © 2004. Virtual Text / www.ergito.com

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The C-terminal region regulates the activity of the receptor in a way that varies for the individual receptor. If the C-terminal domain of the glucocorticoid receptor is deleted, the remaining N-terminal protein is constitutively active: it no longer requires steroids for activity. This suggests that, in the absence of steroid, the steroid-binding domain prevents the receptor from recognizing the GRE; it functions as an internal negative regulator. The addition of steroid inactivates the inhibition, releasing the receptor's ability to bind the GRE and activate transcription. The basis for the repression could be internal, relying on interactions with another part of the receptor. Or it could result from an interaction with some other protein, which is displaced when steroid binds. The interaction between the domains is different in the estrogen receptor. If the hormone-binding domain is deleted, the protein is unable to activate transcription, although it continues to bind to the ERE. This region is therefore required to activate rather than to repress activity. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.22.12

Binding to the response element is activated by ligand-binding | SECTION 5.22.12 © 2004. Virtual Text / www.ergito.com

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5.22.13 Steroid receptors recognize response elements by a combinatorial code Key Concepts

• A steroid response element consists of two short half sites that may be palindromic or directly repeated.

• There are only two types of half sites. • A receptor recognizes its response element by the orientation and spacing of the half sites.

• The sequence of the half site is recognized by the first zinc finger. • The second zinc finger is responsible for dimerization, which determines the distance between the subunits.

• Subunit separation in the receptor determines the recognition of spacing in the response element.

• Some steroid receptors function as homodimers but others form heterodimers. • Homodimers recognize palindromic response elements; heterodimers recognize response elements with directly repeated half sites.

Each receptor recognizes a response element that consists of two short repeats (or half sites). This immediately suggests that the receptor binds as a dimer, so that each half of the consensus is contacted by one subunit (reminiscent of the λ operator-repressor interaction described in Molecular Biology 3.12.12 Repressor uses a helix-turn-helix motif to bind DNA). The half sites may be arranged either as palindromes or as repeats in the same orientation. They are separated by 0-4 base pairs whose sequence is irrelevant. Only two types of half site are used by the various receptors. Their orientation and spacing determine which receptor recognizes the response element. This behavior allows response elements that have restricted consensus sequences to be recognized specifically by a variety of receptors. The rules that govern recognition are not absolute, but may be modified by context, and there are also cases in which palindromic response elements are recognized permissively by more than one receptor (for review see 229). The receptors fall into two groups: • Glucocorticoid (GR), mineralocorticoid (MR), androgen (AR), and progesterone (PR) receptors all form homodimers. They recognize response elements whose half sites have the consensus sequence TGTTCT. Figure 22.20 shows that the Steroid receptors recognize response elements by a combinatorial code | SECTION 5.22.13 © 2004. Virtual Text / www.ergito.com

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half sites are arranged as palindromes, and the spacing between the sites determines the type of element. The estrogen (ER) receptor functions in the same way, but has the half site sequence TGACCT. • The 9-cis-retinoic acid (RXR) receptor forms homodimers and also forms heterodimers with ~15 other receptors, including thyroid (T3R), vitamin D (VDR), and retinoic acid (RAR) (1436). Figure 22.21 shows that the dimers recognize half elements with the sequence TGACCT. The half sites are arranged as direct repeats, and recognition is controlled by spacing between them (1769). Some of the heterodimeric receptors are activated when the ligand binds to the partner for RXR; others can be activated by ligand binding either to this subunit or to the RXR subunit. These receptors can also form homodimers, which recognize palindromic sequences.

Figure 22.20 Response elements formed from the palindromic half site TGTTCT are recognized by several different receptors depending on the spacing between the half sites.

Figure 22.21 Response elements with the direct repeat TGACCT are recognized by heterodimers of which one member is RXR.

Now we are in a position to understand the basis for specificity of recognition. Recall that Figure 22.18 shows how recognition of the sequence of the half site is conferred by the amino acid sequence in the first finger. Specificity for the spacing between half sites is carried by amino acids in the second finger. The structure of the dimer determines the distance between the subunits that sit in successive turns of the major groove, and thus controls the response to the spacing of half sites (679). The exact positions of the residues responsible for dimerization differ in individual pairwise combinations. Steroid receptors recognize response elements by a combinatorial code | SECTION 5.22.13 © 2004. Virtual Text / www.ergito.com

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How do the steroid receptors activate transcription? They do not act directly on the basal apparatus, but function via a coactivating complex. The coactivator includes various activities, including the common component CBP/p300, one of whose functions is to modify the structure of chromatin by acetylating histones (see Figure 23.13). All receptors in the superfamily are ligand-dependent activators of transcription. However, some are also able to repress transcription. The TR and RAR receptors, in the form of heterodimers with RXR, bind to certain loci in the absence of ligand and repress transcription by means of their ability to interact with a corepressor protein. The corepressor functions by the reverse of the mechanism used by coactivators: it inhibits the function of the basal transcription apparatus, one of its actions being the deacetylation of histones (see Figure 23.15). We do not know the relative importance of the repressor activity vis-À-vis the ligand-dependent activation in the physiological response to hormone (660). The effect of ligand binding on the receptor is to convert it from a repressing complex to an activating complex, as shown in Figure 22.22. In the absence of ligand, the receptor is bound to a corepressor complex. The component of the corepressor that binds to the receptor is SMRT. Binding of ligand causes a conformational change that displaces SMRT. This allows the coactivator to bind.

Figure 22.22 TR and RAR bind the SMRT corepressor in the absence of ligand. The promoter is not expressed. When SMRT is displaced by binding of ligand, the receptor binds a coactivator complex. This leads to activation of transcription by the basal apparatus.

Steroid receptors recognize response elements by a combinatorial code | SECTION 5.22.13 © 2004. Virtual Text / www.ergito.com

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Reviews 229. Yamamoto, K. R. (1985). Steroid receptor regulated transcription of specific genes and gene networks. Annu. Rev. Genet. 19, 209-252. 1436. Mangelsdorf, D. J. and Evans, R. (1995). The RXR heterodimers and orphan receptors. Cell 83, 841-850.

References 660. Hurlein, A. J. et al. (1995). Ligand-independent repression by the thyroid hormone receptor mediated by a nuclear receptor corepressor. Nature 377, 397-404. 679. Rastinejad, F., Perlmann, T., Evans, R. M., and Sigler, P. B. (1995). Structural determinants of nuclear receptor assembly on DNA direct repeats. Nature 375, 203-211. 1769. Umesono, K., Murakami, K. K., Thompson, C. C., and Evans, R. M. (1991). Direct repeats as selective response elements for the thyroid hormone, retinoic acid, and vitamin D3 receptors. Cell 65, 1255-1266.


Steroid receptors recognize response elements by a combinatorial code | SECTION 5.22.13 © 2004. Virtual Text / www.ergito.com

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5.22.14 Homeodomains bind related targets in DNA Key Concepts

• The homeodomain is a DNA-binding domain of 60 amino acids that has three α-helices.

• The C-terminal α-helix-3 is 17 amino acids and binds in the major groove of DNA. • The N-terminal arm of the homeodomain projects into the minor groove of DNA. • Proteins containing homeodomains may be either activators or repressors of transcription.

The homeobox is a sequence that codes for a domain of 60 amino acids present in proteins of many or even all eukaryotes. Its name derives from its original identification in Drosophila homeotic loci (whose genes determine the identity of body structures). It is present in many of the genes that regulate early development in Drosophila, and a related motif is found in genes in a wide range of higher eukaryotes. The homeodomain is found in many genes concerned with developmental regulation (see Molecular Biology 6.31.22 The homeobox is a common coding motif in homeotic genes). Sequences related to the homeodomain are found in several types of animal transcription factors. In Drosophila homeotic genes, the homeodomain often (but not always) occurs close to the C-terminal end. Some examples of genes containing homeoboxes are summarized in Figure 22.23. Often the genes have little conservation of sequence except in the homeobox. The conservation of the homeobox sequence varies. A major group of homeobox-containing genes in Drosophila has a well conserved sequence, with 80-90% similarity in pairwise comparisons. Other genes have less closely related homeoboxes. The homeodomain is sometimes combined with other motifs in animal transcription factors. One example is presented by the Oct (octamer-binding) proteins, in which a conserved stretch of 75 amino acids called the Pou region is located close to a region resembling the homeodomain. The homeoboxes of the Pou group of proteins are the least closely related to the original group, and thus comprise the farthest extension of the family.

Homeodomains bind related targets in DNA | SECTION 5.22.14 © 2004. Virtual Text / www.ergito.com

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Figure 22.23 The homeodomain may be the sole DNA-binding motif in a transcriptional regulator or may be combined with other motifs. It represents a discrete (60 residue) part of the protein.

The homeodomain is responsible for binding to DNA, and experiments to swap homeodomains between proteins suggest that the specificity of DNA recognition lies within the homeodomain, but (like the situation with phage repressors) no simple code relating protein and DNA sequences can be deduced. The C-terminal region of the homeodomain shows homology with the helix-turn-helix motif of prokaryotic repressors. We recall from Molecular Biology 3.12.12 Repressor uses a helix-turn-helix motif to bind DNA that the λ repressor has a "recognition helix" ( α-helix-3) that makes contacts in the major groove of DNA, while the other helix ( α-helix-2) lies at an angle across the DNA. The homeodomain can be organized into three potential helical regions; the sequences of three examples are compared in Figure 22.24. The best conserved part of the sequence lies in the third helix. The difference between these structures and the prokaryotic repressor structures lies in the length of the helix that recognizes DNA, helix-3, which is 17 amino acids long in the homeodomain, compared to 9 residues long in the λ repressor.


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Figure 22.24 The homeodomain of the Antennapedia gene represents the major group of genes containing homeoboxes in Drosophila; engrailed (en) represents another type of homeotic gene; and the mammalian factor Oct-2 represents a distantly related group of transcription factors. The homeodomain is conventionally numbered from 1 to 60. It starts with the N-terminal arm, and the three helical regions occupy residues 10-22, 28-38, and 42-58. Amino acids in red are conserved in all three examples.

The structure of the homeodomain of the D. melanogaster Engrailed protein is represented schematically in Figure 22.25. Helix 3 binds in the major groove of DNA and makes the majority of the contacts between protein and nucleic acid. Many of the contacts that orient the helix in the major groove are made with the phosphate backbone, so they are not specific for DNA sequence. They lie largely on one face of the double helix, and flank the bases with which specific contacts are made. The remaining contacts are made by the N-terminal arm of the homeodomain, the sequence that just precedes the first helix. It projects into the minor groove. So the N-terminal and C-terminal regions of the homeodomain are primarily responsible for contacting DNA (678).

Figure 22.25 Helix 3 of the homeodomain binds in the major groove of DNA, with helices 1 and 2 lying outside the double helix. Helix 3 contacts both the phosphate backbone and specific bases. The N-terminal arm lies in the minor groove, and makes additional contacts. Homeodomains bind related targets in DNA | SECTION 5.22.14 © 2004. Virtual Text / www.ergito.com

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A striking demonstration of the generality of this model derives from a comparison of the crystal structure of the homeodomain of engrailed with that of the α2 mating protein of yeast. The DNA-binding domain of this protein resembles a homeodomain, and can form three similar helices: its structure in the DNA groove can be superimposed almost exactly on that of the engrailed homeodomain. These similarities suggest that all homeodomains bind to DNA in the same manner. This means that a relatively small number of residues in helix-3 and in the N-terminal arm are responsible for specificity of contacts with DNA (for review see 239). One group of homeodomain-containing proteins is the set of Hox proteins (see Figure 31.39). They bind to DNA with rather low sequence specificity, and it has been puzzling how these proteins can have different specificities. It turns out that Hox proteins often bind to DNA as heterodimers with a partner (called Exd in flies and Pbx in vertebrates). The heterodimer has a more restricted specificity in vitro than an individual Hox protein; typically it binds the 10 bp sequence TGATNNATNN. Still this is not enough to account for the differences in the specificities of Hox proteins. A third protein, Hth, which is necessary to localize Exd in the nucleus, also forms part of the complex that binds DNA, and may restrict the binding sites further. But since the same partners (Exd and Hth) are present together with each Hox protein in the trimeric complex, it remains puzzling how each Hox protein has sufficient specificity. Homeodomain proteins can be either transcriptional activators or repressors. The nature of the factor depends on the other domain(s) – the homeodomain is responsible solely for binding to DNA. The activator or repressor domains both act by influencing the basal apparatus. Activator domains may interact with coactivators that in turn bind to components of the basal apparatus. Repressor domains also interact with the transcription apparatus (that is, they do not act by blocking access to DNA as such). The repressor Eve, for example, interacts directly with TFIID (674).


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Reviews 239. Gehring, W. J. et al. (1994). Homeodomain-DNA recognition. Cell 78, 211-223.

References 674. Han, K., Levine, M. S., and Manley, J. L. (1989). Synergistic activation and repression of transcription by Drosophila homeobox proteins. Cell 56, 573-583. 678. Wolberger, C. et al. (1991). Crystal structure of a MAT α 2 homeodomain-operator complex suggests a general model for homeodomain-DNA interactions. Cell 67, 517-528.



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5.22.15 Helix-loop-helix proteins interact by combinatorial association Key Terms The helix-loop-helix (HLH) motif is responsible for dimerization of a class of transcription factors called HLH proteins. A bHLH protein has a basic DNA-binding sequence close to the dimerization motif. A bHLH protein has a basic DNA-binding region adjacent to the helix-loop-helix motif. Key Concepts

• Helix-loop-helix proteins have a motif of 40-50 amino acids that comprises two amphipathic α-helices of 15-16 residues separated by a loop.

• The helices are responsible for dimer formation. • bHLH proteins have a basic sequence adjacent to the HLH motif that is responsible for binding to DNA.

• Class A bHLH proteins are ubiquitously expressed. Class B bHLH proteins are tissue specific.

• A class B protein usually forms a heterodimer with a class A protein. • HLH proteins that lack the basic region prevent a bHLH partner in a heterodimer from binding to DNA.

• HLH proteins form combinatorial associations that may be changed during development by the addition or removal of specific proteins.

Two common features in DNA-binding proteins are the presence of helical regions that bind DNA, and the ability of the protein to dimerize. Both features are represented in the group of helix-loop-helix proteins that share a common type of sequence motif: a stretch of 40-50 amino acids contains two amphipathic α-helices separated by a linker region (the loop) of varying length. (An amphipathic helix forms two faces, one presenting hydrophobic amino acids, the other presenting charged amino acids.) The proteins in this group form both homodimers and heterodimers by means of interactions between the hydrophobic residues on the corresponding faces of the two helices (675). The helical regions are 15-16 amino acids long, and each contains several conserved residues. Two examples are compared in Figure 22.26. The ability to form dimers resides with these amphipathic helices, and is common to all HLH proteins. The loop is probably important only for allowing the freedom for the two helical regions to interact independently of one another.

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Figure 22.26 All HLH proteins have regions corresponding to helix 1 and helix 2, separated by a loop of 10-24 residues. Basic HLH proteins have a region with conserved positive charges immediately adjacent to helix 1.

Most HLH proteins contain a region adjacent to the HLH motif itself that is highly basic, and which is needed for binding to DNA. There are ~6 conserved residues in a stretch of 15 amino acids (see Figure 22.26). Members of the group with such a region are called bHLH proteins. A dimer in which both subunits have the basic region can bind to DNA. The HLH domains probably correctly orient the two basic regions contributed by the individual subunits. The bHLH proteins fall into two general groups. Class A consists of proteins that are ubiquitously expressed, including mammalian E12/E47. Class B consists of proteins that are expressed in a tissue-specific manner, including mammalian MyoD, myogenin, and Myf-5 ( a group of activators that are involved in myogenesis [muscle formation]). A common modus operandi for a tissue-specific bHLH protein is to form a heterodimer with a ubiquitous partner. There is also a group of gene products that specify development of the nervous system in D. melanogaster (where Ac-S is the tissue-specific component and da is the ubiquitous component). The Myc proteins (which are the cellular counterparts of oncogene products and are involved in growth regulation) form a separate class of bHLH proteins, whose partners and targets are different. Dimers formed from bHLH proteins differ in their abilities to bind to DNA. For example, E47 homodimers, E12-E47 heterodimers, and MyoD-E47 heterodimers all form efficiently and bind strongly to DNA; E12 homodimerizes well but binds DNA poorly, while MyoD homodimerizes only poorly. So both dimer formation and DNA binding may represent important regulatory points. At this juncture, it is possible to define groups of HLH proteins whose members form various pairwise combinations, but not to predict from the sequences the strengths of dimer formation or DNA binding. All of the dimers in this group that bind DNA recognize the same consensus sequence, but we do not know yet whether different homodimers and heterodimers have preferences for slightly different target sites that are related to their functions. Differences in DNA-binding result from properties of the region in or close to the HLH motif; for example, E12 differs from E47 in possessing an inhibitory region just by the basic region, which prevents DNA binding by homodimers. Some HLH proteins lack the basic region and/or contain proline residues that appear to disrupt its function. The example of the protein Id is shown in Figure 22.26. Proteins of this type have the same capacity to dimerize as bHLH proteins, but a dimer that contains Helix-loop-helix proteins interact by combinatorial association | SECTION 5.22.15 © 2004. Virtual Text / www.ergito.com

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one subunit of this type can no longer bind to DNA specifically. This is a forceful demonstration of the importance of doubling the DNA-binding motif in DNA-binding proteins (670; 671; 675). The importance of the distinction between the nonbasic HLH and bHLH proteins is suggested by the properties of two pairs of HLH proteins: the da-Ac-S/emc pair and the MyoD/Id pair. A model for their functions in forming a regulatory network is illustrated in Figure 22.27.

Figure 22.27 An HLH dimer in which both subunits are of the bHLH type can bind DNA, but a dimer in which one subunit lacks the basic region cannot bind DNA.

In D. melanogaster, the gene emc (extramacrochaetae) is required to establish the normal spatial pattern of adult sensory organs. It functions by suppressing the functions of several genes, including da (daughterless) and the achaete-scute complex (Ac-S). Ac-S and da are genes of the bHLH type. The suppressor emc codes for an HLH protein that lacks the basic region. We suppose that, in the absence of emc function, the da and Ac-S proteins form dimers that activate transcription of appropriate target genes, but the production of emc protein causes the formation of heterodimers that cannot bind to DNA. So production of emc protein in the appropriate cells is necessary to suppress the function of Ac-S/da. The formation of muscle cells is triggered by a change in the transcriptional program that requires several bHLH proteins, including MyoD. MyoD is produced specifically in myogenic cells; and, indeed, overexpression of MyoD in certain other cells can induce them to commence a myogenic program. The trigger for muscle differentiation is probably a heterodimer consisting of MyoD-E12 or MyoD-E47, rather than a MyoD homodimer. Before myogenesis begins, a member of the nonbasic HLH type, the Id protein, may bind to MyoD and/or E12 and E47 to form heterodimers that cannot bind to DNA. It binds to E12/E47 better than to MyoD, and so might function by sequestering the ubiquitous bHLH partner. Overexpression of Id can prevent myogenesis. So the removal of Id could be the trigger that releases Helix-loop-helix proteins interact by combinatorial association | SECTION 5.22.15 © 2004. Virtual Text / www.ergito.com

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MyoD to initiate myogenesis (668; 669; for review see 236). A bHLH activator such as MyoD can be controlled in several ways. It is prevented from binding to DNA when it is sequestered by an HLH partner such as Id. It can activate transcription when bound to bHLH partner such as E12 or E47. It can also act as a site-specific repressor when bound to another partner; the bHLH protein MyoR forms a MyoD-MyoR dimer in proliferating myoblasts that represses transcription (at the same target loci at which MyoD-E12/E47 activate transcription). The behavior of the HLH proteins therefore illustrates two general principles of transcriptional regulation. A small number of proteins form combinatorial associations. Particular combinations have different functions with regard to DNA binding and transcriptional regulation. Differentiation may depend either on the presence or on the removal of particular partners.


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Reviews 236. Weintraub, H. (1991). The MyoD gene family: nodal point during specification of the muscle cell lineage. Science 251, 761-766.

References 668. Davis, R. L. et al. (1987). Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell 51, 987-1000. 669. Davis, R. L. et al. (1990). The MyoD DNA binding domain contains a recognition code for muscle-specific gene activation. Cell 60, 733-746. 670. Benezra, R. et al. (1990). The protein Id: a negative regulator of helix-loop-helix DNA-binding proteins. Cell 61, 49-59. 671. Lassar, A. B. et al. (1991). Functional activity of myogenic HLH proteins requires hetero-oligomerization with E12/E47-like proteins in vitro. Cell 66, 305-315. 675. Murre, C., McCaw, P. S., and Baltimore, D. (1989). A new DNA binding and dimerization motif in immunoglobulin enhancer binding, daughterless, MyoD, and myc proteins. Cell 56, 777-783.



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5.22.16 Leucine zippers are involved in dimer formation Key Terms The leucine zipper is a dimerization motif adjacent to a basic DNA-binding region that is found in a class of transcription factors. A bZIP protein has a basic DNA-binding region adjacent to a leucine zipper dimerization motif. Key Concepts

• The leucine zipper is an amphipathic helix that dimerizes. • The zipper is adjacent to a basic region that binds DNA. • Dimerization forms the bZIP motif in which the two basic regions symmetrically bind inverted repeats in DNA.

Interactions between proteins are a common theme in building a transcription complex, and a motif found in several activators (and other proteins) is involved in both homo- and heteromeric interactions. The leucine zipper is a stretch of amino acids rich in leucine residues that provide a dimerization motif. Dimer formation itself has emerged as a common principle in the action of proteins that recognize specific DNA sequences, and in the case of the leucine zipper, its relationship to DNA binding is especially clear, because we can see how dimerization juxtaposes the DNA-binding regions of each subunit. The reaction is depicted diagrammatically in Figure 22.28 (673).

Figure 22.28 The basic regions of the bZIP motif are held together by the dimerization at the adjacent zipper region when the hydrophobic faces of two leucine zippers interact in parallel orientation. Leucine zippers are involved in dimer formation | SECTION 5.22.16 © 2004. Virtual Text / www.ergito.com

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Molecular Biology An amphipathic α-helix has a structure in which the hydrophobic groups (including leucine) face one side, while charged groups face the other side. A leucine zipper forms an amphipathic helix in which the leucines of the zipper on one protein could protrude from the α-helix and interdigitate with the leucines of the zipper of another protein in parallel to form a coiled coil. The two right-handed helices wind around each other, with 3.5 residues per turn, so the pattern repeats integrally every 7 residues. How is this structure related to DNA binding? The region adjacent to the leucine repeats is highly basic in each of the zipper proteins, and could comprise a DNA-binding site. The two leucine zippers in effect form a Y-shaped structure, in which the zippers comprise the stem, and the two basic regions stick out to form the arms that bind to DNA. This is known as the bZIP structural motif. It explains why the target sequences for such proteins are inverted repeats with no separation (676). Zippers may be used to sponsor formation of homodimers or heterodimers. They are lengthy motifs. Leucine (or another hydrophobic amino acid) occupies every seventh residue in the potential zipper. There are 4 repeats of the zipper (Leu-X6) in the protein C/EBP (a factor that binds as a dimer to both the CAAT box and the SV40 core enhancer), and 5 repeats in the factors Jun and Fos (which form the heterodimeric activator, AP1). AP1 was originally identified by its binding to a DNA sequence in the SV40 enhancer (see Figure 21.24). The active preparation of AP1 includes several polypeptides. A major component is Jun, the product of the gene c-jun, which was identified by its relationship with the oncogene v-jun carried by an avian sarcoma virus (see Molecular Biology 6.30.18 Oncoproteins may regulate gene expression). The mouse genome contains a family of related genes, c-jun (the original isolate) and junB and junD (identified by sequence homology with jun). There are considerable sequence similarities in the three Jun proteins; they have leucine zippers that can interact to form homodimers or heterodimers. The other major component of AP1 is the product of another gene with an oncogenic counterpart. The c-fos gene is the cellular homologue to the oncogene v-fos carried by a murine sarcoma virus. Expression of c-fos activates genes whose promoters or enhancers possess an AP1 target site. The c-fos product is a nuclear phosphoprotein that is one of a group of proteins. The others are described as Fos-related antigens (FRA); they constitute a family of Fos-like proteins. Fos also has a leucine zipper. Fos cannot form homodimers, but can form a heterodimer with Jun. A leucine zipper in each protein is required for the reaction. The ability to form dimers is a crucial part of the interaction of these factors with DNA. Fos cannot by itself bind to DNA, possibly because of its failure to form a dimer. But the Jun-Fos heterodimer can bind to DNA with same target specificity as the Jun-Jun dimer; and this heterodimer binds to the AP1 site with an affinity ~10× that of the Jun homodimer.

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Reviews 676. Vinson, C. R., Sigler, P. B., and McKnight, S. L. (1989). Scissors-grip model for DNA recognition by a family of leucine zipper proteins. Science 246, 911-916.

References 673. Landschulz, W. H., Johnson, P. F., and McKnight, S. L. (1988). The leucine zipper: a hypothetical structure common to a new class of DNA binding proteins. Science 240, 1759-1764.


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5.22.17 Summary Transcription factors include basal factors, activators, and coactivators. Basal factors interact with RNA polymerase at the startpoint. Activators bind specific short response elements (REs) located in promoters or enhancers. Activators function by making protein-protein interactions with the basal apparatus. Some activators interact directly with the basal apparatus; others require coactivators to mediate the interaction. Activators often have a modular construction, in which there are independent domains responsible for binding to DNA and for activating transcription. The main function of the DNA-binding domain may be to tether the activating domain in the vicinity of the initiation complex. Some response elements are present in many genes and are recognized by ubiquitous factors; others are present in a few genes and are recognized by tissue-specific factors. Promoters for RNA polymerase II contain a variety of short cis-acting elements, each of which is recognized by a trans-acting factor. The cis-acting elements are located upstream of the TATA box and may be present in either orientation and at a variety of distances with regard to the startpoint. The upstream elements are recognized by activators that interact with the basal transcription complex to determine the efficiency with which the promoter is used. Some activators interact directly with components of the basal apparatus; others interact via intermediaries called coactivators. The targets in the basal apparatus are the TAFs of TFIID, or TFIIB or TFIIA. The interaction stimulates assembly of the basal apparatus. Several groups of transcription factors have been identified by sequence homologies. The homeodomain is a 60 residue sequence found in genes that regulate development in insects and worms and in mammalian transcription factors. It is related to the prokaryotic helix-turn-helix motif and provides the motif by which the factors bind to DNA. Another motif involved in DNA-binding is the zinc finger, which is found in proteins that bind DNA or RNA (or sometimes both). A finger has cysteine residues that bind zinc. One type of finger is found in multiple repeats in some transcription factors; another is found in single or double repeats in others. Steroid receptors were the first members identified of a group of transcription factors in which the protein is activated by binding a small hydrophobic hormone. The activated factor becomes localized in the nucleus, and binds to its specific response element, where it activates transcription. The DNA-binding domain has zinc fingers. The receptors are homodimers or heterodimers. The homodimers all recognize palindromic response elements with the same consensus sequence; the difference between the response elements is the spacing between the inverted repeats. The heterodimers recognize direct repeats, again being distinguished by the spacing between the repeats. The DNA-binding motif of these receptors includes two zinc fingers; the first determines which consensus sequence is recognized, and the second responds to the spacing between the repeats. The leucine zipper contains a stretch of amino acids rich in leucine that are involved Summary | SECTION 5.22.17 © 2004. Virtual Text / www.ergito.com

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in dimerization of transcription factors. An adjacent basic region is responsible for binding to DNA. HLH (helix-loop-helix) proteins have amphipathic helices that are responsible for dimerization, adjacent to basic regions that bind to DNA. bHLH proteins have a basic region that binds to DNA, and fall into two groups: ubiquitously expressed and tissue-specific. An active protein is usually a heterodimer between two subunits, one from each group. When a dimer has one subunit that does not have the basic region, it fails to bind DNA, so such subunits can prevent gene expression. Combinatorial associations of subunits form regulatory networks. Many transcription factors function as dimers, and it is common for there to be multiple members of a family that form homodimers and heterodimers. This creates the potential for complex combinations to govern gene expression. In some cases, a family includes inhibitory members, whose participation in dimer formation prevents the partner from activating transcription. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.22.17


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CONTROLLING CHROMATIN STRUCTURE

5.23.1 Introduction Key Terms Epigenetic changes influence the phenotype without altering the genotype. They consist of changes in the properties of a cell that are inherited but that do not represent a change in genetic information. A prion is a proteinaceous infectious agent, which behaves as an inheritable trait, although it contains no nucleic acid. Examples are PrPSc, the agent of scrapie in sheep and bovine spongiform encephalopathy, and Psi, which confers an inherited state in yeast.

When transcription is treated in terms of interactions involving DNA and individual transcription factors and RNA polymerases, we get an accurate description of the events that occur in vitro, but this lacks an important feature of transcription in vivo. The cellular genome is organized as nucleosomes, but initiation of transcription generally is prevented if the promoter region is packaged into nucleosomes. In this sense, histones function as generalized repressors of transcription (a rather old idea), although we see in this Chapter that they are also involved in more specific interactions. Activation of a gene requires changes in the state of chromatin: the essential issue is how the transcription factors gain access to the promoter DNA. Local chromatin structure is an integral part of controlling gene expression. Genes may exist in either of two structural conditions. Genes are found in an "active" state only in the cells in which they are expressed. The change of structure precedes the act of transcription, and indicates that the gene is "transcribable." This suggests that acquisition of the "active" structure must be the first step in gene expression. Active genes are found in domains of euchromatin with a preferential susceptibility to nucleases (see Molecular Biology 5.20.16 Domains define regions that contain active genes). Hypersensitive sites are created at promoters before a gene is activated (see Molecular Biology 5.20.15 DNAase hypersensitive sites change chromatin structure). More recently it has turned out that there is an intimate and continuing connection between initiation of transcription and chromatin structure. Some activators of gene transcription directly modify histones; in particular, acetylation of histones is associated with gene activation. Conversely, some repressors of transcription function by deacetylating histones. So a reversible change in histone structure in the vicinity of the promoter is involved in the control of gene expression. This may be part of the mechanism by which a gene is maintained in an active or inactive state. The mechanisms by which local regions of chromatin are maintained in an inactive (silent) state are related to the means by which an individual promoter is repressed. The proteins involved in the formation of heterochromatin act on chromatin via the histones, and modifications of the histones may be an important feature in the interaction. Once established, such changes in chromatin may persist through cell divisions, creating an epigenetic state in which the properties of a gene are Introduction | SECTION 5.23.1 © 2004. Virtual Text / www.ergito.com

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determined by the self-perpetuating structure of chromatin. The name epigenetic reflects the fact that a gene may have an inherited condition (it may be active or may be inactive) which does not depend on its sequence. Yet a further insight into epigenetic properties is given by the self-perpetuating structures of prions (proteinaceous infectious agents). Last updated on 10-2-2001 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.23.1


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5.23.2 Chromatin can have alternative states Key Concepts

• Chromatin structure is stable and cannot be changed by altering the equilibrium of transcription factors and histones.

Two types of model have been proposed to explain how the state of expression of DNA is changed: equilibrium and discontinuous change-of-state. Figure 23.1 shows the equilibrium model. Here the only pertinent factor is the concentration of the repressor or activator protein, which drives an equilibrium between free form and DNA-bound form. When the concentration of the protein is high enough, its DNA-binding site is occupied, and the state of expression of the DNA is affected. (Binding might either repress or activate any particular target sequence.) This type of model explains the regulation of transcription in bacterial cells, where gene expression is determined exclusively by the actions of individual repressor and activator proteins (see Molecular Biology 3.10 The operon). Whether a bacterial gene is transcribed can be predicted from the sum of the concentrations of the various factors that either activate or repress the individual gene. Changes in these concentrations at any time will change the state of expression accordingly. In most cases, the protein binding is cooperative, so that once the concentration becomes high enough, there is a rapid association with DNA, resulting in a switch in gene expression.

Figure 23.1 In an equilibrium model, the state of a binding site on DNA depends on the concentration of the protein that binds to it.

A different situation applies with eukaryotic chromatin. Early in vitro experiments showed that either an active or inactive state can be established, but this is not affected by the subsequent addition of other components. The transcription factor TFIIIA, required for RNA polymerase III to transcribe 5S rRNA genes, cannot activate its target genes in vitro if they are complexed with histones. However, if the factor is presented with free DNA, it forms a transcription complex, and then the addition of histones does not prevent the gene from remaining active. Once the factor has bound, it remains at the site, allowing a succession of RNA polymerase molecules to initiate transcription. Whether the factor or histones get to the control Chromatin can have alternative states | SECTION 5.23.2 © 2004. Virtual Text / www.ergito.com

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site first may be the critical factor (680; for review see 227; 228). Figure 23.2 illustrates the two types of condition that can exist at a eukaryotic promoter. In the inactive state, nucleosomes are present, and they prevent basal factors and RNA polymerase from binding. In the active state, the basal apparatus occupies the promoter, and histone octamers cannot bind to it. Each type of state is stable.

Figure 23.2 If nucleosomes form at a promoter, transcription factors (and RNA polymerase) cannot bind. If transcription factors (and RNA polymerase) bind to the promoter to establish a stable complex for initiation, histones are excluded.

A similar situation is seen with the TFIID complex at promoters for RNA polymerase II. A plasmid containing an adenovirus promoter can be transcribed in vitro by RNA polymerase II in a reaction that requires TFIID and other transcription factors. The template can be assembled into nucleosomes by the addition of histones. If the histones are added before the TFIID, transcription cannot be initiated. But if the TFIID is added first, the template still can be transcribed in its chromatin form. So TFIID can recognize free DNA, but either cannot recognize or cannot function on nucleosomal DNA. Only the TFIID must be added before the histones; the other transcription factors and RNA polymerase can be added later. This suggests that binding of TFIID to the promoter creates a structure to which the other components of the transcription apparatus can bind (681). It is important to note that these in vitro systems use disproportionate quantities of components, which may create unnatural situations. The major importance of these results, therefore, is not that they demonstrate the mechanism used in vivo, but that they establish the principle that transcription factors or nucleosomes may form stable structures that cannot be changed merely by changing the equilibrium with free Chromatin can have alternative states | SECTION 5.23.2 © 2004. Virtual Text / www.ergito.com

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components. Last updated on 5-12-2001

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Reviews 227. Brown, D. D. (1984). The role of stable complexes that repress and activate eukaryotic genes. Cell 37, 359-365. 228. Weintraub, H. (1985). Assembly and propagation of repressed and derepressed chromosomal states. Cell 42, 705-711.

References 680. Bogenhagen, D. F., Wormington, W. M., and Brown, D. D. (1982). Stable transcription complexes of Xenopus 5S RNA genes: a means to maintain the differentiated state. Cell 28, 413-421. 681. Workman, J. L. and Roeder, R. G. (1987). Binding of transcription factor TFIID to the major late promoter during in vitro nucleosome assembly potentiates subsequent initiation by RNA polymerase II. Cell 51, 613-622.


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5.23.3 Chromatin remodeling is an active process Key Terms Chromatin remodeling describes the energy-dependent displacement or reorganization of nucleosomes that occurs in conjunction with activation of genes for transcription. SWI/SNF is a chromatin remodeling complex; it uses hydrolysis of ATP to change the organization of nucleosomes. Key Concepts

• There are several chromatin remodeling complexes that use energy provided by hydrolysis of ATP.

• The SWI/SNF, RSC, and NURF complexes all are very large; there are some common subunits.

• A remodeling complex does not itself have specificity for any particular target site, but must be recruited by a component of the transcription apparatus.

The general process of inducing changes in chromatin structure is called chromatin remodeling. This consists of mechanisms for displacing histones that depend on the input of energy. Many protein-protein and protein-DNA contacts need to be disrupted to release histones from chromatin. There is no free ride: the energy must be provided to disrupt these contacts. Figure 23.3 illustrates the principle of a dynamic model by a factor that hydrolyzes ATP. When the histone octamer is released from DNA, other proteins (in this case transcription factors and RNA polymerase) can bind.

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Figure 23.3 The dynamic model for transcription of chromatin relies upon factors that can use energy provided by hydrolysis of ATP to displace nucleosomes from specific DNA sequences.

Figure 23.4 summarizes the types of remodeling changes in chromatin that can be characterized in vitro:

Figure 23.4 Remodeling complexes can cause nucleosomes to slide along DNA, can displace nucleosomes from DNA, or can reorganize the spacing between nucleosomes.


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• Histone octamers may slide along DNA, changing the relationship between the nucleic acid and protein. This alters the position of a particular sequence on the nucleosomal surface. • The spacing between histone octamers may be changed, again with the result that the positions of individual sequences are altered relative to protein. • And the most extensive change is that an octamer(s) may be displaced entirely from DNA to generate a nucleosome-free gap. The most common use of chromatin remodeling is to change the organization of nucleosomes at the promoter of a gene that is to be transcribed. This is required to allow the transcription apparatus to gain access to the promoter. The remodeling most often takes the form of displacing one or more histone octamers. This can be detected by a change in the micrococcal nuclease ladder where protection against cleavage has been lost. It often results in the creation of a site that is hypersensitive to cleavage with DNAase I (see Molecular Biology 5.20.15 DNAase hypersensitive sites change chromatin structure). Sometimes there are less dramatic changes, for example, involving a change in rotational positioning of a single nucleosome; this may be detected by loss of the DNAaseI 10 base ladder. So changes in chromatin structure may extend from altering the positions of nucleosomes to removing them altogether (for review see 234; 237). Chromatin remodeling is undertaken by large complexes that use ATP hydrolysis to provide the energy for remodeling. The heart of the remodeling complex is its ATPase subunit. Remodeling complexes are usually classified according to the type of ATPase subunit – those with related ATPase subunits are considered to belong to the same family (usually some other subunits are common also). Figure 23.5 keeps the names straight (for review see 3007). The two major types of complex are SWI/SNF and ISW (ISW stands for imitation SWI). Yeast has two complexes of each type. Complexes of both types are also found in fly and in Man (for review see 1969; 2413). Each type of complex may undertake a different range of remodeling activities (for review see 3252; 3432).


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Figure 23.5 Remodeling complexes can be classified by their ATPase subunits.

SWI/SNF was the first remodeling complex to be identified. Its name reflects the fact that many of its subunits are coded by genes originally identified by SWI or SNF mutations in S. cerevisiae. Mutations in these loci are pleiotropic, and the range of defects is similar to those shown by mutants that have lost the CTD tail of RNA polymerase II. These mutations also show genetic interactions with mutations in genes that code for components of chromatin, in particular SIN1, which codes for a nonhistone protein, and SIN2, which codes for histone H3. The SWI and SNF genes are required for expression of a variety of individual loci (~120 or 2% of S. cerevisiae genes are affected). Expression of these loci may require the SWI/SNF complex to remodel chromatin at their promoters (684; 682; 685). SWI/SNF acts catalytically in vitro (1971), and there are only ~150 complexes per yeast cell. All of the genes encoding the SWI/SNF subunits are nonessential, which implies that yeast must also have other ways of remodeling chromatin (1974; 1975). The RSC complex is more abundant and also is essential. It acts at ~ 700 target loci (2495). SWI/SNF complexes can remodel chromatin in vitro without overall loss of histones or can displace histone octamers (1972; 1973). Both types of reaction may pass through the same intermediate in which the structure of the target nucleosome is altered, leading either to reformation of a (remodeled) nucleosome on the original DNA or to displacement of the histone octamer to a different DNA molecule. The SWI/SNF complex alters nucleosomal sensitivity to DNAase I at the target site, and induces changes in protein-DNA contacts that persist after it has been released from the nucleosomes (688; 694). The SWI2 subunit is the ATPase that provides the energy for remodeling by SWI/SNF. There are many contacts between DNA and a histone octamer – 14 are identified in the crystal structure. All of these contacts must be broken for an octamer to be released or for it to move to a new position. How is this achieved? Some obvious mechanisms can be excluded because we know that single-stranded DNA is not generated during remodeling (and there are no helicase activities associated with the Chromatin remodeling is an active process | SECTION 5.23.3 © 2004. Virtual Text / www.ergito.com

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complexes). Present thinking is that remodeling complexes in the SWI and ISW classes use the hydrolysis of ATP to twist DNA on the nucleosomal surface. Indirect evidence suggests that this creates a mechanical force that allows a small region of DNA to be released from the surface and then repositioned (3008). One important reaction catalyzed by remodeling complexes involves nucleosome sliding. It was first observed that the ISW family affects nucleosome positioning without displacing octamers (1974; 1975). This is achieved by a sliding reaction, in which the octamer moves along DNA. Sliding is prevented if the N-terminal tail of histone H4 is removed, but we do not know exactly how the tail functions in this regard (2218). SWI/SNF complexes have the same capacity; the reaction is prevented by the introduction of a barrier in the DNA, which suggests that a sliding reaction is involved, in which the histone octamer moves more or less continuously along DNA without ever losing contact with it (3253). One puzzle about the action of the SWI/SNF complex is its sheer size. It has 11 subunits with a combined molecular weight ~2 × 106. It dwarfs RNA polymerase and the nucleosome, making it difficult to understand how all of these components could interact with DNA retained on the nucleosomal surface. However, a transcription complex with full activity, called RNA polymerase II holoenzyme, can be found that contains the RNA polymerase itself, all the TFII factors except TBP and TFIIA, and the SWI/SNF complex, which is associated with the CTD tail of the polymerase. In fact, virtually all of the SWI/SNF complex may be present in holoenzyme preparations. This suggests that the remodeling of chromatin and recognition of promoters is undertaken in a coordinated manner by a single complex (689). Last updated on 10-16-2002


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Reviews 234. Grunstein, M. (1990). Histone function in transcription. Annu. Rev. Cell Biol. 6, 643-678. 237. Felsenfeld, G. (1992). Chromatin as an essential part of the transcriptional mechanism. Nature 355, 219-224. 2413. Narlikar, G. J., Fan, H. Y., and Kingston, R. E. (2002). Cooperation between complexes that regulate chromatin structure and transcription. Cell 108, 475-487. 3007. Vignali, M., Hassan, A. H., Neely, K. E., and Workman, J. L. (2000). ATP-dependent chromatin-remodeling complexes. Mol. Cell Biol. 20, 1899-1910. 3252. Becker, P. B. and Horz, W. (2002). ATP-dependent nucleosome remodeling. Annu. Rev. Biochem. 71, 247-273. 3432. Tsukiyama, T. (2002). The in vivo functions of ATP-dependent chromatin-remodelling factors. Nat. Rev. Mol. Cell Biol. 3, 422-429.


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References 682. Cairns, B. R., Kim, Y.- J., Sayre, M. H., Laurent, B. C., and Kornberg, R. (1994). A multisubunit complex containing the SWI/ADR6, SWI2/1, SWI3, SNF5, and SNF6 gene products isolated from yeast. Proc. Natl. Acad. Sci. USA 91, 1950-622. 684. Peterson, C. L. and Herskowitz, I. (1992). Characterization of the yeast SWI1, SWI2, and SWI3 genes, which encode a global activator of transcription. Cell 68, 573-583. 685. Tamkun, J. W., Deuring, R., Scott, M. P., Kissinger, M., Pattatucci, A. M., Kaufman, T. C., and Kennison, J. A. (1992). brahma: a regulator of Drosophila homeotic genes structurally related to the yeast transcriptional activator SNF2/SWI2. Cell 68, 561-572. 688. Cote, J., Quinn, J., Workman, J. L., and Peterson, C. L. (1994). Stimulation of GAL4 derivative binding to nucleosomal DNA by the yeast SWI/SNF complex. Science 265, 53-60. 689. Kwon, H., Imbaizano, A. N., Khavari, P. A., Kingston, R. E., and Green, M. R. (1994). Nucleosome disruption and enhancement of activator binding of human SWI/SNF complex. Nature 370, 477-481. 694. Schnitzler, G., Sif, S., and Kingston, R. E. (1998). Human SWI/SNF interconverts a nucleosome between its base state and a stable remodeled state. Cell 94, 17-27. 1969. Kingston, R. E. and Narlikar, G. J. (1999). ATP-dependent remodeling and acetylation as regulators of chromatin fluidity. Genes Dev. 13, 2339-2352. 1971. Logie, C. and Peterson, C. L. (1997). Catalytic activity of the yeast SWI/SNF complex on reconstituted nucleosome arrays. EMBO J. 16, 6772-6782. 1972. Lorch, Y., Cairns, B. R., Zhang, M., and Kornberg, R. D. (1998). Activated RSC-nucleosome complex and persistently altered form of the nucleosome. Cell 94, 29-34. 1973. Lorch, Y., Zhang, M., and Kornberg, R. D. (1999). Histone octamer transfer by a chromatin-remodeling complex. Cell 96, 389-392. 1974. Tsukiyama, T., Daniel, C., Tamkun, J., and Wu, C. (1995). ISWI, a member of the SWI2/SNF2 ATPase family, encodes the 140 kDa subunit of the nucleosome remodeling factor. Cell 83, 1021-1026. 1975. Tsukiyama, T., Palmer, J., Landel, C. C., Shiloach, J., and Wu, C. (1999). Characterization of the imitation switch subfamily of ATP-dependent chromatin-remodeling factors in S. cerevisiae. Genes Dev. 13, 686-697. 2218. Hamiche, A., Kang, J. G., Dennis, C., Xiao, H., and Wu, C. (2001). Histone tails modulate nucleosome mobility and regulate ATP-dependent nucleosome sliding by NURF. Proc. Natl. Acad. Sci. USA 98, 14316-14321. 2495. Robert, F., Young, R. A., and Struhl, K. (2002). Genome-wide location and regulated recruitment of the RSC nucleosome remodeling complex. Genes Dev. 16, 806-819. 3008. Gavin, I., Horn, P. J., and Peterson, C. L. (2001). SWI/SNF chromatin remodeling requires changes in DNA topology. Mol. Cell 7, 97-104. 3253. Whitehouse, I., Flaus, A., Cairns, B. R., White, M. F., Workman, J. L., and Owen-Hughes, T. (1999). Nucleosome mobilization catalysed by the yeast SWI/SNF complex. Nature 400, 784-787.



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5.23.4 Nucleosome organization may be changed at the promoter Key Concepts

• Remodeling complexes are recruited to promoters by sequence-specific activators. • The factor may be released once the remodeling complex has bound. • The MMTV promoter requires a change in rotational positioning of a nucleosome to allow an activator to bind to DNA on the nucleosome.

How are remodeling complexes targeted to specific sites on chromatin? They do not themselves contain subunits that bind specific DNA sequences. This suggests the model shown in Figure 23.6 in which they are recruited by activators or (sometimes) by repressors (1864; 1970).

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Figure 23.6 A remodeling complex binds to chromatin via an activator (or repressor).

The interaction between transcription factors and remodeling complexes gives a key insight into their modus operandi. The transcription factor Swi5p activates the HO locus in yeast. (Note that Swi5p is not a member of the SWI/SNF complex.) Swi5p enters nuclei toward the end of mitosis and binds to the HO promoter. It then recruits SWI/SNF to the promoter. Then Swi5p is released, leaving SWI/SNF at the promoter (1966). This means that a transcription factor can activate a promoter by a "hit and run" mechanism, in which its function is fulfilled once the remodeling complex has bound. The involvement of remodeling complexes in gene activation was discovered because the complexes are necessary for the ability of certain transcription factors to activate their target genes. One of the first examples was the GAGA factor, which activates the hsp70 Drosophila promoter in vitro. Binding of GAGA to four (CT)n-rich sites on the promoter disrupts the nucleosomes, creates a hypersensitive Nucleosome organization may be changed at the promoter | SECTION 5.23.4 © 2004. Virtual Text / www.ergito.com

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region, and causes the adjacent nucleosomes to be rearranged so that they occupy preferential instead of random positions. Disruption is an energy-dependent process that requires the NURF remodeling complex. The organization of nucleosomes is altered so as to create a boundary that determines the positions of the adjacent nucleosomes (691). During this process, GAGA binds to its target sites and DNA, and its presence fixes the remodeled state. The PHO system was one of the first in which it was shown that a change in nucleosome organization is involved in gene activation (for review see 1976). At the PHO5 promoter, the bHLH regulator PHO4 responds to phosphate starvation by inducing the disruption of four precisely positioned nucleosomes. This event is independent of transcription (it occurs in a TATA– mutant) and independent of replication. There are two binding sites for PHO4 at the promoter, one located between nucleosomes, which can be bound by the isolated DNA-binding domain of PHO4, and the other within a nucleosome, which cannot be recognized. Disruption of the nucleosome to allow DNA binding at the second site is necessary for gene activation. This action requires the presence of the transcription-activating domain. The activator sequence of VP16 can substitute for the PHO4 activator sequence in nucleosome disruption. This suggests that disruption occurs by protein-protein interactions that involve the same region that makes protein-protein contacts to activate transcription (605). In this case, it is not known which remodeling complex is involved in executing the effects. It is not always the case, however, that nucleosomes must be excluded in order to permit initiation of transcription. Some activators can bind to DNA on a nucleosomal surface. Nucleosomes appear to be precisely positioned at some steroid hormone response elements in such a way that receptors can bind. Receptor binding may alter the interaction of DNA with histones, and even lead to exposure of new binding sites. The exact positioning of nucleosomes could be required either because the nucleosome "presents" DNA in a particular rotational phase or because there are protein-protein interactions between the activators and histones or other components of chromatin. So we have now moved some way from viewing chromatin exclusively as a repressive structure to considering which interactions between activators and chromatin can be required for activation. The MMTV promoter presents an example of the need for specific nucleosomal organization. It contains an array of 6 partly palindromic sites, each bound by one dimer of hormone receptor (HR), which constitute the HRE. It also has a single binding site for the factor NF1, and two adjacent sites for the factor OTF. HR and NF1 cannot bind simultaneously to their sites in free DNA. Figure 23.7 shows how the nucleosomal structure controls binding of the factors.


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Figure 23.7 Hormone receptor and NF1 cannot bind simultaneously to the MMTV promoter in the form of linear DNA, but can bind when the DNA is presented on a nucleosomal surface.

The HR protects its binding sites at the promoter when hormone is added, but does not affect the micrococcal nuclease-sensitive sites that mark either side of the nucleosome. This suggests that HR is binding to the DNA on the nucleosomal surface. However, the rotational positioning of DNA on the nucleosome prior to hormone addition allows access to only two of the four sites. Binding to the other two sites requires a change in rotational positioning on the nucleosome. This can be detected by the appearance of a sensitive site at the axis of dyad symmetry (which is in the center of the binding sites that constitute the HRE). NF1 can be footprinted on the nucleosome after hormone induction, so these structural changes may be necessary to allow NF1 to bind, perhaps because they expose DNA and abolish the steric hindrance by which HR blocks NF1 binding to free DNA (687; 690). Last updated on 9-5-2001


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Reviews 1976. Lohr, D. (1997). Nucleosome transactions on the promoters of the yeast GAL and PHO genes. J. Biol. Chem. 272, 26795-26798.

References 605. Schmid, V. M., Fascher, K.-D., and Horz, W. (1992). Nucleosome disruption at the yeast PHO5 promoter upon PHO5 induction occurs in the absence of DNA replication. Cell 71, 853-864. 687. McPherson, C. E., Shim, E.-Y., Friedman, D. S., and Zaret, K. S. (1993). An active tissue-specific enhancer and bound transcription factors existing in a precisely positioned nucleosomal array. Cell 75, 387-398. 690. Truss, M., Barstch, J., Schelbert, A., Hache, R. J. G., and Beato, M. (1994). Hormone induces binding of receptors and transcription factors to a rearranged nucleosome on the MMTV promoter in vitro. EMBO J. 14, 1737-1751. 691. Tsukiyama, T., Becker, P. B., and Wu, C. (1994). ATP-dependent nucleosome disruption at a heat shock promoter mediated by binding of GAGA transcription factor. Nature 367, 525-532. 1864. Kadam, S., McAlpine, G. S., Phelan, M. L., Kingston, R. E., Jones, K. A., and Emerson, B. M. (2000). Functional selectivity of recombinant mammalian SWI/SNF subunits. Genes Dev. 14, 2441-2451. 1966. Cosma, M. P., Tanaka, T., and Nasmyth, K. (1999). Ordered recruitment of transcription and chromatin remodeling factors to a cell cycle- and developmentally regulated promoter. Cell 97, 299-311. 1970. Yudkovsky, N., Logie, C., Hahn, S., and Peterson, C. L. (1999). Recruitment of the SWI/SNF chromatin remodeling complex by transcriptional activators. Genes Dev. 13, 2369-2374.



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5.23.5 Histone modification is a key event Key Terms Silencing describes the repression of gene expression in a localized region, usually as the result of a structural change in chromatin. Heterochromatin describes regions of the genome that are highly condensed, are not transcribed, and are late-replicating. Heterochromatin is divided into two types, which are called constitutive and facultative.

Whether a gene is expressed depends on the structure of chromatin both locally (at the promoter) and in the surrounding domain. Chromatin structure correspondingly can be regulated by individual activation events or by changes that affect a wide chromosomal region. The most localized events concern an individual target gene, where changes in nucleosomal structure and organization occur in the immediate vicinity of the promoter. More general changes may affect regions as large as a whole chromosome. Changes that affect large regions control the potential of a gene to be expressed. The term silencing is used to refer to repression of gene activity in a local chromosomal region. The term heterochromatin is used to describe chromosomal regions that are large enough to be seen to have a physically more compact structure in the microscope. The basis for both types of change is the same: additional proteins bind to chromatin and either directly or indirectly prevent transcription factors and RNA polymerase from activating promoters in the region. Changes at an individual promoter control whether transcription is initiated for a particular gene. These changes may be either activating or repressing. All of these events depend on interactions with histones. Changes in chromatin structure are initiated by modifying the N-terminal tails of the histones, especially H3 and H4. The histone tails consist of the N-terminal 20 amino acids, and extend from the nucleosome between the turns of DNA (see Figure 20.25 in Molecular Biology 5.20.8 Organization of the histone octamer). Figure 23.8 shows that they can be modified at several sites, by methylation, acetylation, or phosphorylation (see Molecular Biology 5.20.9 The N-terminal tails of histones are modified). The modifications reduce positive charge. The histone modifications may directly affect nucleosome structure or create binding sites for the attachment of nonhistone proteins that change the properties of chromatin.

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Figure 23.8 The N-terminal tails of histones H3 and H4 can be acetylated, methylated, or phosphorylated at several positions.

The range of nucleosomes that is targeted for modification can vary. Modification can be a local event, for example, restricted to nucleosomes at the promoter. Or it can be a general event, extending for example to an entire chromosome. Figure 23.9 shows that there is a general correlation in which acetylation is associated with active chromatin while methylation is associated with inactive chromatin. However, this is not a simple rule, and the particular sites that are modified, as well as combinations of specific modifications may be important, so there are certainly exceptions in which (for example) histones methylated at a certain position are found in active chromatin. Mutations in one of the histone acetylase complexes of yeast have the opposite effect from usual (they prevent silencing of some genes), emphasizing the lack of a uniform effect of acetylation (2228).

Figure 23.9 Acetylation of H3 and H4 is associated with active chromatin, while methylation is associated with inactive chromatin.

The specificity of the modifications is indicated by the fact that many of the modifying enzymes have individual target sites in specific histones. Figure 23.10 summarizes the effects of some of the modifications. Most modified sites are subject to only a single type of modification. In some cases, modification of one site may activate or inhibit modification of another site. The idea that combinations of signals may be used to define chromatin types has sometimes been called the histone code (for review see 2033). Histone modification is a key event | SECTION 5.23.5 © 2004. Virtual Text / www.ergito.com

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Figure 23.10 Most modified sites in histones have a single, specific type of modification, but some sites can have more than one type of modification. Individual functions can be associated with some of the modifications.



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Reviews 2033. Jenuwein, T. and Allis, C. D. (2001). Translating the histone code. Science 293, 1074-1080.

References 2228. Osada, S., Sutton, A., Muster, N., Brown, C. E., Yates, J. R., Sternglanz, R., and Workman, J. L. (2001). The yeast SAS (something about silencing) protein complex contains a MYST-type putative acetyltransferase and functions with chromatin assembly factor ASF1. Genes Dev. 15, 3155-3168.



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5.23.6 Histone acetylation occurs in two circumstances Key Concepts

• Histone acetylation occurs transiently at replication. • Histone acetylation is associated with activation of gene expression.

All the core histones can be acetylated. The major targets for acetylation are lysines in the N-terminal tails of histones H3 and H4. Acetylation occurs in two different circumstances: • during DNA replication; • and when genes are activated. When chromosomes are replicated, during the S phase of the cell cycle, histones are transiently acetylated (1980; for review see 1978). Figure 23.11 shows that this acetylation occurs before the histones are incorporated into nucleosomes. We know that histones H3 and H4 are acetylated at the stage when they are associated with one another in the H32·H42 tetramer. The tetramer is then incorporated into nucleosomes. Quite soon after, the acetyl groups are removed.

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Figure 23.11 Acetylation at replication occurs on histones before they are incorporated into nucleosomes.

The importance of the acetylation is indicated by the fact that preventing acetylation of both histones H3 and H4 during replication causes loss of viability in yeast (1977). The two histones are redundant as substrates, since yeast can manage perfectly well so long as they can acetylate either one of these histones during S phase. There are two possible roles for the acetylation: it could be needed for the histones to be recognized by factors that incorporate them into nucleosomes; or it could be required for the assembly and/or structure of the new nucleosome. The factors that are known to be involved in chromatin assembly do not distinguish between acetylated and nonacetylated histones, suggesting that the modification is more likely to be required for subsequent interactions (1979). It has been thought for a long time that acetylation might be needed to help control protein-protein interactions that occur as histones are incorporated into nucleosomes. Some evidence for such a role is that the yeast SAS histone acetylase complex binds to chromatin assembly complexes at the replication fork, where it acetylates 16Lys of histone H4 (2008; 2007). This may be part of the system that establishes the histone acetylation patterns after replication. Outside of S phase, acetylation of histones in chromatin is generally correlated with the state of gene expression. The correlation was first noticed because histone acetylation is increased in a domain containing active genes, and acetylated chromatin is more sensitive to DNAase I and (possibly) to micrococcal nuclease. Figure 23.12 shows that this involves the acetylation of histone tails in nucleosomes. We now know that this occurs largely because of acetylation of the nucleosomes in the vicinity of the promoter when a gene is activated. Histone acetylation occurs in two circumstances | SECTION 5.23.6 © 2004. Virtual Text / www.ergito.com

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Figure 23.12 Acetylation associated with gene activation occurs by directly modifying histones in nucleosomes.

In addition to events at individual promoters, widescale changes in acetylation occur on sex chromosomes. This is part of the mechanism by which the activities of genes on the X chromosome are altered to compensate for the presence of two X chromosomes in one species but only one X chromosome (in addition to the Y chromosome) in the other species (see Molecular Biology 5.23.17 X chromosomes undergo global changes). The inactive X chromosome in female mammals has underacetylated H4. The super-active X chromosome in Drosophila males has increased acetylation of H4 (616). This suggests that the presence of acetyl groups may be a prerequisite for a less condensed, active structure. In male Drosophila, the X chromosome is acetylated specifically at 16Lys of histone H4. The HAT that is responsible is an enzyme called MOF that is recruited to the chromosome as part of a large protein complex (1229). This "dosage compensation" complex is responsible for introducing general changes in the X chromosome that enable it to be more highly expressed. The increased acetylation is only one of its activities. Last updated on 12-17-2001


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Reviews 1978. Verreault, A. (2000). De novo nucleosome assembly: new pieces in an old puzzle. Genes Dev. 14, 1430-1438. 2007. Hirose, Y. and Manley, J. L. (2000). RNA polymerase II and the integration of nuclear events. Genes Dev. 14, 1415-1429.

References 616. Turner, B. M., Birley, A. J., and Lavender, J. (1992). Histone H4 isoforms acetylated at specific lysine residues define individual chromosomes and chromatin domains in Drosophila polytene nuclei. Cell 69, 375-384. 1229. Akhtar, A. and Becker, P. B. (2000). Activation of transcription through histone H4 acetylation by MOF, an acetyltransferase essential for dosage compensation in Drosophila. Mol. Cell 5, 367-375. 1977. Ling, X., Harkness, T. A., Schultz, M. C., Fisher-Adams, G., and Grunstein, M. (1996). Yeast histone H3 and H4 amino termini are important for nucleosome assembly in vivo and in vitro: redundant and position-independent functions in assembly but not in gene regulation. Genes Dev. 10, 686-699. 1979. Shibahara, K., Verreault, A., and Stillman, B. (2000). The N-terminal domains of histones H3 and H4 are not necessary for chromatin assembly factor-1- mediated nucleosome assembly onto replicated DNA in vitro. Proc. Natl. Acad. Sci. USA 97, 7766-7771. 1980. Jackson, V., Shires, A., Tanphaichitr, N., and Chalkley, R. (1976). Modifications to histones immediately after synthesis. J. Mol. Biol. 104, 471-483. 2008. 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, 5350-5354.



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5.23.7 Acetylases are associated with activators Key Terms Histone acetyltransferase (HAT) enzymes modify histones by addition of acetyl groups; some transcriptional coactivators have HAT activity. A deacetylase is an enzyme that removes acetyl groups from proteins. Histone deacetyltransferase (HDAC) enzymes remove acetyl groups from histones; they may be associated with repressors of transcription. Key Concepts

• Deacetylated chromatin may have a more condensed structure. • Transcription activators are associated with histone acetylase activities in large complexes.

• Histone acetylases vary in their target specificity. • Acetylation could affect transcription in a quantitative or qualitative way.

Acetylation is reversible. Each direction of the reaction is catalyzed by a specific type of enzyme. Enzymes that can acetylate histones are called histone acetyltransferases or HATs; the acetyl groups are removed by histone deacetylases or HDACs. There are two groups of HAT enzymes: group A describes those that are involved with transcription; group B describes those involved with nucleosome assembly. Two inhibitors have been useful in analyzing acetylation. Trichostatin and butyric acid inhibit histone deacetylases, and cause acetylated nucleosomes to accumulate. The use of these inhibitors has supported the general view that acetylation is associated with gene expression; in fact, the ability of butyric acid to cause changes in chromatin resembling those found upon gene activation was one of the first indications of the connection between acetylation and gene activity. The breakthrough in analyzing the role of histone acetylation was provided by the characterization of the acetylating and deacetylating enzymes, and their association with other proteins that are involved in specific events of activation and repression. A basic change in our view of histone acetylation was caused by the discovery that HATs are not necessarily dedicated enzymes associated with chromatin: rather it turns out that known activators of transcription have HAT activity. The connection was established when the catalytic subunit of a group A HAT was identified as a homologue of the yeast regulator protein GCN5. Then it was shown that GCN5 itself has HAT activity (with histones H3 and H4 as substrates). GCN5 is part of an adaptor complex that is necessary for the interaction between certain enhancers and their target promoters. Its HAT activity is required for activation of Acetylases are associated with activators | SECTION 5.23.7 © 2004. Virtual Text / www.ergito.com

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the target gene (693). This enables us to redraw our picture for the action of coactivators as shown in Figure 23.13, where RNA polymerase is bound at a hypersensitive site and coactivators are acetylating histones on the nucleosomes in the vicinity (692). Many examples are now known of interactions of this type.

Figure 23.13 Coactivators may have HAT activities that acetylate the tails of nucleosomal histones.

GCN5 leads us into one of the most important acetylase complexes. In yeast, GCN5 is part of the 1.8 MDa SAGA complex, which contains several proteins that are involved in transcription (for review see 1969). Among these proteins are several TAFIIs (696). Also, the TAFII145 subunit of TFIID is an acetylase. There are some functional overlaps between TFIID and SAGA, most notably that yeast can manage with either TAFII145 or GCN5, but is damaged by the deletion of both. This suggests that an acetylase activity is essential for gene expression, but can be provided by either TFIID or SAGA (1062). As might be expected from the size of the SAGA complex, acetylation is only one of its functions, although its other functions in gene activation are less well characterized. One of the first general activators to be characterized as an HAT was p300/CBP. (Actually, p300 and CBP are different proteins, but they are so closely related that they are often referred to as a single type of activity.) p300/CBP is a coactivator that links an activator to the basal apparatus (see Figure 22.8). p300/CBP interacts with various activators, including hormone receptors, AP-1 (c-Jun and c-Fos), and MyoD. The interaction is inhibited by the viral regulator proteins adenovirus E1A and SV40 T antigen, which bind to p300/CBP to prevent the interaction with transcription factors; this explains how these viral proteins inhibit cellular transcription. (This inhibition is important for the ability of the viral proteins to contribute to the tumorigenic state; see Molecular Biology 6.30.18 Oncoproteins may regulate gene Acetylases are associated with activators | SECTION 5.23.7 © 2004. Virtual Text / www.ergito.com

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expression). p300/CBP acetylates the N-terminal tails of H4 in nucleosomes. Another coactivator, called PCAF, preferentially acetylates H3 in nucleosomes. p300/CBP and PCAF form a complex that functions in transcriptional activation. In some cases yet another HAT is involved: the coactivator ACTR, which functions with hormone receptors, is itself an HAT that acts on H3 and H4, and also recruits both p300/CBP and PCAF to form a coactivating complex. One explanation for the presence of multiple HAT activities in a coactivating complex is that each HAT has a different specificity, and that multiple different acetylation events are required for activation. A general feature of acetylation is that an HAT is part of a large complex. Figure 23.14 shows a simplified model for their behavior. Typically the complex will contain a targeting subunit(s) that determines the binding sites on DNA. This determines the target for the HAT. The complex also contains effector subunits that affect chromatin structure or act directly on transcription. Probably at least some of the effectors require the acetylation event in order to act. Deacetylation, catalyzed by an HDAC, may work in a similar way.

Figure 23.14 Complexes that modify chromatin structure or activity have targeting subunits that determine their sites of action, HAT or HDAC enzymes that acetylate or deacetylate histones, and effector subunits that have other actions on chromatin or DNA.

Acetylation occurs at both replication (when it is transient) and at transcription (when it is maintained while the gene is active). Is it playing the same role in each case? One possibility is that the important effect is on nucleosome structure. Acetylation may be necessary to "loosen" the nucleosome core. At replication, acetylation of histones could be necessary to allow them to be incorporated into new cores more easily. At transcription, a similar effect could be necessary to allow a related change in structure, possibly even to allow the histone core to be displaced from DNA. Alternatively, acetylation could generate binding sites for other proteins that are required for transcription. In either case, deacetylation would reverse the effect. Is the effect of acetylation quantitative or qualitative? One possibility is that a certain number of acetyl groups are required to have an effect, and the exact positions at which they occur are largely irrelevant. An alternative is that individual acetylation events have specific effects. We might interpret the existence of complexes containing multiple HAT activities in either way – if individual enzymes have different specificities, we may need multiple activities either to acetylate a sufficient Acetylases are associated with activators | SECTION 5.23.7 © 2004. Virtual Text / www.ergito.com

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number of different positions or because the individual events are necessary for different effects upon transcription. At replication, it appears, at least with respect to histone H4, that acetylation at any two of three available positions is adequate, favoring a quantitative model in this case. Where chromatin structure is changed to affect transcription, acetylation at specific positions may be important (see Molecular Biology 5.23.15 Heterochromatin depends on interactions with histones).

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References 692. Chen, H. et al. (1997). Nuclear receptor coactivator ACTR is a novel histoneacetyltransferase and forms a multimeric activation complex with P/CAF and CP/p300. Cell 90, 569-580. 693. Brownell, J. E. et al. (1996). Tetrahymena histone acetyltransferase A: a homologue to yeast Gcn5p linking histone acetylation to gene activation. Cell 84, 843-851. 696. Grant, P. A. et al. (1998). A subset of TAFIIs are integral components of the SAGA complex required for nucleosome acetylation and transcriptional stimulation. Cell 94, 45-53. 1062. Lee, T. I., Causton, H. C., Holstege, F. C., Shen, W. C., Hannett, N., Jennings, E. G., Winston, F., Green, M. R., and Young, R. A. (2000). Redundant roles for the TFIID and SAGA complexes in global transcription. Nature 405, 701-704. 1969. Kingston, R. E. and Narlikar, G. J. (1999). ATP-dependent remodeling and acetylation as regulators of chromatin fluidity. Genes Dev. 13, 2339-2352.


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5.23.8 Deacetylases are associated with repressors Key Concepts

• Deacetylation is associated with repression of gene activity. • Deacetylases are present in complexes with repressor activity.

In yeast, mutations in SIN3 and Rpd3 behave as though these loci repress a variety of genes. The proteins form a complex with the DNA-binding protein Ume6, which binds to the URS1 element. The complex represses transcription at the promoters containing URS1, as illustrated in Figure 23.15 (3312). Rpd3 has histone deacetylase activity; we do not know whether the function of Sin3 is just to bring Rpd3 to the promoter or whether it has an additional role in repression.

Figure 23.15 A repressor complex contains three components: a DNA binding subunit, a corepressor, and a histone deacetylase.

A similar system for repression is found in mammalian cells (3313; 3314). The bHLH family of transcription regulators includes activators that function as heterodimers, including MyoD (see Molecular Biology 5.22.15 Helix-loop-helix proteins interact by combinatorial association). It also includes repressors, in particular the heterodimer Mad:Max, where Mad can be any one of a group of closely related proteins. The Mad:Max heterodimer (which binds to specific DNA sites) interacts with a homologue of Sin3 (called mSin3 in mouse and hSin3 in man). mSin3 is part of a repressive complex that includes histone binding proteins and the histone deacetylases HDAC1 and HDAC2. Deacetylase activity is required for repression. The modular nature of this system is emphasized by other means of Deacetylases are associated with repressors | SECTION 5.23.8 © 2004. Virtual Text / www.ergito.com

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employment: a corepressor (SMRT), which enables retinoid hormone receptors to repress certain target genes, functions by binding mSin3, which in turns brings the HDAC activities to the site. Another means of bringing HDAC activities to the site may be a connection with MeCP2, a protein that binds to methylated cytosines (see Molecular Biology 5.21.19 CpG islands are regulatory targets). Absence of histone acetylation is also a feature of heterochromatin. This is true of both constitutive heterochromatin (typically involving regions of centromeres or telomeres) and facultative heterochromatin (regions that are inactivated in one cell although they may be active in another). Typically the N-terminal tails of histones H3 and H4 are not acetylated in heterochromatic regions (for review see 2414). Last updated on 9-5-2001

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Reviews 2414. Richards, E. J., Elgin, S. C., and Richards, S. C. (2002). Epigenetic codes for heterochromatin formation and silencing: rounding up the usual suspects. Cell 108, 489-500.

References 3312. Kadosh, D. and Struhl, K. (1997). Repression by Ume6 involves recruitment of a complex containing Sin3 corepressor and Rpd3 histone deacetylase to target promoters. Cell 89, 365-371. 3313. Ayer, D. E., Lawrence, Q. A., and Eisenman, R. N. (1995). Mad-Max transcriptional repression is mediated by ternary complex formation with mammalian homologs of yeast repressor Sin3. Cell 80, 767-776. 3314. Schreiber-Agus, N., Chin, L., Chen, K., Torres, R., Rao, G., Guida, P., Skoultchi, A. I., and DePinho, R. A. (1995). An amino-terminal domain of Mxi1 mediates anti-Myc oncogenic activity and interacts with a homolog of the yeast transcriptional repressor SIN3. Cell 80, 777-786.


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5.23.9 Methylation of histones and DNA is connected Key Concepts

• Methylation of both DNA and histones is a feature of inactive chromatin. • The two types of methylation event may be connected.

Methylation of both histones and DNA is associated with inactivity (for review see 2414). Sites that are methylated in histones include two lysines in the tail of H3 and an arginine in the tail of H4. Methylation of H3 9Lys is a feature of condensed regions of chromatin, including heterochromatin as seen in bulk and also smaller regions that are known not to be expressed. The histone methyltransferase enzyme that targets this lysine is called SUV39H1 (2014). (We see the origin of this peculiar name in Molecular Biology 5.23.14 Some common motifs are found in proteins that modify chromatin). Its catalytic site has a region called the SET domain. Other histone methyltransferases act on arginine (for review see 2237). In addition, methylation may occur on 79Lys in the globular core region of H3; this may be necessary for the formation of heterochromatin at telomeres (3215). Most of the methylation sites in DNA are CpG islands (see Molecular Biology 5.21.19 CpG islands are regulatory targets). CpG sequences in heterochromatin are usually methylated. Conversely, it is necessary for the CpG islands located in promoter regions to be unmethylated in order for a gene to be expressed (see Molecular Biology 5.21.18 Gene expression is associated with demethylation). Methylation of DNA and methylation of histones may be connected. Some histone methyltransferase enzymes contain potential binding sites for the methylated CpG doublet, raising the possibility that a methylated DNA sequence may cause a histone methyltransferase to bind. A possible connection in the opposite direction is indicated by the fact that in the fungus Neurospora, the methylation of DNA is prevented by a mutation in a gene coding for a histone methylase that acts on 9Lys of histone H3 (2183). This suggests that methylation of the histone is a signal involved in recruiting the DNA methylase to chromatin. The important point is not the detailed order of events – which remains to be worked out – but the fact that one type of modification can be the trigger for another. Last updated on 1-3-2003

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Reviews 2237. Zhang, Y. and Reinberg, D. (2001). Transcription regulation by histone methylation: interplay between different covalent modifications of the core histone tails. Genes Dev. 15, 2343-2360. 2414. Richards, E. J., Elgin, S. C., and Richards, S. C. (2002). Epigenetic codes for heterochromatin formation and silencing: rounding up the usual suspects. Cell 108, 489-500.

References 2014. Rea, S., Eisenhaber, F., O'Carroll, D., Strahl, B. D., Sun, Z. W., Sun, M., Opravil, S., Mechtler, K., Ponting, C. P., Allis, C. D., and Jenuwein, T. (2000). Regulation of chromatin structure by site-specific histone H3 methyltransferases. Nature 406, 593-599. 2183. Tamaru, H. and Selker, E. U. (2001). A histone H3 methyltransferase controls DNA methylation in Neurospora crassa. Nature 414, 277-283. 3215. Ng, H. H., Feng, Q., Wang, H., Erdjument-Bromage, H., Tempst, P., Zhang, Y., and Struhl, K. (2002). Lysine methylation within the globular domain of histone H3 by Dot1 is important for telomeric silencing and Sir protein association. Genes Dev. 16, 1518-1527.


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5.23.10 Chromatin states are interconverted by modification Key Concepts

• Acetylation of histones is associated with gene activation. • Methylation of DNA and of histones is associated with heterochromatin.

Figure 23.16 summarizes three types of differences that are found between active chromatin and inactive chromatin:

Figure 23.16 Acetylation of histones activates chromatin, and methylation of DNA and histones inactivates chromatin.

• Active chromatin is acetylated on the tails of histones H3 and H4. • Inactive chromatin is methylated on 9Lys of histone H3. • Inactive chromatin is methylated on cytosines of CpG doublets. The reverse types of events occur if we compare the activation of a promoter with the generation of heterochromatin. The actions of the enzymes that modify chromatin Chromatin states are interconverted by modification | SECTION 5.23.10 © 2004. Virtual Text / www.ergito.com

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ensure that activating events are mutually exclusive with inactivating events. Methylation of H3 9Lys and acetylation of H3 14Lys are mutually antagonistic. Acetylases and deacetylases may trigger the initiating events. Deacetylation allows methylation to occur, which causes formation of a heterochromatic complex (see Molecular Biology 5.23.15 Heterochromatin depends on interactions with histones). Acetylation marks a region as active (see Molecular Biology 5.23.11 Promoter activation involves an ordered series of events). This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.23.10

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5.23.11 Promoter activation involves an ordered series of events Key Concepts

• The remodeling complex may recruit the acetylating complex. • Acetylation of histones may be the event that maintains the complex in the activated state.

How are acetylases (or deacetylases) recruited to their specific targets? As we have seen with remodeling complexes, the process is likely to be indirect. A sequence-specific activator (or repressor) may interact with a component of the acetylase (or deacetylase) complex to recruit it to a promoter. There may also be direct interactions between remodeling complexes and histone-modifying complexes. Binding by the SWI/SNF remodeling complex may lead in turn to binding by the SAGA acetylase complex (1966). Acetylation of histones may then in fact stabilize the association with the SWI/SNF complex, making a mutual reinforcement of the changes in the components at the promoter (1967). We can connect all of the events at the promoter into the series summarized in Figure 23.17. The initiating event is binding of a sequence-specific component (which is able to find its target DNA sequence in the context of chromatin). This recruits a remodeling complex. Changes occur in nucleosome structure. An acetylating complex binds, and the acetylation of target histones provides a covalent mark that the locus has been activated.

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Figure 23.17 Promoter activation involves binding of a sequence-specific activator, recruitment and action of a remodeling complex, and recruitment and action of an acetylating complex.

Modification of DNA also occurs at the promoter. Methylation of cytosine at CpG doublets is associated with gene inactivity (see Molecular Biology 5.21.18 Gene expression is associated with demethylation). The basis for recognition of DNA as a target for methylation is not very well established (see Molecular Biology 5.23.20 DNA methylation is responsible for imprinting). It is clear that chromatin remodeling at the promoter requires a variety of changes that affect nucleosomes, including acetylation, but what changes are required within the gene to allow an RNA polymerase to traverse it? We know that RNA polymerase can transcribe DNA in vitro at rates comparable to the in vivo rate (~25 nucleotides per second) only with template of free DNA. Several proteins have been characterized for their abilities to improve the speed with which RNA polymerase transcribes chromatin in vivo (1427; 1428; 1429). The common feature is that they Promoter activation involves an ordered series of events | SECTION 5.23.11 © 2004. Virtual Text / www.ergito.com

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act on chromatin (for review see 1426). A current model for their action is that they associate with RNA polymerase and travel with it along the template, modifying nucleosome structure by acting on histones. Among these factors are histone acetylases. One possibility is that the first RNA polymerase to transcribe a gene is a pioneer polymerase carrying factors that change the structure of the transcription unit so as to make it easier for subsequent polymerases. Last updated on 12-5-2001


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Reviews 1426. Orphanides, G. and Reinberg, D. (2000). RNA polymerase II elongation through chromatin. Nature 407, 471-475.

References 1427. Orphanides, G., LeRoy, G., Chang, C. H., Luse, D. S., and Reinberg, D. (1998). FACT, a factor that facilitates transcript elongation through nucleosomes. Cell 92, 105-116. 1428. Wada, T., Takagi, T., Yamaguchi, Y., Ferdous, A., Imai, T., Hirose, S., Sugimoto, S., Yano, K., Hartzog, G. A., Winston, F., Buratowski, S., and Handa, H. (1998). DSIF, a novel transcription elongation factor that regulates RNA polymerase II processivity, is composed of human Spt4 and Spt5 homologs. Genes Dev. 12, 343-356. 1429. Bortvin, A. and Winston, F. (1996). Evidence that Spt6p controls chromatin structure by a direct interaction with histones. Science 272, 1473-1476. 1966. Cosma, M. P., Tanaka, T., and Nasmyth, K. (1999). Ordered recruitment of transcription and chromatin remodeling factors to a cell cycle- and developmentally regulated promoter. Cell 97, 299-311. 1967. Hassan, A. H., Neely, K. E., and Workman, J. L. (2001). Histone acetyltransferase complexes stabilize swi/snf binding to promoter nucleosomes. Cell 104, 817-827.



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5.23.12 Histone phosphorylation affects chromatin structure Key Concepts

• At least two histones are targets for phosphorylation, possibly with opposing effects.

Histones are phosphorylated in two circumstances: • cyclically during the cell cycle; • and in association with chromatin remodeling. It is has been known for a very long time that histone H1 is phosphorylated at mitosis, and more recently it was discovered that H1 is an extremely good substrate for the Cdc2 kinase that controls cell division. This led to speculations that the phosphorylation might be connected with the condensation of chromatin, but so far no direct effect of this phosphorylation event has been demonstrated, and we do not know whether it plays a role in cell division (see Molecular Biology 6.29.7 Protein phosphorylation and dephosphorylation control the cell cycle). Loss of a kinase that phosphorylates histone H3 on 10Ser has devastating effects on chromatin structure. Figure 23.18 compares the usual extended structure of the polytene chromosome set of D. melanogaster (upper photograph) with the structure that is found in a null mutant that has no JIL-1 kinase (lower photograph). The absence of JIL-1 is lethal, but the chromosomes can be visualized in the larvae before they die (2192).

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Figure 23.18 Polytene chromosomes of flies that have no JIL-1 kinase have abnormal polytene chromosomes that are condensed instead of extended. Photograph kindly provided by Kristen M. Johansen.

The cause of the disruption of structure is most likely the failure to phosphorylate histone H3 (of course, JIL-1 may also have other targets). This suggests that H3 phosphorylation is required to generate the more extended chromosome structure of euchromatic regions. Evidence supporting the idea that JIL-1 acts directly on chromatin is that it associates with the complex of proteins that binds to the X chromosome to increase its gene expression in males (see Molecular Biology 5.23.17 X chromosomes undergo global changes). This leaves us with somewhat conflicting impressions of the roles of histone phosphorylation. If it is important in the cell cycle, it is likely to be as a signal for condensation. Its effect in chromatin remodeling appears to be the opposite. It is of course possible that phosphorylation of different histones, or even of different amino acid residues in one histone, has opposite effects on chromatin structure. Last updated on 12-10-2001


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References 2192. Wang, Y., Zhang, W., Jin, Y., Johansen, J., and Johansen, K. M. (2001). The JIL-1 tandem kinase mediates histone H3 phosphorylation and is required for maintenance of chromatin structure in Drosophila. Cell 105, 433-443.



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5.23.13 Heterochromatin propagates from a nucleation event Key Terms Epigenetic changes influence the phenotype without altering the genotype. They consist of changes in the properties of a cell that are inherited but that do not represent a change in genetic information. Position effect variegation (PEV) is silencing of gene expression that occurs as the result of proximity to heterochromatin. Telomeric silencing describes the repression of gene activity that occurs in the vicinity of a telomere. Key Concepts

• Heterochromatin is nucleated at a specific sequence and the inactive structure propagates along the chromatin fiber.

• Genes within regions of heterochromatin are inactivated. • Because the length of the inactive region varies from cell to cell, inactivation of genes in this vicinity causes position effect variegation.

• Similar spreading effects occur at telomeres and at the silent cassettes in yeast mating type.

An interphase nucleus contains both euchromatin and heterochromatin. The condensation state of heterochromatin is close to that of mitotic chromosomes. Heterochromatin is inert. It remains condensed in interphase, is transcriptionally repressed, replicates late in S phase, and may be localized to the nuclear periphery. Centromeric heterochromatin typically consists of satellite DNAs. However, the formation of heterochromatin is not rigorously defined by sequence. When a gene is transferred, either by a chromosomal translocation or by transfection and integration, into a position adjacent to heterochromatin, it may become inactive as the result of its new location, implying that it has become heterochromatic. Such inactivation is the result of an epigenetic effect (see Molecular Biology 5.23.22 Epigenetic effects can be inherited). It may differ between individual cells in an animal, and results in the phenomenon of position effect variegation (PEV), in which genetically identical cells have different phenotypes. This has been well characterized in Drosophila. Figure 23.19 shows an example of position effect variegation in the fly eye, in which some regions lack color while others are red, because the white gene is inactivated by adjacent heterochromatin in some cells, while it remained active in other cells.

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Figure 23.19 Position effect variegation in eye color results when the white gene is integrated near heterochromatin. Cells in which white is inactive give patches of white eye, while cells in which white is active give red patches. The severity of the effect is determined by the closeness of the integrated gene to heterochromatin. Photograph kindly provided by Steve Henikoff.

The explanation for this effect is shown in Figure 23.20. Inactivation spreads from heterochromatin into the adjacent region for a variable distance. In some cells it goes far enough to inactivate a nearby gene, but in others it does not. This happens at a certain point in embryonic development, and after that point the state of the gene is inherited by all the progeny cells. Cells descended from an ancestor in which the gene was inactivated form patches corresponding to the phenotype of loss-of-function (in the case of white, absence of color).


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Figure 23.20 Extension of heterochromatin inactivates genes. The probability that a gene will be inactivated depends on its distance from the heterochromatin region.

The closer a gene lies to heterochromatin, the higher the probability that it will be inactivated. This suggests that the formation of heterochromatin may be a two-stage process: a nucleation event occurs at a specific sequence; and then the inactive structure propagates along the chromatin fiber. The distance for which the inactive structure extends is not precisely determined, and may be stochastic, being influenced by parameters such as the quantities of limiting protein components. One factor that may affect the spreading process is the activation of promoters in the region; an active promoter may inhibit spreading (2191). Genes that are closer to heterochromatin are more likely to be inactivated, and will therefore be inactive in a greater proportion of cells. On this model, the boundaries of a heterochromatic region might be terminated by exhausting the supply of one of the proteins that is required. The effect of telomeric silencing in yeast is analogous to position effect variegation in Drosophila; genes translocated to a telomeric location show the same sort of variable loss of activity. This results from a spreading effect that propagates from the telomeres. A second form of silencing occurs in yeast. Yeast mating type is determined by the activity of a single active locus (MAT), but the genome contains two other copies of the mating type sequences (HML and HMR), which are maintained in an inactive form. The silent loci HML and HMR share many properties with heterochromatin, and could be regarded as constituting regions of heterochromatin in miniature (see Molecular Biology 4.18.7 Silent cassettes at HML and HMR are repressed). Heterochromatin propagates from a nucleation event | SECTION 5.23.13 © 2004. Virtual Text / www.ergito.com

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References 2191. Ahmad, K. and Henikoff, S. (2001). Modulation of a transcription factor counteracts heterochromatic gene silencing in Drosophila. Cell 104, 839-847.



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5.23.14 Some common motifs are found in proteins that modify chromatin Key Concepts

• The chromo domain is found in several chromatin proteins that have either activating or repressing effects on gene expression

• The SET domain is part of the catalytic site of protein methyltransferases.

Our insights into the molecular mechanisms for controlling the structure of chromatin start with mutants that affect position effect variegation. Some 30 genes have been identified in Drosophila. They are named systematically as Su(var) for genes whose products act to suppress variegation and E(var) for genes whose products enhance variegation. Remember that the genes were named for the behavior of the mutant loci. Su(var) mutations lie in genes whose products are needed for the formation of heterochromatin. They include enzymes that act on chromatin, such as histone deacetylases, and proteins that are localized to heterochromatin. E(var) mutations lie in genes whose products are needed to activate gene expression. They include members of the SWI/SNF complex. We see immediately from these properties that modification of chromatin structure is important for controlling the formation of heterochromatin. The universality of these mechanisms is indicated by the fact that many of these loci have homologues in yeast that display analogous properties. Some of the homologues in S. pombe are clr (cryptic loci regulator) genes, in which mutations affect silencing. Many of the Su(var) and E(var) proteins have a common protein motif of 60 amino acids called the chromo domain. The fact that this domain is found in proteins of both groups suggests that it represents a motif that participates in protein-protein interactions with targets in chromatin (for summary see 2026). Chromo domain(s) are mostly responsible for targeting proteins to heterochromatin. They function by recognizing methylated lysines in histone tails (see Molecular Biology 5.23.15 Heterochromatin depends on interactions with histones and Molecular Biology 5.23.16 Polycomb and trithorax are antagonistic repressors and activators) Su(var)3-9 has a chromo domain and also a SET domain, a motif that is found in several Su(var) proteins. Its mammalian homologues localize to centromeric heterochromatin. It is the histone methyltransferase that acts on 9Lys of histone H3 (see Molecular Biology 5.23.9 Methylation of histones and DNA is connected). The SET domain is part of the active site, and in fact is a marker for the methylase activity. The bromo domain is found in a variety of proteins that interact with chromatin, including histone acetylases. The crystal structure shows that it has a binding site for acetylated lysine (2028). The bromo domain itself recognizes only a very short sequence of 4 amino acids including the acetylated lysine, so specificity for target Some common motifs are found in proteins that modify chromatin | SECTION 5.23.14 © 2004. Virtual Text / www.ergito.com

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recognition must depend on interactions involving other regions (2032). Besides the acetylases, the bromo domain is found in a range of proteins that interact with chromatin, including components of the transcription apparatus. This implies that it is used to recognize acetylated histones, which means that it is likely to be found in proteins that are involved with gene activation. Although there is a general correlation in which active chromatin is acetylated while inactive chromatin is methylated on histones, there are some exceptions to the rule. The best characterized is that acetylation of 12Lys of H4 is associated with heterochromatin (616). Multiple modifications may occur on the same histone tail, and one modification may influence another. Phosphorylation of a lysine at one position may be necessary for acetylation of a lysine at another position. Figure 23.21 shows the situation in the tail of H3, which can exist in either of two alternative states. The inactive state has Methyl-9Lys. The active state has Acetyl-9Lys and Phospho-10Ser. These states can be maintained over extended regions of chromatin (2025). The phosphorylation of 10 Ser and the methylation of 9Lysare mutually inhibitory, suggesting the order of events shown in the figure. This situation may cause the tail to flip between the active and active states.

Figure 23.21 Multiple modifications in the H3 tail affect chromatin activity.


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References 616. Turner, B. M., Birley, A. J., and Lavender, J. (1992). Histone H4 isoforms acetylated at specific lysine residues define individual chromosomes and chromatin domains in Drosophila polytene nuclei. Cell 69, 375-384. 2025. Litt, M. D., Simpson, M., Gaszner, M., Allis, C. D., and Felsenfeld, G. (2001). Correlation between histone lysine methylation and developmental changes at the chicken beta-globin locus. Science 293, 2453-2455. 2026. Koonin, E. V., Zhou, S., and Lucchesi, J. C. (1995). The chromo superfamily: new members, duplication of the chromo domain and possible role in delivering transcription regulators to chromatin. Nucleic Acids Res. 23, 4229-4233. 2028. Dhalluin, C., Carlson, J. E., Zeng, L., He, C., Aggarwal, A. K., and Zhou, M. M. (1999). Structure and ligand of a histone acetyltransferase bromodomain. Nature 399, 491-496. 2032. Owen, D. J., Ornaghi, P., Yang, J. C., Lowe, N., Evans, P. R., Ballario, P., Neuhaus, D., Filetici, P., and Travers, A. A. (2000). The structural basis for the recognition of acetylated histone H4 by the bromodomain of histone acetyltransferase Gcn5p. EMBO J. 19, 6141-6149.


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5.23.15 Heterochromatin depends on interactions with histones Key Concepts

• HP1 is the key protein in forming mammalian heterochromatin, and acts by binding to methylated H3 histone.

• RAP1 initiates formation of heterochromatin in yeast by binding to specific target sequences in DNA.

• The targets of RAP1 include telomeric repeats and silencers at HML and HMR. • RAP1 recruits SIR3/SIR4, which interact with the N-terminal tails of H3 and H4.

Inactivation of chromatin occurs by the addition of proteins to the nucleosomal fiber. The inactivation may be due to a variety of effects, including condensation of chromatin to make it inaccessible to the apparatus needed for gene expression, addition of proteins that directly block access to regulatory sites, or proteins that directly inhibit transcription. Two systems that have been characterized at the molecular level involve HP1 in mammals and the SIR complex in yeast (for review see 2420). Although there are no detailed similarities between the proteins involved in each system, the general mechanism of reaction is similar: the points of contact in chromatin are the N-terminal tails of the histones. HP1 (heterochromatin protein 1) is one of most important Su(var) proteins. This was originally identified as a protein that is localized to heterochromatin by staining polytene chromosomes with an antibody directed against the protein (2021). It was later shown to be the product of the gene Su(var)2-5 (2022). Its homologue in the yeast S. pombe is coded by swi6. The original protein identified as HP1 is now called HP1 α, since two related proteins, HP1 β and HP1 γ, have since been found. HP1 contains a chromo domain near the N-terminus, and another domain that is related to it, called the chromo-shadow domain, at the C-terminus (see Figure 23.23). The importance of the chromo domain is indicated by the fact that it is the location of many of the mutations in HP1 (2023). Mutation of a deacetylase that acts on the H3 Ac-14Lys prevents the methylation at Lys (2238). H3 that is methylated at 9Lys binds the protein HP1 via the chromo domain (2016, 2015, (4178, 4180). This suggests the model for initiating formation of heterochromatin shown in Figure 23.22. First the deacetylase acts to remove the modification at 14Lys. Then the SUV39H1 methylase acts on the histone H3 tail to create the methylated signal to which HP1 will bind. Figure 23.23 expands the reaction to show that the interaction occurs between the chromo domain and the methylated lysine. This is a trigger for forming inactive chromatin. Figure 23.24 9

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shows that the inactive region may then be extended by the ability of further HP1 molecules to interact with one another (for review see 2237).

Figure 23.22 SUV39H1 is a histone methyltransferase that acts on 9Lys of histone H3. HP1 binds to the methylated histone.

Figure 23.23 Methylation of histone H3 creates a binding site for HP1.


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Figure 23.24 Binding of HP1 to methylated histone H3 forms a trigger for silencing because further molecules of HP1 aggregate on the nucleosome chain.

The existence of a common basis for silencing in yeast is suggested by its reliance on a common set of genetic loci. Mutations in any one of a number of genes cause HML and HMR to become activated, and also relieve the inactivation of genes that have been integrated near telomeric heterochromatin. The products of these loci therefore function to maintain the inactive state of both types of heterochromatin (for review see 210, 4530). Figure 23.25 proposes a model for actions of these proteins. Only one of them is a sequence-specific DNA-binding protein. This is RAP1, which binds to the C1-3A repeats at the telomeres, and also binds to the cis-acting silencer elements that are needed for repression of HML and HMR (619). The proteins SIR3 and SIR4 interact with RAP1 and also with one another (they may function as a heteromultimer). SIR3/SIR4 interact with the N-terminal tails of the histones H3 and H4. [In fact, the first evidence that histones might be involved directly in formation of heterochromatin was provided by the discovery that mutations abolishing silencing at HML/HMR map to genes coding for H3 and H4 (620; for review see 209)].


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Figure 23.25 Formation of heterochromatin is initiated when RAP1 binds to DNA. SIR3/4 bind to RAP1 and also to histones H3/H4. The complex polymerizes along chromatin and may connect telomeres to the nuclear matrix.

RAP1 has the crucial role of identifying the DNA sequences at which heterochromatin forms. It recruits SIR3/SIR4, and they interact directly with the histones H3/H4 (622; 1214). Once SIR3/SIR4 have bound to histones H3/H4, the complex may polymerize further, and spread along the chromatin fiber. This may inactivate the region, either because coating with SIR3/SIR4 itself has an inhibitory effect, or because binding to histones H3/H4 induces some further change in structure. We do not know what limits the spreading of the complex. The C-terminus of SIR3 has a similarity to nuclear lamin proteins (constituents of the nuclear matrix) and may be responsible for tethering heterochromatin to the nuclear periphery (621; 623). A similar series of events forms the silenced regions at HMR and HML (see also Molecular Biology 4.18.7 Silent cassettes at HML and HMR are repressed). Three sequence-specific factors are involved in triggering formation of the complex: RAP1, ABF1 (a transcription factor), and ORC (the origin replication complex). In this case, SIR1 binds to a sequence-specific factor and recruits SIR2,3,4 to form the repressive structure. SIR2 is a histone deacetylase (2416; 2418; 2419). The deacetylation Heterochromatin depends on interactions with histones | SECTION 5.23.15 © 2004. Virtual Text / www.ergito.com

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reaction is necessary to maintain binding of the SIR complex to chromatin (for review see 2420). How does a silencing complex repress chromatin activity? It could condense chromatin so that regulator proteins cannot find their targets. The simplest case would be to suppose that the presence of a silencing complex is mutually incompatible with the presence of transcription factors and RNA polymerase. The cause could be that silencing complexes block remodeling (and thus indirectly prevent factors from binding) or that they directly obscure the binding sites on DNA for the transcription factors. However, the situation may not be this simple, because transcription factors and RNA polymerase can be found at promoters in silenced chromatin (1963). This could mean that the silencing complex prevents the factors from working rather than from binding as such. In fact, there may be competition between gene activators and the repressing effects of chromatin, so that activation of a promoter inhibits spread of the silencing complex (2191). Another specialized chromatin structure forms at the centromere. Its nature is suggested by the properties of an S. cerevisiae mutation, cse4, that disrupts the structure of the centromere. Cse4p is a protein that is related to histone H3. A mammalian centromeric protein, CENP-A, has a related sequence. Genetic interactions between cse4 and CDE-II, and between cse4 and a mutation in the H4 histone gene, suggest that a histone octamer may form around a core of Cse4p-H4, and then the centromeric complexes CBF1 and CBF3 may attach to form the centromere (591; 624). The centromere may then be associated with the formation of heterochromatin in the region. In human cells, the centromere-specific protein CENP-B is required to initiate modifications of histone H3 (deacetylation of 9Lys and 14Lys, followed by methylation of 9Lys) that trigger an association with the protein Swi6 that leads to the formation of heterochromatin in the region (3218). Last updated on 9-24-2003


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Reviews 209. Thompson, J. S., Hecht, A., and Grunstein, M. (1993). Histones and the regulation of heterochromatin in yeast. Cold Spring Harbor Symp. Quant. Biol. 58, 247-256. 210. Loo, S. and Rine, J. (1995). Silencing and heritable domains of gene expression. Annu. Rev. Cell Dev. Biol. 11, 519-548. 2237. Zhang, Y. and Reinberg, D. (2001). Transcription regulation by histone methylation: interplay between different covalent modifications of the core histone tails. Genes Dev. 15, 2343-2360. 2420. Moazed, D. (2001). Common themes in mechanisms of gene silencing. Mol. Cell 8, 489-498. 4530. Rusche, L. N., Kirchmaier, A. L., and Rine, J. (2003). The establishment, inheritance, and function of silenced chromatin in Saccharomyces cerevisiae. Annu. Rev. Biochem. 72, 481-516.


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References 591. Bloom, K. S. and Carbon, J. (1982). Yeast centromere DNA is in a unique and highly ordered structure in chromosomes and small circular minichromosomes. Cell 29, 305-317. 619. Shore, D. and Nasmyth, K. (1987). Purification and cloning of a DNA-binding protein from yeast that binds to both silencer and activator elements. Cell 51, 721-732. 620. Kayne, P. S., Kim, U. J., Han. M., Mullen, R. J., Yoshizaki, F., and Grunstein, M. (1988). Extremely conserved histone H4 N terminus is dispensable for growth but essential for repressing the silent mating loci in yeast. Cell 55, 27-39. 621. Palladino, F., Laroche, T., Gilson, E., Axelrod, A., Pillus, L., and Gasser, S. M. (1993). SIR3 and SIR4 proteins are required for the positioning and integrity of yeast telomeres. Cell 75, 543-555. 622. Moretti, P., Freeman, K., Coodly, L., and Shore, D. (1994). Evidence that a complex of SIR proteins interacts with the silencer and telomere-binding protein RAP1. Genes Dev. 8, 2257-2269. 623. Hecht, A., Laroche, T., Strahl-Bolsinger, S., Gasser, S. M., and Grunstein, M. (1995). Histone H3 and H4 N-termini interact with the silent information regulators SIR3 and SIR4: a molecular model for the formation of heterochromatin in yeast. Cell 80, 583-592. 624. Meluh, P. B. et al. (1998). Cse4p is a component of the core centromere of S. cerevisiae. Cell 94, 607-613. 1214. Manis, J. P., Gu, Y., Lansford, R., Sonoda, E., Ferrini, R., Davidson, L., Rajewsky, K., and Alt, F. W. (1998). Ku70 is required for late B cell development and immunoglobulin heavy chain class switching. J. Exp. Med. 187, 2081-2089. 1963. Sekinger, E. A. and Gross, D. S. (2001). Silenced chromatin is permissive to activator binding and PIC recruitment. Cell 105, 403-414. 2015. Bannister, A. J., Zegerman, P., Partridge, J. F., Miska, E. A., Thomas, J. O., Allshire, R. C., and Kouzarides, T. (2001). Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature 410, 120-124. 2016. Lachner, M., O'Carroll, D., Rea, S., Mechtler, K., and Jenuwein, T. (2001). Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature 410, 116-120. 2021. James, T. C. and Elgin, S. C. (1986). Identification of a nonhistone chromosomal protein associated with heterochromatin in D. melanogaster and its gene. Mol. Cell Biol. 6, 3862-3872. 2022. Eissenberg, J. C., Morris, G. D., Reuter, G., and Hartnett, T. (1992). The heterochromatin-associated protein HP-1 is an essential protein in Drosophila with dosage-dependent effects on position-effect variegation. Genetics 131, 345-352. 2023. Platero, J. S., Hartnett, T., and Eissenberg, J. C. (1995). Functional analysis of the chromo domain of HP1. EMBO J. 14, 3977-3986. 2191. Ahmad, K. and Henikoff, S. (2001). Modulation of a transcription factor counteracts heterochromatic gene silencing in Drosophila. Cell 104, 839-847. 2238. Nakayama , J., Rice, J. C., Strahl, B. D., Allis, C. D., and Grewal, S. I. (2001). Role of histone H3 lysine 9 methylation in epigenetic control of heterochromatin assembly. Science 292, 110-113. 2416. Imai, S., Armstrong, C. M., Kaeberlein, M., and Guarente, L. (2000). Transcriptional silencing and longevity protein Sir2 is an NAD-dependent histone deacetylase. Nature 403, 795-800. 2418. Smith, J. S., Brachmann, C. B., Celic, I., Kenna, M. A., Muhammad, S., Starai, V. J., Avalos, J. L., Escalante-Semerena, J. C., Grubmeyer, C., Wolberger, C., and Boeke, J. D. (2000). A phylogenetically conserved NAD+-dependent protein deacetylase activity in the Sir2 protein


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family. Proc. Natl. Acad. Sci. USA 97, 6658-6663. 2419. Landry, J., Sutton, A., Tafrov, S. T., Heller, R. C., Stebbins, J., Pillus, L., and Sternglanz, R. (2000). The silencing protein SIR2 and its homologs are NAD-dependent protein deacetylases. Proc. Natl. Acad. Sci. USA 97, 5807-5811. 3218. Nakagawa, H., Lee, J. K., Hurwitz, J., Allshire, R. C., Nakayama, J., Grewal, S. I., Tanaka, K., and Murakami, Y. (2002). Fission yeast CENP-B homologs nucleate centromeric heterochromatin by promoting heterochromatin-specific histone tail modifications. Genes Dev. 16, 1766-1778. 4178. Cheutin, T., McNairn, A. J., Jenuwein, T., Gilbert, D. M., Singh, P. B., and Misteli, T. (2003). Maintenance of stable heterochromatin domains by dynamic HP1 binding. Science 299, 721-725. 4180. Schotta, G., Ebert, A., Krauss, V., Fischer, A., Hoffmann, J., Rea, S., Jenuwein, T., Dorn, R., and Reuter, G. (2002). Central role of Drosophila SU(VAR)3-9 in histone H3-K9 methylation and heterochromatic gene silencing. EMBO J. 21, 1121-1131.



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5.23.16 Polycomb and trithorax are antagonistic repressors and activators Key Concepts

• Polycomb group proteins (Pc-G) perpetuate a state of repression through cell divisions.

• The PRE is a DNA sequence that is required for the action of Pc-G. • The PRE provides a nucleation center from which Pc-G proteins propagate an inactive structure.

• No individual Pc-G protein has yet been found that can bind the PRE. • Trithorax group proteins antagonize the actions of the Pc-G.

Heterochromatin provides one example of the specific repression of chromatin. Another is provided by the genetics of homeotic genes in Drosophila, which have led to the identification of a protein complex that may maintain certain genes in a repressed state. Pc mutants show transformations of cell type that are equivalent to gain-of-function mutations in the genes Antennapedia (Antp) or Ultrabithorax, because these genes are expressed in tissues in which usually they are repressed. This implicates Pc in regulating transcription. Furthermore, Pc is the prototype for a class of loci called the Pc group (Pc-G); mutations in these genes generally have the same result of derepressing homeotic genes, suggesting the possibility that the group of proteins has some common regulatory role. A connection between chromatin remodeling and repression is indicated by the properties of brahma, a fly counterpart to SWI2, which codes for component of the SWI/SNF remodeling complex. Loss of brahma function suppresses mutations in Polycomb. Consistent with the pleiotropy of Pc mutations, Pc is a nuclear protein that can be visualized at ~80 sites on polytene chromosomes. These sites include the Antp gene. Another member of the Pc-G, polyhomeotic, is visualized at a set of polytene chromosome bands that are identical with those bound by Pc. The two proteins coimmunoprecipitate in a complex of ~2.5 × 106 D that contains 10-15 polypeptides. The relationship between these proteins and the products of the ~30 Pc-G genes remains to be established. One possibility is that some of these gene products form a general repressive complex,and then some of the other proteins associate with it to determine its specificity (683; 697). The Pc-G proteins are not conventional repressors. They are not responsible for determining the initial pattern of expression of the genes on which they act. In the absence of Pc-G proteins, these genes are initially repressed as usual, but later in development the repression is lost without Pc-G group functions. This suggests that the Pc-G proteins in some way recognize the state of repression when it is established, and they then act to perpetuate it through cell division of the daughter Polycomb and trithorax are antagonistic repressors and activators | SECTION 5.23.16 © 2004. Virtual Text / www.ergito.com

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cells. Figure 23.26 shows a model in which Pc-G proteins bind in conjunction with a repressor, but the Pc-G proteins remain bound after the repressor is no longer available. This is necessary to maintain repression, so that if Pc-G proteins are absent, the gene becomes activated (698).

Figure 23.26 Pc-G proteins do not initiate repression, but are responsible for maintaining it.

A region of DNA that is sufficient to enable the response to the Pc-G genes is called a PRE (Polycomb response element). It can be defined operationally by the property that it maintains repression in its vicinity throughout development. The assay for a PRE is to insert it close to a reporter gene that is controlled by an enhancer that is repressed in early development, and then to determine whether the reporter becomes expressed subsequently in the descendants. An effective PRE will prevent such re-expression (700). The PRE is a complex structure, ~10 kb. No individual member of the Pc-G proteins has yet been shown to bind to specific sequences in the PRE, so the basis for the assembly of the complex is still unknown. When a locus is repressed by Pc-G proteins, however, the proteins appear to be present over a much larger length of DNA than the PRE itself. Polycomb is found locally over a few kilobases of DNA surrounding a PRE. This suggests that the PRE may provide a nucleation center, from which a structural state depending on Pc-G proteins may propagate. This model is supported by the observation of effects related to position effect variegation (see Figure 23.20), that Polycomb and trithorax are antagonistic repressors and activators | SECTION 5.23.16 © 2004. Virtual Text / www.ergito.com

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is, a gene near to a locus whose repression is maintained by Pc-G may become heritably inactivated in some cells but not others. In one typical situation, crosslinking experiments in vivo showed that Pc protein is found over large regions of the bithorax complex that are inactive, but the protein is excluded from regions that contain active genes. The idea that this could be due to cooperative interactions within a multimeric complex is supported by the existence of mutations in Pc that change its nuclear distribution and abolish the ability of other Pc-G members to localize in the nucleus. The role of Pc-G proteins in maintaining, as opposed to establishing, repression must mean that the formation of the complex at the PRE also depends on the local state of gene expression (699). A connection between the Pc-G complex and more general structural changes in chromatin is suggested by the inclusion of a chromo domain in Pc. (In fact, the chromo domain was first identified as a region of homology between Pc and the protein HP1 found in heterochromatin.) Since variegation is caused by the spreading of inactivity from constitutive heterochromatin, it is likely that the chromo domain is used by Pc and HP1 in a similar way to induce the formation of heterochromatic or inactive structures (see Molecular Biology 5.23.14 Some common motifs are found in proteins that modify chromatin). The chromo domain of Pc binds to 27Lys on H3 (4179) (analogous to HP1's use of its chromo domain to bind to 9Lys). This model implies that similar mechanisms are used to repress individual loci or to create heterochromatin. This is probably how the PRC-1 (Polycomb-repressive complex)works. Another complex that contains Polycomb, the Esc-E(z) complex, has a histone methyltransferase activity, carried by E(z), which may target the complex by methylating H3 in the appropriate chromatin locations. The trithorax group (trxG) of proteins have the opposite effect to the Pc-G proteins: they act to maintain genes in an active state. There may be some similarities in the actions of the two groups: mutations in some loci prevent both Pc-G and trx from functioning, suggesting that they could rely on common components. A factor coded by the trithorax-like gene, called GAGA because it binds to GA-rich consensus sequences, has binding sites in the PRE. In fact, the sites where Pc binds to DNA coincide with the sites where GAGA factor binds (704, 702). What does this mean? GAGA is probably needed for activating factors, including trxG members, to bind to DNA. Is it also needed for PcG proteins to bind and exercise repression? This is not yet clear, but such a model would demand that something other than GAGA determines which of the alternative types of complex subsequently assemble at the site. Last updated on 9-24-2003

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References 683. Franke, A., DeCamillis, M., Zink, D., Cheng, N., Brock, H. W., and Paro, R. (1992). Polycomb and polyhomeotic are constituents of a multimeric protein complex in chromatin ofD. melanogaster. EMBO J. 11, 2941-29. 697. Zink, B. and Paro, R. (1989). In vivo binding patterns of a trans-regulator of the homeotic genes in D. melanogaster. Nature 337, 468-471. 698. Eissenberg, J. C., James, T. C., Fister-Hartnett, D. M., Hartnett, T., Ngan, V., and Elgin, S. C. R. (1990). Mutation in a heterochromatin-specific chromosomal protein is associated with suppression of position-effect variegation in D. melanogaster. Proc. Natl. Acad. Sci. USA 87, 9923-9927. 699. Orlando, V. and Paro, R. (1993). Mapping Polycomb-repressed domains in the bithorax complex using in vivo formaldehyde cross-linked chromatin. Cell 75, 1187-1198. 700. Chan, C.-S., Rastelli, L., and Pirrotta, V. (1994). A Polycomb response element in the Ubx gene that determines an epigenetically inherited state of repression. EMBO J. 13, 2553-2564. 702. Strutt, H., Cavalli, G., and Paro, R. (1997). Colocalization of Polycomb protein and GAGA factor on regulatory elements responsible for the maintenance of homeotic gene expression. EMBO J. 16, 3621-3632. 704. Geyer, P. K. and Corces, V. G. (1992). DNA position-specific repression of transcription by a Drosophila zinc finger protein. Genes Dev. 6, 1865-1873. 4179. Fischle, W., Wang, Y., Jacobs, S. A., Kim, Y., Allis, C. D., and Khorasanizadeh, S. (2003). Molecular basis for the discrimination of repressive methyl-lysine marks in histone H3 by Polycomb and HP1 chromodomains. Genes Dev. 17, 1870-1881.


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5.23.17 X chromosomes undergo global changes Key Terms Dosage compensation describes mechanisms employed to compensate for the discrepancy between the presence of two X chromosomes in one sex but only one X chromosome in the other sex. Constitutive heterochromatin describes the inert state of permanently nonexpressed sequences, usually satellite DNA. Facultative heterochromatin describes the inert state of sequences that also exist in active copies-for example, one mammalian X chromosome in females. The single X hypothesis describes the inactivation of one X chromosome in female mammals. The n-1 rule states that only one X chromosome is active in female mammalian cells; any other(s) are inactivated. Key Concepts

• One of the two X chromosomes is inactivated at random in each cell during embryogenesis of eutherian mammals.

• In exceptional cases where there are >2 X chromosomes, all but one are inactivated.

• The Xic (X inactivation center) is a cis-acting region on the X chromosome that is necessary and sufficient to ensure that only one X chromosome remains active.

• Xic includes the Xist gene which codes for an RNA that is found only on inactive X chromosomes.

• The mechanism that is responsible for preventing Xist RNA from accumulating on the active chromosome is unknown.

Sex presents an interesting problem for gene regulation, because of the variation in the number of X chromosomes. If X-linked genes were expressed equally well in each sex, females would have twice as much of each product as males. The importance of avoiding this situation is shown by the existence of dosage compensation, which equalizes the level of expression of X-linked genes in the two sexes. Mechanisms used in different species are summarized in Figure 23.27:

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Figure 23.27 Different means of dosage compensation are used to equalize X chromosome expression in male and female.

• In mammals, one of the two female X chromosomes is inactivated completely. The result is that females have only one active X chromosome, which is the same situation found in males. The active X chromosome of females and the single X chromosome of males are expressed at the same level. • In Drosophila, the expression of the single male X chromosome is doubled relative to the expression of each female X chromosome. • In C. elegans, the expression of each female X chromosome is halved relative to the expression of the single male X chromosome. The common feature in all these mechanisms of dosage compensation is that the entire chromosome is the target for regulation. A global change occurs that quantitatively affects all of the promoters on the chromosome. We know most about the inactivation of the X chromosome in mammalian females, where the entire chromosome becomes heterochromatic. The twin properties of heterochromatin are its condensed state and associated inactivity. It can be divided into two types: • Constitutive heterochromatin contains specific sequences that have no coding function. Typically these include satellite DNAs, and are often found at the centromeres. These regions are invariably heterochromatic because of their intrinsic nature. • Facultative heterochromatin takes the form of entire chromosomes that are inactive in one cell lineage, although they can be expressed in other lineages. The example par excellence is the mammalian X chromosome. The inactive X chromosome is perpetuated in a heterochromatic state, while the active X chromosome is part of the euchromatin. So identical DNA sequences are involved in both states. Once the inactive state has been established, it is inherited by descendant cells. This is an example of epigenetic inheritance, because it does not depend on the DNA sequence. Our basic view of the situation of the female mammalian X chromosomes was formed by the single X hypothesis in 1961 (see Great Experiments 11.10 The X chromosomes undergo global changes | SECTION 5.23.17 © 2004. Virtual Text / www.ergito.com

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discovery of X-chromosome inactivation). Female mice that are heterozygous for X-linked coat color mutations have a variegated phenotype in which some areas of the coat are wild-type, but others are mutant. Figure 23.28 shows that this can be explained if one of the two X chromosomes is inactivated at random in each cell of a small precursor population. Cells in which the X chromosome carrying the wild-type gene is inactivated give rise to progeny that express only the mutant allele on the active chromosome. Cells derived from a precursor where the other chromosome was inactivated have an active wild-type gene. In the case of coat color, cells descended from a particular precursor stay together and thus form a patch of the same color, creating the pattern of visible variegation. In other cases, individual cells in a population will express one or the other of X-linked alleles; for example, in heterozygotes for the X-linked locus G6PD, any particular red blood cell will express only one of the two allelic forms. [Random inactivation of one X chromosome occurs in eutherian mammals. In marsupials, the choice is directed: it is always the X chromosome inherited from the father that is inactivated (625).]

Figure 23.28 X-linked variegation is caused by the random inactivation of one X chromosome in each precursor cell. Cells in which the + allele is on the active chromosome have wild phenotype; but cells in which the – allele is on the active chromosome have mutant phenotype.

Inactivation of the X chromosome in females is governed by the n-1 rule: however many X chromosomes are present, all but one will be inactivated. In normal females there are of course 2 X chromosomes, but in rare cases where nondisjunction has generated a 3X or greater genotype, only one X chromosome remains active. This suggests a general model in which a specific event is limited to one X chromosome and protects it from an inactivation mechanism that applies to all the others.


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A single locus on the X chromosome is sufficient for inactivation. When a translocation occurs between the X chromosome and an autosome, this locus is present on only one of the reciprocal products, and only that product can be inactivated. By comparing different translocations, it is possible to map this locus, which is called the Xic (X-inactivation center). A cloned region of 450 kb contains all the properties of the Xic. When this sequence is inserted as a transgene on to an autosome, the autosome becomes subject to inactivation (in a cell culture system) (627). Xic is a cis-acting locus that contains the information necessary to count X chromosomes and inactivate all copies but one. Inactivation spreads from Xic along the entire X chromosome. When Xic is present on an X chromosome-autosome translocation, inactivation spreads into the autosomal regions (although the effect is not always complete). Xic contains a gene, called Xist, that is expressed only on the inactive X chromosome. The behavior of this gene is effectively the opposite from all other loci on the chromosome, which are turned off. Deletion of Xist prevents an X chromosome from being inactivated. However, it does not interfere with the counting mechanism (because other X chromosomes can be inactivated). So we can distinguish two features of Xic: an unidentified element(s) required for counting; and the Xist gene required for inactivation. Figure 23.29 illustrates the role of Xist RNA in X-inactivation (for review see 3237). Xist codes for an RNA that lacks open reading frames. The Xist RNA "coats" the X chromosome from which it is synthesized, suggesting that it has a structural role. Prior to X-inactivation, it is synthesized by both female X chromosomes. Following inactivation, the RNA is found only on the inactive X chromosome. The transcription rate remains the same before and after inactivation, so the transition depends on post-transcriptional events (626).


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Figure 23.29 X-inactivation involves stabilization of Xist RNA, which coats the inactive chromosome.

Prior to X-inactivation, Xist RNA decays with a half life of ~2 hr. X-inactivation is mediated by stabilizing the Xist RNA on the inactive X chromosome. The Xist RNA shows a punctate distribution along the X chromosome, suggesting that association with proteins to form particulate structures may be the means of stabilization. We do not know yet what other factors may be involved in this reaction and how the Xist RNA is limited to spreading in cis along the chromosome. The characteristic features of the inactive X chromosome, which include a lack of acetylation of histone H4, and methylation of CpG sequences (see Molecular Biology 5.21.19 CpG islands are regulatory targets), presumably occur later as part of the mechanism of inactivation (628; 617). The n–1 rule suggests that stabilization of Xist RNA is the "default," and that some blocking mechanism prevents stabilization at one X chromosome (which will be the active X). This means that, although Xic is necessary and sufficient for a chromosome to be inactivated, the products of other loci may be necessary for the establishment of an active X chromosome. Silencing of Xist expression is necessary for the active X. Deletion of the gene for DNA methyltransferase prevents silencing of Xist, probably because methylation at the Xist promoter is necessary for cessation of transcription.


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Reviews 3237. Plath, K., Mlynarczyk-Evans, S., Nusinow, D. A., and Panning, B. (2002). Xist RNA and the mechanism of x chromosome inactivation. Annu. Rev. Genet. 36, 233-278.

References 617. Jeppesen, P. and Turner, B. M. (1993). The inactive X chromosome in female mammals is distinguished by a lack of histone H4 acetylation, a cytogenetic marker for gene expression. Cell 74, 281-289. 625. Lyon, M. F. (1961). Gene action in the X chromosome of the mouse. Nature 190, 372-373. 626. Penny, G. D. et al. (1996). Requirement for Xist in X chromosome inactivation. Nature 379, 131-137. 627. Lee, J. T. et al. (1996). A 450 kb transgene displays properties of the mammalian X-inactivation center. Cell 86, 83-94. 628. Panning, B., Dausman, J., and Jaenisch, R. (1997). X chromosome inactivation is mediated by Xist RNA stabilization. Cell 90, 907-916.



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5.23.18 Chromosome condensation is caused by condensins Key Terms Structural maintenance of chromosomes (SMC) describes a group of proteins that include the cohesins, which hold sister chromatids together, and the condensins, which are involved in chromosome condensation. Condensin proteins are components of a complex that binds to chromosomes to cause condensation for meiosis or mitosis. They are members of the SMC family of proteins. Cohesin proteins form a complex that holds sister chromatids together. They include some SMC proteins. Key Concepts

• SMC proteins are ATPases that include the condensins and the cohesins. • A heterodimer of SMC proteins associates with other subunits. • The condensins cause chromatin to be more tightly coiled by introducing positive supercoils into DNA.

• Condensins are responsible for condensing chromosomes at mitosis. • Chromosome-specific condensins are responsible for condensing inactive X chromosomes in C. elegans.

The structures of entire chromosomes are influenced by interactions with proteins of the SMC (structural maintenance of chromosome) family. They are ATPases that fall into two functional groups (for review see 3445). Condensins are involved with the control of overall structure, and are responsible for the condensation into compact chromosomes at mitosis. Cohesins are concerned with connections between sister chromatids that must be released at mitosis (see Molecular Biology 6.29.19 Cohesins hold sister chromatids together). Both consist of dimers formed by SMC proteins. Condensins form complexes that have a core of the heterodimer SMC2-SMC4 associated with other (non SMC) proteins. Cohesins have a similar organization based on the heterodimeric core of SMC1-SMC3. Figure 23.31 shows that an SMC protein has a coiled-coil structure in its center, interrupted by a flexible hinge region. Both the amino and carboxyl termini have ATP- and DNA-binding motifs. Different models have been proposed for the actions of these proteins depending on whether they dimerize by intra- or inter-molecular interactions.

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Figure 23.31 An SMP protein has a "Walker module" with an ATP-binding motif and DNA-binding site at each end, connected by coiled coils that are linked by a hinge region.

Experiments with the bacterial homologues of the SMC proteins suggest that a dimer is formed by an antiparallel interaction between the coiled coils, so that the N-terminus of one subunit bonds to the C-terminus of the other subunit. The existence of a flexible hinge region could allow cohesins and condensins to depend on a different mode of action by the dimer. Figure 23.33 shows that cohesins have a V-shaped structure, with the arms separated by an 86° angle, whereas condensins are more sharply bent back, with only 6° between the arms. This enables cohesins to hold sister chromatids together, while condensins instead condense an individual chromosome (for review see 2323; 2262; 2375). Figure 23.30 shows that a cohesin could take the form of an extended dimer that cross-links two DNA molecules. Figure 23.32 shows that a condensin could take the form of a V-shaped dimer – essentially bent at the hinge – that pulls together distant sites on the same DNA molecule, causing it to condense.


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Figure 23.30 SMC proteins dimerize by anti-parallel interactions between the central coiled coils. Both terminal regions of each subunit have ATP- and DNA-binding motifs. Cohesins may form an extended structure that allows two different DNA molecules to be linked.

Figure 23.32 Condensins may form a compact structure by bending at the hinge, causing DNA to become compacted.

Figure 23.33 The two halves of a condensin are folded back at an angle of 6°. Cohesins have a more open conformation with an angle of 86° between the two halves.


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An alternative model is suggested by experiments to suggest that the yeast proteins dimerize by intramolecular interactions, that is, a homodimer is formed solely by interaction between two identical subunits (2860). Dimers of two different proteins (in this case, SMC1 and SMC3) may then interact at both their head and hinge regions to form a circular structure as illustrated in Figure 23.35. Instead of binding directly to DNA, a structure of this type could hold DNA molecules together by encircling them (for review see 2859).

Figure 23.35 Cohesins may dimerize by intramolecular connections, then forming multimers that are connected at the heads and at the hinge. Such a structure could hold two molecules of DNA together by surrounding them.

Visualization of mitotic chromosomes shows that condensins are located all along the length of the chromosome, as can be seen in Figure 23.34. (By contrast, cohesins are found at discrete locations; see Figure 29.34).


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Figure 23.34 Condensins are located along the entire length of a mitotic chromosome. DNA is red; condensins are yellow. Photograph kindly provided by Ana Losada and Tatsuya Hirano.

The condensin complex was named for its ability to cause chromatin to condense in vitro (2463). It has an ability to introduce positive supercoils into DNA in an action that uses hydrolysis of ATP and depends on the presence of topoisomerase I. This ability is controlled by the phosphorylation of the non-SMC subunits, which occurs at mitosis. We do not know yet how this connects with other modifications of chromatin, for example, the phosphorylation of histones. The activation of the condensin complex specifically at mitosis makes it questionable whether it is also involved in the formation of interphase heterochromatin. Global changes occur in other types of dosage compensation. In Drosophila, a complex of proteins is found in males, where it localizes on the X chromosome. In C. elegans, a protein complex associates with both X chromosomes in XX embryos, but the protein components remain diffusely distributed in the nuclei of XO embryos. The protein complex contains an SMC core, and is similar to the condensin complexes that are associated with mitotic chromosomes in other species. This suggests that it has a structural role in causing the chromosome to take up a more condensed, inactive state. Multiple sites on the X chromosome may be needed for the complex to be fully distributed along it. The complex binds to these sites, and then spreads along the vhromosome to cover it more thoroughly (4844). Changes affecting all the genes on a chromosome, either negatively (mammals and C. elegans) or positively (Drosophila) are therefore a common feature of dosage compensation. However, the components of the dosage compensation apparatus may vary as well as the means by which it is localized to the chromosome, and of course its mechanism of action is different in each case. Last updated on March 10, 2004


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Reviews 2262. Hirano, T. (2000). Chromosome cohesion, condensation, and separation. Annu. Rev. Biochem. 69, 115-144. 2323. Hirano, T. (1999). SMC-mediated chromosome mechanics: a conserved scheme from bacteria to vertebrates? Genes Dev. 13, 11-19. 2375. Hirano, T. (2002). The ABCs of SMC proteins: two-armed ATPases for chromosome condensation, cohesion, and repair. Genes Dev. 16, 399-414. 2859. Nasmyth, K. (2002). Segregating sister genomes: the molecular biology of chromosome separation. Science 297, 559-565. 3445. Jessberger, R. (2002). The many functions of SMC proteins in chromosome dynamics. Nat. Rev. Mol. Cell Biol. 3, 767-778.

References 2463. Kimura, K., Rybenkov, V. V., Crisona, N. J., Hirano, T., and Cozzarelli, N. R. (1999). 13S condensin actively reconfigures DNA by introducing global positive writhe: implications for chromosome condensation. Cell 98, 239-248. 2860. Haering, C. H., Lowe, J.,Hochwage, A., and Nasmyth, K. (2002). Molecular architecture of SMC proteins and the yeast cohesin complex. Mol. Cell 9, 773-788. 4844. Csankovszki, G., McDonel, P., and Meyer, B. J. (2004). Recruitment and spreading of the C. elegans dosage compensation complex along X chromosomes. Science 303, 1182-1185.



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5.23.19 DNA methylation is perpetuated by a maintenance methylase Key Terms A fully methylated site is a palindromic sequence that is methylated on both strands of DNA. A hemi-methylated site is a palindromic sequence that is methylated on only one strand of DNA. A demethylase is a casual name for an enzyme that removes a methyl group, typically from DNA, RNA, or protein. A methyltransferase (Methylase) is an enzyme that adds a methyl group to a substrate, which can be a small molecule, a protein, or a nucleic acid. A de novo methylase adds a methyl group to an unmethylated target sequence on DNA. A maintenance methylase adds a methyl group to a target site that is already hemimethylated. Key Concepts

• Most methyl groups in DNA are found on cytosine on both strands of the CpG doublet.

• Replication converts a fully methylated site to a hemi-methylated site. • Hemi-methylated sites are converted to fully methylated sites by a maintenance methylase.

Methylation of DNA occurs at specific sites. In bacteria, it is associated with identifying the particular bacterial strain, and also with distinguishing replicated and nonreplicated DNA (see Molecular Biology 4.15.24 Controlling the direction of mismatch repair ). In eukaryotes, its principal known function is connected with the control of transcription; methylation is associated with gene inactivation (see Molecular Biology 5.21.18 Gene expression is associated with demethylation). From 2-7% of the cytosines of animal cell DNA are methylated (the value varies with the species). Most of the methyl groups are found in CG "doublets," and, in fact, the majority of the CG sequences are methylated. Usually the C residues on both strands of this short palindromic sequence are methylated, giving the structure

Such a site is described as fully methylated. But consider the consequences of DNA methylation is perpetuated by a maintenance methylase | SECTION 5.23.19 © 2004. Virtual Text / www.ergito.com

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replicating this site. Figure 23.36 shows that each daughter duplex has one methylated strand and one unmethylated strand. Such a site is called hemi-methylated (for review see 217).

Figure 23.36 The state of methylated sites could be perpetuated by an enzyme that recognizes only hemimethylated sites as substrates.

The perpetuation of the methylated site now depends on what happens to hemimethylated DNA. If methylation of the unmethylated strand occurs, the site is restored to the fully methylated condition. However, if replication occurs first, the hemimethylated condition will be perpetuated on one daughter duplex, but the site will become unmethylated on the other daughter duplex. Figure 23.37 shows that the state of methylation of DNA is controlled by methylases, which add methyl groups to the 5 position of cytosine, and demethylases, which remove the methyl groups. (The more formal name for the enzymes uses methyltransferase as the description.)

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Figure 23.37 The state of methylation is controlled by three types of enzyme. De novo and perpetuation methylases are known, but demethylases have not been identified.

There are two types of DNA methylase, whose actions are distinguished by the state of the methylated DNA. To modify DNA at a new position requires the action of the de novo methylase, which recognizes DNA by virtue of a specific sequence. It acts only on nonmethylated DNA, to add a methyl group to one strand. There are two de novo methylases (Dnmt3A and Dnmt3B) in mouse; they have different target sites, and both are essential for development (941). A maintenance methylase acts constitutively only on hemimethylated sites to convert them to fully methylated sites. Its existence means that any methylated site is perpetuated after replication. There is one maintenance methylase (Dnmt1) in mouse, and it is essential: mouse embryos in which its gene has been disrupted do not survive past early embryogenesis (942). Maintenance methylation is virtually 100% efficient, ensuring that the situation shown on the left of Figure 23.36 usually prevails in vivo. The result is that, if a de novo methylation occurs on one allele but not on the other, this difference will be perpetuated through ensuing cell divisions, maintaining a difference between the alleles that does not depend on their sequences. Methylation has various types of targets. Gene promoters are the most common target. The promoters are methylated when the gene is inactive, but unmethylated when it is active. The absence of Dnmt1 in mouse causes widespread demethylation DNA methylation is perpetuated by a maintenance methylase | SECTION 5.23.19 © 2004. Virtual Text / www.ergito.com

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at promoters, and we assume this is lethal because of the uncontrolled gene expression. Satellite DNA is another target. Mutations in Dnmt3B prevent methylation of satellite DNA, which causes centromere instability at the cellular level. Mutations in the corresponding human gene cause a disease (1964). The importance of methylation is emphasized by another human disease, which is caused by mutation of the gene for the protein McCp2 that binds methylated CpG sequences (1965). The methylases are conventional enzymes that act on a DNA target. However, there may also be a methylation system that uses a short RNA sequence to target a corresponding DNA sequence for methylation (see Molecular Biology 3.11.18 Antisense RNA can be used to inactivate gene expression) Nothing is known about the mechanism of operation of this system (for review see 2077; 2078). How are demethylated regions established and maintained? If a DNA site has not been methylated, a protein that recognizes the unmethylated sequence could protect it against methylation. Once a site has been methylated, there are two possible ways to generate demethylated sites. One is to block the maintenance methylase from acting on the site when it is replicated. After a second replication cycle, one of the daughter duplexes will be unmethylated (as shown on the right side of Figure 23.36). The other is actively to demethylate the site, as shown in Figure 23.38, either by removing the methyl group directly from cytosine, or by excising the methylated cytosine or cytidine from DNA for replacement by a repair system. We know that active demethylation can occur to the paternal genome soon after fertilization, but we do not know what mechanism is used (for review see 2424).


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Figure 23.38 DNA could be demethylated by removing the methyl group, the base, or the nucleotide. Removal of the base or nucleotide would require its replacement by a repair system.



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Reviews 217. Bird, A. P. (1986). A fraction of the mouse genome that is derived from islands of nonmethylated, CpG-rich DNA. Nature 321, 209-213. 2077. Sharp, P. A. (2001). RNA interference--2001. Genes Dev. 15, 485-490. 2078. Matzke, M., Matzke, A. J., and Kooter, J. M. (2001). RNA: guiding gene silencing. Science 293, 1080-1083. 2424. Bird, A. (2002). DNA methylation patterns and epigenetic memory. Genes Dev. 16, 6-21.

References 941. Okano, M., Bell, D. W., Haber, D. A., and Li. E. (1999). DNA methyltransferases Dnmt3a and Dnmt3b are essential for de novo methylation and mammalian development. Cell 99, 247-257. 942. Li, E., Bestor, T. H., and Jaenisch, R. (1992). Targeted mutation of the DNA methyltransferase gene results in embryonic lethality. Cell 69, 915-926. 1964. Xu, G. L., Bestor, T. H., Bourc'his, D., Hsieh, C. L., Tommerup, N., Bugge, M., Hulten, M., Qu, X., Russo, J. J., and Viegas-Paquignot, E. (1999). Chromosome instability and immunodeficiency syndrome caused by mutations in a DNA methyltransferase gene. Nature 402, 187-191. 1965. Amir, R. E., Van den Veyver, I. B., Wan, M., Tran, C. Q., Francke, U., and Zoghbi, H. Y. (1999). Rett syndrome is caused by mutations in X-linked MECP2, encoding methyl-CpG-binding protein 2. Nat. Genet. 23, 185-188.



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5.23.20 DNA methylation is responsible for imprinting Key Terms Imprinting describes a change in a gene that occurs during passage through the sperm or egg with the result that the paternal and maternal alleles have different properties in the very early embryo. May be caused by methylation of DNA. Key Concepts

• Paternal and maternal alleles may have different patterns of methylation at fertilization.

• Methylation is usually associated with inactivation of the gene. • When genes are differentially imprinted, survival of the embryo may require that the functional allele is provided by the parent with the unmethylated allele.

• Survival of heterozygotes for imprinted genes is different depending on the direction of the cross.

• Imprinted genes occur in clusters and may depend on a local control site where de novo methylation occurs unless specifically prevented.

The pattern of methylation of germ cells is established in each sex during gametogenesis by a two stage process: first the existing pattern is erased by a genome-wide demethylation; then the pattern specific for each sex is imposed. All allelic differences are lost when primordial germ cells develop in the embryo; irrespective of sex, the previous patterns of methylation are erased, and a typical gene is then unmethylated. In males, the pattern develops in two stages. The methylation pattern that is characteristic of mature sperm is established in the spermatocyte. But further changes are made in this pattern after fertilization. In females, the maternal pattern is imposed during oogenesis, when oocytes mature through meiosis after birth. As may be expected from the inactivity of genes in gametes, the typical state is to be methylated. However, there are cases of differences between the two sexes, where a locus is unmethylated in one sex. A major question is how the specificity of methylation is determined in the male and female gametes. Systematic changes occur in early embryogenesis. Some sites will continue to be methylated, but others will be specifically unmethylated in cells in which a gene is expressed. From the pattern of changes, we may infer that individual sequence-specific demethylation events occur during somatic development of the organism as particular genes are activated (629). DNA methylation is responsible for imprinting | SECTION 5.23.20 © 2004. Virtual Text / www.ergito.com

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The specific pattern of methyl groups in germ cells is responsible for the phenomenon of imprinting, which describes a difference in behavior between the alleles inherited from each parent. The expression of certain genes in mouse embryos depends upon the sex of the parent from which they were inherited. For example, the allele coding for IGF-II (insulin-like growth factor II) that is inherited from the father is expressed, but the allele that is inherited from the mother is not expressed. The IGF-II gene of oocytes is methylated, but the IGF-II gene of sperm is not methylated, so that the two alleles behave differently in the zygote. This is the most common pattern, but the dependence on sex is reversed for some genes. In fact, the opposite pattern (expression of maternal copy) is shown for IGF-IIR, the receptor for IGF-II (for review see 215). This sex-specific mode of inheritance requires that the pattern of methylation is established specifically during each gametogenesis. The fate of a hypothetical locus in a mouse is illustrated in Figure 23.39. In the early embryo, the paternal allele is nonmethylated and expressed, and the maternal allele is methylated and silent. What happens when this mouse itself forms gametes? If it is a male, the allele contributed to the sperm must be nonmethylated, irrespective of whether it was originally methylated or not. So when the maternal allele finds itself in a sperm, it must be demethylated. If the mouse is a female, the allele contributed to the egg must be methylated; so if it was originally the paternal allele, methyl groups must be added.

Figure 23.39 The typical pattern for imprinting is that a methylated locus is inactive. If this is the maternal allele, only the paternal allele is active, and will be essential for viability. The methylation pattern is reset when gametes are formed, so that all sperm have the paternal type, and all oocytes have the maternal type.

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The consequence of imprinting is that an embryo requires a paternal allele for this gene. So in the case of a heterozygous cross where the allele of one parent has an inactivating mutation, the embryo will survive if the wild-type allele comes from the father, but will die if the wild-type allele is from the mother. This type of dependence on the directionality of the cross (in contrast with Mendelian genetics) is an example of epigenetic inheritance, where some factor other than the sequences of the genes themselves influences their effects (see Molecular Biology 5.23.22 Epigenetic effects can be inherited). Although the paternal and maternal alleles have identical sequences, they display different properties, depending on which parent provided them. These properties are inherited through meiosis and the subsequent somatic mitoses. Imprinted genes are sometimes clustered. More than half of the 17 known imprinted genes in mouse are contained in two particular regions, each containing both maternally and paternally expressed genes. This suggests the possibility that imprinting mechanisms may function over long distances. Some insights into this possibility come from deletions in the human population that cause the Prader-Willi and Angelman diseases. Most cases are caused by the same 4 Mb deletion, but the syndromes are different, depending on which parent contributed the deletion. The reason is that the deleted region contains at least one gene that is paternally imprinted and at least one that is maternally imprinted. There are some rare cases, however, with much smaller deletions. Prader-Willi syndrome can be caused by a 20 kb deletion that silences genes that are distant on either side of it. The basic effect of the deletion is to prevent a father from resetting the paternal mode to a chromosome inherited from his mother. The result is that these genes remain in maternal mode, so that the paternal as well as maternal alleles are silent in the offspring. The inverse effect is found in some small deletions that cause Angelman's syndrome. The implication is that this region comprises some sort of "imprint center" that acts at a distance to switch one parental type to the other. Last updated on 4-24-2002


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Reviews 215. Bartolomei, M. S. and Tilghman, S. (1997). Genomic imprinting in mammals. Annu. Rev. Genet. 31, 493-525.

References 629. Chaillet, J. R., Vogt, T. F., Beier, D. R., and Leder, P. (1991). Parental-specific methylation of an imprinted transgene is established during gametogenesis and progressively changes during embryogenesis. Cell 66, 77-83.



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5.23.21 Oppositely imprinted genes can be controlled by a single center Key Concepts

• Imprinted genes are controlled by methylation of cis-acting sites. • Methylation may be responsible for either inactivating or activating a gene.

Imprinting is determined by the state of methylation of a cis-acting site near a target gene or genes. These regulatory sites are known as DMDs (differentially methylated domains) or ICRs (imprinting control regions). Deletion of these sites removes imprinting, and the target loci then behave the same in both maternal and paternal genomes. The behavior of a region containing two genes, Igf2 and H19, illustrates the ways in which methylation can control gene activity. Figure 23.40 shows that these two genes react oppositely to the state of methylation at a site located between them, called the ICR. The ICR is methylated on the paternal allele. H19 shows the typical response of inactivation. However, Igf2 is expressed. The reverse situation is found on a maternal allele, where the ICR is not methylated. H19 now becomes expressed, but Igf2 is inactivated.

Figure 23.40 ICR is methylated on the paternal allele, where Igf2 is active and H19 is inactive. ICR is unmethylated on the maternal allele, where Igf2 is inactive and H19 is active.

The control of Igf2 is exercised by an insulator function of the ICR. Figure 23.41 shows that when the ICR is unmethylated, it binds the protein CTCF. This creates an insulator function that blocks an enhancer from activating the Igf2 promoter (2048; 2052). This is an unusual effect in which methylation indirectly activates a gene by blocking an insulator. Oppositely imprinted genes can be controlled by a single center | SECTION 5.23.21 © 2004. Virtual Text / www.ergito.com

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Figure 23.41 The ICR is an insulator that prevents an enhancer from activating Igf2. The insulator functions only when it binds CTCF to unmethylated DNA.

The regulation of H19 shows the more usual direction of control in which methylation creates an inactive imprinted state. This could reflect a direct effect of methylation on promoter activity. Last updated on 4-24-2002

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References 2048. Bell, A. C. and Felsenfeld, G. (2000). Methylation of a CTCF-dependent boundary controls imprinted expression of the Igf2 gene. Nature 405, 482-485. 2052. Hark, A. T., Schoenherr, C. J., Katz, D. J., Ingram, R. S., Levorse, J. M., and Tilghman, S. M. (2000). CTCF mediates methylation-sensitive enhancer-blocking activity at the H19/Igf2 locus. Nature 405, 486-489.


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5.23.22 Epigenetic effects can be inherited Key Terms Epigenetic changes influence the phenotype without altering the genotype. They consist of changes in the properties of a cell that are inherited but that do not represent a change in genetic information. Key Concepts

• Epigenetic effects can result from modification of a nucleic acid after it has been synthesized or by the perpetuation of protein structures.

Epigenetic inheritance describes the ability of different states, which may have different phenotypic consequences, to be inherited without any change in the sequence of DNA. How can this occur? We can divide epigenetic mechanisms into two general classes: • DNA may be modified by the covalent attachment of a moiety that is then perpetuated. Two alleles with the same sequence may have different states of methylation that confer different properties. • Or a self perpetuating protein state may be established. This might involve assembly of a protein complex, modification of specific protein(s), or establishment of an alternative protein conformation. Methylation establishes epigenetic inheritance so long as the maintenance methylase acts constitutively to restore the methylated state after each cycle of replication, as shown in Figure 23.36. A state of methylation can be perpetuated through an indefinite series of somatic mitoses. This is probably the "default" situation. Methylation can also be perpetuated through meiosis: for example, in the fungus Ascobolus there are epigenetic effects that can be transmitted through both mitosis and meiosis by maintaining the state of methylation. In mammalian cells, epigenetic effects are created by resetting the state of methylation differently in male and female meioses. Situations in which epigenetic effects appear to be maintained by means of protein states are less well understood in molecular terms. Position effect variegation shows that constitutive heterochromatin may extend for a variable distance, and the structure is then perpetuated through somatic divisions. Since there is no methylation of DNA in Saccharomyces and a vanishingly small amount in Drosophila, the inheritance of epigenetic states of position effect variegation or telomeric silencing in these organisms is likely to be due to the perpetuation of protein structures. Figure 23.42 considers two extreme possibilities for the fate of a protein complex at Epigenetic effects can be inherited | SECTION 5.23.22 © 2004. Virtual Text / www.ergito.com

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replication.

Figure 23.42 What happens to protein complexes on chromatin during replication?

• A complex could perpetuate itself if it splits symmetrically, so that half complexes associate with each daughter duplex. If the half complexes have the capacity to nucleate formation of full complexes, the original state will be restored. This is basically analogous to the maintenance of methylation. The problem with this model is that there is no evident reason why protein complexes should behave in this way. • A complex could be maintained as a unit and segregate to one of the two daughter duplexes. The problem with this model is that it requires a new complex to be assembled de novo on the other daughter duplex, and it is not evident why this should happen. Consider now the need to perpetuate a heterochromatic structure consisting of protein complexes. Suppose that a protein is distributed more or less continuously along a stretch of heterochromatin, as implied in Figure 23.20. If individual subunits are distributed at random to each daughter duplex at replication, the two daughters will continue to be marked by the protein, although its density will be reduced to half of the level before replication. If the protein has a self-assembling property that causes new subunits to associate with it, the original situation may be restored. Basically, the existence of epigenetic effects forces us to the view that a protein responsible for such a situation must have some sort of self-templating or Epigenetic effects can be inherited | SECTION 5.23.22 © 2004. Virtual Text / www.ergito.com

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self-assembling capacity. In some cases, it may be the state of protein modification, rather than the presence of the protein per se, that is responsible for an epigenetic effect. There is a general correlation between the activity of chromatin and the state of acetylation of the histones, in particular the acetylation of histones H3 and H4, which occurs on their N-terminal tails. Activation of transcription is associated with acetylation in the vicinity of the promoter; and repression of transcription is associated with deacetylation (see Molecular Biology 5.23.7 Acetylases are associated with activators). The most dramatic correlation is that the inactive X chromosome in mammalian female cells is underacetylated on histone H4. The inactivity of constitutive heterochromatin may require that the histones are not acetylated. If a histone acetyltransferase is tethered to a region of telomeric heterochromatin in yeast, silenced genes become active. When yeast is exposed to trichostatin (an inhibitor of deacetylation), centromeric heterochromatin becomes acetylated, and silenced genes in centromeric regions may become active. The effect may persist even after trichostatin has been removed. In fact, it may be perpetuated through mitosis and meiosis. This suggests that an epigenetic effect has been created by changing the state of histone acetylation. How might the state of acetylation be perpetuated? Suppose that the H32·H42 tetramer is distributed at random to the two daughter duplexes. This creates the situation shown in Figure 23.43, in which each daughter duplex contains some histone octamers that are fully acetylated on the H3 and H4 tails, while others are completely unacetylated. To account for the epigenetic effect, we could suppose that the presence of some fully acetylated histone octamers provides a signal that causes the unacetylated octamers to be acetylated.

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Figure 23.43 Acetylated cores are conserved and distributed at random to the daughter chromatin fibers at replication. Each daughter fiber has a mixture of old (acetylated) cores and new (unacetylated) cores.

(The actual situation is probably more complicated than shown in the figure, because transient acetylations occur during replication. If they are simply reversed following deposition of histones into nucleosomes, they may be irrelevant. An alternative possibility is that the usual deacetylation is prevented, instead of, or as well as, inducing acetylation.) This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.23.22

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5.23.23 Yeast prions show unusual inheritance Key Terms A prion is a proteinaceous infectious agent, which behaves as an inheritable trait, although it contains no nucleic acid. Examples are PrPSc, the agent of scrapie in sheep and bovine spongiform encephalopathy, and Psi, which confers an inherited state in yeast. Key Concepts

• The Sup35 protein in its wild-type soluble form is a termination factor for translation.

• It can also exist in an alternative form of oligomeric aggregates, in which it is not active in protein synthesis.

• The presence of the oligomeric form causes newly synthesized protein to acquire the inactive structure.

• Conversion between the two forms is influenced by chaperones. – • The wild-type form has the+ recessive genetic state psi and the mutant form has the

dominant genetic state PSI .

One of the clearest cases of the dependence of epigenetic inheritance on the condition of a protein is provided by the behavior of prions – proteinaceous infectious agents. They have been characterized in two circumstances: by genetic effects in yeast; and as the causative agents of neurological diseases in mammals, including man. A striking epigenetic effect is found in yeast, where two different states can be inherited that map to a single genetic locus, although the sequence of the gene is the same in both states. The two different states are [psi–] and [PSI+]. A switch in condition occurs at a low frequency as the result of a spontaneous transition between the states (for review see 211; 212). The psi genotype maps to the locus sup35, which codes for a translation termination factor. Figure 23.44 summarizes the effects of the Sup35 protein in yeast. In wild-type cells, which are characterized as [psi–], the gene is active, and Sup35 protein terminates protein synthesis. In cells of the mutant [PSI+] type, the factor does not function, causing a failure to terminate protein synthesis properly. (This was originally detected by the lethal effects of the enhanced efficiency of suppressors of ochre codons in [PSI+] strains.)

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Figure 23.44 The state of the Sup35 protein determines whether termination of translation occurs.

[PSI+] strains have unusual genetic properties. When a [psi–] strain is crossed with a [PSI+] strain, all of the progeny are [PSI+]. This is a pattern of inheritance that would be expected of an extrachromosomal agent, but the [PSI+] trait cannot be mapped to any such nucleic acid. The [PSI+] trait is metastable, which means that, although it is inherited by most progeny, it is lost at a higher rate than is consistent with mutation. Similar behavior is shown also by the locus URE2, which codes for a protein required for nitrogen-mediated repression of certain catabolic enzymes. When a yeast strain is converted into an alternative state, called [URE3], the Ure2 protein is no longer functional (637). The [PSI+] state is determined by the conformation of the Sup35 protein. In a wild-type [psi–] cell, the protein displays its normal function. But in a [PSI+] cell, the protein is present in an alternative conformation in which its normal function has been lost. To explain the unilateral dominance of [PSI+] over [psi–] in genetic crosses, we must suppose that the presence of protein in the [PSI+] state causes all the protein in the cell to enter this state. This requires an interaction between the [PSI+] protein and newly synthesized protein, probably reflecting the generation of an oligomeric state in which the [PSI+] protein has a nucleating role, as illustrated in Figure 23.45 (for review see 997).


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Figure 23.45 Newly synthesized Sup35 protein is converted into the [PSI+] state by the presence of pre-existing [PSI+] protein.

A feature common to both the Sup35 and Ure2 proteins is that each consists of two domains that function independently. The C-terminal domain is sufficient for the activity of the protein. The N-terminal domain is sufficient for formation of the structures that make the protein inactive. So yeast in which the N-terminal domain of Sup35 has been deleted cannot acquire the [PSI+] state; and the presence of an [PSI+] N-terminal domain is sufficient to maintain Sup35 protein in the [PSI+] condition (639). The critical feature of the N-terminal domain is that it is rich in glutamine and asparagine residues. Loss of function in the [PSI+] state is due to the sequestration of the protein in an oligomeric complex. Sup35 protein in [PSI+] cells is clustered in discrete foci, whereas the protein in [psi–] cells is diffused in the cytosol. Sup35 protein from [PSI+] cells forms amyloid fibers in vitro – these have a characteristic high content of β sheet structures (640). The involvement of protein conformation (rather than covalent modification) is suggested by the effects of conditions that affect protein structure. Denaturing treatments cause loss of the [PSI+] state. And in particular, the chaperone Hsp104 is Yeast prions show unusual inheritance | SECTION 5.23.23 © 2004. Virtual Text / www.ergito.com

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Figure 23.46 Purified protein can convert the[[psi-] state of yeast to [PSI+].


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Reviews 211. Wickner, R. B. (1996). Prions and RNA viruses of S. cerevisiae. Annu. Rev. Genet. 30, 109-139. 212. Lindquist, S. (1997). Mad cows meet psi-chotic yeast: the expansion of the prion hypothesis. Cell 89, 495-498. 213. Horwich, A. L. and Weissman, J. S. (1997). Deadly conformations: protein misfolding in prion disease. Cell 89, 499-510. 997. Serio, T. R. and Lindquist, S. L. (1999). [PSI+]: an epigenetic modulator of translation termination efficiency. Annu. Rev. Cell Dev. Biol. 15, 661-703.

References 637. Wickner, R. B. (1994). [URE3] as an altered URE2 protein: evidence for a prion analog in S. cerevisiae. Science 264, 566-569. 638. Chernoff, Y. O. et al. (1995). Role of the chaperone protein Hsp104 in propagation of the yeast prion-like factor [PSI+]. Science 268, 880-884. 639. Masison, D. C. and Wickner, R. B. (1995). Prion-inducing domain of yeast Ure2p and protease resistance of Ure2p in prion-containing cells. Science 270, 93-95. 640. Glover, J. R. et al. (1997). Self-seeded fibers formed by Sup35, the protein determinant of [PSI+], a heritable prion-like factor of S. cerevisiae. Cell 89, 811-819. 1069. Sparrer, H E, Santoso, A, Szoka, F C, and Weissman, J S (2000). Evidence for the prion hypothesis: induction of the yeast [PSI+] factor by in vitro-converted Sup35 protein. Science 289, 595-599. 1953. Derkatch, I. L., Bradley, M.E., Masse, S. V., Zadorsky, S.P., Polozkov, G. V., Inge-Vechtomov, S. G., Liebman S. W. (2000). Dependence and independence of [PSI(+)] and [PIN(+)]: a two-prion system in yeast? EMBO J. 19, 1942-1952. 1954. Derkatch, I. L., Bradley, M. E., Hong, J. Y., and Liebman, S. W. (2001). Prions affect the appearance of other prions: the story of [PIN(+)]. Cell 106, 171-182. 1955. Osherovich, L. Z. and Weissman, J. S. (2001). Multiple gln/asn-rich prion domains confer susceptibility to induction of the yeast. Cell 106, 183-194.



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5.23.24 Prions cause diseases in mammals Key Terms Scrapie is a infective agent made of protein. Kuru is a human neurological disease caused by prions. It may be caused by eating infected brains. Key Concepts responsible for scrapie exists in two forms, the wild-type noninfectious • The protein C Sc form PrP which is susceptible to proteases, and the disease-causing form PrP which is resistant to proteases.

• The neurological disease can be transmitted to mice by injecting the purified PrP

Sc

protein into mice.

• The recipient mouse must have a copy of the PrP gene coding for the mouse protein.

Sc itself by causing the newly synthesized PrP • The PrP protein can perpetuate Sc C

protein to take up the PrP form instead of the PrP form.

Sc • Multiple strains of PrP may have different conformations of the protein.

Prion diseases have been found in sheep and Man, and, more recently, in cows. The basic phenotype is an ataxia – a neurodegenerative disorder that is manifested by an inability to remain upright. The name of the disease in sheep, scrapie, reflects the phenotype: the sheep rub against walls in order to stay upright. Scrapie can be perpetuated by inoculating sheep with tissue extracts from infected animals. The disease kuru was found in New Guinea, where it appeared to be perpetuated by cannibalism, in particular the eating of brains. Related diseases in Western populations with a pattern of genetic transmission include Gerstmann-Straussler syndrome; and the related Creutzfeldt-Jakob disease (CJD) occurs sporadically. Most recently, a disease resembling CJD appears to have been transmitted by consumption of meat from cows suffering from "mad cow" disease (634). When tissue from scrapie-infected sheep is inoculated into mice, the disease occurs in a period ranging from 75-150 days. The active component is a protease-resistant protein. The protein is coded by a gene that is normally expressed in brain. The form of the protein in normal brain, called PrPC, is sensitive to proteases. Its conversion to the resistant form, called PrpSc, is associated with occurrence of the disease. The infectious preparation has no detectable nucleic acid, is sensitive to UV irradiation at wave lengths that damage protein, and has a low infectivity (1 infectious unit / 105 PrPSc proteins). This corresponds to an epigenetic inheritance in which there is no change in genetic information, because normal and diseased cells have the same PrP gene sequence, but the PrPSc form of the protein is the infectious agent, whereas PrPC is harmless (383; 384; 385; for review see 203). Prions cause diseases in mammals | SECTION 5.23.24 © 2004. Virtual Text / www.ergito.com

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Molecular Biology The basis for the difference between the PrPSc and PrpC forms appears to lie with a change in conformation rather than with any covalent alteration. Both proteins are glycosylated and linked to the membrane by a GPI-linkage. No changes in these modifications have been found. The PrPSc form has a high content of β sheets, which is absent from the PrPC form. The assay for infectivity in mice allows the dependence on protein sequence to be tested. Figure 23.47 illustrates the results of some critical experiments. In the normal situation, PrPSc protein extracted from an infected mouse will induce disease (and ultimately kill) when it is injected into a recipient mouse. If the PrP gene is "knocked out", a mouse becomes resistant to infection. This experiment demonstrates two things. First, the endogenous protein is necessary for an infection, presumably because it provides the raw material that is converted into the infectious agent. Second, the cause of disease is not the removal of the PrPC form of the protein, because a mouse with no PrPC survives normally: the disease is caused by a gain-of-function in PrPSc (386).

Figure 23.47 A PrpSc protein can only infect an animal that has the same type of endogenous PrPC protein.

The existence of species barriers allows hybrid proteins to be constructed to delineate the features required for infectivity. The original preparations of scrapie were perpetuated in several types of animal, but these cannot always be transferred readily. For example, mice are resistant to infection from prions of hamsters. This means that hamster-PrPSc cannot convert mouse-PrPC to PrPSc. However, the situation changes if the mouse PrP gene is replaced by a hamster PrP gene. (This can be done by introducing the hamster PrP gene into the PrP knockout mouse.) A mouse with a hamster PrP gene is sensitive to infection by hamster PrPSc. This suggests that the conversion of cellular PrPC protein into the Sc state requires that the PrPSc and PrPC proteins have matched sequences. Prions cause diseases in mammals | SECTION 5.23.24 © 2004. Virtual Text / www.ergito.com

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Molecular Biology There are different "strains" of PrPSc, which are distinguished by characteristic incubation periods upon inoculation into mice. This implies that the protein is not restricted solely to alternative states of PrPC and PrPSc, but that there may be multiple Sc states. These differences must depend on some self-propagating property of the protein other than its sequence. If conformation is the feature that distinguishes PrPSc from PrPC, then there must be multiple conformations, each of which has a self-templating property when it converts PrPC (636; for review see 214). The probability of conversion from PrPC to PrPSc is affected by the sequence of PrP. Gerstmann-Straussler syndrome in man is caused by a single amino acid change in PrP. This is inherited as a dominant trait. If the same change is made in the mouse PrP gene, mice develop the disease. This suggests that the mutant protein has an increased probability of spontaneous conversion into the Sc state. Similarly, the sequence of the PrP gene determines the susceptibility of sheep to develop the disease spontaneously; the combination of amino acids at three positions (codons 136, 154, and 171) determines susceptibility. The prion offers an extreme case of epigenetic inheritance, in which the infectious agent is a protein that can adopt multiple conformations, each of which has a self-templating property. This property is likely to involve the state of aggregation of the protein.

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Reviews 203. Prusiner, S. (1982). Novel proteinaceous infectious particles cause scrapie. Science 216, 136-144. 214. Prusiner, S. B. and Scott, M. R. (1997). Genetics of prions. Annu. Rev. Genet. 31, 139-175.

References 383. McKinley, M. P., Bolton, D. C., and Prusiner, S. B. (1983). A protease-resistant protein is a structural component of the scrapie prion. Cell 35, 57-62. 384. Oesch, B. et al. (1985). A cellular gene encodes scrapie PrP27-30 protein. Cell 40, 735-746. 385. Basler, K., Oesch, B., Scott, M., Westaway, D., Walchli, M., Groth, D. F., McKinley, M. P., Prusiner, S. B., and Weissmann, C. (1986). Scrapie and cellular PrP isoforms are encoded by the same chromosomal gene. Cell 46, 417-428. 386. Bueler, H. et al. (1993). Mice devoid of PrP are resistant to scrapie. Cell 73, 1339-1347. 634. Hsiao, K. et al. (1989). Linkage of a prion protein missense variant to Gerstmann-Straussler syndrome. Nature 338, 342-345. 636. Scott, M. et al. (1993). Propagation of prions with artificial properties in transgenic mice expressing chimeric PrP genes. Cell 73, 979-988.


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5.23.25 Summary The existence of a preinitiation complex signals that the gene is in an "active" state, ready to be transcribed. The complex is stable, and may remain in existence through many cycles of replication. The ability to form a preinitiation complex could be a general regulatory mechanism. By binding to a promoter to make it possible for RNA polymerase in turn to bind, the factor in effect switches the gene on. The variety of situations in which hypersensitive sites occur suggests that their existence reflects a general principle. Sites at which the double helix initiates an activity are kept free of nucleosomes. A transcription factor, or some other nonhistone protein concerned with the particular function of the site, modifies the properties of a short region of DNA so that nucleosomes are excluded. The structures formed in each situation need not necessarily be similar (except that each, by definition, creates a site hypersensitive to DNAase I). Genes whose control regions are organized in nucleosomes usually are not expressed. In the absence of specific regulatory proteins, promoters and other regulatory regions are organized by histone octamers into a state in which they cannot be activated. This may explain the need for nucleosomes to be precisely positioned in the vicinity of a promoter, so that essential regulatory sites are appropriately exposed. Some transcription factors have the capacity to recognize DNA on the nucleosomal surface, and a particular positioning of DNA may be required for initiation of transcription. Active chromatin and inactive chromatin are not in equilibrium. Sudden, disruptive events are needed to convert one to the other. Chromatin remodeling complexes have the ability to displace histone octamers by a mechanism that involves hydrolysis of ATP. Remodeling complexes are large and are classified according to the type of the ATPase subunit. Two common types are SWI/SNF and ISW. A typical form of this chromatin remodeling is to displace one or more histone octamers from specific sequences of DNA, creating a boundary that results in the precise or preferential positioning of adjacent nucleosomes. Chromatin remodeling may also involve changes in the positions of nucleosomes, sometimes involving sliding of histone octamers along DNA. Acetylation of histones occurs at both replication and transcription and could be necessary to form a less compact chromatin structure. Some coactivators, which connect transcription factors to the basal apparatus, have histone acetylase activity. Conversely, repressors may be associated with deacetylases. The modifying enzymes are usually specific for particular amino acids in particular histones. The most common sites for modification are located in the N-terminal tails of histones H3 and H4, which extrude from nucleosomes between the turns of DNA. The activating (or repressing) complexes are usually large and often contain several activities that undertake different modifications of chromatin. Some common motifs found in proteins that modify chromatin are the chromo domain (concerned with protein-protein interactions). the bromo domain (which targets acetylated lysine), and the SET domain (part of the active sites of histone methyltransferases). Summary | SECTION 5.23.25 © 2004. Virtual Text / www.ergito.com

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The formation of heterochromatin occurs by proteins that bind to specific chromosomal regions (such as telomeres) and that interact with histones. The formation of an inactive structure may propagate along the chromatin thread from an initiation center. Similar events occur in silencing of the inactive yeast mating type loci. Repressive structures that are required to maintain the inactive states of particular genes are formed by the Pc-G protein complex in Drosophila. They share with heterochromatin the property of propagating from an initiation center. Formation of heterochromatin may be initiated at certain sites and then propagated for a distance that is not precisely determined. When a heterochromatic state has been established, it is inherited through subsequent cell divisions. This gives rise to a pattern of epigenetic inheritance, in which two identical sequences of DNA may be associated with different protein structures, and therefore have different abilities to be expressed. This explains the occurrence of position effect variegation in Drosophila. Modification of histone tails is a trigger for chromatin reorganization. Acetylation is generally associated with gene activation. Histones acetylases are found in activating complexes, and histone deacetylases are found in inactivating complexes. Histone methylation is associated with gene inactivation. Some histone modifications may be exclusive or synergistic with others. Inactive chromatin at yeast telomeres and silent mating type loci appears to have a common cause, and involves the interaction of certain proteins with the N-terminal tails of histones H3 and H4. Formation of the inactive complex may be initiated by binding of one protein to a specific sequence of DNA; the other components may then polymerize in a cooperative manner along the chromosome. Inactivation of one X chromosome in female (eutherian) mammals occurs at random. The Xic locus is necessary and sufficient to count the number of X chromosomes. The n-1 rule ensures that all but one X chromosome are inactivated. Xic contains the gene Xist, which codes for an RNA that is expressed only on the inactive X chromosome. Stabilization of Xist RNA is the mechanism by which the inactive X chromosome is distinguished. Methylation of DNA is inherited epigenetically. Replication of DNA creates hemimethylated products, and a maintenance methylase restores the fully methylated state. Some methylation events depend on parental origin. Sperm and eggs contain specific and different patterns of methylation, with the result that paternal and maternal alleles are differently expressed in the embryo. This is responsible for imprinting, in which the nonmethylated allele inherited from one parent is essential because it is the only active allele; the allele inherited from the other parent is silent. Patterns of methylation are reset during gamete formation in every generation. Prions are proteinaceous infectious agents that are responsible for the disease of scrapie in sheep and for related diseases in man. The infectious agent is a variant of a normal cellular protein. The PrPSc form has an altered conformation that is self-templating: the normal PrPC form does not usually take up this conformation, but does so in the presence of PrPSc. A similar effect is responsible for inheritance of the PSI element in yeast.


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RNA SPLICING AND PROCESSING

5.24.1 Introduction Key Terms Pre-mRNA is used to describe the nuclear transcript that is processed by modification and splicing to give an mRNA. RNA splicing is the process of excising the sequences in RNA that correspond to introns, so that the sequences corresponding to exons are connected into a continuous mRNA. Heterogeneous nuclear RNA (hnRNA) comprises transcripts of nuclear genes made by RNA polymerase II; it has a wide size distribution and low stability. An hnRNP is the ribonucleoprotein form of hnRNA (heterogeneous nuclear RNA), in which the hnRNA is complexed with proteins. Since pre-mRNAs are not exported until processing is complete, hnRNPs are found only in the nucleus.

Interrupted genes are found in all classes of organisms. They represent a minor proportion of the genes of the very lowest eukaryotes, but the vast majority of genes in higher eukaryotic genomes. Genes vary widely according to the numbers and lengths of introns, but a typical mammalian gene has 7-8 exons spread out over ~16 kb. The exons are relatively short (~100-200 bp), and the introns are relatively long (>1 kb) (see Molecular Biology 1.2.7 Genes show a wide distribution of sizes). The discrepancy between the interrupted organization of the gene and the uninterrupted organization of its mRNA requires processing of the primary transcription product. The primary transcript has the same organization as the gene, and is sometimes called the pre-mRNA. Removal of the introns from pre-mRNA leaves a typical messenger of ~2.2 kb. The process by which the introns are removed is called RNA splicing. Removal of introns is a major part of the production of RNA in all eukaryotes. (Although interrupted genes are relatively rare in lower eukaryotes such as yeast, the overall proportion underestimates the importance of introns, because most of the genes that are interrupted code for relatively abundant proteins. Splicing is therefore involved in the production of a greater proportion of total mRNA than would be apparent from analysis of the genome, perhaps as much as 50%.) One of the first clues about the nature of the discrepancy in size between nuclear genes and their products in higher eukaryotes was provided by the properties of nuclear RNA. Its average size is much larger than mRNA, it is very unstable, and it has a much greater sequence complexity. Taking its name from its broad size distribution, it was called heterogeneous nuclear RNA (hnRNA). It includes pre-mRNA, but could also include other transcripts (that is, which are not ultimately processed to mRNA; for review see 17). The physical form of hnRNA is a ribonucleoprotein particle (hnRNP), in which the Introduction | SECTION 5.24.1 © 2004. Virtual Text / www.ergito.com

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hnRNA is bound by proteins. As characterized in vitro, an hnRNP particle takes the form of beads connected by a fiber. The structure is summarized in Figure 24.1. The most abundant proteins in the particle are the core proteins, but other proteins are present at lower stoichiometry, making a total of ~20 proteins. The proteins typically are present at ~108 copies per nucleus, compared with ~106 molecules of hnRNA. Some of the proteins may have a structural role in packaging the hnRNA; several are known to shuttle between the nucleus and cytoplasm, and play roles in exporting the RNA or otherwise controlling its activity (for review see 249; 3428).

Figure 24.1 hnRNA exists as a ribonucleoprotein particle organized as a series of beads.

Splicing occurs in the nucleus, together with the other modifications that are made to newly synthesized RNAs. The process of expressing an interrupted gene is reviewed in Figure 24.2. The transcript is capped at the 5 ′ end (see Molecular Biology 2.5.9 The 5 ′ end of eukaryotic mRNA is capped), has the introns removed, and is polyadenylated at the 3 ′ end (see Molecular Biology 2.5.10 The 3 ′ terminus is polyadenylated). The RNA is then transported through nuclear pores to the cytoplasm, where it is available to be translated.


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Figure 24.2 RNA is modified in the nucleus by additions to the 5 ′ and 3 ′ ends and by splicing to remove the introns. The splicing event requires breakage of the exon-intron junctions and joining of the ends of the exons. Mature mRNA is transported through nuclear pores to the cytoplasm, where it is translated.

With regard to the various processing reactions that occur in the nucleus, we should like to know at what point splicing occurs vis-À-vis the other modifications of RNA. Does splicing occur at a particular location in the nucleus; and is it connected with other events, for example, nucleocytoplasmic transport? Does the lack of splicing make an important difference in the expression of uninterrupted genes? With regard to the splicing reaction itself, one of the main questions is how its specificity is controlled. What ensures that the ends of each intron are recognized in pairs so that the correct sequence is removed from the RNA? Are introns excised from a precursor in a particular order? Is the maturation of RNA used to regulate gene expression by discriminating among the available precursors or by changing the pattern of splicing? We can identify several types of splicing systems: • Introns are removed from the nuclear pre-mRNAs of higher eukaryotes by a Introduction | SECTION 5.24.1 © 2004. Virtual Text / www.ergito.com

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system that recognizes only short consensus sequences conserved at exon-intron boundaries and within the intron. This reaction requires a large splicing apparatus, which takes the form of an array of proteins and ribonucleoproteins that functions as a large particulate complex (the spliceosome). The mechanism of splicing involves transesterifications, and the catalytic center includes RNA as well as proteins. • Certain RNAs have the ability to excise their introns autonomously. Introns of this type fall into two groups, as distinguished by secondary/tertiary structure. Both groups use transesterification reactions in which the RNA is the catalytic agent (see Molecular Biology 5.26 Catalytic RNA). • The removal of introns from yeast nuclear tRNA precursors involves enzymatic activities that handle the substrate in a way resembling the tRNA processing enzymes, in which a critical feature is the conformation of the tRNA precursor. These splicing reactions are accomplished by enzymes that use cleavage and ligation.


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Reviews 17.

Lewin, B. (1975). Units of transcription and translation: sequence components of hnRNA and mRNA. Cell 4, 77-93.

249. Dreyfuss, G. et al. (1993). hnRNP proteins and the biogenesis of mRNA. Annu. Rev. Biochem. 62, 289-321. 3428. Dreyfuss, G., Kim, V. N., and Kataoka, N. (2002). Messenger-RNA-binding proteins and the messages they carry. Nat. Rev. Mol. Cell Biol. 3, 195-205.



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5.24.2 Nuclear splice junctions are short sequences Key Terms Splice sites are the sequences immediately surrounding the exon-intron boundaries. The GT-AG rule describes the presence of these constant dinucleotides at the first two and last two positions of introns of nuclear genes. Key Concepts

• Splice sites are the sequences immediately surrounding the exon-intron boundaries. They are named for their positions relative to the intron.

• The 5 ′ splice site at the 5 ′ (left) end of the intron includes the consensus sequence GU.

• The 3 ′ splice site at the 3 ′ (right) end of the intron includes the consensus sequence AG.

• The GU-AG rule (originally called the GT-AG rule in terms of DNA sequence)

describes the requirement for these constant dinucleotides at the first two and last two positions of introns in pre-mRNAs.

To focus on the molecular events involved in nuclear intron splicing, we must consider the nature of the splice sites, the two exon-intron boundaries that include the sites of breakage and reunion. By comparing the nucleotide sequence of mRNA with that of the structural gene, the junctions between exons and introns can be assigned. There is no extensive homology or complementarity between the two ends of an intron. However, the junctions have well conserved, though rather short, consensus sequences. It is possible to assign a specific end to every intron by relying on the conservation of exon-intron junctions. They can all be aligned to conform to the consensus sequence given in Figure 24.3.

Figure 24.3 The ends of nuclear introns are defined by the GU-AG rule.

Nuclear splice junctions are short sequences | SECTION 5.24.2 © 2004. Virtual Text / www.ergito.com

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The subscripts indicate the percent occurrence of the specified base at each consensus position. High conservation is found only immediately within the intron at the presumed junctions. This identifies the sequence of a generic intron as: GU##AG Because the intron defined in this way starts with the dinucleotide GU and ends with the dinucleotide AG, the junctions are often described as conforming to the GT-AG rule. (This reflects the fact that the sequences were originally analyzed in terms of DNA, but of course the GT in the coding strand sequence of DNA becomes a GU in the RNA.) Note that the two sites have different sequences and so they define the ends of the intron directionally. They are named proceeding from left to right along the intron as the 5 ′ splice site (sometimes called the left or donor site) and the 3 ′ splice site (also called the right or acceptor site). The consensus sequences are implicated as the sites recognized in splicing by point mutations that prevent splicing in vivo and in vitro (for review see 242; 243).


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Reviews 242. Padgett, R. A. (1986). Splicing of messenger RNA precursors. Annu. Rev. Biochem. 55, 1119-1150. 243. Sharp, P. A. (1987). Splicing of mRNA precursors. Science 235, 766-771.



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5.24.3 Splice junctions are read in pairs Key Concepts

• Splicing depends only on recognition of pairs of splice junctions. • All 5 ′ splice sites are functionally equivalent, and all 3 ′ splice sites are functionally equivalent.

A typical mammalian mRNA has many introns. The basic problem of pre-mRNA splicing results from the simplicity of the splice sites, and is illustrated in Figure 24.4: what ensures that the correct pairs of sites are spliced together? The corresponding GU-AG pairs must be connected across great distances (some introns are >10 kb long). We can imagine two types of principle that might be responsible for pairing the appropriate 5 ′ and 3 ′ sites:

Figure 24.4 Splicing junctions are recognized only in the correct pairwise combinations.

• It could be an intrinsic property of the RNA to connect the sites at the ends of a particular intron. This would require matching of specific sequences or structures. • Or all 5 ′ sites may be functionally equivalent and all 3 ′ sites may be similarly indistinguishable, but splicing could follow rules that ensure a 5 ′ site is always connected to the 3 ′ site that comes next in the RNA. Neither the splice sites nor the surrounding regions have any sequence complementarity, which excludes models for complementary base pairing between Splice junctions are read in pairs | SECTION 5.24.3 © 2004. Virtual Text / www.ergito.com

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Molecular Biology intron ends. And experiments using hybrid RNA precursors show that any 5 ′ splice site can in principle be connected to any 3 ′ splice site. For example, when the first exon of the early SV40 transcription unit is linked to the third exon of mouse β globin, the hybrid intron can be excised to generate a perfect connection between the SV40 exon and the β-globin exon. Indeed, this interchangeability is the basis for the exon-trapping technique described previously in Figure 2.12. Such experiments make two general points: • Splice sites are generic: they do not have specificity for individual RNA precursors, and individual precursors do not convey specific information (such as secondary structure) that is needed for splicing. • The apparatus for splicing is not tissue specific; an RNA can usually be properly spliced by any cell, whether or not it is usually synthesized in that cell. (We discuss exceptions in which there are tissue-specific alternative splicing patterns in Molecular Biology 5.24.12 Alternative splicing involves differential use of splice junctions.) Here is a paradox. Probably all 5 ′ splice sites look similar to the splicing apparatus, and all 3 ′ splice sites look similar to it. In principle any 5 ′ splice site may be able to react with any 3 ′ splice site. But in the usual circumstances splicing occurs only between the 5 ′ and 3 ′ sites of the same intron. What rules ensure that recognition of splice sites is restricted so that only the 5 ′ and 3 ′ sites of the same intron are spliced? Are introns removed in a specific order from a particular RNA? Using RNA blotting, we can identify nuclear RNAs that represent intermediates from which some introns have been removed. Figure 24.5 shows a blot of the precursors to ovomucoid mRNA. There is a discrete series of bands, which suggests that splicing occurs via definite pathways. (If the seven introns were removed in an entirely random order, there would be more than 300 precursors with different combinations of introns, and we should not see discrete bands.)

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Figure 24.5 Northern blotting of nuclear RNA with an ovomucoid probe identifies discrete precursors to mRNA. The contents of the more prominent bands are indicated. Photograph kindly provided by Bert O#Malley.

There does not seem to be a unique pathway, since intermediates can be found in which different combinations of introns have been removed. However, there is evidence for a preferred pathway or pathways. When only one intron has been lost, it is virtually always 5 or 6. But either can be lost first. When two introns have been lost, 5 and 6 are again the most frequent, but there are other combinations. Intron 3 is never or very rarely lost at one of the first three splicing steps. From this pattern, we see that there is a preferred pathway in which introns are removed in the order 5/6, 7/4, 2/1, 3. But there are other pathways, since (for example), there are some molecules in which 4 or 7 is lost last. A caveat in interpreting these results is that we do not have proof that all these intermediates actually lead to mature mRNA. The general conclusion suggested by this analysis is that the conformation of the RNA influences the accessibility of the splice sites. As particular introns are removed, the conformation changes, and new pairs of splice sites become available. Splice junctions are read in pairs | SECTION 5.24.3 © 2004. Virtual Text / www.ergito.com

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But the ability of the precursor to remove its introns in more than one order suggests that alternative conformations are available at each stage. Of course, the longer the molecule, the more structural options become available; and when we consider larger genes, it becomes difficult to see how specific secondary structures could control the reaction. One important conclusion of this analysis is that the reaction does not proceed sequentially along the precursor. A simple model to control recognition of splice sites would be for the splicing apparatus to act in a processive manner. Having recognized a 5 ′ site, the apparatus might scan the RNA in the appropriate direction until it meets the next 3 ′ site. This would restrict splicing to adjacent sites. But this model is excluded by experiments that show that splicing can occur in trans as an intermolecular reaction under special circumstances (see Molecular Biology 5.24.13 trans-splicing reactions use small RNAs) or in RNA molecules in which part of the nucleotide chain is replaced by a chemical linker. This means that there cannot be a requirement for strict scanning along the RNA from the 5 ′ splice site to the 3 ′ splice site. Another problem with the scanning model is that it cannot explain the existence of alternative splicing patterns, where (for example) a common 5 ′ site is spliced to more than one 3 ′ site. The basis for proper recognition of correct splice site pairs remains incompletely defined. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.24.3

Splice junctions are read in pairs | SECTION 5.24.3 © 2004. Virtual Text / www.ergito.com

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5.24.4 pre-mRNA splicing proceeds through a lariat Key Terms The lariat is an intermediate in RNA splicing in which a circular structure with a tail is created by a 5 ′ -2 ′ bond. The branch site is a short sequence just before the end of an intron at which the lariat intermediate is formed in splicing by joining the 5 ′ nucleotide of the intron to the 2 ′ position of an Adenosine. A transesterification reaction breaks and makes chemical bonds in a coordinated transfer so that no energy is required. Key Concepts

• Splicing requires the 5 ′ and 3 ′ splice sites and a branch site just upstream of the 3 ′ splice site.

• The branch sequence is conserved in yeast but less well conserved in higher eukaryotes.

• A lariat is formed when the intron is cleaved at the 5 ′ splice site, and the 5 ′ end is joined to a 2 ′ position at an A at the branch site in the intron.

• The intron is released as a lariat when it is cleaved at the 3 ′ splice site, and the left and right exons are then ligated together.

• The reactions occur by transesterifications in which a bond is transferred from one location to another.

The mechanism of splicing has been characterized in vitro, using systems in which introns can be removed from RNA precursors. Nuclear extracts can splice purified RNA precursors, which shows that the action of splicing is not linked to the process of transcription. Splicing can occur to RNAs that are neither capped nor polyadenylated. However, although the splicing reaction as such is independent of transcription or modification to the RNA, these events normally occur in a coordinated manner, and the efficiency of splicing may be influenced by other processing events. The stages of splicing in vitro are illustrated in the pathway of Figure 24.6. We discuss the reaction in terms of the individual RNA species that can be identified, but remember that in vivo the species containing exons are not released as free molecules, but remain held together by the splicing apparatus (for review see 253).

pre-mRNA splicing proceeds through a lariat | SECTION 5.24.4 © 2004. Virtual Text / www.ergito.com

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Figure 24.6 Splicing occurs in two stages. First the 5 ′ exon is cleaved off; then it is joined to the 3 ′ exon.

The first step is to make a cut at the 5 ′ splice site, separating the left exon and the right intron-exon molecule. The left exon takes the form of a linear molecule. The right intron-exon molecule forms a lariat, in which the 5 ′ terminus generated at the end of the intron becomes linked by a 5 ′ –2 ′ bond to a base within the intron. The target base is an A in a sequence that is called the branch site (712). Cutting at the 3 ′ splice site releases the free intron in lariat form, while the right exon is ligated (spliced) to the left exon. The cleavage and ligation reactions are shown separately in the figure for illustrative purposes, but actually occur as one coordinated transfer. The lariat is then "debranched" to give a linear excised intron, which is rapidly degraded. The sequences needed for splicing are the short consensus sequences at the 5 ′ and 3 ′ splice sites and at the branch site. Together with the knowledge that most of the sequence of an intron can be deleted without impeding splicing, this indicates that there is no demand for specific conformation in the intron (or exon). pre-mRNA splicing proceeds through a lariat | SECTION 5.24.4 © 2004. Virtual Text / www.ergito.com

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Molecular Biology The branch site plays an important role in identifying the 3 ′ splice site. The branch site in yeast is highly conserved, and has the consensus sequence UACUAAC. The branch site in higher eukaryotes is not well conserved, but has a preference for purines or pyrimidines at each position and retains the target A nucleotide (see Figure 24.6) (717). The branch site lies 18-40 nucleotides upstream of the 3 ′ splice site. Mutations or deletions of the branch site in yeast prevent splicing. In higher eukaryotes, the relaxed constraints in its sequence result in the ability to use related sequences (called cryptic sites) when the authentic branch is deleted. Proximity to the 3 ′ splice site appears to be important, since the cryptic site is always close to the authentic site. A cryptic site is used only when the branch site has been inactivated. When a cryptic branch sequence is used in this manner, splicing otherwise appears to be normal; and the exons give the same products as wild type. The role of the branch site therefore is to identify the nearest 3 ′ splice site as the target for connection to the 5 ′ splice site (713). This can be explained by the fact that an interaction occurs between protein complexes that bind to these two sites. The bond that forms the lariat goes from the 5 ′ position of the invariant G that was at the 5 ′ end of the intron to the 2 ′ position of the invariant A in the branch site. This corresponds to the third A residue in the yeast UACUAAC box. The chemical reactions proceed by transesterification: a bond is in effect transferred from one location to another. Figure 24.7 shows that the first step is a nucleophilic attack by the 2 ′ –OH of the invariant A of the UACUAAC sequence on the 5 ′ splice site. In the second step, the free 3 ′ –OH of the exon that was released by the first reaction now attacks the bond at the 3 ′ splice site. Note that the number of phosphodiester bonds is conserved. There were originally two 5 ′ –3 ′ bonds at the exon-intron splice sites; one has been replaced by the 5 ′ –3 ′ bond between the exons, and the other has been replaced by the 5 ′ –2 ′ bond that forms the lariat (for review see 251).


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Figure 24.7 Nuclear splicing occurs by two transesterification reactions in which an OH group attacks a phosphodiester bond.


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Reviews 251. Weiner, A. (1993). mRNA splicing and autocatalytic introns: distant cousins or the products of chemical determinism. Cell 72, 161-164. 253. Sharp, P.A. (1994). Split genes and RNA splicing. Cell 77, 805-815.

References 712. Reed, R. and Maniatis, T. (1985). Intron sequences involved in lariat formation during pre-mRNA splicing. Cell 41, 95-105. 713. Reed, R. and Maniatis, T. (1986). A role for exon sequences and splice-site proximity in splice-site selection. Cell 46, 681-690. 717. Zhuang, Y. A., Goldstein, A. M., and Weiner, A. M. (1989). UACUAAC is the preferred branch site for mammalian mRNA splicing. Proc. Natl. Acad. Sci. USA 86, 2752-2756.



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5.24.5 snRNAs are required for splicing Key Terms A small nuclear RNA (snRNA) is one of many small RNA species confined to the nucleus; several of the snRNAs are involved in splicing or other RNA processing reactions. Small cytoplasmic RNAs (scRNA) are present in the cytoplasm and (sometimes are also found in the nucleus). snRNPs (snurp) are small nuclear ribonucleoproteins (snRNAs associated with proteins). scRNPs (scyrp) are small cytoplasmic ribonucleoproteins (scRNAs associated with proteins). The spliceosome is a complex formed by the snRNPs that are required for splicing together with additional protein factors. Anti-Sm is an autoimmune antiserum that defines the Sm epitope that is common to a group of proteins found in snRNPs that are involved in RNA splicing. Key Concepts

• The five snRNPs involved in splicing are U1, U2, U5, U4, and U6. • Together with some additional proteins, the snRNPs form the spliceosome. • All the snRNPs except U6 contain a conserved sequence that binds the Sm proteins that are recognized by antibodies generated in autoimmune disease.

The 5 ′ and 3 ′ splice sites and the branch sequence are recognized by components of the splicing apparatus that assemble to form a large complex. This complex brings together the 5 ′ and 3 ′ splice sites before any reaction occurs, explaining why a deficiency in any one of the sites may prevent the reaction from initiating. The complex assembles sequentially on the pre-mRNA, and several intermediates can be recognized by fractionating complexes of different sizes. Splicing occurs only after all the components have assembled (719). The splicing apparatus contains both proteins and RNAs (in addition to the pre-mRNA). The RNAs take the form of small molecules that exist as ribonucleoprotein particles. Both the nucleus and cytoplasm of eukaryotic cells contain many discrete small RNA species. They range in size from 100-300 bases in higher eukaryotes, and extend in length to ~1000 bases in yeast. They vary considerably in abundance, from 105-106 molecules per cell to concentrations too low to be detected directly. Those restricted to the nucleus are called small nuclear RNAs (snRNA); those found in the cytoplasm are called small cytoplasmic RNAs (scRNA). In their natural state, they exist as ribonucleoprotein particles (snRNP and scRNP). snRNAs are required for splicing | SECTION 5.24.5 © 2004. Virtual Text / www.ergito.com

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Colloquially, they are sometimes known as snurps and scyrps. There is also a class of small RNAs found in the nucleolus, called snoRNAs, which are involved in processing ribosomal RNA (see Molecular Biology 5.24.22 Small RNAs are required for rRNA processing). The snRNPs involved in splicing, together with many additional proteins, form a large particulate complex, called the spliceosome. Isolated from the in vitro splicing systems, it comprises a 50-60S ribonucleoprotein particle. The spliceosome may be formed in stages as the snRNPs join, proceeding through several "presplicing complexes." The spliceosome is a large body, greater in mass than the ribosome. Figure 24.8 summarizes the components of the spliceosome (3210). The 5 snRNAs account for more than a quarter of the mass; together with their 45 associated proteins, they account for almost half of the mass. Some 70 other proteins found in the spliceosome are described as splicing factors. They include proteins required for assembly of the spliceosome, proteins required for it to bind to the RNA substrate, and proteins involved in the catalytic process. In addition to these proteins, another ~30 proteins associated with the spliceosome have been implicated in acting at other stages of gene expression, suggesting that the spliceosome may serve as a coordinating apparatus.

Figure 24.8 The spliceosome is ~12 MDa. 5 snRNAPs account for almost half of the mass. The remaining proteins include known splicing factors and also proteins that are involved in other stages of gene expression.

The spliceosome forms on the intact precursor RNA and passes through an intermediate state in which it contains the individual 5 ′ exon linear molecule and the right lariat-intron-exon. Little spliced product is found in the complex, which suggests that it is usually released immediately following the cleavage of the 3 ′ site and ligation of the exons. snRNAs are required for splicing | SECTION 5.24.5 © 2004. Virtual Text / www.ergito.com

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We may think of the snRNP particles as being involved in building the structure of the spliceosome. Like the ribosome, the spliceosome depends on RNA-RNA interactions as well as protein-RNA and protein-protein interactions. Some of the reactions involving the snRNPs require their RNAs to base pair directly with sequences in the RNA being spliced; other reactions require recognition between snRNPs or between their proteins and other components of the spliceosome. The importance of snRNA molecules can be tested directly in yeast by making mutations in their genes. Mutations in 5 snRNA genes are lethal and prevent splicing. All of the snRNAs involved in splicing can be recognized in conserved forms in animal, bird, and insect cells. The corresponding RNAs in yeast are often rather larger, but conserved regions include features that are similar to the snRNAs of higher eukaryotes. The snRNPs involved in splicing are U1, U2, U5, U4, and U6. They are named according to the snRNAs that are present. Each snRNP contains a single snRNA and several (98% of splicing junctions in the human genome). 9 proteins and is called the EJC (exon junction complex).

Splicing is connected to export of mRNA | SECTION 5.24.10 © 2004. Virtual Text / www.ergito.com

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Figure 24.16 The EJC (exon junction complex) binds to RNA by recognizing the splicing complex.

The EJC is involved in several functions of spliced mRNAs (for review see 3428). Some of the proteins of the EJC are directly involved in these functions, and others recruit additional proteins for particular functions. The first contact in assembling the EJC is made with one of the splicing factors (2072; 2073; 2074). Then after splicing, the EJC remains attached to the mRNA just upstream of the exon-exon junction (2071; 2075; 3232; 3233). The EJC is not associated with RNAs transcribed from genes that lack introns, so its involvement in the process is unique for spliced products. If introns are deleted from a gene, its RNA product is exported much more slowly to the cytoplasm (2069). This suggests that the intron may provide a signal for attachment of the export apparatus. We can now account for this phenomenon in terms of a series of protein interactions, as shown in Figure 24.17. The EJC includes a group of proteins called the REF family (the best characterized member is called Aly) (2070). The REF proteins in turn interact with a transport protein (variously called TAP and Mex) which has direct responsibility for interaction with the nuclear pore (for review see 2422).


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Figure 24.17 A REF protein binds to a splicing factor and remains with the spliced RNA product. REF binds to an export factor that binds to the nuclear pore.

A similar system may be used to identify a spliced RNA so that nonsense mutations prior to the last exon trigger its degradation in the cytoplasm (see Molecular Biology 2.5.14 Nonsense mutations trigger a surveillance system). Last updated on 1-6-2003


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Reviews 2422. Reed, R. and Hurt, E. (2002). A conserved mRNA export machinery coupled to pre-mRNA splicing. Cell 108, 523-531. 3428. Dreyfuss, G., Kim, V. N., and Kataoka, N. (2002). Messenger-RNA-binding proteins and the messages they carry. Nat. Rev. Mol. Cell Biol. 3, 195-205.

References 2069. Luo, M. J. and Reed, R. (1999). Splicing is required for rapid and efficient mRNA export in metazoans. Proc. Natl. Acad. Sci. USA 96, 14937-14942. 2070. Rodrigues, J. P., Rode, M., Gatfield, D., Blencowe, B., Blencowe, M., and Izaurralde, E. (2001). REF proteins mediate the export of spliced and unspliced mRNAs from the nucleus. Proc. Natl. Acad. Sci. USA 98, 1030-1035. 2071. Le Hir, H., Izaurralde, E., Maquat, L. E., and Moore, M. J. (2000). The spliceosome deposits multiple proteins 20-24 nucleotides upstream of mRNA exon-exon junctions. EMBO J. 19, 6860-6869. 2072. Kataoka, N., Yong, J., Kim, V. N., Velazquez, F., Perkinson, R. A., Wang, F., and Dreyfuss, G. (2000). Pre-mRNA splicing imprints mRNA in the nucleus with a novel RNA-binding protein that persists in the cytoplasm. Mol. Cell 6, 673-682. 2073. Luo, M. L., Zhou, Z., Magni, K., Christoforides, C., Rappsilber, J., Mann, M., and Reed, R. (2001). Pre-mRNA splicing and mRNA export linked by direct interactions between UAP56 and Aly. Nature 413, 644-647. 2074. Strasser, K. and Hurt, E. (2001). Splicing factor Sub2p is required for nuclear mRNA export through its interaction with Yra1p. Nature 413, 648-652. 2075. Zhou, Z., Luo, M. J., Straesser, K., Katahira, J., Hurt, E., and Reed, R. (2000). The protein Aly links pre-messenger-RNA splicing to nuclear export in metazoans. Nature 407, 401-405. 3232. Le Hir, H., Gatfield, D., Izaurralde, E., and Moore, M. J. (2001). The exon-exon junction complex provides a binding platform for factors involved in mRNA export and nonsense-mediated mRNA decay. EMBO J. 20, 4987-4997. 3233. Reichert, V. L., Le Hir, H., Jurica, M. S., and Moore, M. J. (2002). 5 ′ exon interactions within the human spliceosome establish a framework for exon junction complex structure and assembly. Genes Dev. 16, 2778-2791.



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5.24.11 Group II introns autosplice via lariat formation Key Terms Autosplicing (Self-splicing) describes the ability of an intron to excise itself from an RNA by a catalytic action that depends only on the sequence of RNA in the intron. Key Concepts

• Group II introns excise themselves from RNA by an autocatalytic splicing event. • The splice junctions and mechanism of splicing of group II introns are similar to splicing of nuclear introns.

• A group II intron folds into a secondary structure that generates a catalytic site resembling the structure of U6-U2-nuclear intron.

Introns in protein-coding genes (in fact, in all genes except nuclear tRNA-coding genes) can be divided into three general classes. Nuclear pre-mRNA introns are identified only by the possession of the GU...AG dinucleotides at the 5 ′ and 3 ′ ends and the branch site/pyrimidine tract near the 3 ′ end. They do not show any common features of secondary structure. Group I and group II introns are found in organelles and in bacteria. (Group I introns are found also in the nucleus in lower eukaryotes.) Group I and group II introns are classified according to their internal organization. Each can be folded into a typical type of secondary structure. The group I and group II introns have the remarkable ability to excise themselves from an RNA. This is called autosplicing. Group I introns are more common than group II introns. There is little relationship between the two classes, but in each case the RNA can perform the splicing reaction in vitro by itself, without requiring enzymatic activities provided by proteins; however, proteins are almost certainly required in vivo to assist with folding (see Molecular Biology 5.26 Catalytic RNA). Figure 24.18 shows that three classes of introns are excised by two successive transesterifications (shown previously for nuclear introns in Figure 24.6). In the first reaction, the 5 ′ exon-intron junction is attacked by a free hydroxyl group (provided by an internal 2 ′ –OH position in nuclear and group II introns, and by a free guanine nucleotide in group I introns). In the second reaction, the free 3 ′ –OH at the end of the released exon in turn attacks the 3 ′ intron-exon junction.

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Figure 24.18 Three classes of splicing reactions proceed by two transesterifications. First, a free OH group attacks the exon 1–intron junction. Second, the OH created at the end of exon 1 attacks the intron–exon 2 junction.

There are parallels between group II introns and pre-mRNA splicing. Group II mitochondrial introns are excised by the same mechanism as nuclear pre-mRNAs, via a lariat that is held together by a 5 ′ –2 ′ bond. An example of a lariat produced by splicing a group II intron is shown in Figure 24.19. When an isolated group II RNA is incubated in vitro in the absence of additional components, it is able to perform the splicing reaction. This means that the two transesterification reactions shown in Figure 24.18 can be performed by the group II intron RNA sequence itself. Because the number of phosphodiester bonds is conserved in the reaction, an external supply of energy is not required; this could have been an important feature in the evolution of splicing (for review see 260).


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Figure 24.19 Splicing releases a mitochondrial group II intron in the form of a stable lariat. Photograph kindly provided by Leslie Grivell and Annika Arnberg.

A group II intron forms into a secondary structure that contains several domains formed by base-paired stems and single-stranded loops. Domain 5 is separated by 2 bases from domain 6, which contains an A residue that donates the 2 ′ –OH group for the first transesterification. This constitutes a catalytic domain in the RNA. Figure 24.20 compares this secondary structure with the structure formed by the combination of U6 with U2 and of U2 with the branch site. The similarity suggests that U6 may have a catalytic role.


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Figure 24.20 Nuclear splicing and group II splicing involve the formation of similar secondary structures. The sequences are more specific in nuclear splicing; group II splicing uses positions that may be occupied by either purine (R) or either pyrimidine (Y).

The features of group II splicing suggest that splicing evolved from an autocatalytic reaction undertaken by an individual RNA molecule, in which it accomplished a controlled deletion of an internal sequence. Probably such a reaction requires the RNA to fold into a specific conformation, or series of conformations, and would occur exclusively in cis conformation. The ability of group II introns to remove themselves by an autocatalytic splicing event stands in great contrast to the requirement of nuclear introns for a complex splicing apparatus. We may regard the snRNAs of the spliceosome as compensating for the lack of sequence information in the intron, and providing the information required to form particular structures in RNA. The functions of the snRNAs may Group II introns autosplice via lariat formation | SECTION 5.24.11 © 2004. Virtual Text / www.ergito.com

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have evolved from the original autocatalytic system. These snRNAs act in trans upon the substrate pre-mRNA; we might imagine that the ability of U1 to pair with the 5 ′ splice site, or of U2 to pair with the branch sequence, replaced a similar reaction that required the relevant sequence to be carried by the intron. So the snRNAs may undergo reactions with the pre-mRNA substrate and with one another that have substituted for the series of conformational changes that occur in RNAs that splice by group II mechanisms. In effect, these changes have relieved the substrate pre-mRNA of the obligation to carry the sequences needed to sponsor the reaction. As the splicing apparatus has become more complex (and as the number of potential substrates has increased), proteins have played a more important role.


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Reviews 260. Michel, F. and Ferat, J.-L. (1995). Structure and activities of group II introns. Annu. Rev. Biochem. 64, 435-461.



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5.24.12 Alternative splicing involves differential use of splice junctions Key Terms Alternative splicing describes the production of different RNA products from a single product by changes in the usage of splicing junctions. Key Concepts

• Specific exons may be excluded or included in the RNA product by using or failing to use a pair of splicing junctions.

• Exons may be extended by changing one of the splice junctions to use an alternative junction.

• Sex determination in Drosophila involves a series of alternative splicing events in genes coding for successive products of a pathway.

• P elements of Drosophila show germline-specific alternative splicing.

When an interrupted gene is transcribed into an RNA that gives rise to a single type of spliced mRNA, there is no ambiguity in assignment of exons and introns. But the RNAs of some genes follow patterns of alternative splicing, when a single gene gives rise to more than one mRNA sequence. In some cases, the ultimate pattern of expression is dictated by the primary transcript, because the use of different startpoints or the generation of alternative 3 ′ ends alters the pattern of splicing. In other cases, a single primary transcript is spliced in more than one way, and internal exons are substituted, added, or deleted. In some cases, the multiple products all are made in the same cell, but in others the process is regulated so that particular splicing patterns occur only under particular conditions (for review see 246). One of the most pressing questions in splicing is to determine what controls the use of such alternative pathways. Proteins that intervene to bias the use of alternative splice sites have been identified in two ways. In some mammalian systems, it has been possible to characterize alternative splicing in vitro, and to identify proteins that are required for the process. In D. melanogaster, aberrations in alternative splicing may be caused either by mutations in the genes that are alternatively spliced or in the genes whose products are necessary for the reaction. Figure 24.21 shows examples of alternative splicing in which one splice site remains constant, but the other varies. The large T/ small t antigens of SV40 and the products of the adenovirus E1A region are generated by connecting a varying 5 ′ site to a constant 3 ′ site. In the case of the T/t antigens, the 5 ′ site used for T antigen removes a termination codon that is present in the t antigen mRNA, so that T antigen is larger than t antigen. In the case of the E1A transcripts, one of the 5 ′ sites connects to the last exon in a different reading frame, again making a significant change in the C-terminal part of the protein. In these examples, all the relevant Alternative splicing involves differential use of splice junctions | SECTION 5.24.12 © 2004. Virtual Text / www.ergito.com

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splicing events take place in every cell in which the gene is expressed, so all the protein products are made.

Figure 24.21 Alternative forms of splicing may generate a variety of protein products from an individual gene. Changing the splice sites may introduce termination codons (shown by asterisks) or change reading frames.

There are differences in the ratios of T/t antigens in different cell types. A protein extracted from cells that produce relatively more small t antigen can cause preferential production of small t RNA in extracts from other cell types. This protein, which was called ASF (alternative splicing factor), turns out to be the same as the splicing factor SF2, which is required for early steps in spliceosome assembly and for the first cleavage-ligation reaction (see Figure 24.13). ASF/SF2 is an RNA-binding protein in the SR family. When a pre-mRNA has more than one 5 ′ splice site preceding a single 3 ′ splice site, increased concentrations of ASF/SF2 promote use of the 5 ′ site nearest to the 3 ′ site at the expense of the other site (3316; 3317). This effect of ASF/SF2 can be counteracted by another splicing factor, SF5.

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The exact molecular roles of the factors in controlling splice utilization are not yet known, but we see in general terms that alternative splicing involving different 5 ′ sites may be influenced by proteins involved in spliceosome assembly. In the case of T/t antigens, the effect probably rests on increased binding of the SR proteins to the site that is preferentially used. Alternative splicing also may be influenced by repression of one site. Exons 2 and 3 of the mouse troponin T gene are mutually exclusive; exon 2 is used in smooth muscle, but exon 3 is used in other tissues. Smooth muscle contains proteins that bind to repeated elements located on either side of exon 3, and which prevent use of the 3 ′ and 5 ′ sites that are needed to include it. The pathway of sex determination in D. melanogaster involves interactions between a series of genes in which alternative splicing events distinguish male and female. The pathway takes the form illustrated in Figure 24.22, in which the ratio of X chromosomes to autosomes determines the expression of sxl, and changes in expression are passed sequentially through the other genes to dsx, the last in the pathway.

Figure 24.22 Sex determination in D. melanogaster involves a pathway in which different splicing events occur in females. Blocks at any stage of the pathway result in male development.

The pathway starts with sex-specific splicing of sxl. Exon 3 of the sxl gene contains a termination codon that prevents synthesis of functional protein. This exon is included in the mRNA produced in males, but is skipped in females. (Exon skipping Alternative splicing involves differential use of splice junctions | SECTION 5.24.12 © 2004. Virtual Text / www.ergito.com

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illustrated for another example in Figure 24.23.) As a result, only females produce Sxl protein. The protein has a concentration of basic amino acids that resembles other RNA-binding proteins. The presence of Sxl protein changes the splicing of the transformer (tra) gene. Figure 24.21 shows that this involves splicing a constant 5 ′ site to alternative 3 ′ sites. One splicing pattern occurs in both males and females, and results in an RNA that has an early termination codon. The presence of Sxl protein inhibits usage of the normal 3 ′ splice site by binding to the polypyrimidine tract at its branch site (3319). When this site is skipped, the next 3 ′ site is used. This generates a female-specific mRNA that codes for a protein. So tra produces a protein only in females; this protein is a splicing regulator. tra2 has a similar function in females (but is also expressed in the male germline). The Tra and Tra2 proteins are SR splicing factors that act directly upon the target transcripts. Tra and Tra2 cooperate (in females) to affect the splicing of dsx. Figure 24.23 shows examples of cases in which splice sites are used to add or to substitute exons or introns, again with the consequence that different protein products are generated. In the doublesex (dsx) gene, females splice the 5 ′ site of intron 3 to the 3 ′ site of that intron; as a result translation terminates at the end of exon 4. Males splice the 5 ′ site of intron 3 directly to the 3 ′ site of intron 4, thus omitting exon 4 from the mRNA, and allowing translation to continue through exon 6. The result of the alternative splicing is that different proteins are produced in each sex: the male product blocks female sexual differentiation, while the female product represses expression of male-specific genes.


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Figure 24.23 Alternative splicing events that involve both sites may cause exons to be added or substituted.

Alternative splicing of dsx RNA is controlled by competition between 3 ′ splice sites. dsx RNA has an element downstream of the leftmost 3 ′ splice site that is bound by Tra2; Tra and SR proteins associate with Tra2 at the site, which becomes an enhancer that assists binding of U2AF at the adjacent pyrimidine tract (3320; 3321). This commits the formation of the spliceosome to use this 3 ′ site in females rather than the alternative 3 ′ site. The proteins recognize the enhancer cooperatively, possibly relying on formation of some secondary structure as well as sequence per se. Sex determination therefore has a pleasing symmetry: the pathway starts with a female-specific splicing event that causes omission of an exon that has a termination codon, and ends with a female-specific splicing event that causes inclusion of an exon that has a termination codon. The events have different molecular bases. At the first control point, Sxl inhibits the default splicing pattern. At the last control point, Tra and Tra2 cooperate to promote the female-specific splice. The Tra and Tra2 proteins are not needed for normal splicing, because in their absence flies develop normally (as males). As specific regulators, they need not necessarily participate in the mechanics of the splicing reaction; in this respect they differ from SF2, which is a factor required for general splicing, but can also Alternative splicing involves differential use of splice junctions | SECTION 5.24.12 © 2004. Virtual Text / www.ergito.com

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influence choice of alternative splice sites. P elements of D. melanogaster show a tissue-specific splicing pattern. In somatic cells, there are two splicing events, but in germline an additional splicing event removes another intron. Because a termination codon lies in the germline-specific intron, a longer protein (with different properties) is produced in germline. We discuss the consequences for control of transposition in Molecular Biology 4.16.15 P elements are activated in the germline, and note for now that the tissue specificity results from differences in the splicing apparatus. The default splicing pathway of the P element pre-mRNA when the RNA is subjected to a heterologous (human) splicing extract is the germline pattern, in which intron 3 is excised. But extracts of somatic cells of D. melanogaster contain a protein that inhibits excision of this intron. The protein binds to sequences in exon 3; if these sequences are deleted, the intron is excised. The function of the protein is therefore probably to repress association of the spliceosome with the 5 ′ site of intron 3.


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Reviews 246. Green, M. R. (1991). Biochemical mechanisms of constitutive and regulated pre-mRNA splicing. Annu. Rev. Cell Biol. 7, 559-599.

References 3316. Wu, J. Y. and Maniatis, T. (1993). Specific interactions between proteins implicated in splice site selection and regulated alternative splicing. Cell 75, 1061-1070. 3317. Sun, Q., Mayeda, A., Hampson, R. K., Krainer, A. R., and Rottman, F. M. (1993). General splicing factor SF2/ASF promotes alternative splicing by binding to an exonic splicing enhancer. Genes Dev. 7, 2598-2608. 3319. Handa, N., Nureki, O., Kurimoto, K., Kim, I., Sakamoto, H., Shimura, Y., Muto, Y., and Yokoyama, S. (1999). Structural basis for recognition of the tra mRNA precursor by the Sex-lethal protein. Nature 398, 579-585. 3320. Tian, M. and Maniatis, T. (1993). A splicing enhancer complex controls alternative splicing of doublesex pre-mRNA. Cell 74, 105-114. 3321. Lynch, K. W. and Maniatis, T. (1996). Assembly of specific SR protein complexes on distinct regulatory elements of the Drosophila doublesex splicing enhancer. Genes Dev. 10, 2089-2101.



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5.24.13 trans-splicing reactions use small RNAs Key Terms SL RNA (Spliced leader RNA) is a small RNA that donates an exon in the trans-splicing reaction of trypanosomes and nematodes. Key Concepts

• Splicing reactions usually occur only in cis between splice junctions on the same molecule of RNA.

• trans-splicing occurs in trypanosomes and worms where a short sequence (SL RNA) is spliced to the 5 ′ ends of many precursor mRNAs.

• SL RNA has a structure resembling the Sm-binding site of U snRNAs and may play an analogous role in the reaction.

In both mechanistic and evolutionary terms, splicing has been viewed as an intramolecular reaction, amounting essentially to a controlled deletion of the intron sequences at the level of RNA. In genetic terms, splicing occurs only in cis. This means that only sequences on the same molecule of RNA can be spliced together. The upper part of Figure 24.24 shows the normal situation. The introns can be removed from each RNA molecule, allowing the exons of that RNA molecule to be spliced together, but there is no intermolecular splicing of exons between different RNA molecules. We cannot say that trans splicing never occurs between pre-mRNA transcripts of the same gene, but we know that it must be exceedingly rare, because if it were prevalent the exons of a gene would be able to complement one another genetically instead of belonging to a single complementation group.

trans-splicing reactions use small RNAs | SECTION 5.24.13 © 2004. Virtual Text / www.ergito.com

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Figure 24.24 Splicing usually occurs only in cis between exons carried on the same physical RNA molecule, but trans splicing can occur when special constructs are made that support base pairing between introns.

Some manipulations can generate trans-splicing. In the example illustrated in the lower part of Figure 24.24, complementary sequences were introduced into the introns of two RNAs. Base pairing between the complements should create an H-shaped molecule. This molecule could be spliced in cis, to connect exons that are covalently connected by an intron, or it could be spliced in trans, to connect exons of the juxtaposed RNA molecules. Both reactions occur in vitro. Another situation in which trans-splicing is possible in vitro occurs when substrate RNAs are provided in the form of one containing a 5 ′ splice site and the other containing a 3 ′ splice site together with appropriate downstream sequences (which may be either the next 5 ′ splice site or a splicing enhancer). In effect, this mimics splicing by exon definition (see the right side of Figure 24.12), and shows that in vitro it is not necessary for the left and right splice sites to be on the same RNA molecule. These results show that there is no mechanistic impediment to trans-splicing. They exclude models for splicing that require processive movement of a spliceosome along the RNA. It must be possible for a spliceosome to recognize the 5 ′ and 3 ′ splice sites of different RNAs when they are in close proximity. Although trans-splicing is rare, it occurs in vivo in some special situations. One is revealed by the presence of a common 35 base leader sequence at the end of numerous mRNAs in the trypanosome. But the leader sequence is not coded trans-splicing reactions use small RNAs | SECTION 5.24.13 © 2004. Virtual Text / www.ergito.com

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upstream of the individual transcription units. Instead it is transcribed into an independent RNA, carrying additional sequences at its 3 ′ end, from a repetitive unit located elsewhere in the genome. Figure 24.25 shows that this RNA carries the 35 base leader sequence followed by a 5 ′ splice site sequence. The sequences coding for the mRNAs carry a 3 ′ splice site just preceding the sequence found in the mature mRNA (730).

Figure 24.25 The SL RNA provides an exon that is connected to the first exon of an mRNA by trans-splicing. The reaction involves the same interactions as nuclear cis-splicing, but generates a Y-shaped RNA instead of a lariat.

When the leader and the mRNA are connected by a trans-splicing reaction, the 3 ′ region of the leader RNA and the 5 ′ region of the mRNA in effect comprise the 5 ′ and 3 ′ halves of an intron. When splicing occurs, a 5 ′ –2 ′ link forms by the usual reaction between the GU of the 5 ′ intron and the branch sequence near the AG of the 3 ′ intron. Because the two parts of the intron are not covalently linked, this generates a Y-shaped molecule instead of a lariat (729). A similar situation is presented by the expression of actin genes in C. elegans. Three actin mRNAs (and some other RNAs) share the same 22 base leader sequence at the 5 ′ terminus. The leader sequence is not coded in the actin gene, but is transcribed independently as part of a 100 base RNA coded by a gene elsewhere. trans-splicing also occurs in chloroplasts (731). The RNA that donates the 5 ′ exon for trans splicing is called the SL RNA (spliced leader RNA). The SL RNAs found in several species of trypanosomes and also in the nematode (C. elegans) have some common features. They fold into a common secondary structure that has three stem-loops and a single-stranded region that trans-splicing reactions use small RNAs | SECTION 5.24.13 © 2004. Virtual Text / www.ergito.com

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resembles the Sm-binding site. The SL RNAs therefore exist as snRNPs that count as members of the Sm snRNP class. Trypanosomes possess the U2, U4, and U6 snRNAs, but do not have U1 or U5 snRNAs. The absence of U1 snRNA can be explained by the properties of the SL RNA, which can carry out the functions that U1 snRNA usually performs at the 5 ′ splice site; thus SL RNA in effect consists of an snRNA sequence possessing U1 function, linked to the exon-intron site that it recognizes. There are two types of SL RNA in C. elegans. SL1 RNA (the first to be discovered) is used for splicing to coding sequences that are preceded only by 5 ′ nontranslated regions (the most common situation). SL2 RNA is used in cases in which a pre-mRNA contains two coding sequences; it is spliced to the second sequence, thus releasing it from the first, and allowing it to be used as an independent mRNA (732; 733; for review see 250). About 15% of all genes in C. elegans are organized in transcription units that include more than one gene (most often 2-3 genes) (2862). The significance of this form of organization for control of gene expression is not clear. These transcription units do not generally resemble operons where the genes function coordinately in a pathway. The trans-splicing reaction of the SL RNA may represent a step towards the evolution of the pre-mRNA splicing apparatus. The SL RNA provides in cis the ability to recognize the 5 ′ splice site, and this probably depends upon the specific conformation of the RNA. The remaining functions required for splicing are provided by independent snRNPs. The SL RNA can function without participation of proteins like those in U1 snRNP, which suggests that the recognition of the 5 ′ splice site depends directly on RNA. Last updated on 8-22-2002


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Reviews 250. Nilsen, T. (1993). trans-splicing of nematode pre-mRNA. Annu. Rev. Immunol. 47, 413-440.

References 729. Murphy, W. J., Watkins, K. P., and Agabian, N. (1986). Identification of a novel Y branch structure as an intermediate in trypanosome mRNA processing: evidence for trans-splicing. Cell 47, 517-525. 730. Sutton, R. and Boothroyd, J. C. (1986). Evidence for trans-splicing in trypanosomes. Cell 47, 527-535. 731. Krause, M. and Hirsh, D. (1987). A trans-spliced leader sequence on actin mRNA in C. elegans. Cell 49, 753-761. 732. Huang, X. Y. and Hirsh, D. (1989). A second trans-spliced RNA leader sequence in the nematode C. elegans. Proc. Natl. Acad. Sci. USA 86, 8640-8644. 733. Hannon, G. J. et al. (1990). trans-splicing of nematode pre-mRNA in vitro. Cell 61, 1247-1255. 2862. Blumenthal, T., Evans, D., Link, C. D., Guffanti, A., Lawson, D., Thierry-Mieg, J., Thierry-Mieg, D., Chiu, W. L., Duke, K., Kiraly, M., and Kim, S. K. (2002). A global analysis of C. elegans operons. Nature 417, 851-854.



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5.24.14 Yeast tRNA splicing involves cutting and rejoining Key Terms An RNA ligase is an enzyme that functions in tRNA splicing to make a phosphodiester bond between the two exon sequences that are generated by cleavage of the intron. Key Concepts

• tRNA splicing occurs by successive cleavage and ligation reactions.

Most splicing reactions depend on short consensus sequences and occur by transesterification reactions in which breaking and making of bonds is coordinated. The splicing of tRNA genes is achieved by a different mechanism that relies upon separate cleavage and ligation reactions. Some 59 of the 272 nuclear tRNA genes in the yeast S. cerevisiae are interrupted. Each has a single intron, located just one nucleotide beyond the 3 ′ side of the anticodon. The introns vary in length from 14-60 bp. Those in related tRNA genes are related in sequence, but the introns in tRNA genes representing different amino acids are unrelated. There is no consensus sequence that could be recognized by the splicing enzymes. This is also true of interrupted nuclear tRNA genes of plants, amphibians, and mammals. All the introns include a sequence that is complementary to the anticodon of the tRNA. This creates an alternative conformation for the anticodon arm in which the anticodon is base paired to form an extension of the usual arm. An example is drawn in Figure 24.26. Only the anticodon arm is affected – the rest of the molecule retains its usual structure.

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Figure 24.26 The intron in yeast tRNAPhe base pairs with the anticodon to change the structure of the anticodon arm. Pairing between an excluded base in the stem and the intron loop in the precursor may be required for splicing.

The exact sequence and size of the intron is not important. Most mutations in the intron do not prevent splicing. Splicing of tRNA depends principally on recognition of a common secondary structure in tRNA rather than a common sequence of the intron. Regions in various parts of the molecule are important, including the stretch between the acceptor arm and D arm, in the T ψ C arm, and especially the anticodon arm. This is reminiscent of the structural demands placed on tRNA for protein synthesis (see Molecular Biology 2.6 Protein synthesis). The intron is not entirely irrelevant, however. Pairing between a base in the intron loop and an unpaired base in the stem is required for splicing. Mutations at other positions that influence this pairing (for example, to generate alternative patterns for pairing) influence splicing. The rules that govern availability of tRNA precursors for splicing resemble the rules that govern recognition by aminoacyl-tRNA synthetases (see Molecular Biology 2.7.9 tRNAs are charged with amino acids by synthetases). In a temperature-sensitive mutant of yeast that fails to remove the introns, the interrupted precursors accumulate in the nucleus. The precursors can be used as substrates for a cell-free system extracted from wild-type cells. The splicing of the precursor can be followed by virtue of the resulting size reduction. This is seen by the change in position of the band on gel electrophoresis, as illustrated in Figure 24.27. The reduction in size can be accounted for by the appearance of a band representing the intron.


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Figure 24.27 Splicing of yeast tRNA in vitro can be followed by assaying the RNA precursor and products by gel electrophoresis.

The cell-free extract can be fractionated by assaying the ability to splice the tRNA. The in vitro reaction requires ATP. Characterizing the reactions that occur with and without ATP shows that the two separate stages of the reaction are catalyzed by different enzymes. • The first step does not require ATP. It involves phosphodiester bond cleavage by an atypical nuclease reaction. It is catalyzed by an endonuclease. • The second step requires ATP and involves bond formation; it is a ligation reaction, and the responsible enzyme activity is described as an RNA ligase. Last updated on 8-29-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.24.14


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5.24.15 The splicing endonuclease recognizes tRNA Key Concepts

• An endonuclease cleaves the tRNA precursors at both ends of the intron. • The yeast endonuclease is a heterotetramer, with two (related) catalytic subunits. • It uses a measuring mechanism to determine the sites of cleavage by their positions relative to a point in the tRNA structure.

• The archaeal nuclease has a simpler structure and recognizes a bulge-helix-bulge structural motif in the substrate.

The endonuclease is responsible for the specificity of intron recognition. It cleaves the precursor at both ends of the intron. The yeast endonuclease is a heterotetrameric protein. Its activities are illustrated in Figure 24.28. The related subunits Sen34 and Sen2 cleave the 3 ′ and 5 ′ splice sites, respectively. Subunit Sen54 may determine the sites of cleavage by "measuring" distance from a point in the tRNA structure. This point is in the elbow of the (mature) L-shaped structure (2878). The role of subunit Sen15 is not known, but its gene is essential in yeast. The base pair that forms between the first base in the anticodon loop and the base preceding the 3 ′ splice site is required for 3 ′ splice site cleavage (735; 736; 737; 2880).

Figure 24.28 The 3 ′ and 5 ′ cleavages in S. cerevisiae pre-tRNA are catalyzed by different subunits of the endonuclease. Another subunit may determine location of the cleavage sites by measuring distance from the mature structure. The AI base pair is also important.

An interesting insight into the evolution of tRNA splicing is provided by the endonucleases of archaea. These are homodimers or homotetramers, in which each subunit has an active site (although only two of the sites function in the tetramer) that The splicing endonuclease recognizes tRNA | SECTION 5.24.15 © 2004. Virtual Text / www.ergito.com

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cleaves one of the splice sites (2879). The subunit has sequences related to the sequences of the active sites in the Sen34 and Sen2 subunits of the yeast enzyme. However, the archaeal enzymes recognize their substrates in a different way. Instead of measuring distance from particular sequences, they recognize a structural feature, called the bulge-helix-bulge. Figure 24.29 shows that cleavage occurs in the two bulges (2877; 2876).

Figure 24.29 Archaeal tRNA splicing endonuclease cleaves each strand at a bulge in a bulge-helix-bulge motif.

So the origin of splicing of tRNA precedes the separation of the archaea and the eukaryotes. If it originated by insertion of the intron into tRNAs, this must have been a very ancient event. Last updated on 8-29-2002

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References 735. Reyes, V. M. and Abelson, J. (1988). Substrate recognition and splice site determination in yeast tRNA splicing. Cell 55, 719-730. 736. Mattoccia, E. et al. (1988). Site selection by the tRNA splicing endonuclease of X. laevis. Cell 55, 731-738. 737. Baldi, I. M. et al. (1992). Participation of the intron in the reaction catalyzed by the Xenopus tRNA splicing endonuclease. Science 255, 1404-1408. 2876. Diener, J. L. and Moore, P. B. (1998). Solution structure of a substrate for the archaeal pre-tRNA splicing endonucleases: the bulge-helix-bulge motif. Mol. Cell 1, 883-894. 2877. Lykke-Andersen, J. and Garrett, R. A. (1997). RNA-protein interactions of an archaeal homotetrameric splicing endoribonuclease with an exceptional evolutionary history. EMBO J. 16, 6290-6300. 2878. Trotta, C. R., Miao, F., Arn, E. A., Stevens, S. W., Ho, C. K., Rauhut, R., and Abelson, J. N. (1997). The yeast tRNA splicing endonuclease: a tetrameric enzyme with two active site subunits homologous to the archaeal tRNA endonucleases. Cell 89, 849-858. 2879. Kleman-Leyer, K., Armbruster, D. W., and Daniels, C. J. (2000). Properties of H. volcanii tRNA intron endonuclease reveal a relationship between the archaeal and eucaryal tRNA intron processing systems. Cell 89, 839-847. 2880. Di Nicola Negri, E., Fabbri, S., Bufardeci, E., Baldi, M. I., Mattoccia, E., and Tocchini-Valentini, G. P. (1997). The eucaryal tRNA splicing endonuclease recognizes a tripartite set of RNA elements. Cell 89, 859-866.


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5.24.16 tRNA cleavage and ligation are separate reactions Key Concepts

• Release of the intron generates two half-tRNAs that pair to form the mature structure.

• The halves have the unusual ends 5 ′ hydroxyl and 2 ′ –3 ′ cyclic phosphate. • The 5 ′ –OH end is phosphorylated by a polynucleotide kinase, the cyclic

phosphate group is opened by phosphodiesterase to generate a 2 ′ –phosphate terminus and 3 ′ –OH group, exon ends are joined by an RNA ligase, and the 2 ′ –phosphate is removed by a phosphatase.

The overall tRNA splicing reaction is summarized in Figure 24.30. The products of cleavage are a linear intron and two half-tRNA molecules. These intermediates have unique ends. Each 5 ′ terminus ends in a hydroxyl group; each 3 ′ terminus ends in a 2 ′ ,3 ′ –cyclic phosphate group. (All other known RNA splicing enzymes cleave on the other side of the phosphate bond.)

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Figure 24.30 Splicing of tRNA requires separate nuclease and ligase activities. The exon-intron boundaries are cleaved by the nuclease to generate 2 ′ -3 ′ cyclic phosphate and 5 ′ OH termini. The cyclic phosphate is opened to generate 3 ′ -OH and 2 ′ phosphate groups. The 5 ′ -OH is phosphorylated. After releasing the intron, the tRNA half molecules fold into a tRNA-like structure that now has a 3 ′ -OH, 5 ′ -P break. This is sealed by a ligase.

The two half-tRNAs base pair to form a tRNA-like structure. When ATP is added, the second reaction occurs. Both of the unusual ends generated by the endonuclease must be altered. The cyclic phosphate group is opened to generate a 2 ′ –phosphate terminus. This reaction requires cyclic phosphodiesterase activity. The product has a 2 ′ –phosphate group and a 3 ′ –OH group. The 5 ′ –OH group generated by the nuclease must be phosphorylated to give a 5 ′ –phosphate. This generates a site in which the 3 ′ –OH is next to the 5 ′ –phosphate. Covalent integrity of the polynucleotide chain is then restored by ligase activity. All three activities – phosphodiesterase, polynucleotide kinase, and adenylate synthetase (which provides the ligase function) – are arranged in different functional domains on a single protein. They act sequentially to join the two tRNA halves. The spliced molecule is now uninterrupted, with a 5 ′ –3 ′ phosphate linkage at the site of splicing, but it also has a 2 ′ –phosphate group marking the event. The surplus group must be removed by a phosphatase. Generation of a 2 ′ ,3 ′ –cyclic phosphate also occurs during the tRNA-splicing reaction in plants and mammals. The reaction in plants seems to be the same as in tRNA cleavage and ligation are separate reactions | SECTION 5.24.16 © 2004. Virtual Text / www.ergito.com

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yeast, but the detailed chemical reactions are different in mammals. The yeast tRNA precursors also can be spliced in an extract obtained from the germinal vesicle (nucleus) of Xenopus oocytes. This shows that the reaction is not species-specific. Xenopus must have enzymes able to recognize the introns in the yeast tRNAs. The ability to splice the products of tRNA genes is therefore well conserved, but is likely to have a different origin from the other splicing reactions (such as that of nuclear pre-mRNA). The tRNA-splicing reaction uses cleavage and synthesis of bonds and is determined by sequences that are external to the intron. Other splicing reactions use transesterification, in which bonds are transferred directly, and the sequences required for the reaction lie within the intron. Last updated on 8-29-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.24.16

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5.24.17 The unfolded protein response is related to tRNA splicing Key Concepts

• Ire1p is an inner nuclear membrane protein with its N-terminal domain in the ER lumen, and its C-terminal domain in the nucleus.

• Binding of an unfolded protein to the N-terminal domain activates the C-terminal nuclease by autophosphorylation.

• The activated nuclease cleaves Hac1 mRNA to release an intron and generate exons that are ligated by a tRNA ligase.

• The spliced Hac1 mRNA codes for a transcription factor that activates genes coding for chaperones that help to fold unfolded proteins.

An unusual splicing system that is related to tRNA splicing mediates the response to unfolded proteins in yeast. The accumulation of unfolded proteins in the lumen of the ER triggers a response pathway that leads to increased transcription of genes coding for chaperones that assist protein folding in the ER. A signal must therefore be transmitted from the lumen of the ER to the nucleus. The sensor that activates the pathway is the protein Ire1p. It is an integral membrane protein (Ser/Thr) kinase that has domains on each side of the ER membrane. The N-terminal domain in the lumen of the ER detects the presence of unfolded proteins, presumably by binding to exposed motifs. This causes aggregation of monomers and activates the C-terminal domain on the other side of the membrane by autophosphorylation. Genes that are activated by this pathway have a common promoter element, the UPRE (unfolded protein response element). The transcription factor Hac1p binds to the UPRE, and is produced in response to accumulation of unfolded proteins. The trigger for production of Hac1p is the action of Ire1p on Hac1 mRNA. The operation of the pathway is summarized in Figure 24.31. Under normal conditions, when the pathway is not activated, Hac1 mRNA is translated into a protein that is rapidly degraded. The activation of Ire1p results in the splicing of the Hac1 mRNA to change the sequence of the protein to a more stable form. This form provides the functional transcription factor that activates genes with the UPRE.

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Figure 24.31 The unfolded protein response occurs by activating special splicing of HAC1 mRNA to produce a transcription factor that recognizes the UPRE.

Unusual splicing components are involved in this reaction. Ire1P has an endonuclease activity that acts directly on Hac1 mRNA to cleave the two splicing junctions (739). The two junctions are ligated by the tRNA ligase that acts in the tRNA splicing pathway (738). The endonuclease reaction resembles the cleavage of tRNA during splicing (3322). Where does the modification of Hac1 mRNA occur? Ire1p is probably located in the inner nuclear membrane, with the N-terminal sensor domain in the ER lumen, and the C-terminal kinase/nuclease domain in the nucleus. This would it enable it to act directly on Hac1 RNA before it is exported to the cytoplasm. It also would allow easy access by the tRNA ligase. There is no apparent relationship between the Ire1p nuclease activity and the tRNA splicing endonuclease, so it is not obvious how this specialized system would have evolved.


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References 738. Sidrauski, C., Cox, J. S., and Walter, P. (1996). tRNA ligase is required for regulated mRNA splicing in the unfolded protein response. Cell 87, 405-413. 739. Sidrauski, C. and Walter, P. (1997). The transmembrane kinase Ire1p is a site-specific endonuclease that initiates mRNA splicing in the unfolded protein response. Cell 90, 1031-1039. 3322. Gonzalez, T. N., Sidrauski, C., Dorfler, S., and Walter, P. (1999). Mechanism of non-spliceosomal mRNA splicing in the unfolded protein response pathway. EMBO J. 18, 3119-3132.



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5.24.18 The 3 ′ ends of polI and polIII transcripts are generated by termination Key Concepts

• RNA polymerase I terminates transcription at an 18 base terminator sequence. • RNA polymerase III terminates transcription in poly(U)4 sequence embedded in a G·C-rich sequence.

3 ′ ends of RNAs can be generated in two ways. Some RNA polymerases terminate transcription at a defined (terminator) sequence in DNA, as shown in Figure 24.32. Other RNA polymerases do not show discrete termination, but continue past the site corresponding to the 3 ′ end, which is generated by cleavage of the RNA by an endonuclease, as shown in Figure 24.33.

Figure 24.32 When a 3 ′ end is generated by termination, RNA polymerase and RNA are released at a discrete (terminator) sequence in DNA.

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Figure 24.33 When a 3 ′ end is generated by cleavage, RNA polymerase continues transcription while an endonuclease cleaves at a defined sequence in the RNA.

Information about the termination reaction for eukaryotic RNA polymerases is less detailed than our knowledge of initiation. RNA polymerases I and III have discrete termination events (like bacterial RNA polymerase), but it is not clear whether RNA polymerase II usually terminates in this way. For RNA polymerase I, the sole product of transcription is a large precursor that contains the sequences of the major rRNA. The precursor is subjected to extensive processing. Termination occurs at a discrete site >1000 bp downstream of the mature 3 ′ end, which is generated by cleavage. Termination involves recognition of an 18 base terminator sequence by an ancillary factor. With RNA polymerase III, transcription in vitro generates molecules with the same 5 ′ and 3 ′ ends as those synthesized in vivo. The termination reaction resembles intrinsic termination by bacterial RNA polymerase (see Molecular Biology 3.9.21 There are two types of terminators in E. coli). Termination usually occurs at the second U within a run of 4 U bases, but there is heterogeneity, with some molecules ending in 3 or even 4 U bases. The same heterogeneity is seen in molecules synthesized in vivo, so it seems to be a bona fide feature of the termination reaction. Just like the prokaryotic terminators, the U run is embedded in a G·C-rich region. Although sequences of dyad symmetry are present, they are not needed for termination, since mutations that abolish the symmetry do not prevent the normal completion of RNA synthesis. Nor are any sequences beyond the U run necessary, since all distal sequences can be replaced without any effect on termination. The U run itself is not sufficient for termination, because regions of 4 successive U residues exist within transcription units read by RNA polymerase III. (However, there are no internal U5 runs, which fits with the greater efficiency of termination when the terminator is a U5 rather than U4 sequence.) The critical feature in termination must therefore be the recognition of a U4 sequence in a context that is The 3 ′ ends of polI and polIII transcripts are generated by termination | SECTION 5.24.18 © 2004. Virtual Text / www.ergito.com

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rich in G·C base pairs. How does the termination reaction occur? It cannot rely on the weakness of the rU-dA RNA-DNA hybrid region that lies at the end of the transcript, because often only the first two U residues are transcribed. Perhaps the G·C-rich region plays a role in slowing down the enzyme, but there does not seem to be a counterpart to the hairpin involved in prokaryotic termination. We remain puzzled how the enzyme can respond so specifically to such a short signal. And in contrast with the initiation reaction, which RNA polymerase III cannot accomplish alone, termination seems to be a function of the enzyme itself. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.24.18

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5.24.19 The 3 ′ ends of mRNAs are generated by cleavage and polyadenylation Key Terms Cordycepin is 3 ′ deoxyadenosine, an inhibitor of polyadenylation of RNA. Endonucleases cleave bonds within a nucleic acid chain; they may be specific for RNA or for single-stranded or double-stranded DNA. Poly(A) polymerase is the enzyme that adds the stretch of polyadenylic acid to the 3 ′ of eukaryotic mRNA. It does not use a template. Key Concepts

• The sequence AAUAAA is a signal for cleavage to generate a 3 ′ end of mRNA that is polyadenylated.

• The reaction requires a protein complex that contains a specificity factor, an endonuclease, and poly(A) polymerase.

• The specificity factor and endonuclease cleave RNA downstream of AAUAAA. • The specificity factor and poly(A) polymerase add ~200 A residues processively to the 3 ′ end.

• A·U-rich sequences in the 3 ′ tail control cytoplasmic polyadenylation or deadenylation during Xenopus embryonic development.

It is not clear whether RNA polymerase II actually engages in a termination event at a specific site. It is possible that its termination is only loosely specified. In some transcription units, termination occurs >1000 bp downstream of the site corresponding to the mature 3 ′ end of the mRNA (which is generated by cleavage at a specific sequence). Instead of using specific terminator sequences, the enzyme ceases RNA synthesis within multiple sites located in rather long "terminator regions." The nature of the individual termination sites is not known. The 3 ′ ends of mRNAs are generated by cleavage followed by polyadenylation. Addition of poly(A) to nuclear RNA can be prevented by the analog 3 ′ –deoxyadenosine, also known as cordycepin. Although cordycepin does not stop the transcription of nuclear RNA, its addition prevents the appearance of mRNA in the cytoplasm. This shows that polyadenylation is necessary for the maturation of mRNA from nuclear RNA. Generation of the 3 ′ end is illustrated in Figure 24.34. RNA polymerase transcribes past the site corresponding to the 3 ′ end, and sequences in the RNA are recognized as targets for an endonucleolytic cut followed by polyadenylation. A single processing complex undertakes both the cutting and polyadenylation. The polyadenylation stabilizes the mRNA against degradation from the 3 ′ end. Its 5 ′ end The 3 ′ ends of mRNAs are generated by cleavage and polyadenylation | SECTION 5.24.19 © 2004. Virtual Text / www.ergito.com

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is already stabilized by the cap. RNA polymerase continues transcription after the cleavage, but the 5 ′ end that is generated by the cleavage is unprotected. As a result, the rest of the transcript is rapidly degraded. This makes it difficult to determine what is happening beyond the point of cleavage.

Figure 24.34 The sequence AAUAAA is necessary for cleavage to generate a 3 ′ end for polyadenylation.

A common feature of mRNAs in higher eukaryotes (but not in yeast) is the presence of the highly conserved sequence AAUAAA in the region from 11-30 nucleotides upstream of the site of poly(A) addition. Deletion or mutation of the AAUAAA hexamer prevents generation of the polyadenylated 3 ′ end. The signal is needed for both cleavage and polyadenylation (744; 745; for review see 248). The development of a system in which polyadenylation occurs in vitro opened the route to analyzing the reactions. The formation and functions of the complex that undertakes 3 ′ processing are illustrated in Figure 24.35. Generation of the proper 3 ′ terminal structure requires an endonuclease (consisting of the components CFI and CFII) to cleave the RNA, a poly(A) polymerase (PAP) to synthesize the poly(A) tail, and a specificity component (CPSF) that recognizes the AAUAAA sequence and directs the other activities. A stimulatory factor, CstF, binds to a G-U-rich sequence that is downstream from the cleavage site itself (746).

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Figure 24.35 The 3 ′ processing complex consists of several activities. CPSF and CstF each consist of several subunits; the other components are monomeric. The total mass is >900 kD.

The specificity factor contains 4 subunits, which together bind specifically to RNA containing the sequence AAUAAA. The individual subunits are proteins that have common RNA-binding motifs, but which by themselves bind nonspecifically to RNA. Protein-protein interactions between the subunits may be needed to generate the specific AAUAAA-binding site. CPSF binds strongly to AAUAAA only when CstF is also present to bind to the G-U-rich site. The specificity factor is needed for both the cleavage and polyadenylation reactions. It exists in a complex with the endonuclease and poly(A) polymerase, and this complex usually undertakes cleavage followed by polyadenylation in a tightly coupled manner. The two components CFI and CFII (cleavage factors I and II), together with specificity factor, are necessary and sufficient for the endonucleolytic cleavage. The 3 ′ ends of mRNAs are generated by cleavage and polyadenylation | SECTION 5.24.19 © 2004. Virtual Text / www.ergito.com

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The poly(A) polymerase has a nonspecific catalytic activity. When it is combined with the other components, the synthetic reaction becomes specific for RNA containing the sequence AAUAAA. The polyadenylation reaction passes through two stages. First, a rather short oligo(A) sequence (~10 residues) is added to the 3 ′ end. This reaction is absolutely dependent on the AAUAAA sequence, and poly(A) polymerase performs it under the direction of the specificity factor. In the second phase, the oligo(A) tail is extended to the full ~200 residue length. This reaction requires another stimulatory factor that recognizes the oligo(A) tail and directs poly(A) polymerase specifically to extend the 3 ′ end of a poly(A) sequence. The poly(A) polymerase by itself adds A residues individually to the 3 ′ position. Its intrinsic mode of action is distributive; it dissociates after each nucleotide has been added. However, in the presence of CPSF and PABP (poly(A)-binding protein), it functions processively to extend an individual poly(A) chain. The PABP is a 33 kD protein that binds stoichiometrically to the poly(A) stretch. The length of poly(A) is controlled by the PABP, which in some way limits the action of poly(A) polymerase to ~200 additions of A residues. The limit may represent the accumulation of a critical mass of PABP on the poly(A) chain. PABP binds to the translation initiation factor eIF4G, thus generating a closed loop in which a protein complex contains both the 5 ′ and 3 ′ ends of the mRNA (see Figure 6.20 in Molecular Biology 2.6.9 Eukaryotes use a complex of many initiation factors). Polyadenylation is an important determinant of mRNA function. It may affect both stability and initiation of translation (see Molecular Biology 2.5.10 The 3 ′ terminus is polyadenylated). In embryonic development in some organisms, the presence of poly(A) is used to control translation, and pre-existing mRNAs may either be polyadenylated (to stimulate translation) or deadenylated (to terminate translation). During Xenopus embryonic development, polyadenylation of mRNA in the cytoplasm in Xenopus depends on a specific cis-acting element (the CPE) in the 3 ′ tail. This is another AU-rich sequence, UUUUUAU (2313; 2314). In Xenopus embryos at least two type of cis-acting sequences found in the 3 ′ tail can trigger deadenylation. EDEN (embryonic deadenylation element) is a 17 nucleotide sequence (2310). ARE elements are AU-rich, usually containing tandem repeats of AUUUA (2311). There is a poly(A)-specific RNAase (PARN) that could be involved in the degradation (2312). Of course, deadenylation is not always triggered by specific elements; in some situations (including the normal degradation of mRNA as it ages), poly(A) is degraded unless it is specifically stabilized. Last updated on 1-22-2002


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Reviews 248. Wahle, E. and Keller, W. (1992). The biochemistry of 3 ′ -end cleavage and polyadenylation of messenger RNA precursors. Annu. Rev. Biochem. 61, 419-440.

References 744. Conway, L. and Wickens, M. (1985). A sequence downstream of AAUAAA is required for formation of SV40 late mRNA 3 ′ termini in frog oocytes. Proc. Natl. Acad. Sci. USA 82, 3949-3953. 745. Gil, A. and Proudfoot, N. (1987). Position-dependent sequence elements downstream of AAUAAA are required for efficient rabbit β -globin mRNA 3 ′ end formation. Cell 49, 399-406. 746. Takagaki, Y., Ryner, L. C., and Manley, J. L. (1988). Separation and characterization of a poly(A) polymerase and a cleavage/specificity factor required for pre-mRNA polyadenylation. Cell 52, 731-742. 2310. Bouvet, P., Omilli, F., Arlot-Bonnemains, Y., Legagneux, V., Roghi, C., Bassez, T., and Osborne, H. B. (1994). The deadenylation conferred by the 3 ′ untranslated region of a developmentally controlled mRNA in Xenopus embryos is switched to polyadenylation by deletion of a short sequence element. Mol. Cell Biol. 14, 1893-1900. 2311. Voeltz, G. K. and Steitz, J. A. (1998). AUUUA sequences direct mRNA deadenylation uncoupled from decay during Xenopus early development. Mol. Cell Biol. 18, 7537-7545. 2312. Karner, C. G., Wormington, M., Muckenthaler, M., Schneider, S., Dehlin, E., and Wahle, E. (1998). The deadenylating nuclease (DAN) is involved in poly(A) tail removal during the meiotic maturation of Xenopus oocytes. EMBO J. 17, 5427-5437. 2313. Fox, C. A., Sheets, M. D., and Wickens, M. P. (1989). Poly(A) addition during maturation of frog oocytes: distinct nuclear and cytoplasmic activities and regulation by the sequence UUUUUAU. Genes Dev. 3, 2151-2162. 2314. McGrew, L. L., Dworkin-Rastl, E., Dworkin, M. B., and Richter, J. D. (1989). Poly(A) elongation during Xenopus oocyte maturation is required for translational recruitment and is mediated by a short sequence element. Genes Dev. 3, 803-815.



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5.24.20 Cleavage of the 3 ′ end of histone mRNA may require a small RNA Key Concepts

• Histone mRNAs are not polyadenylated; their 3 ′ ends are generated by a cleavage reaction that depends on the structure of the mRNA.

• The cleavage reaction requires the SLBP to bind to a stem-loop structure, and the U7 snRNA to pair with an adjacent single-stranded region.

Some mRNAs are not polyadenylated. The formation of their 3 ′ ends is therefore different from the coordinated cleavage/polyadenylation reaction. The most prominent members of this mRNA class are the mRNAs coding for histones that are synthesized during DNA replication. Formation of their 3 ′ ends depends upon secondary structure. The structure at the 3' terminus is a highly conserved stem-loop structure, with a stem of 6 bp and a loop of 4 nucleotides. Cleavage occurs 4-5 bases downstream of the stem-loop. Two factors are required for the cleavage reaction: the stem-loop binding protein (SLBP) recognizes the structure (3324); and the U7 snRNA pairs with a purine-rich sequence (the histone downstream element, or HDE) located ~10 nucleotides downstream of the cleavage site (743; 3325). Mutations that prevent formation of the duplex stem of the stem-loop prevent formation of the end of the RNA. Secondary mutations that restore duplex structure (though not necessarily the original sequence) behave as revertants. This suggests that formation of the secondary structure is more important than the exact sequence. The SLBP binds to the stem-loop and then interacts with U7 snRNP to enhance its interaction with the downstream binding site for U7 snRNA (3326). U7 snRNP is a minor snRNP consisting of the 63 nucleotide U7 snRNA and a set of several proteins (including Sm proteins; see Molecular Biology 5.24.5 snRNAs are required for splicing). The reaction between histone H3 mRNA and U7 snRNA is drawn in Figure 24.36. The upstream hairpin and the HDE that pairs with U7 snRNA are conserved in histone H3 mRNAs of several species. The U7 snRNA has sequences towards its 5 ′ end that pair with the histone mRNA consensus sequences. 3 ′ processing is inhibited by mutations in the HDE that reduce ability to pair with U7 snRNA. Compensatory mutations in U7 snRNA that restore complementarity also restore 3 ′ processing (3327). This suggests that U7 snRNA functions by base pairing with the histone mRNA. The sequence of the HDE varies among the various histone mRNAs, with the result that binding of snRNA is not by itself necessarily stable, but requires also the interaction with SLBP

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Figure 24.36 Generation of the 3 ′ end of histone H3 mRNA depends on a conserved hairpin and a sequence that base pairs with U7 snRNA.

Cleavage to generate a 3 ′ terminus occurs a fixed distance from the site recognized by U7 snRNA, which suggests that the snRNA is involved in defining the cleavage site (for review see 241). However, the factor(s) actually responsible for cleavage have not yet been identified. Last updated on 1-20-2003


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Reviews 241. Birnstiel, M. L. (1985). Transcription termination and 3 ′ processing: the end is in site. Cell 41, 349-359.

References 743. Galli, G. et al. (1983). Biochemical complementation with RNA in the Xenopus oocyte: a small RNA is required for the generation of 3 ′ histone mRNA termini. Cell 34, 823-828. 3324. Wang, Z. F., Whitfield, M. L., Ingledue, T. C., Dominski, Z., 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. 3325. Mowry, K. L. and Steitz, J. A. (1987). Identification of the human U7 snRNP as one of several factors involved in the 3 ′ end maturation of histone premessenger RNA''''s. Science 238, 1682-1687. 3326. Dominski, Z., Erkmann, J. A., Greenland, J. A., and Marzluff, W. F. (2001). Mutations in the RNA binding domain of stem-loop binding protein define separable requirements for RNA binding and for histone pre-mRNA processing. Mol. Cell. Biol. 21, 2008-2017. 3327. Bond, U. M., Yario, T. A., and Steitz, J. A. (1991). Multiple processing-defective mutations in a mammalian histone pre-mRNA are suppressed by compensatory changes in U7 RNA both in vitro and in vitro. Genes Dev. 5, 1709-1722.



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5.24.21 Production of rRNA requires cleavage events Key Terms 45S RNA is a precursor that contains the sequences of both major ribosomal RNAs (28S and 18S rRNAs). Key Concepts

• The large and small rRNAs are released by cleavage from a common precursor RNA.

The major rRNAs are synthesized as part of a single primary transcript that is processed to generate the mature products. The precursor contains the sequences of the 18S, 5.8S, and 28S rRNAs. In higher eukaryotes, the precursor is named for its sedimentation rate as 45S RNA. In lower eukaryotes, it is smaller (35S in yeast). The mature rRNAs are released from the precursor by a combination of cleavage events and trimming reactions (for review see 980). Figure 24.37 shows the general pathway in yeast. There can be variations in the order of events, but basically similar reactions are involved in all eukaryotes. Most of the 5 ′ ends are generated directly by a cleavage event. Most of the 3 ′ ends are generated by cleavage followed by a 3 ′ –5 ′ trimming reaction.

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Figure 24.37 Mature eukaryotic rRNAs are generated by cleavage and trimming events from a primary transcript.

Many ribonucleases have been implicated in processing rRNA, including the exosome, an assembly of several exonucleases that also participates in mRNA degradation (see Molecular Biology 2.5.13 mRNA degradation involves multiple activities). Mutations in individual enzymes usually do not prevent processing, suggesting that their activities are redundant and that different combinations of cleavages can be used to generate the mature molecules. There are always multiple copies of the transcription unit for the rRNAs. The copies are organized as tandem repeats (see Molecular Biology 1.4.9 The repeated genes for rRNA maintain constant sequence). 5S RNA is transcribed from separate genes by RNA polymerase III. Usually the 5S genes are clustered, but are separate from the genes for the major rRNAs. (In the case of yeast, a 5S gene is associated with each major transcription unit, but is transcribed independently.) There is a difference in the organization of the precursor in bacteria. The sequence corresponding to 5.8S rRNA forms the 5 ′ end of the large (23S) rRNA, that is, there is no processing between these sequences. Figure 24.38 shows that the precursor also contains the 5S rRNA and one or two tRNAs. In E. coli, the 7 rrn operons are Production of rRNA requires cleavage events | SECTION 5.24.21 © 2004. Virtual Text / www.ergito.com

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dispersed around the genome; four rrn loci contain one tRNA gene between the 16S and 23S rRNA sequences, and the other rrn loci contain two tRNA genes in this region. Additional tRNA genes may or may not be present between the 5S sequence and the 3 ′ end. So the processing reactions required to release the products depend on the content of the particular rrn locus.

Figure 24.38 The rrn operons in E. coli contain genes for both rRNA and tRNA. The exact lengths of the transcripts depend on which promoters (P) and terminators (t) are used. Each RNA product must be released from the transcript by cuts on either side.

In both prokaryotic and eukaryotic rRNA processing, ribosomal proteins (and possibly also other proteins) bind to the precursor, so that the substrate for processing is not the free RNA but is a ribonucleoprotein complex.


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Reviews 980. Venema, J. and Tollervey, D. (1999). Ribosome synthesis in S. cerevisiae. Annu. Rev. Genet. 33, 261-311.



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5.24.22 Small RNAs are required for rRNA processing Key Terms A snoRNA is a small nuclear RNA that is localized in the nucleolus. Key Concepts

• The C/D group of snoRNAs is required for modifying the 2 ′ position of ribose with a methyl group.

• The H/ACA group of snoRNAs is required for converting uridine to pseudouridine. • In each case the snoRNA base pairs with a sequence of rRNA that contains the target base to generate a typical structure that is the substrate for modification.

Processing and modification of rRNA requires a class of small RNAs called snoRNAs (small nucleolar RNAs). There are 71 snoRNAs in the yeast (S. cerevisiae) genome. They are associated with the protein fibrillarin, which is an abundant component of the nucleolus (the region of the nucleus where the rRNA genes are transcribed). Some snoRNAs are required for cleavage of the precursor to rRNA; one example is U3 snoRNA, which is required for the first cleavage event in both yeast and Xenopus (740). We do not know what role the snoRNA plays in cleavage. It could be required to pair with the rRNA sequence to form a secondary structure that is recognized by an endonuclease. Two groups of snoRNAs are required for the modifications that are made to bases in the rRNA (see Great Experiments 4.6 Small nucleolar RNAs guide rRNA modification). The members of each group are identified by very short conserved sequences and common features of secondary structure (1216; 1217). The C/D group of snoRNAs is required for adding a methyl group to the 2 ′ position of ribose. There are >100 2 ′ -O-methyl groups at conserved locations in vertebrate rRNAs. This group takes its name from two short conserved sequences motifs called boxes C and D. Each snoRNA contains a sequence near the D box that is complementary to a region of the 18S or 28S rRNA that is methylated. Loss of a particular snoRNA prevents methylation in the rRNA region to which it is complementary. Figure 24.39 suggests that the snoRNA base pairs with the rRNA to create the duplex region that is recognized as a substrate for methylation. Methylation occurs within the region of complementarity, at a position that is fixed 5 bases on the 5 ′ side of the D box (741; 1220). Probably each methylation event is specified by a different snoRNA; ~40 snoRNAs have been characterized so far. The methylase(s) have not been characterized; one possibility is that the snoRNA itself provides part of the methylase activity. Small RNAs are required for rRNA processing | SECTION 5.24.22 © 2004. Virtual Text / www.ergito.com

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Figure 24.39 A snoRNA base pairs with a region of rRNA that is to be methylated.

Another group of snoRNAs is involved in the synthesis of pseudouridine. There are 43 ψ residues in yeast rRNAs and ~100 in vertebrate rRNAs. The synthesis of pseudouridine involves the reaction shown in Figure 24.40 in which the N1 bond from uridylic acid to ribose is broken, the base is rotated, and C5 is rejoined to the sugar.

Figure 24.40 Uridine is converted to pseudouridine by replacing the N1-sugar bond with a C5-sugar bond and rotating the base relative to the sugar.

Pseudouridine formation in rRNA requires the H/ACA group of ~20 snoRNAs. They are named for the presence of an ACA triplet 3 nucleotides from the 3 ′ end and a partially conserved sequence (the H box) that lies between two stem-loop hairpin structures. Each of these snoRNAs has a sequence complementary to rRNA within the stem of each hairpin. Figure 24.41shows the structure that would be produced by Small RNAs are required for rRNA processing | SECTION 5.24.22 © 2004. Virtual Text / www.ergito.com

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pairing with the rRNA. Within each pairing region, there are two unpaired bases, one of which is a uridine that is converted to pseudouridine (742; 1218).

Figure 24.41 H/ACA snoRNAs have two short conserved sequences and two hairpin structures, each of which has regions in the stem that are complementary to rRNA. Pseudouridine is formed by converting an unpaired uridine within the complementary region of the rRNA.

The H/ACA snoRNAs are associated with a nucleolar protein called Gar1p, which is required for pseudouridine formation, but its function is unknown (1219). The known pseudouridine synthases are proteins that function without an RNA cofactor. Synthases that could be involved in snoRNA-mediated pseudouridine synthesis have not been identified. The involvement of the U7 snRNA in 3 ′ end generation, and the role of snoRNAs in rRNA processing and modification, is consistent with the view we develop in Molecular Biology 5.26 Catalytic RNA that many – perhaps all – RNA processing events depend on RNA-RNA interactions. As with splicing reactions, the snRNA probably functions in the form of a ribonucleoprotein particle containing proteins as well as the RNA. It is common (although not the only mechanism of action) for the RNA of the particle to base pair with a short sequence in the substrate RNA. Last updated on 10-30-2000

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References 740. Kass, S. et al. (1990). The U3 small nucleolar ribonucleoprotein functions in the first step of preribosomal RNA processing. Cell 60, 897-908. 741. Kiss-Laszlo, Z. et al. (1996). Site-specific ribose methylation of preribosomal RNA: a novel function for small nucleolar RNAs. Cell 85, 1077-1068. 742. Ni, J., Tien, A. L., and Fournier, M. J. (1997). Small nucleolar RNAs direct site-specific synthesis of pseudouridine in rRNA. Cell 89, 565-573. 1216. Balakin, A. G., Smith, L., and Fournier, M. J. (1996). The RNA world of the nucleolus: two major families of small RNAs defined by different box elements with related functions. Cell 86, 823-834. 1217. Ganot, P., Caizergues-Ferrer, M., and Kiss, T. (1997). The family of box ACA small nucleolar RNAs is defined by an evolutionarily conserved secondary structure and ubiquitous sequence elements essential for RNA accumulation. Genes Dev. 11, 941-956. 1218. Ganot, P., Bortolin, M. L., and Kiss, T. (1997). Site-specific pseudouridine formation in preribosomal RNA is guided by small nucleolar RNAs. Cell 89, 799-809. 1219. Bousquet-Antonelli, C., Henry, Y., G'elugne, J. P., Caizergues-Ferrer, M., and Kiss, T. (1997). A small nucleolar RNP protein is required for pseudouridylation of eukaryotic ribosomal RNAs. EMBO J. 16, 4770-4776. 1220. Kiss-Laszlo, Z., Henry, Y., and Kiss, T. (1998). Sequence and structural elements of methylation guide snoRNAs essential for site-specific ribose methylation of pre-rRNA. EMBO J. 17, 797-807.


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5.24.23 Summary Splicing accomplishes the removal of introns and the joining of exons into the mature sequence of RNA. There are at least four types of reaction, as distinguished by their requirements in vitro and the intermediates that they generate. The systems include eukaryotic nuclear introns, group I and group II introns, and tRNA introns. Each reaction involves a change of organization within an individual RNA molecule, and is therefore a cis-acting event. pre-mRNA splicing follows preferred but not obligatory pathways. Only very short consensus sequences are necessary; the rest of the intron appears irrelevant. All 5 ′ splice sites are probably equivalent, as are all 3 ′ splice sites. The required sequences are given by the GU-AG rule, which describes the ends of the intron. The UACUAAC branch site of yeast, or a less well conserved consensus in mammalian introns, is also required. The reaction with the 5 ′ splice site involves formation of a lariat that joins the GU end of the intron via a 5 ′ –2 ′ linkage to the A at position 6 of the branch site. Then the 3 ′ –OH end of the exon attacks the 3 ′ splice site, so that the exons are ligated and the intron is released as a lariat. Both reactions are transesterifications in which bonds are conserved. Several stages of the reaction require hydrolysis of ATP, probably to drive conformational changes in the RNA and/or protein components. Lariat formation is responsible for choice of the 3 ′ splice site. Alternative splicing patterns are caused by protein factors that either stimulate use of a new site or that block use of the default site. pre-mRNA splicing requires formation of a spliceosome, a large particle that assembles the consensus sequences into a reactive conformation. The spliceosome most often forms by the process of intron definition, involving recognition of the 5 ′ splice site, branch site, and 3 ′ splice site. An alternative pathway involves exon definition, which involves initial recognition of the 5 ′ splice sites of both the substrate intron and the next intron. Its formation passes through a series of stages from the E (commitment) complex that contains U1 snRNP and splicing factors, through the A and B complexes as additional components are added. The spliceosome contains the U1, U2, U4/U6, and U5 snRNPs and some additional splicing factors. The U1, U2, and U5 snRNPs each contain a single snRNA and several proteins; the U4/U6 snRNP contains 2 snRNAs and several proteins. Some proteins are common to all snRNP particles. The snRNPs recognize consensus sequences. U1 snRNA base pairs with the 5 ′ splice site, U2 snRNA base pairs with the branch sequence, U5 snRNP acts at the 5 ′ splice site. When U4 releases U6, the U6 snRNA base pairs with U2, and this may create the catalytic center for splicing. An alternative set of snRNPs provides analogous functions for splicing the U12-dependent subclass of introns. The snRNA molecules may have catalytic-like roles in splicing and other processing reactions. In the nucleolus, two groups of snoRNAs are responsible for pairing with rRNAs at sites that are modified; group C/D snoRNAs indicate target sites for methylation, and group ACA snoRNAs identify sites where uridine is converted to pseudouridine. Summary | SECTION 5.24.23 © 2004. Virtual Text / www.ergito.com

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Splicing is usually intramolecular, but trans-(intermolecular) splicing occurs in trypanosomes and nematodes. It involves a reaction between a small SL RNA and the pre-mRNA. The SL RNA resembles U1 snRNA and may combine the role of providing the exon and the functions of U1. In worms there are two types of SL RNA, one used for splicing to the 5 ′ end of an mRNA, the other for splicing to an internal site. Group II introns share with nuclear introns the use of a lariat as intermediate, but are able to perform the reaction as a self-catalyzed property of the RNA. These introns follow the GT-AG rule, but form a characteristic secondary structure that holds the reacting splice sites in the appropriate apposition. Yeast tRNA splicing involves separate endonuclease and ligase reactions. The endonuclease recognizes the secondary (or tertiary) structure of the precursor and cleaves both ends of the intron. The two half-tRNAs released by loss of the intron can be ligated in the presence of ATP. The termination capacity of RNA polymerase II has not been characterized, and 3 ′ ends of its transcripts are generated by cleavage. The sequence AAUAAA, located 11-30 bases upstream of the cleavage site, provides the signal for both cleavage and polyadenylation. An endonuclease and the poly(A) polymerase are associated in a complex with other factors that confer specificity for the AAUAAA signal. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.24.23


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IMMUNE DIVERSITY

5.25.1 Introduction Key Terms An immune response is an organism's reaction, mediated by components of the immune system, to an antigen. An antigen is any foreign substance whose entry into an organism provokes an immune response by stimulating the synthesis of an antibody (an immunoglobulin protein that can bind to the antigen). A B cell is a lymphocyte that produces antibodies. B cells development occurs primarily in bone marrow. T cells are lymphocytes of the T (thymic) lineage; may be subdivided into several functional types. They carry TcR (T-cell receptor) and are involved in the cell-mediated immune response. The humoral response is an immune response that is mediated primarily by antibodies. It is defined as immunity that can be transferred from one organism to another by serum antibody. An immunoglobulin (Antibody) is a class of protein that is produced by B cells in response to antigen. An antibody is a protein (immunoglobulin) produced by B lymphocyte cells that recognizes a particular 'foreign antigen', and thus triggers the immune response. A helper T cell is a T lymphocyte that activates macrophages and stimulates B cell proliferation and antibody production. Helper T cells usually express cell surface CD4 but not CD8. Two mutants are said to complement each other when a diploid that is heterozygous for each mutation produces the wild type phenotype. The cell-mediated response is the immune response that is mediated primarily by T lymphocytes. It is defined based on immunity that cannot be transferred from one organism to another by serum antibody. A cytotoxic T cell is a T lymphocyte (usually CD8+) that can be stimulated to kill cells containing intracellular pathogens, such as viruses. The T cell receptor (TCR) is the antigen receptor on T lymphocytes. It is clonally expressed and binds to a complex of MHC class I or class II protein and antigen-derived peptide. The major histocompatibility complex (MHC) is a chromosomal region containing genes that are involved in the immune response. The genes encode proteins for antigen presentation, cytokines, and complement proteins. The MHC is highly polymorphic. Tolerance is the lack of an immune response to an antigen (either self antigen or foreign antigen) due to clonal deletion. An autoimmune disease is a pathological condition in which the immune response is directed to self antigen.


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It is an axiom of genetics that the genetic constitution created in the zygote by the combination of sperm and egg is inherited by all somatic cells of the organism. We look to differential control of gene expression, rather than to changes in DNA content, to explain the different phenotypes of particular somatic cells. Yet there are exceptional situations in which the reorganization of certain DNA sequences is used to regulate gene expression or to create new genes. The immune system provides a striking and extensive case in which the content of the genome changes, when recombination creates active genes in lymphocytes. Other cases are represented by the substitution of one sequence for another to change the mating type of yeast or to generate new surface antigens by trypanosomes (see Molecular Biology 4.18 Rearrangement of DNA). The immune response of vertebrates provides a protective system that distinguishes foreign proteins from the proteins of the organism itself. Foreign material (or part of the foreign material) is recognized as comprising an antigen. Usually the antigen is a protein (or protein-attached moiety) that has entered the bloodstream of the animal – for example, the coat protein of an infecting virus. Exposure to an antigen initiates production of an immune response that specifically recognizes the antigen and destroys it. Immune reactions are the responsibility of white blood cells – the B and T lymphocytes, and macrophages. The lymphocytes are named after the tissues that produce them. In mammals, B cells mature in the bone marrow, while T cells mature in the thymus. Each class of lymphocyte uses the rearrangement of DNA as a mechanism for producing the proteins that enable it to participate in the immune response. The immune system has many ways to destroy an antigenic invader, but it is useful to consider them in two general classes. Which type of response the immune system mounts when it encounters a foreign structure depends partly on the nature of the antigen. The response is defined according to whether it is executed principally by B cells or T cells. The humoral response depends on B cells. It is mediated by the secretion of antibodies, which are immunoglobulin proteins. Production of an antibody specific for a foreign molecule is the primary event responsible for recognition of an antigen. Recognition requires the antibody to bind to a small region or structure on the antigen. The function of antibodies is represented in Figure 25.1. Foreign material circulating in the bloodstream, for example, a toxin or pathogenic bacterium, has a surface that presents antigens. The antigen(s) are recognized by the antibodies, which form an Introduction | SECTION 5.25.1 © 2004. Virtual Text / www.ergito.com

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antigen-antibody complex. This complex then attracts the attention of other components of the immune system.

Figure 25.1 Humoral immunity is conferred by the binding of free antibodies to antigens to form antigen-antibody complexes that are removed from the bloodstream by macrophages or that are attacked directly by the complement proteins.

The humoral response depends on these other components in two ways. First, B cells need signals provided by T cells to enable them to secrete antibodies. These T cells are called helper T cells, because they assist the B cells. Second, antigen-antibody formation is a trigger for the antigen to be destroyed. The major pathway is provided by the action of complement, a component whose name reflects its ability to "complement" the action of the antibody itself. Complement consists of a set of ~20 proteins that function through a cascade of proteolytic actions. If the target antigen is part of a cell, for example, an infecting bacterium, the action of complement culminates in lysing the target cell. The action of complement also provides a means of attracting macrophages, which scavenge the target cells or their products. Alternatively, the antigen-antibody complex may be taken up directly by macrophages (scavenger cells) and destroyed. The cell-mediated response is executed by a class of T lymphocytes called cytotoxic T cells (also called killer T cells). The basic function of the T cell in recognizing a target antigen is indicated in Figure 25.2. A cell-mediated response typically is elicited by an intracellular parasite, such as a virus that infects the body's own cells. As a result of the viral infection, fragments of foreign (viral) antigens are displayed on the surface of the cell. These fragments are recognized by the T cell receptor (TCR), which is the T cells' equivalent of the antibody produced by a B Introduction | SECTION 5.25.1 © 2004. Virtual Text / www.ergito.com

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cell.

Figure 25.2 In cell-mediated immunity, killer T cells use the T-cell receptor to recognize a fragment of the foreign antigen which is presented on the surface of the target cell by the MHC protein.

A crucial feature of this recognition reaction is that the antigen must be presented by a cellular protein that is a member of the MHC (major histocompatibility complex). The MHC protein has a groove on its surface that binds a peptide fragment derived from the foreign antigen. The combination of peptide fragment and MHC protein is recognized by the T cell receptor. Every individual has a characteristic set of MHC proteins. They are important in graft reactions; a graft of tissue from one individual to another is rejected because of the difference in MHC proteins between the donor and recipient, an issue of major medical importance. The demand that the T lymphocytes recognize both foreign antigen and MHC protein ensures that the cell-mediated response acts only on host cells that have been infected with a foreign antigen. (We discuss the division of MHC proteins into the general types of class I and class II later in Molecular Biology 5.25.20 The major histocompatibility locus codes for many genes of the immune system.) The purpose of each type of immune response is to attack a foreign target. Target recognition is the prerogative of B-cell immunoglobulins and T cell receptors. A crucial aspect of their function lies in the ability to distinguish "self" from "nonself." Proteins and cells of the body itself must never be attacked. Foreign targets must be destroyed entirely. The property of failing to attack "self" is called tolerance. Loss of Introduction | SECTION 5.25.1 © 2004. Virtual Text / www.ergito.com

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this ability results in an autoimmune disease, in which the immune system attacks its own body, often with disastrous consequences. What prevents the lymphocyte pool from responding to "self" proteins? Tolerance probably arises early in lymphocyte cell development when B and T cells that recognize "self" antigens are destroyed. This is called clonal deletion. In addition to this negative selection, there is also positive selection for T cells carrying certain sets of T cell receptors. A corollary of tolerance is that it can be difficult to obtain antibodies against proteins that are closely related to those of the organism itself. As a practical matter, therefore, it may be difficult to use (for example) mice or rabbits to obtain antibodies against human proteins that have been highly conserved in mammalian evolution. The tolerance of the mouse or rabbit for its own protein may extend to the human protein in such cases. Each of the three groups of proteins required for the immune response – immunoglobulins, T cell receptors, MHC proteins – is diverse. Examining a large number of individuals, we find many variants of each protein. Each protein is coded by a large family of genes; and in the case of antibodies and the T cell receptors, the diversity of the population is increased by DNA rearrangements that occur in the relevant lymphocytes. Immunoglobulins and T cell receptors are direct counterparts, each produced by its own type of lymphocyte. The proteins are related in structure, and their genes are related in organization. The sources of variability are similar. The MHC proteins also share some common features with the antibodies, as do other lymphocyte-specific proteins. In dealing with the genetic organization of the immune system, we are therefore concerned with a series of related gene families, indeed a superfamily that may have evolved from some common ancestor representing a primitive immune response. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.25.1


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IMMUNE DIVERSITY

5.25.2 Clonal selection amplifies lymphocytes that respond to individual antigens Key Terms The clonal selection theory proposed that each lymphocyte expresses a single antigen receptor specificity and that only those lymphocytes that bind to a given antigen are stimulated to proliferate and to function in eliminating that antigen. Thus, the antigen "selects" the lymphocytes to be activated. Clonal selection is now an established principle in immunology. The primary immune response is an organism's immune response upon first exposure to a given antigen. It is characterized by a relatively shorter duration and lower affinity antibodies than in the secondary immune response. A memory cell is a lymphocyte that has been stimulated during the primary immune response to antigen and that is rapidly activated upon subsequent exposure to that antigen. Memory cells respond more rapidly to antigen than naive cells. The secondary immune response is an organism's immune response upon a second exposure to a given antigen. This second exposure is also referred to as a "booster". The secondary immune response is characterized by a more rapid induction, greater magnitude, and higher affinity antibodies than the primary immune response. A hapten is a small molecule that acts as an antigen when conjugated to a protein. An antigenic determinant is the portion of an antigen that is recognized by the antigen receptor on lymphocytes. It is also called an epitope. An epitope is the portion of an antigen that is recognized by the antigen receptor on lymphocytes. It is also called an antigenic determinant. Key Concepts

• Each B lymphocyte expresses a single immunoglobulin and each T lymphocyte expresses a single T cell receptor.

• There is a very large variety of immunoglobulins and T cell receptors. • Antigen binding to an immunoglobulin or T cell receptor triggers clonal multiplication of the cell.

The name of the immune response describes one of its central features. After an organism has been exposed to an antigen, it becomes immune to the effects of a new infection. Before exposure to a particular antigen, the organism lacks adequate capacity to deal with any toxic effects. This ability is acquired during the immune response. After the infection has been defeated, the organism retains the ability to respond rapidly in the event of a re-infection. These features are accommodated by the clonal selection theory illustrated in Figure Clonal selection amplifies lymphocytes that respond to individual antigens | SECTION 5.25.2 © 2004. Virtual Text / www.ergito.com

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25.3. The pool of lymphocytes contains B cells and T cells carrying a large variety of immunoglobulins or T cell receptors. But any individual B lymphocyte produces one immunoglobulin, which is capable of recognizing only a single antigen; similarly any individual T lymphocyte produces only one particular T cell receptor.

Figure 25.3 The pool of immature lymphocytes contains B cells and T cells making antibodies and receptors with a variety of specificities. Reaction with an antigen leads to clonal expansion of the lymphocyte with the antibody (B cell) or receptor (T cell) that can recognize the antigen.

In the pool of immature lymphocytes, the unstimulated B cells and T cells are morphologically indistinguishable. But on exposure to antigen, a B cell whose antibody is able to bind the antigen, or a T cell whose receptor can recognize it, is stimulated to divide, probably by some feedback from the surface of the cell, where the antibody/receptor-antigen reaction occurs. The stimulated cells then develop into mature B or T lymphocytes, which includes morphological changes involving (for example) an increase in cell size (especially pronounced for B cells). Clonal selection amplifies lymphocytes that respond to individual antigens | SECTION 5.25.2 © 2004. Virtual Text / www.ergito.com

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The initial expansion of a specific B- or T cell population upon first exposure to an antigen is called the primary immune response. Large numbers of B or T lymphocytes with specificity for the target antigen are produced. Each population represents a clone of the original responding cell. Antibody is secreted from the B cells in large quantities, and it may even come to dominate the antibody population. After a successful primary immune response has been mounted, the organism retains B cells and T cells carrying the corresponding antibody or receptor. These memory cells represent an intermediate state between the immature cell and the mature cell. They have not acquired all of the features of the mature cell, but they are long-lived, and can rapidly be converted to mature cells. Their presence allows a secondary immune response to be mounted rapidly if the animal is exposed to the same antigen again. The pool of immature lymphocytes in a mammal contains ~1012 cells. This pool contains some lymphocytes that have unique specificities (because a corresponding antigen has never been encountered), while others are represented by up to 106 cells (because clonal selection has expanded the pool to respond to an antigen). What features are recognized in an antigen? Antigens are usually macromolecular. Although small molecules may have antigenic determinants and can be recognized by antibodies, usually they are not effective in provoking an immune response (because of their small size). But they do provoke a response when conjugated with a larger carrier molecule (usually a protein). A small molecule that is used to provoke a response by such means is called a hapten. Only a small part of the surface of a macromolecular antigen is actually recognized by any one antibody. The binding site consists of only 5-6 amino acids. Of course, any particular protein may have more than one such binding site, in which case it provokes antibodies with specificities for different regions. The region provoking a response is called an antigenic determinant or epitope. When an antigen contains several epitopes, some may be more effective than others in provoking the immune response; in fact, they may be so effective that they entirely dominate the response. How do lymphocytes find target antigens and where does their maturation take place? Lymphocytes are peripatetic cells. They develop from immature stem cells that are located in the adult bone marrow. They migrate to the peripheral lymphoid tissues (spleen, lymph nodes) either directly via the bloodstream (if they are B cells) or via the thymus (where they become T cells). The lymphocytes recirculate between blood and lymph; the process of dispersion ensures that an antigen will be exposed to lymphocytes of all possible specificities. When a lymphocyte encounters an antigen that binds its antibody or receptor, clonal expansion begins the immune response. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.25.2

Clonal selection amplifies lymphocytes that respond to individual antigens | SECTION 5.25.2 © 2004. Virtual Text / www.ergito.com

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5.25.3 Immunoglobulin genes are assembled from their parts in lymphocytes Key Terms The immunoglobulin light chain is one of two types of subunits in an antibody tetramer. Each antibody contains two light chains. The N-terminus of the light chain forms part of the antigen recognition site. The immunoglobulin heavy chain is one of two types of subunits in an antibody tetramer. Each antibody contains two heavy chains. The N-terminus of the heavy chain forms part of the antigen recognition site, whereas the C-terminus determines the subclass (isotype). The variable region (V region) of an immunoglobulin chain is coded by the V gene and varies extensively when different chains are compared, as the result of multiple (different) genomic copies and changes introduced during construction of an active immunoglobulin. Constant regions (C region) of immunoglobulins are coded by C genes and are the parts of the chain that vary least. Those of heavy chains identify the type of immunoglobulin. A V gene is sequence coding for the major part of the variable (N-terminal) region of an immunoglobulin chain. C genes code for the constant regions of immunoglobulin protein chains. Somatic recombination describes the process of joining a C gene to a C gene in a lymphocyte to generate an immunoglobulin or T cell receptor. Key Concepts

• An immunoglobulin is a tetramer of two light chains and two heavy chains. • Light chains fall into the lambda and kappa families; heavy chains form a single family.

• Each chain has an N-terminal variable region (V) and a C-terminal constant region (C).

• The V domain recognizes antigen and the C domain provides the effector response. • V domains and C domains are separately coded by V gene segments and C gene segments.

• A gene coding for an intact immunoglobulin chain is generated by somatic recombination to join a V gene segment with a C gene segment.

A remarkable feature of the immune response is an animal's ability to produce an appropriate antibody whenever it is exposed to a new antigen. How can the organism be prepared to produce antibody proteins each designed specifically to recognize an Immunoglobulin genes are assembled from their parts in lymphocytes | SECTION 5.25.3 © 2004. Virtual Text / www.ergito.com

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antigen whose structure cannot be anticipated? For practical purposes, we usually reckon that a mammal has the ability to produce 106-108 different antibodies. Each antibody is an immunoglobulin tetramer consisting of two identical light chains (L) and two identical heavy chains (H). If any light chain can associate with any heavy chain, to produce 106–108 potential antibodies requires 103–104 different light chains and 103–104 different heavy chains. There are 2 types of light chain and ~10 types of heavy chain. Different classes of immunoglobulins have different effector functions. The class is determined by the heavy chain constant region, which exercises the effector function (see Figure 25.17). The structure of the immunoglobulin tetramer is illustrated in Figure 25.4. Light chains and heavy chains share the same general type of organization in which each protein chain consists of two principal regions: the N-terminal variable region (V region); and the C-terminal constant region (C region). They were defined originally by comparing the amino acid sequences of different immunoglobulin chains. As the names suggest, the variable regions show considerable changes in sequence from one protein to the next, while the constant regions show substantial homology.

Figure 25.4 Heavy and light chains combine to generate an immunoglobulin with several discrete domains.

Corresponding regions of the light and heavy chains associate to generate distinct domains in the immunoglobulin protein. The variable (V) domain is generated by association between the variable regions of the light chain and heavy chain. The V domain is responsible for recognizing the antigen. An immunoglobulin has a Y-shaped structure in which the arms of the Y are identical, and each arm has a copy of the V domain. Production of V domains of different specificities creates the ability to respond to diverse antigens. The total Immunoglobulin genes are assembled from their parts in lymphocytes | SECTION 5.25.3 © 2004. Virtual Text / www.ergito.com

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number of variable regions for either light- or heavy-chain proteins is measured in hundreds. So the protein displays the maximum versatility in the region responsible for binding the antigen. The number of constant regions is vastly smaller than the number of variable regions – typically there are only 1-10 C regions for any particular type of chain. The constant regions in the subunits of the immunoglobulin tetramer associate to generate several individual C domains. The first domain results from association of the single constant region of the light chain (CL) with the CH1 part of the heavy-chain constant region. The two copies of this domain complete the arms of the Y-shaped molecule. Association between the C regions of the heavy chains generates the remaining C domains, which vary in number depending on the type of heavy chain. Comparing the characteristics of the variable and constant regions, we see the central dilemma in immunoglobulin gene structure. How does the genome code for a set of proteins in which any individual polypeptide chain must have one of 10 D segments lies on the chromosome between the VH segments and the 4 JH segments. V-D-J joining takes place in two stages, as illustrated in Figure 25.7. First one of the D segments recombines with a JH segment; then a VH segment recombines with the DJH combined segment. The reconstruction leads to expression of the adjacent CH segment (which consists of several exons). (We discuss the use of different CH gene segments in Molecular Biology 5.25.12 Class switching is caused by DNA recombination; now we will just consider the reaction in terms of the connection to one of several J segments that precede a CH gene segment.)

Heavy chains are assembled by two recombinations | SECTION 5.25.5 © 2004. Virtual Text / www.ergito.com

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Figure 25.7 Heavy genes are assembled by sequential joining reactions. First a D segment is joined to a J segment; then a V gene segment is joined to the D segment.

The D segments are organized in a tandem array. The mouse heavy-chain locus contains 12 D segments of variable length; the human locus has ~30 D segments (not all necessarily active). Some unknown mechanism must ensure that the same D segment is involved in the D-J joining and V-D joining reactions. (When we discuss joining of V and C gene segments for heavy chains, we assume the process has been completed by V-D and D-J joining reactions.) The V gene segments of all three immunoglobulin families are similar in organization. The first exon codes for the signal sequence (involved in membrane attachment), and the second exon codes for the major part of the variable region itself (1000 chains by combining 300 V genes with 4-5 C genes.

• An H locus can produce >4000 chains by combining 300 V genes, 20 D segments, and 4 J segments.

Now we must examine the different types of V and C gene segments to see how much diversity can be accommodated by the variety of the coding regions carried in the germline. In each light Ig gene family, many V gene segments are linked to a much smaller number of C gene segments. Figure 25.8 shows that the λ locus has ~6 C gene segments, each preceded by its own J segment. The λ locus in mouse is much less diverse than the human locus. The main difference is that in mouse there are only two V λ gene segments; each is linked to two J-C regions. Of the 4 C λ gene segments, one is inactive. At some time in the past, the mouse suffered a catastrophic deletion of most of its germline V λ gene segments.

Figure 25.8 The lambda family consists of V gene segments linked to a small number of J-C gene segments.

Figure 25.9 shows that the κ locus has only one C gene segment, although it is preceded by 5 J segments (one of them inactive). The V κ gene segments occupy a large cluster on the chromosome, upstream of the constant region. The human cluster has two regions. Just preceding the C κ gene segment, a region of 600 kb contains the 5 J κ segments and 40 V κ gene segments. A gap of 800 kb separates this region from another group of 36 V κ gene segments.

Recombination generates extensive diversity | SECTION 5.25.6 © 2004. Virtual Text / www.ergito.com

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Figure 25.9 The human and mouse kappa families consist of V gene segments linked to 5 J segments connected to a single C gene segment.

The V κ gene segments can be subdivided into families, defined by the criterion that members of a family have >80% amino acid identity. The mouse family is unusually large, ~1000 genes, and there are ~18 V κ families, varying in size from 2-100 members. Like other families of related genes, therefore, related V gene segments form subclusters, generated by duplication and divergence of individual ancestral members. However, many of the V segments are inactive pseudogenes, and 95% of T lymphocytes, and TCR γ δ is found on 2 Mb. MHC proteins of classes I and II are coded by two separate regions. The class III region is defined as the segment between them. The extended regions describe segments that are syntenic on either end of the cluster. The major difference between mouse and Man is the presence of H2 class I genes in the extended region on the left. The murine locus is located on chromosome 17, and the human locus on chromosome 6.

There are several hundred genes in the MHC regions of mammals, but it is possible for MHC functions to be provided by far fewer genes, as in the case of the chicken, where the MHC region is 92 Kb and has only 9 genes (910). As in comparisons of other gene families, we find differences in the exact numbers of genes devoted to each function. Because the MHC locus shows extensive variation between individuals, the number of genes may be different in different individuals. As a general rule, however, a mouse genome has fewer active H2 genes than a human genome. The class II genes are unique to mammals (except for one subgroup), and as a rule, birds and fish have different genes in their place. There are ~8 functional class I genes in Man and ~30 in the mouse. The class I region also includes many other genes. The class III regions are closely similar in Man and mouse. All MHC proteins are dimers located in the plasma membrane, with a major part of the protein protruding on the extracellular side. The structures of class I and class II MHC proteins are related, although they have different components, as summarized in Figure 25.34.

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Figure 25.34 Class I and class II histocompatibility antigens have a related structure. Class I antigens consist of a single ( α ) polypeptide, with three external domains ( α 1, α 2, α 3), that interacts with β 2 microglobulin ( β 2 m). Class II antigens consist of two ( α and β ) polypeptides, each with two domains ( α 1 & α 2, β 1 & β 2) with a similar overall structure.

Class II antigens consist of two chains, α and β, whose combination generates an overall structure in which there are two extracellular domains. All class I MHC proteins consist of a dimer between the class I chain itself and the β2-microglobulin protein. The class I chain is a 45 kD transmembrane component that has three external domains (each ~90 amino acids long, one of which interacts with β2 microglobulin), a transmembrane region of ~40 residues, and a short cytoplasmic domain of ~30 residues that resides within the cell. The β2 microglobulin is a secreted protein of 12 kD. It is needed for the class I chain to be transported to the cell surface. Mice that lack the β2 microglobulin gene have no MHC class I antigen on the cell surface. The organization of class I genes summarized in Figure 25.35 coincides with the protein structure. The first exon codes for a signal sequence (cleaved from the protein during membrane passage). The next three exons code for each of the external domains. The fifth exon codes for the transmembrane domain. And the last three rather small exons together code for the cytoplasmic domain. The only difference in the genes for human transplantation antigens is that their cytoplasmic The major histocompatibility locus codes for many genes of the immune system | SECTION 5.25.20 © 2004. Virtual Text / www.ergito.com

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domain is coded by only two exons.

Figure 25.35 Each class of MHC genes has a characteristic organization, in which exons represent individual protein domains

The exon coding for the third external domain of the class I genes is highly conserved relative to the other exons. The conserved domain probably represents the region that interacts with β2 microglobulin, which explains the need for constancy of structure. This domain also exhibits homologies with the constant region domains of immunoglobulins. What is responsible for generating the high degree of polymorphism in these genes? Most of the sequence variation between alleles occurs in the first and second external domains, sometimes taking the form of a cluster of base substitutions in a small region. One mechanism involved in their generation is gene conversion between class I genes. Pseudogenes are present as well as functional genes. The gene for β2 microglobulin is located on a separate chromosome. It has four exons, the first coding for a signal sequence, the second for the bulk of the protein (from amino acids 3 to 95), the third for the last four amino acids and some of the nontranslated trailer, and the last for the rest of the trailer. The length of β2 microglobulin is similar to that of an immunoglobulin V gene; there are certain similarities in amino acid constitution; and there are some (limited) homologies of nucleotide sequence between β2 microglobulin and Ig constant domains or type I gene third external domains. All the groups of genes that we have discussed in this chapter may have descended from a common ancestor that coded for a primitive domain. Last updated on 4-15-2003


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Reviews 262. Steinmetz, M. and Hood, L. (1983). Genes of the MHC complex in mouse and man. Science 222, 727-732. 265. Flavell, R. A., Allen, H., Burkly, L. C., Sherman, D. H., Waneck, G. L., and Widera, G. (1986). Molecular biology of the H-2 histocompatibility complex. Science 233, 437-443. 3772. Kumnovics, A., Takada, T., and Lindahl, K. F. (2003). Genomic organization of the Mammalian MHC. Annu. Rev. Immunol. 21, 629-657.

References 910. Kaufman, J. et al. (1999). The chicken B locus is a minimal essential major histocompatibility complex. Nature 401, 923-925.



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5.25.21 Innate immunity utilizes conserved signaling pathways Key Terms Adaptive immunity is the response mediated by lymphocytes that are activated by their specific interaction with antigen. The adaptive immune response develops over several days as lymphocytes with antigen-specific receptors are stimulated to proliferate and become effector cells. It is responsible for immunological memory. Acquired immunity is another term for adaptive immunity. Innate immunity is the rapid response mediated by cells with non-varying (germline-encoded) receptors that recognize pathogen. The cells of the innate immune response act to eliminate the pathogen and initiate the adaptive immune response. A pathogen-associated molecular pattern is (PAMP) a molecular structure on the surface of a pathogen. A given PAMP may be conserved across a large number of pathogens. During an immune response, PAMPs may be recognized by receptors on cells that mediate innate immunity. A lipopolysaccharide is (LPS) a molecule containing both lipid and sugar components. It is present in the outer membrane of Gram-negative bacteria. It is also an endotoxin responsible for inducing septic shock during an infection. An endotoxin is a toxin that is present on the surface of Gram-negative bacteria (as opposed to exotoxins, which are secreted). LPS is an example of an endotoxin. A Toll-like receptor is (TLR) a plasma membrane receptor that is expressed on phagocytes and other cells and is involved in signaling during the innate immune response. TLRs are related to IL-1 receptors. The leucine-rich region is (LLR) a motif found in the extracellular domains of some surface receptor proteins in animal and plant cells. Key Concepts

• Innate immunity is triggered by receptors that recognize motifs (PAMPs) that are highly conserved in bacteria or other infective agents.

• Toll-like receptors are commonly used to activate the response pathway. • The pathways are highly conserved from invertebrates to vertebrates, and an analogous pathway is found in plants.

The immune response described in this chapter comprises a set of reactions that respond to a pathogen by selecting lymphocytes (B cells or T cells) whose receptors (antibodies or TCR) have a high affinity for the pathogen. The basis for this selective process is the generation of a very large number of receptors so as to create a high possibility of recognizing a foreign molecule. Receptors that recognize the body's Innate immunity utilizes conserved signaling pathways | SECTION 5.25.21 © 2004. Virtual Text / www.ergito.com

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own proteins are screened out early in the process. Activation of the receptors on B cells triggers the pathways of the humoral response; activation of the receptors on T cells triggers the pathways of the cell-mediated response. The overall response to an antigen via selection of receptors on the lymphocytes is called adaptive immunity (or acquired immunity). The response typically is mounted over several days, following the initial activation of B cells or T cells that recognize the foreign pathogen. The organism retains a memory of the response, which enables it to respond more rapidly if it is exposed again to the same pathogen. The principles of adaptive immunity are similar through the vertebrate kingdoms, although details vary. Another sort of immune response occurs more quickly and is found in a greater range of animals (including those that do not have adaptive immunity). Innate immunity depends on the recognition of certain, pre-defined patterns in foreign pathogens. These patterns are motifs that are conserved in the pathogens because they have an essential role in their function, but they are not found in higher eukaryotes. The motif is typically recognized by a receptor dedicated to the purpose of triggering the innate response upon an infection. Receptors that trigger the innate response are found on cells such as neutrophils and macrophages, and cause the pathogen to be phagocytosed and killed. The response is rapid, because the set of receptors is already present on the cells and does not have to be amplified by selection. It is widely conserved and is found in organisms ranging from flies to Man. When the innate response is able to deal effectively with an infection, the adaptive response will not be triggered. There is some overlap between the responses in that they activate some of the same pathways, so cells activated by the innate response may subsequently participate in the adaptive response. The motifs that trigger innate immunity are sometimes called pathogen-associated molecular patterns (PAMPs). Figure 25.36 shows that they are widely distributed across broad ranges of organisms. Formyl-methionine is used to initiate most bacterial proteins, but is not found in eukaryotes. The peptidoglycan of the cell wall is unique to bacteria. Lipopolysaccharide (LPS) is a component of the outer membrane of most gram-negative bacteria; also known as endotoxin, it is responsible for septic shock syndrome.

Figure 25.36 Pathogen-associated molecular patterns (PAMPs) are compounds that are common to large ranges of bacteria or yeasts and are exposed when an infection occurs. Innate immunity utilizes conserved signaling pathways | SECTION 5.25.21 © 2004. Virtual Text / www.ergito.com

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A key insight into the nature of innate immunity was the discovery of the involvement of Toll-like receptors (TLR) (for review see 2382). The receptor Toll, which is related to mammalian IL1 receptor, triggers the pathway in Drosophila that controls dorsal-ventral development (see Molecular Biology 6.31.9 Dorsal-ventral development uses localized receptor-ligand interactions). This leads to activation of the transcription factor dorsal, a member of the Rel family, which is related to the mammalian factor NF- κB. The pathway of innate immunity is parallel to the Toll pathway, with similar components. In fact, one of the first indications of the nature of innate immunity in flies was the discovery of the transcription factor Dif (dorsal-related immunity factor), which is activated by one of the pathways (2378). Flies have no system of adaptive immunity, but are resistant to microbial infections (for review see 2385). This is because their innate immune systems trigger synthesis of potent antimicrobial peptides. Seven distinct peptides have been identified in Drosophila, where they are synthesized in the fat body (the equivalent organ to the liver). Two of the peptides are antifungal, five act largely on bacteria. The general mode of action is to kill the target organism by permeabilizing its membrane. All of these peptides are coded by genes whose promoters respond to transcription factors of the Rel family. Figure 25.37 summarizes the components of the innate pathways in Drosophila.

Figure 25.37 Innate immunity is triggered by PAMPs. In flies, they cause the production of peptides that activate Toll-like receptors. The receptors lead to a pathway that activates a transcription factor of the Rel family. Target genes for this factor include bactericidal and anti-fungal peptides. The peptides act by permeabilizing the membrane of the pathogenic organism.

Two innate response pathways function in Drosophila, one responding principally to Innate immunity utilizes conserved signaling pathways | SECTION 5.25.21 © 2004. Virtual Text / www.ergito.com

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fungi, the other principally to gram-negative bacteria. Gram-positive bacteria may be able to trigger both pathways. Figure 25.38 outlines the steps in each pathway. Fungi and gram-positive bacteria activate a proteolytic cascade that generates peptides that activate a Toll-like receptor. This is the NF- κB-like pathway. The dToll receptor activates the transcription factor Dif (a relative of NF- κB), leading ultimately to activation of the anti-fungal peptide drosomycin (2379; 2381). Gram-negative bacteria trigger a pathway via a different receptor that activates the transcription factor Relish, leading to production of the bactericidal peptide attacin (2380). This pathway is called the Imd pathway after one of its components, a protein that has a "death domain" related to those found in the pathways for apoptosis.

Figure 25.38 One of the Drosophila innate immunity pathways is closely related to the mammalian pathway for activating NF-kB; the other has components related to those of apoptosis pathways.

The key agents in responding to the bacteria are proteins called PGRPs because of their high affinities for bacterial peptidoglycans. There are two types of these proteins. PGRP-SAs are short extracellular proteins. They probably function by activating the proteases that trigger the Toll pathway. PGRP-LCs are transmembrane proteins with an extracellular PGRP domain. Their exact role has to be determined. The innate immune response is highly conserved (for review see 2459, 4381). Mice that are resistant to septic shock when they are treated with LPS have mutations in the Toll-like receptor TLR4 (2383). A human homologue of the Toll receptor can activate some immune-response genes, suggesting that the pathway of innate immunity may also function in Man (2384). The pathway downstream of the Innate immunity utilizes conserved signaling pathways | SECTION 5.25.21 © 2004. Virtual Text / www.ergito.com

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Toll-like receptors is generally similar in all cases, typically leading to activation of the transcription factor NF- κB. We do not yet know whether the upstream pathway is conserved, in particular whether the PAMPs function by generating ligands that in turn activate the Toll-like receptors or whether they might interact directly with them. The pathway upstream of the Toll-like receptors is different in mammals and flies, because the pathogens directly activate mammalian Toll-like receptors. In the case of LPS, the pathogen binds to the surface protein CD14; this enables CD14 to activate TLR4, triggering the innate response pathway (2383). There are ~20 receptors in the TLR (Toll-like receptor) class in the human genome, which gives some indication of how many pathogens can trigger the innate response. Plants have extensive defense mechanisms, among which are pathways analogous to the innate response in animals (for review see 2387). The same principle applies that PAMPs are the motifs that identify the infecting agent as a pathogen. The proteins that respond to the pathogens are coded by a class of genes called the disease resistance genes. Many of these genes code for receptors that share a property with the TLR class of animal receptors: the extracellular domain has a motif called the leucine-rich region (LLR). The response mechanism is different from animal cells, and is directed to activating a MAPK cascade. Many different pathogens activate the same cascade, which suggests that a variety of pathogen-receptor interactions converge at or before the activation of the first MAPK (2386). Last updated on November 10, 2003

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Reviews 2382. Aderem, A. and Ulevitch, R. J. (2000). Toll-like receptors in the induction of the innate immune response. Nature 406, 782-787. 2385. Hoffmann, J. A., Kafatos, F. C., Janeway, C. A., and Ezekowitz, R. A. (1999). Phylogenetic perspectives in innate immunity. Science 284, 1313-1318. 2387. Dangl, J. L. and Jones, J. D. (2001). Plant pathogens and integrated defence responses to infection. Nature 411, 826-833. 2459. Janeway, C. A. and Medzhitov, R. (2002). Innate immune recognition. Annu. Rev. Immunol. 20, 197-216.

References 2378. Ip, Y. T., Reach, M., Engstrom, Y., Kadalayil, L., Cai, H., GonzÃ¡lez-Crespo, S., Tatei, K., and Levine, M. (1993). Dif, a dorsal-related gene that mediates an immune response in Drosophila. Cell 75, 753-763. 2379. Lemaitre, B., Nicolas, E., Michaut, L., Reichhart, J. M., and Hoffmann, J. A. (1996). The dorsoventral regulatory gene cassette spÃtzle/Toll/cactus controls the potent antifungal response in Drosophila adults. Cell 86, 973-983. 2380. Williams, M. J., Rodriguez, A., Kimbrell, D. A., and Eldon, E. D. (1997). The 18-wheeler mutation reveals complex antibacterial gene regulation in Drosophila host defense. EMBO J. 16, 6120-6130. 2381. Rutschmann, S., Jung, A. C., Hetru, C., Reichhart, J. M., Hoffmann, J. A., and Ferrandon, D. (2000). The Rel protein DIF mediates the antifungal but not the antibacterial host defense in Drosophila. Immunity 12, 569-580. 2383. Poltorak, A., He, X., Smirnova, I., Liu, M. Y., Huffel, C. V., Du, X., Birdwell, D., Alejos, E., Silva, M., Galanos, C., Freudenberg, M., Ricciardi-Castagnoli, P., Layton, B., and Beutler, B. (1998). Defective LPS signaling in C3H/HeJ and C57BL/10ScCr mice: mutations in Tlr4 gene. Science 282, 2085-2088. 2384. Medzhitov, R., Preston-Hurlburt, P., and Janeway, C. A. (1997). A human homologue of the Drosophila Toll protein signals activation of adaptive immunity. Nature 388, 394-397. 2386. Asai, T., Tena, G., Plotnikova, J., Willmann, M. R., Chiu, W. L., Gomez-Gomez, L., Boller, T., Ausubel, F. M., and Sheen, J. (2002). MAP kinase signalling cascade in Arabidopsis innate immunity. Nature 415, 977-983. 4381. Hoffmann, J. A. (2003). The immune response of Drosophila. Nature 426, 33-38.


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5.25.22 Summary Immunoglobulins and T cell receptors are proteins that play analogous functions in the roles of B cells and T cells in the immune system. An Ig or TCR protein is generated by rearrangement of DNA in a single lymphocyte; exposure to an antigen recognized by the Ig or TCR leads to clonal expansion to generate many cells which have the same specificity as the original cell. Many different rearrangements occur early in the development of the immune system, creating a large repertoire of cells of different specificities. Each immunoglobulin protein is a tetramer containing two identical light chains and two identical heavy chains. A TCR is a dimer containing two different chains. Each polypeptide chain is expressed from a gene created by linking one of many V segments via D and J segments to one of a few C segments. Ig L chains (either κ or λ) have the general structure V-J-C, Ig H chains have the structure V-D-J-C, TCR α and γ have components like Ig L chains, and TCR δ and β are like Ig H chains. Each type of chain is coded by a large cluster of V genes separated from the cluster of D, J, and C segments. The numbers of each type of segment, and their organization, are different for each type of chain, but the principle and mechanism of recombination appear to be the same. The same nonamer and heptamer consensus sequences are involved in each recombination; the reaction always involves joining of a consensus with 23 bp spacing to a consensus with 12 bp spacing. The cleavage reaction is catalyzed by the RAG1 and RAG2 proteins, and the joining reaction is catalyzed by the same NHEJ pathway that repairs double-strand breaks in cells. The mechanism of action of the RAG proteins is related to the action of site-specific recombination catalyzed by resolvases. Although considerable diversity is generated by joining different V, D, J segments to a C segment, additional variations are introduced in the form of changes at the junctions between segments during the recombination process. Changes are also induced in immunoglobulin genes by somatic mutation, which requires the actions of cytidine deaminase and uracil glycosylase. Mutations induced by cytidine deaminase probably lead to removal of uracil by uracil glycosylase, followed by the induction of mutations at the sites where bases are missing. Allelic exclusion ensures that a given lymphocyte synthesizes only a single Ig or TCR. A productive rearrangement inhibits the occurrence of further rearrangements. Although the use of the V region is fixed by the first productive rearrangement, B cells switch use of CH genes from the initial µ chain to one of the H chains coded farther downstream. This process involves a different type of recombination in which the sequences between the VDJ region and the new CH gene are deleted. More than one switch occurs in CH gene usage. Class switching requires the same cytidine deaminase that is required for somatic mutation, but its role is not known. At an earlier stage of Ig production, switches occur from synthesis of a membrane-bound version of the protein to a secreted version. These switches are accomplished by alternative splicing of the transcript. Summary | SECTION 5.25.22 © 2004. Virtual Text / www.ergito.com

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Innate immunity is a response triggered by receptors whose specificity is predefined for certain common motifs found in bacteria and other infective agents. The receptor that triggers the pathway is typically a member of the Toll-like class, and the pathway resembles the pathway triggered by Toll receptors during embryonic development. The pathway culminates in activation of transcription factors that cause genes to be expressed whose products inactivate the infective agent, typically by permeabilizing its membrane. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.25.22


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CATALYTIC RNA

5.26.1 Introduction Key Terms A ribozyme is an RNA that has catalytic activity. RNA editing describes a change of sequence at the level of RNA following transcription.

The idea that only proteins have enzymatic activity was deeply rooted in biochemistry. (Yet devotées of protein function once thought that only proteins could have the versatility to be the genetic material!) A rationale for the identification of enzymes with proteins lies in the view that only proteins, with their varied three-dimensional structures and variety of side-groups, have the flexibility to create the active sites that catalyze biochemical reactions. But the characterization of systems involved in RNA processing has shown this view to be an over-simplification. Several types of catalytic reactions are now known to reside in RNA. Ribozyme has become a general term used to describe an RNA with catalytic activity, and it is possible to characterize the enzymatic activity in the same way as a more conventional enzyme. Some RNA catalytic activities are directed against separate substrates, while others are intramolecular (which limits the catalytic action to a single cycle). Introns of the group I and group II classes possess the ability to splice themselves out of the pre-mRNA that contains them. Engineering of group I introns has generated RNA molecules that have several other catalytic activities related to the original activity. The enzyme ribonuclease P is a ribonucleoprotein that contains a single RNA molecule bound to a protein. The RNA possesses the ability to catalyze cleavage in a tRNA substrate, while the protein component plays an indirect role, probably to maintain the structure of the catalytic RNA. The common theme of these reactions is that the RNA can perform an intramolecular or intermolecular reaction that involves cleavage or joining of phosphodiester bonds in vitro. Although the specificity of the reaction and the basic catalytic activity is provided by RNA, proteins associated with the RNA may be needed for the reaction to occur efficiently in vivo. RNA splicing is not the only means by which changes can be introduced in the informational content of RNA. In the process of RNA editing, changes are introduced at individual bases, or bases are added at particular positions within an mRNA. The insertion of bases (most commonly uridine residues) occurs for several genes in the mitochondria of certain lower eukaryotes; like splicing, it involves the breakage and reunion of bonds between nucleotides, but also requires a template for Introduction | SECTION 5.26.1 © 2004. Virtual Text / www.ergito.com

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coding the information of the new sequence. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.26.1


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CATALYTIC RNA

5.26.2 Group I introns undertake self-splicing by transesterification Key Terms Autosplicing (Self-splicing) describes the ability of an intron to excise itself from an RNA by a catalytic action that depends only on the sequence of RNA in the intron. Key Concepts

• The only factors required for autosplicing in vitro by group I introns are a monovalent cation, a divalent cation, and a guanine nucleotide.

• Splicing occurs by two transesterifications, without requiring input of energy. • The 3 ′ –OH end of the guanine cofactor attacks the 5 ′ end of the intron in the first transesterification.

• The 3 ′ –OH end generated at the end of the first exon attacks the junction between the intron and second exon in the second transesterification.

• The intron is released as a linear molecule that circularizes when its 3 ′ –OH terminus attacks a bond at one of two internal positions.

414 16 • The G –A internal bond of the intron can also be attacked by other nucleotides

in a trans-splicing reaction.

Group I introns are found in diverse locations. They occur in the genes coding for rRNA in the nuclei of the lower eukaryotes Tetrahymena thermophila (a ciliate) and Physarum polycephalum (a slime mold). They are common in the genes of fungal mitochondria. They are present in three genes of phage T4 and also are found in bacteria. Group I introns have an intrinsic ability to splice themselves. This is called self-splicing or autosplicing. (This property is found also in the group II introns discussed in Molecular Biology 5.24.11 Group II introns autosplice via lariat formation. (749)) Self-splicing was discovered as a property of the transcripts of the rRNA genes in T. thermophila (for an account of the discovery see Great Experiments 4.4 RNA catalysis). The genes for the two major rRNAs follow the usual organization, in which both are expressed as part of a common transcription unit. The product is a 35S precursor RNA with the sequence of the small rRNA in the 5 ′ part, and the sequence of the larger (26S) rRNA toward the 3 ′ end. In some strains of T. thermophila, the sequence coding for 26S rRNA is interrupted by a single, short intron. When the 35S precursor RNA is incubated in vitro, splicing occurs as an autonomous reaction. The intron is excised from the precursor and accumulates as a linear fragment of 400 bases, which is subsequently converted to a Group I introns undertake self-splicing by transesterification | SECTION 5.26.2 © 2004. Virtual Text / www.ergito.com

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circular RNA. These events are summarized in Figure 26.1.

Figure 26.1 Splicing of the Tetrahymena 35S rRNA precursor can be followed by gel electrophoresis. The removal of the intron is revealed by the appearance of a rapidly moving small band. When the intron becomes circular, it electrophoreses more slowly, as seen by a higher band.

The reaction requires only a monovalent cation, a divalent cation, and a guanine nucleotide cofactor. No other base can be substituted for G; but a triphosphate is not needed; GTP, GDP, GMP, and guanosine itself all can be used, so there is no net energy requirement. The guanine nucleotide must have a 3 ′ –OH group (747). The fate of the guanine nucleotide can be followed by using a radioactive label. The radioactivity initially enters the excised linear intron fragment. The G residue becomes linked to the 5 ′ end of the intron by a normal phosphodiester bond. Figure 26.2 shows that three transfer reactions occur. In the first transfer, the guanine nucleotide behaves as a cofactor that provides a free 3 ′ –OH group that attacks the 5 ′ end of the intron. This reaction creates the G–intron link and generates a 3 ′ –OH group at the end of the exon. The second transfer involves a similar chemical reaction, in which this 3 ′ –OH then attacks the second exon. The two transfers are connected; no free exons have been observed, so their ligation may occur as part of the same reaction that releases the intron. The intron is released as a linear molecule, but the third transfer reaction converts it to a circle.

Group I introns undertake self-splicing by transesterification | SECTION 5.26.2 © 2004. Virtual Text / www.ergito.com

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Figure 26.2 Self-splicing occurs by transesterification reactions in which bonds are exchanged directly. The bonds that have been generated at each stage are indicated by the shaded boxes.

Each stage of the self-splicing reaction occurs by a transesterification, in which one phosphate ester is converted directly into another, without any intermediary hydrolysis. Bonds are exchanged directly, and energy is conserved, so the reaction does not require input of energy from hydrolysis of ATP or GTP. (There is a parallel for the transfer of bonds without net input of energy in the DNA nicking-closing enzymes discussed in Molecular Biology 4.15 Recombination and repair.) If each of the consecutive transesterification reactions involves no net change of energy, why does the splicing reaction proceed to completion instead of coming to equilibrium between spliced product and nonspliced precursor? The concentration of GTP is high relative to that of RNA, and therefore drives the reaction forward; and a change in secondary structure in the RNA prevents the reverse reaction. The in vitro system includes no protein so the ability to splice is intrinsic to the RNA. Group I introns undertake self-splicing by transesterification | SECTION 5.26.2 © 2004. Virtual Text / www.ergito.com

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The RNA forms a specific secondary/tertiary structure in which the relevant groups are brought into juxtaposition so that a guanine nucleotide can be bound to a specific site and then the bond breakage and reunion reactions shown in Figure 26.2 can occur. Although a property of the RNA itself, the reaction is assisted in vivo by proteins, which stabilize the RNA structure (748). The ability to engage in these transfer reactions resides with the sequence of the intron, which continues to be reactive after its excision as a linear molecule. Figure 26.3 summarizes its activities:

Figure 26.3 The excised intron can form circles by using either of two internal sites for reaction with the 5 ′ end, and can reopen the circles by reaction with water or oligonucleotides.

The intron can circularize when the 3 ′ terminal G attacks either of two positions near the 5 ′ end. The internal bond is broken and the new 5 ′ end is transferred to the 3 ′ –OH end of the intron. The primary cyclization usually involves reaction between the terminal G414 and the A16. This is the most common reaction (shown as the third Group I introns undertake self-splicing by transesterification | SECTION 5.26.2 © 2004. Virtual Text / www.ergito.com

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Reviews 255. Cech, T. R. (1985). Self-splicing RNA: implications for evolution. Int. Rev. Cytol. 93, 3-22. 256. Cech, T. R. (1987). The chemistry of self-splicing RNA and RNA enzymes. Science 236, 1532-1539.

References 747. Cech, T. R. et al. (1981). In vitro splicing of the rRNA precursor of Tetrahymena: involvement of a guanosine nucleotide in the excision of the intervening sequence. Cell 27, 487-496. 748. Kruger, K., Grabowski, P. J., Zaug, A. J., Sands, J., Gottschling, D. E., and Cech, T. R. (1982). Self-splicing RNA: autoexcision and autocyclization of the ribosomal RNA intervening sequence of Tetrahymena. Cell 31, 147-157. 749. Belfort, M., Pedersen-Lane, J., West, D., Ehrenman, K., Maley, G., Chu, F., and Maley, F. (1985). Processing of the intron-containing thymidylate synthase (td) gene of phage T4 is at the RNA level. Cell 41, 375-382. 750. Been, M. D. and Cech, T. R. (1986). One binding site determines sequence specificity of Tetrahymena pre-rRNA self-splicing, trans-splicing, and RNA enzyme activity. Cell 47, 207-216. 2512. Myers, C. A., Kuhla, B., Cusack, S., and Lambowitz, A. M. (2002). tRNA-like recognition of group I introns by a tyrosyl-tRNA synthetase. Proc. Natl. Acad. Sci. USA 99, 2630-2635.



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CATALYTIC RNA

5.26.3 Group I introns form a characteristic secondary structure Key Concepts

• Group I introns form a secondary structure with 9 duplex regions. • The core of regions P3, P4, P6, P7 has catalytic activity. • Regions P4 and P7 are both formed by pairing between conserved consensus sequences.

• A sequence adjacent to P7 base pairs with the sequence that contains the reactive G.

All group I introns can be organized into a characteristic secondary structure, with 9 helices (P1-P9). Figure 26.4 shows a model for the secondary structure of the Tetrahymena intron. Two of the base-paired regions are generated by pairing between conserved sequence elements that are common to group I introns. P4 is constructed from the sequences P and Q;, P7 is formed from sequences R and S. The other base-paired regions vary in sequence in individual introns. Mutational analysis identifies an intron "core," containing P3, P4, P6, and P7, which provides the minimal region that can undertake a catalytic reaction. The lengths of group I introns vary widely, and the consensus sequences are located a considerable distance from the actual splice junctions.

Group I introns form a characteristic secondary structure | SECTION 5.26.3 © 2004. Virtual Text / www.ergito.com

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Figure 26.4 Group I introns have a common secondary structure that is formed by 9 base paired regions. The sequences of regions P4 and P7 are conserved, and identify the individual sequence elements P, Q, R, and S. P1 is created by pairing between the end of the left exon and the IGS of the intron; a region between P7 and P9 pairs with the 3 ′ end of the intron.

Some of the pairing reactions are directly involved in bringing the splice junctions into a conformation that supports the enzymatic reaction. P1 includes the 3 ′ end of the left exon. The sequence within the intron that pairs with the exon is called the IGS, or internal guide sequence. (Its name reflects the fact that originally the region immediately 3 ′ to the IGS sequence shown in the figure was thought to pair with the 3 ′ splice junction, thus bringing the two junctions together. This interaction may occur, but does not seem to be essential.) A very short sequence, sometimes as short as 2 bases, between P7 and P9, base pairs with the sequence that immediately precedes the reactive G (position 414 in Tetrahymena) at the 3 ′ end of the intron (752). The importance of base pairing in creating the necessary core structure in the RNA is emphasized by the properties of cis-acting mutations that prevent splicing of group I introns. Such mutations have been isolated for the mitochondrial introns through mutants that cannot remove an intron in vivo, and they have been isolated for the Tetrahymena intron by transferring the splicing reaction into a bacterial environment. The construct shown in Figure 26.5 allows the splicing reaction to be followed in E. coli. The self-splicing intron is placed at a location that interrupts the tenth codon of the β-galactosidase coding sequence. The protein can therefore be successfully translated from an RNA only after the intron has been removed.


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Figure 26.5 Placing the Tetrahymena intron within the β -galactosidase coding sequence creates an assay for self-splicing in E. coli. Synthesis of β -galactosidase can be tested by adding a compound that is turned blue by the enzyme. The sequence is carried by a bacteriophage, so the presence of blue plaques (containing infected bacteria) indicates successful splicing.

The synthesis of β-galactosidase in this system indicates that splicing can occur in conditions quite distant from those prevailing in Tetrahymena or even in vitro. One interpretation of this result is that self-splicing can occur in the bacterial cell. Another possibility is that there are bacterial proteins that assist the reaction. Using this assay, we can introduce mutations into the intron to see whether they prevent the reaction. Mutations in the group I consensus sequences that disrupt their base pairing stop splicing. The mutations can be reverted by making compensating changes that restore base pairing. Mutations in the corresponding consensus sequences in mitochondrial group I introns have similar effects. A mutation in one consensus sequence may be reverted by a mutation in the complementary consensus sequence to restore pairing; for example, mutations in the R consensus can be compensated by mutations in the S consensus. Together these results suggest that the group I splicing reaction depends on the formation of secondary structure between pairs of consensus sequences within the intron. The principle established by this work is that sequences distant from the splice junctions themselves are required to form the active site that makes self-splicing possible (751).


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References 751. Burke, J. M. et al. (1986). Role of conserved sequence elements 9L and 2 in self-splicing of the Tetrahymena ribosomal RNA precursor. Cell 45, 167-176. 752. Michel, F. and Wetshof, E. (1990). Modeling of the three-dimensional architecture of group I catalytic introns based on comparative sequence analysis. J. Mol. Biol. 216, 585-610.



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CATALYTIC RNA

5.26.4 Ribozymes have various catalytic activities Key Concepts

• By changing the substrate binding-site of a group I intron, it is possible to introduce alternative sequences that interact with the reactive G.

• The reactions follow classical enzyme kinetics with a low catalytic rate. • Reactions using 2 ′ –OH bonds could have been the basis for evolving the original catalytic activities in RNA.

The catalytic activity of group I introns was discovered by virtue of their ability to autosplice, but they are able to undertake other catalytic reactions in vitro. All of these reactions are based on transesterifications. We analyze these reactions in terms of their relationship to the splicing reaction itself. The catalytic activity of a group I intron is conferred by its ability to generate a particular secondary and tertiary structure that creates active sites, equivalent to the active sites of a conventional (proteinaceous) enzyme. Figure 26.6 illustrates the splicing reaction in terms of these sites (this is the same series of reactions shown previously in Figure 26.2).

Ribozymes have various catalytic activities | SECTION 5.26.4 © 2004. Virtual Text / www.ergito.com

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Figure 26.6 Excision of the group I intron in Tetrahymena rRNA occurs by successive reactions between the occupants of the guanosine-binding site and substrate-binding site. The left exon is red, and the right exon is purple.

The substrate-binding site is formed from the P1 helix, in which the 3 ′ end of the first intron base pairs with the IGS in an intermolecular reaction. A guanosine-binding site is formed by sequences in P7. This site may be occupied either by a free guanosine nucleotide or by the G residue in position 414. In the first transfer reaction, it is used by free guanosine nucleotide; but it is subsequently occupied by G414. The second transfer releases the joined exons. The third transfer creates the circular intron. Binding to the substrate involves a change of conformation; before substrate binding, the 5 ′ end of the IGS is close to P2 and P8, but after binding, when it forms the P1 Ribozymes have various catalytic activities | SECTION 5.26.4 © 2004. Virtual Text / www.ergito.com

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helix, it is close to conserved bases that lie between P4 and P5. The reaction is visualized by contacts that are detected in the secondary structure in Figure 26.7. In the tertiary structure, the two sites alternatively contacted by P1 are 37 Å apart, which implies a substantial movement in the position of P1.

Figure 26.7 The position of the IGS in the tertiary structure changes when P1 is formed by substrate binding.

The L-19 RNA is generated by opening the circular intron (shown as the last stage of the intramolecular rearrangements shown in Figure 26.3). It still retains enzymatic abilities. These resemble the activities involved in the original splicing reaction, and we may consider ribozyme function in terms of the ability to bind an intramolecular sequence complementary to the IGS in the substrate-binding site, while binding either the terminal G414 or a free G-nucleotide in the G-binding site. Figure 26.8 illustrates the mechanism by which the oligonucleotide C5 is extended to generate a C6 chain. The C5 oligonucleotide binds in the substrate-binding site, while G414 occupies the G-binding site. By transesterification reactions, a C is transferred from C5 to the 3 ′ –terminal G, and then back to a new C5 molecule. Further transfer reactions lead to the accumulation of longer cytosine oligonucleotides. The reaction is a true catalysis, because the L-19 RNA remains unchanged, and is available to catalyze multiple cycles. The ribozyme is behaving as a nucleotidyl transferase.


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Figure 26.8 The L-19 linear RNA can bind C in the substrate-binding site; the reactive G-OH 3 ′ end is located in the G-binding site, and catalyzes transfer reactions that convert 2 C5 oligonucleotides into a C4 and a C6 oligonucleotide.

Some further enzymatic reactions are characterized in Figure 26.9. The ribozyme can function as a sequence-specific endoribonuclease by utilizing the ability of the IGS to bind complementary sequences. In this example, it binds an external substrate containing the sequence CUCU, instead of binding the analogous sequence that is usually contained at the end of the left exon. A guanine-containing nucleotide is present in the G-binding site, and attacks the CUCU sequence in precisely the same way that the exon is usually attacked in the first transfer reaction. This cleaves the target sequence into a 5 ′ molecule that resembles the left exon, and a 3 ′ molecule that bears a terminal G residue. By mutating the IGS element, it is possible to change the specificity of the ribozyme, so that it recognizes sequences complementary to the new sequence at the IGS region (for review see 257).


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Figure 26.9 Catalytic reactions of the ribozyme involve transesterifications between a group in the substrate-binding site and a group in the G-binding site.

Altering the IGS, so that the specificity of the substrate-binding site is changed to enable other RNA targets to enter, can be used to generate a ligase activity. An RNA terminating in a 3 ′ –OH is bound in the substrate site, and an RNA terminating in a 5 ′ –G residue is bound in the G-binding site. An attack by the hydroxyl on the phosphate bond connects the two RNA molecules, with the loss of the G residue. The phosphatase reaction is not directly related to the splicing transfer reactions. An oligonucleotide sequence that is complementary to the IGS and terminates in a 3 ′ –phosphate can be attacked by the G414. The phosphate is transferred to the G414, and an oligonucleotide with a free 3 ′ –OH end is then released. The phosphate can then be transferred either to an oligonucleotide terminating in 3 ′ –OH (effectively reversing the reaction) or indeed to water (releasing inorganic phosphate and completing an authentic phosphatase reaction). The reactions catalyzed by RNA can be characterized in the same way as classical enzymatic reactions in terms of Michaelis-Menten kinetics. Figure 26.10 analyzes the reactions catalyzed by RNA. The KM values for RNA-catalyzed reactions are Ribozymes have various catalytic activities | SECTION 5.26.4 © 2004. Virtual Text / www.ergito.com

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low, and therefore imply that the RNA can bind its substrate with high specificity. The turnover numbers are low, which reflects a low catalytic rate. In effect, the RNA molecules behave in the same general manner as traditionally defined for enzymes, although they are relatively slow compared to protein catalysts (where a typical range of turnover numbers is 103–106).

Figure 26.10 Reactions catalyzed by RNA have the same features as those catalyzed by proteins, although the rate is slower. The KM gives the concentration of substrate required for half-maximum velocity; this is an inverse measure of the affinity of the enzyme for substrate. The turnover number gives the number of substrate molecules transformed in unit time by a single catalytic site.

How does RNA provide a catalytic center? Its ability seems reasonable if we think of an active center as a surface that exposes a series of active groups in a fixed relationship. In a protein, the active groups are provided by the side chains of the amino acids, which have appreciable variety, including positive and negative ionic groups and hydrophobic groups. In an RNA, the available moieties are more restricted, consisting primarily of the exposed groups of bases. Short regions are held in a particular structure by the secondary/tertiary conformation of the molecule, providing a surface of active groups able to maintain an environment in which bonds can be broken and made in another molecule. It seems inevitable that the interaction between the RNA catalyst and the RNA substrate will rely on base pairing to create the environment. Divalent cations (typically Mg2+) play an important role in structure, typically being present at the active site where they coordinate the positions of the various groups. They play a direct role in the endonucleolytic activity of virusoid ribozymes (see Molecular Biology 5.26.9 Viroids have catalytic activity). The evolutionary implications of these discoveries are intriguing. The split personality of the genetic apparatus, in which RNA is present in all components, but proteins undertake catalytic reactions, has always been puzzling. It seems unlikely that the very first replicating systems could have contained both nucleic acid and protein. But suppose that the first systems contained only a self-replicating nucleic acid with primitive catalytic activities, just those needed to make and break phosphodiester bonds. If we suppose that the involvement of 2 ′ –OH bonds in current splicing reactions is derived from these primitive catalytic activities, we may argue that the original nucleic acid was RNA, since DNA lacks the 2 ′ –OH group and therefore Ribozymes have various catalytic activities | SECTION 5.26.4 © 2004. Virtual Text / www.ergito.com

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could not undertake such reactions. Proteins could have been added for their ability to stabilize the RNA structure. Then the greater versatility of proteins could have allowed them to take over catalytic reactions, leading eventually to the complex and sophisticated apparatus of modern gene expression.


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Reviews 257. Cech, T. R. (1990). Self-splicing of group I introns. Annu. Rev. Biochem. 59, 543-568.



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CATALYTIC RNA

5.26.5 Some group I introns code for endonucleases that sponsor mobility Key Terms Intron homing describes the ability of certain introns to insert themselves into a target DNA. The reaction is specific for a single target sequence. Key Concepts

• Mobile introns are able to insert themselves into new sites. • Mobile group I introns code for an endonuclease that makes a double-strand break at a target site.

• The intron transposes into the site of the double-strand break by a DNA-mediated replicative mechanism.

Certain introns of both the group I and group II classes contain open reading frames that are translated into proteins. Expression of the proteins allows the intron (either in its original DNA form or as a DNA copy of the RNA) to be mobile: it is able to insert itself into a new genomic site. Introns of both groups I and II are extremely widespread, being found in both prokaryotes and eukaryotes. Group I introns migrate by DNA-mediated mechanisms, whereas group II introns migrate by RNA-mediated mechanisms. Intron mobility was first detected by crosses in which the alleles for the relevant gene differ with regard to their possession of the intron. Polymorphisms for the presence or absence of introns are common in fungal mitochondria. This is consistent with the view that these introns originated by insertion into the gene. Some light on the process that could be involved is cast by an analysis of recombination in crosses involving the large rRNA gene of the yeast mitochondrion. This gene has a group I intron that contains a coding sequence. The intron is present in some strains of yeast (called ω+) but absent in others ( ω–). Genetic crosses between ω+ and ω– are polar: the progeny are usually ω+. If we think of the ω+ strain as a donor and the ω– strain as a recipient, we form the view that in ω+× ω– crosses a new copy of the intron is generated in the ω– genome. As a result, the progeny are all ω+. Mutations can occur in either parent to abolish the polarity. Mutants show normal segregation, with equal numbers of ω+ and ω– progeny. The mutations indicate the nature of the process. Mutations in the ω– strain occur close to the site where the intron would be inserted. Mutations in the ω+ strain lie in the reading frame of the intron and prevent production of the protein. This suggests the model of Figure 26.11, in which the protein coded by the intron in an ω+ strain recognizes the site Some group I introns code for endonucleases that sponsor mobility | SECTION 5.26.5 © 2004. Virtual Text / www.ergito.com

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Figure 26.11 An intron codes for an endonuclease that makes a double-strand break in DNA. The sequence of the intron is duplicated and then inserted at the break.

What is the action of the protein? The product of the ω intron is an endonuclease that recognizes the ω– gene as a target for a double-strand break. The endonuclease recognizes an 18 bp target sequence that contains the site where the intron is inserted. The target sequence is cleaved on each strand of DNA 2 bases to the 3 ′ side of the insertion site. So the cleavage sites are 4 bp apart, and generate overhanging single strands. This type of cleavage is related to the cleavage characteristic of transposons when they migrate to new sites (see Molecular Biology 4.16 Transposons). The double-strand break probably initiates a gene conversion process in which the sequence of the ω+ gene is copied to replace the sequence of the ω– gene. The reaction involves transposition by a duplicative mechanism, and occurs solely at the level of DNA. Insertion of the intron interrupts the sequence recognized by the endonuclease, thus ensuring stability. Many group I introns code for endonucleases that make them mobile. Several different families of endonucleases are found; one common feature is the presence of the amino acid sequence LAGLIDADG near the active site (for review see 4515). Similar introns often carry quite different endonucleases. There are differences in the details of insertion; for example, the endonuclease coded by the phage T4 td intron cleaves a target site that is 24 bp upstream of the site at which the intron is itself inserted. The dissociation between the intron sequence and the endonuclease sequence is emphasized by the fact that the same endonuclease sequences are found in inteins (sequences that code for self-splicing proteins; see Molecular Some group I introns code for endonucleases that sponsor mobility | SECTION 5.26.5 © 2004. Virtual Text / www.ergito.com

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Biology 5.26.12 Protein splicing is autocatalytic). The variation in the endonucleases means that there is no homology between the sequences of their target sites. The target sites are among the longest and therefore the most specific known for any endonucleases (with a range of 14-40 bp). The specificity ensures that the intron perpetuates itself only by insertion into a single target site and not elsewhere in the genome. This is called intron homing. Introns carrying sequences that code for endonucleases are found in a variety of bacteria and lower eukaryotes. These results strengthen the view that introns carrying coding sequences originated as independent elements. Last updated on December 24, 2003

Some group I introns code for endonucleases that sponsor mobility | SECTION 5.26.5 © 2004. Virtual Text / www.ergito.com

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Reviews 4515. Belfort, M. and Roberts, R. J. (1997). Homing endonucleases: keeping the house in order. Nucleic Acids Res. 25, 3379-3388.


Some group I introns code for endonucleases that sponsor mobility | SECTION 5.26.5 © 2004. Virtual Text / www.ergito.com

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CATALYTIC RNA

5.26.6 Some group II introns code for reverse transcriptases Key Concepts

• Some group II introns code for a reverse transcriptase that generates a DNA copy of the RNA sequence that transposes by a retroposon-like mechanism.

Most of the open reading frames contained in group II introns have regions that are related to reverse transcriptases. Introns of this type are found in organelles of lower eukaryotes and also in some bacteria. The reverse transcriptase activity is specific for the intron, and is involved in homing. The reverse transcriptase generates a DNA copy of the intron from the pre-mRNA, and thus allows the intron to become mobile by a mechanism resembling that of retroviruses (see Molecular Biology 4.17.2 The retrovirus life cycle involves transposition-like events). The type of retrotransposition involved in this case resembles that of a group of retroposons that lack LTRs, and which generate the 3 ′ –OH needed for priming by making a nick in the target (see Figure 17.20 in Molecular Biology 4.17.12 LINES use an endonuclease to generate a priming end). The best characterized mobile group II introns code for a single protein in a region of the intron beyond its catalytic core. The typical protein contains an N-terminal reverse transcriptase activity, a central domain associated with maturase activity, and a C-terminal endonuclease domain. The endonuclease initiates the transposition reaction, and thus plays the same role in homing as its counterpart in a group I intron. The reverse transcriptase generates a DNA copy of the intron that is inserted at the homing site. The endonuclease also cleaves target sites that resemble, but are not identical to the homing site, at much lower frequency, leading to insertion of the intron at new locations (3330). Figure 26.12 illustrates the transposition reaction for a typical group II intron. The endonuclease makes a double-strand break at the target site. A 3 ′ end is generated at the site of the break, and provides a primer for the reverse transcriptase. The intron RNA provides the template for the synthesis of cDNA. Because the RNA includes exon sequences on either side of the intron, the cDNA product is longer than the region of the intron itself, so that it can span the double-strand break, allowing the cDNA to repair the break. The result is the insertion of the intron (754; for review see 259).

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Figure 26.12 Reverse transcriptase coded by an intron allows a copy of the RNA to be inserted at a target site generated by a double-strand break.



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Reviews 259. Lambowitz, A. M. and Belfort, M. (1993). Introns as mobile genetic elements. Annu. Rev. Biochem. 62, 587-622.

References 754. Zimmerly, S. et al. (1995). Group II intron mobility occurs by target DNA-primed reverse transcription. Cell 82, 545-554. 3330. Dickson, L., Huang, H. R., Liu, L., Matsuura, M., Lambowitz, A. M., and Perlman, P. S. (2001). Retrotransposition of a yeast group II intron occurs by reverse splicing directly into ectopic DNA sites. Proc. Natl. Acad. Sci. USA 98, 13207-13212.



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5.26.7 Some autosplicing introns require maturases Key Terms A maturase is a protein coded by a group I or group II intron that is needed to assist the RNA to form the active conformation that is required for self-splicing.

Both types of autosplicing intron may code for proteins invovled in their perpetuation and homing, such as endonucelases in group I and reverse transcriptases in group II. In addition, both tyeps of intron may code for maturase activities that are required to assist the splicing reaction (753). The maturase activity is part of the single open reading frame coded by the intron. In the example of introns that code for homing endonucleases, the single protein product has both endonuclease and maturase activity. Mutational analysis shows that the two activities are independent (4516). Structural analysis shows that the endonuclease and maturase activities are provided by different active sites in the protein, each coded by a separate domain (4517). The endonuclease site binds to DNA, but the maturase site binds to the intron RNA. Figure 26.13 shows the structure of one such protein bound to DNA. A characteristic feature of the endonuclease is the presence of parallel α helices, containing the hallmark LAGLIDADG sequences, leading to the two catalytic amino acids. The maturase activity is located some distance away on the surface of the protein.

Figure 26.13 A homing intron codes for and endonuclease of the LAGLIDADG family that also has maturase activity. The LAGLIDADG sequences are part of the two α helices that terminate in the catalytic amino acids close to the DNA duplex. The maturase active site is identified by an arginine residue elsewhere on the surface of the protein.

Introns that code for maturases may be unable to splice themselves effectively in the absence of the protein activity. The maturase is in effect a splicing factor that is Some autosplicing introns require maturases | SECTION 5.26.7 © 2004. Virtual Text / www.ergito.com

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required specifically for splicing of the sequence that codes for it. It functions to assist the folding of the catalytic core to form an active site (3329). The coexistence of endonuclease and maturase activities in the same protein suggests a route for the evolution of the intron. Figure 26.14 suggests that the intron originated in an independent autosplicing element. The insertion into this element of a sequence coding for an endonuclease gave it mobility. However, the insertion might well disruupt the ability of the RNA sequence to fold into the active structure. This would create pressure for assistance from proteins that could restore folding ability. The incorporation of such a sequence into the intron would maintain its independence.

Figure 26.14 The intron originated as independent sequence coding for a self-splicing RNA. The insertion of the endonuclease sequence created a homing intron that was mobile. Then the insertion of the maturase sequence enhanced the ability of the itnron sequewnces to fold into the active structure for splicing.

Some group II introns that do not code for maturase activities may use comparable proteins that are coded by sequences in the host genome. This suggests a possible Some autosplicing introns require maturases | SECTION 5.26.7 © 2004. Virtual Text / www.ergito.com

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route for the evolution of general splicing factors. The factor may have originated as a maturase that specifically assisted the splicing of a particular intron. The coding sequence became isolated from the intron in the host genome, and then it evolved to function with a wider range of substrates that the original intron sequence. The catalytic core of the intron could have evolved into an snRNA. Last updated on January 6, 2004

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References 753. Carignani, G. et al. (1983). An RNA maturase is encoded by the first intron of the mitochondrial gene for the subunit I of cytochrome oxidase in S. cerevisiae. Cell 35, 733-742. 3329. Matsuura, M., Noah, J. W., and Lambowitz, A. M. (2001). Mechanism of maturase-promoted group II intron splicing. EMBO J. 20, 7259-7270. 4516. Henke, R. M., Butow, R. A., and Perlman, P. S. (1995). Maturase and endonuclease functions depend on separate conserved domains of the bifunctional protein encoded by the group I intron aI4 alpha of yeast mitochondrial DNA. EMBO J. 14, 5094-5099. 4517. Bolduc et al. (2003). Structural and biochemical analyses of DNA and RNA binding by a bifunctional homing endopnuclease and group I splicing factor. Genes. Dev. 17, 2875-2888.


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CATALYTIC RNA

5.26.8 The catalytic activity of RNAase P is due to RNA Key Concepts

• Ribonuclease P is a ribonucleoprotein in which the RNA has catalytic activity.

One of the first demonstrations of the capabilities of RNA was provided by the dissection of ribonuclease P, an E. coli tRNA-processing endonuclease. Ribonuclease P can be dissociated into its two components, the 375 base RNA and the 20 kD polypeptide. Under the conditions initially used to characterize the enzyme activity in vitro, both components were necessary to cleave the tRNA substrate. But a change in ionic conditions, an increase in the concentration of Mg2+, renders the protein component superfluous. The RNA alone can catalyze the reaction! Analyzing the results as though the RNA were an enzyme, each "enzyme" catalyzes the cleavage of multiple substrates. Although the catalytic activity resides in the RNA, the protein component greatly increases the speed of the reaction, as seen in the increase in turnover number (see Figure 26.10). Because mutations in either the gene for the RNA or the gene for protein can inactivate RNAase P in vivo, we know that both components are necessary for natural enzyme activity. Originally it had been assumed that the protein provided the catalytic activity, while the RNA filled some subsidiary role, for example, assisting in the binding of substrate (it has some short sequences complementary to exposed regions of tRNA). But these roles are reversed! This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.26.8

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CATALYTIC RNA

5.26.9 Viroids have catalytic activity Key Terms A viroid is a small infectious nucleic acid that does not have a protein coat. A virusoid (Satellite RNA) is a small infectious nucleic acid that is encapsidated by a plant virus together with its own genome. Key Concepts

• Viroids and virusoids form a hammerhead structure that has an self-cleaving activity.

• Similar structures can be generated by pairing a substrate strand that is cleaved by an enzyme strand.

• When an enzyme strand is introduced into a cell, it can pair with a substrate strand target that is then cleaved.

Another example of the ability of RNA to function as an endonuclease is provided by some small plant RNAs (~350 bases) that undertake a self-cleavage reaction. As with the case of the Tetrahymena group I intron, however, it is possible to engineer constructs that can function on external substrates (756). These small plant RNAs fall into two general groups: viroids and virusoids. The viroids are infectious RNA molecules that function independently, without enacapsidation by any protein coat. The virusoids are similar in organization, but are encapsidated by plant viruses, being packaged together with a viral genome. The virusoids cannot replicate independently, but require assistance from the virus. The virusoids are sometimes called satellite RNAs. Viroids and virusoids both replicate via rolling circles (see Figure 13.16). The strand of RNA that is packaged into the virus is called the plus strand. The complementary strand, generated during replication of the RNA, is called the minus strand. Multimers of both plus and minus strands are found. Both types of monomer are generated by cleaving the tail of a rolling circle; circular plus strand monomers are generated by ligating the ends of the linear monomer. Both plus and minus strands of viroids and virusoids undergo self-cleavage in vitro. The cleavage reaction is promoted by divalent metal cations; it generates 5 ′ –OH and 2 ′ –3 ′ –cyclic phosphodiester termini. Some of the RNAs cleave in vitro under physiological conditions. Others do so only after a cycle of heating and cooling; this suggests that the isolated RNA has an inappropriate conformation, but can generate an active conformation when it is denatured and renatured. The viroids and virusoids that undergo self-cleavage form a "hammerhead" secondary structure at the cleavage site, as drawn in the upper part of Figure 26.15. Viroids have catalytic activity | SECTION 5.26.9 © 2004. Virtual Text / www.ergito.com

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The sequence of this structure is sufficient for cleavage. When the surrounding sequences are deleted, the need for a heating-cooling cycle is obviated, and the small RNA self-cleaves spontaneously. This suggests that the sequences beyond the hammerhead usually interfere with its formation.

Figure 26.15 Self-cleavage sites of viroids and virusoids have a consensus sequence and form a hammerhead secondary structure by intramolecular pairing. Hammerheads can also be generated by pairing between a substrate strand and an "enzyme" strand.

The active site is a sequence of only 58 nucleotides. The hammerhead contains three stem-loop regions whose position and size are constant, and 13 conserved nucleotides, mostly in the regions connecting the center of the structure. The conserved bases and duplex stems generate an RNA with the intrinsic ability to cleave (757). An active hammerhead can also be generated by pairing an RNA representing one side of the structure with an RNA representing the other side. The lower part of Figure 26.15 shows an example of a hammerhead generated by hybridizing a 19 base molecule with a 24 base molecule. The hybrid mimics the hammerhead structure, with the omission of loops I and III. When the 19 base RNA is added to the 24 base RNA, cleavage occurs at the appropriate position in the hammerhead. We may regard the top (24 base) strand of this hybrid as comprising the "substrate," and the bottom (19 base) strand as comprising the "enzyme." When the 19 base RNA is mixed with an excess of the 24 base RNA, multiple copies of the 24 base RNA are Viroids have catalytic activity | SECTION 5.26.9 © 2004. Virtual Text / www.ergito.com

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cleaved. This suggests that there is a cycle of 19 base–24 base pairing, cleavage, dissociation of the cleaved fragments from the 19 base RNA, and pairing of the 19 base RNA with a new 24 base substrate. The 19 base RNA is therefore a ribozyme with endonuclease activity. The parameters of the reaction are similar to those of other RNA-catalyzed reactions (see Figure 26.10; for review see 258; 2270). The crystal structure of a hammerhead shows that it forms a compact V-shape, in which the catalytic center lies in a turn, as indicated diagrammatically in Figure 26.16. An Mg2+ ion located in the catalytic site plays a crucial role in the reaction. It is positioned by the target cytidine and by the cytidine at the base of stem 1; it may also be connected to the adjacent uridine. It extracts a proton from the 2 ′ –OH of the target cytidine, and then directly attacks the labile phosphodiester bond. Mutations in the hammerhead sequence that affect the transition state of the cleavage reaction occur in both the active site and other locations, suggesting that there may be a substantial rearrangement of structure prior to cleavage (758).

Figure 26.16 A hammerhead ribozyme forms a V-shaped tertiary structure in which stem 2 is stacked upon stem 3. The catalytic center lies between stem 2/3 and stem 1. It contains a magnesium ion that initiates the hydrolytic reaction.

It is possible to design enzyme-substrate combinations that can form hammerhead structures, and these have been used to demonstrate that introduction of the appropriate RNA molecules into a cell can allow the enzymatic reaction to occur in vivo. A ribozyme designed in this way essentially provides a highly specific restriction-like activity directed against an RNA target. By placing the ribozyme under control of a regulated promoter, it can be used in the same way as (for example) antisense constructs specifically to turn off expression of a target gene under defined circumstances.

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Reviews 258. Symons, R. H. (1992). Small catalytic RNAs. Annu. Rev. Biochem. 61, 641-71. 2270. Doherty, E. A. and Doudna, J. A. (2000). Ribozyme structures and mechanisms. Annu. Rev. Biochem. 69, 597-615.

References 756. Guerrier-Takada, C., Gardiner, K., Marsh, T., Pace, N., and Altman, S. (1983). The RNA moiety of ribonuclease P is the catalytic subunit of the enzyme. Cell 35, 849-857. 757. Forster, A. C. and Symons, R. H. (1987). Self-cleavage of virusoid RNA is performed by the proposed 55-nucleotide active site. Cell 50, 9-16. 758. Scott, W. G., Finch, J. T., and Klug, A. (1995). The crystal structure of an all-RNA hammerhead ribozyme: a proposed mechanism for RNA catalytic cleavage. Cell 81, 991-1002.


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CATALYTIC RNA

5.26.10 RNA editing occurs at individual bases Key Terms RNA editing describes a change of sequence at the level of RNA following transcription. Key Concepts

• Apolipoprotein-B and glutamate receptors have site specific deaminations

catalyzed by cytidine and adenosine deaminases that change the coding sequence.

A prime axiom of molecular biology is that the sequence of an mRNA can only represent what is coded in the DNA. The central dogma envisaged a linear relationship in which a continuous sequence of DNA is transcribed into a sequence of mRNA that is in turn directly translated into protein. The occurrence of interrupted genes and the removal of introns by RNA splicing introduces an additional step into the process of gene expression: the coding sequences (exons) in DNA must be reconnected in RNA. But the process remains one of information transfer, in which the actual coding sequence in DNA remains unchanged. Changes in the information coded by DNA occur in some exceptional circumstances, most notably in the generation of new sequences coding for immunoglobulins in mammals and birds. These changes occur specifically in the somatic cells (B lymphocytes) in which immunoglobulins are synthesized (see Molecular Biology 5.25 Immune diversity). New information is generated in the DNA of an individual during the process of reconstructing an immunoglobulin gene; and information coded in the DNA is changed by somatic mutation. The information in DNA continues to be faithfully transcribed into RNA. RNA editing is a process in which information changes at the level of mRNA. It is revealed by situations in which the coding sequence in an RNA differs from the sequence of DNA from which it was transcribed. RNA editing occurs in two different situations, with different causes. In mammalian cells there are cases in which a substitution occurs in an individual base in mRNA, causing a change in the sequence of the protein that is coded. In trypanosome mitochondria, more widespread changes occur in transcripts of several genes, when bases are systematically added or deleted. Figure 26.17 summarizes the sequences of the apolipoprotein-B gene and mRNA in mammalian intestine and liver. The genome contains a single (interrupted) gene whose sequence is identical in all tissues, with a coding region of 4563 codons. This gene is transcribed into an mRNA that is translated into a protein of 512 kD representing the full coding sequence in the liver.

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Figure 26.17 The sequence of the apo-B gene is the same in intestine and liver, but the sequence of the mRNA is modified by a base change that creates a termination codon in intestine.

A shorter form of the protein, ~250 kD, is synthesized in intestine. This protein consists of the N-terminal half of the full-length protein. It is translated from an mRNA whose sequence is identical with that of liver except for a change from C to U at codon 2153. This substitution changes the codon CAA for glutamine into the ochre codon UAA for termination (759). What is responsible for this substitution? No alternative gene or exon is available in the genome to code for the new sequence, and no change in the pattern of splicing can be discovered. We are forced to conclude that a change has been made directly in the sequence of the transcript. Another example is provided by glutamate receptors in rat brain. Editing at one position changes a glutamine codon in DNA into a codon for arginine in RNA; the change affects the conductivity of the channel and therefore has an important effect on controlling ion flow through the neurotransmitter. At another position in the receptor, an arginine codon is converted to a glycine codon (760). The editing event in apo-B causes C2153 to be changed to U; both changes in the glutamate receptor are from A to I (inosine). These events are deaminations in which the amino group on the nucleotide ring is removed. Such events are catalyzed by enzymes called cytidine and adenosine deaminases, respectively. This type of editing appears to occur largely in the nervous system. There are 16 (potential) targets for cytosine deaminase in D. melanogaster, and all are genes involved in neurotransmission. In many cases, the editing event changes an amino acid at a RNA editing occurs at individual bases | SECTION 5.26.10 © 2004. Virtual Text / www.ergito.com

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functionally important position in the protein. What controls the specificity of an editing reaction? Enzymes that undertake deamination as such often have broad specificity – for example, the best characterized adenosine deaminase acts on any A residue in a duplex RNA region. Editing enzymes are related to the general deaminases, but have other regions or additional subunits that control their specificity. In the case of apoB editing, the catalytic subunit of an editing complex is related to bacterial cytidine deaminase, but has an additional RNA-binding region that helps to recognize the specific target site for editing. A special adenosine deaminase enzyme recognizes the target sites in the glutamate receptor RNA, and similar events occur in a serotonin receptor RNA (762). The complex may recognize a particular region of secondary structure in a manner analogous to tRNA-modifying enzymes or could directly recognize a nucleotide sequence. The development of an in vitro system for the apo-B editing event suggests that a relatively small sequence (~26 bases) surrounding the editing site provides a sufficient target. Figure 26.18 shows that in the case of the GluR-B RNA, a base-paired region that is necessary for recognition of the target site is formed between the edited region in the exon and a complementary sequence in the downstream intron. A pattern of mispairing within the duplex region is necessary for specific recognition. So different editing systems may have different types of requirement for sequence specificity in their substrates (761).

Figure 26.18 Editing of mRNA occurs when a deaminase acts on an adenine in an imperfectly paired RNA duplex region.



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References 759. Powell, L. M., Wallis, S. C., Pease, R. J., Edwards, Y. H., Knott, T. J., and Scott, J. (1987). A novel form of tissue-specific RNA processing produces apolipoprotein-B48 in intestine. Cell 50, 831-840. 760. Sommer, B. et al. (1991). RNA editing in brain controls a determinant of ion flow in glutamate-gated channels. Cell 67, 11-19. 761. Higuchi, M. et al. (1993). RNA editing of AMPA receptor subunit GluR-B: a base-paired intron-exon structure determines position and efficiency. Cell 75, 1361-1370. 762. Navaratnam, N. et al. (1995). Evolutionary origins of apoB mRNA editing: catalysis by a cytidine deaminase that has acquired a novel RNA-binding motif at its active site. Cell 81, 187-195.



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CATALYTIC RNA

5.26.11 RNA editing can be directed by guide RNAs Key Terms A guide RNA is a small RNA whose sequence is complementary to the sequence of an RNA that has been edited. It is used as a template for changing the sequence of the pre-edited RNA by inserting or deleting nucleotides. Key Concepts

• Extensive RNA editing in trypanosome mitochondria occurs by insertions or deletions of uridine.

• The substrate RNA base pairs with a guide RNA on both sides of the region to be edited.

• The guide RNA provides the template for addition (or less often deletion) of uridines.

• Editing is catalyzed by a complex of endonuclease, terminal uridyltransferase activity, and RNA ligase.

Another type of editing is revealed by dramatic changes in sequence in the products of several genes of trypanosome mitochondria. In the first case to be discovered, the sequence of the cytochrome oxidase subunit II protein has a frameshift relative to the sequence of the coxII gene. The sequences of the gene and protein given in Figure 26.19 are conserved in several trypanosome species. How does this gene function?

Figure 26.19 The mRNA for the trypanosome coxII gene has a frameshift relative to the DNA; the correct reading frame is created by the insertion of 4 uridines.

The coxII mRNA has an insert of an additional four nucleotides (all uridines) around the site of frameshift (see Great Experiments 4.7 RNA editing). The insertion restores the proper reading frame; it inserts an extra amino acid and changes the amino acids on either side. No second gene with this sequence can be discovered, and we are forced to conclude that the extra bases are inserted during or after transcription (764). A similar discrepancy between mRNA and genomic sequences is found in genes of the SV5 and measles paramyxoviruses, in these cases involving the addition of G RNA editing can be directed by guide RNAs | SECTION 5.26.11 © 2004. Virtual Text / www.ergito.com

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residues in the mRNA. Similar editing of RNA sequences occurs for other genes, and includes deletions as well as additions of uridine. The extraordinary case of the coxIII gene of T. brucei is summarized in Figure 26.20.

Figure 26.20 Part of the mRNA sequence of T. brucei coxIII shows many uridines that are not coded in the DNA (shown in red) or that are removed from the RNA (shown as T).

More than half of the residues in the mRNA consist of uridines that are not coded in the gene. Comparison between the genomic DNA and the mRNA shows that no stretch longer than 7 nucleotides is represented in the mRNA without alteration; and runs of uridine up to 7 bases long are inserted (1277). What provides the information for the specific insertion of uridines? A guide RNA contains a sequence that is complementary to the correctly edited mRNA. Figure 26.21 shows a model for its action in the cytochrome b gene of Leishmania (1280).

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Figure 26.21 Pre-edited RNA base pairs with a guide RNA on both sides of the region to be edited. The guide RNA provides a template for the insertion of uridines. The mRNA produced by the insertions is complementary to the guide RNA.

The sequence at the top shows the original transcript, or pre-edited RNA. Gaps show where bases will be inserted in the editing process. 8 uridines must be inserted into this region to create the valid mRNA sequence. The guide RNA is complementary to the mRNA for a significant distance including and surrounding the edited region. Typically the complementarity is more extensive on the 3 ′ side of the edited region and is rather short on the 5 ′ side. Pairing between the guide RNA and the pre-edited RNA leaves gaps where unpaired A residues in the guide RNA do not find complements in the pre-edited RNA. The guide RNA provides a template that allows the missing U residues to be inserted at these positions. When the reaction is completed, the guide RNA separates from the mRNA, which becomes available for translation. Specification of the final edited sequence can be quite complex; in this example, a lengthy stretch of the transcript is edited by the insertion altogether of 39 U residues, and this appears to require two guide RNAs that act at adjacent sites. The first guide RNA pairs at the 3 ′ –most site, and the edited sequence then becomes a substrate for further editing by the next guide RNA. The guide RNAs are encoded as independent transcription units. Figure 26.22 shows a map of the relevant region of the Leishmania mitochondrial DNA. It includes the "gene" for cytochrome b, which codes for the pre-edited sequence, and two regions RNA editing can be directed by guide RNAs | SECTION 5.26.11 © 2004. Virtual Text / www.ergito.com

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that specify guide RNAs. Genes for the major coding regions and for their guide RNAs are interspersed.

Figure 26.22 The Leishmania genome contains genes coding for pre-edited RNAs interspersed with units that code for the guide RNAs required to generate the correct mRNA sequences. Some genes have multiple guide RNAs.

In principle, a mutation in either the "gene" or one of its guide RNAs could change the primary sequence of the mRNA, and thus of the protein. By genetic criteria, each of these units could be considered to comprise part of the "gene." Since the units are independently expressed, they should of course complement in trans. If mutations were available, we should therefore find that 3 complementation groups were needed to code for the primary sequence of a single protein. The characterization of intermediates that are partially edited suggests that the reaction proceeds along the pre-edited RNA in the 3 ′ –5 ′ direction. The guide RNA determines the specificity of uridine insertions by its pairing with the pre-edited RNA. Editing of uridines is catalyzed by a 20S enzyme complex that contains an endonuclease, a terminal uridyltransferase (TUTase), and an RNA ligase, as illustrated in Figure 26.23 (2399). It binds the guide RNA and uses it to pair with the pre-edited mRNA. The substrate RNA is cleaved at a site that is (presumably) identified by the absence of pairing with the guide RNA, a uridine is inserted or deleted to base pair with the guide RNA, and then the substrate RNA is ligated. UTP provides the source for the uridyl residue. It is added by the TUTase activity; it is not clear whether this activity, or a separate exonuclease, is responsible for deletion. [At one time it was thought that a stretch of U residues at the end of guide RNA might provide the source for added U residues or a sink for deleted residues, but transfer of U residues to guide RNAs appears to be an aberrant reaction that is not responsible for editing (767).]


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Figure 26.23 Addition or deletion of U residues occurs by cleavage of the RNA, removal or addition of the U, and ligation of the ends. The reactions are catalyzed by a complex of enzymes under the direction of guide RNA.

The structures of partially edited molecules suggest that the U residues are added one a time, and not in groups. It is possible that the reaction proceeds through successive cycles in which U residues are added, tested for complementarity with the guide RNA, retained if acceptable and removed if not, so that the construction of the correct edited sequence occurs gradually. We do not know whether the same types of reaction are involved in editing reactions that add C residues. Last updated on 4-17-2002


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References 764. Benne, R., Van den Burg J., Brakenhoff, J. P., Sloof, P., Van Boom, J. H., and Tromp, M. C. (1986). Major transcript of the frameshifted coxII gene from trypanosome mitochondria contains four nucleotides that are not encoded in the DNA. Cell 46, 819-826. 767. Seiwert, S. D., Heidmann, S. and Stuart, K. (1996). Direct visualization of uridylate deletion in vitro suggests a mechanism for kinetoplastid editing. Cell 84, 831-841. 1277. Feagin, J. E., Abraham, J. M., and Stuart, K. (1988). Extensive editing of the cytochrome c oxidase III transcript in Trypanosoma brucei. Cell 53, 413-422. 1280. Blum, B., Bakalara, N., and Simpson, L. (1990). A model for RNA editing in kinetoplastid mitochondria: "guide" RNA molecules transcribed from maxicircle DNA provide the edited information. Cell 60, 189-198. 2399. Aphasizhev, R., Sbicego, S., Peris, M., Jang, S. H., Aphasizheva, I., Simpson, A. M., Rivlin, A., and Simpson, L. (2002). Trypanosome mitochondrial 3 ′ terminal uridylyl transferase (TUTase): the key enzyme in U-insertion/deletion RNA editing. Cell 108, 637-648.



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CATALYTIC RNA

5.26.12 Protein splicing is autocatalytic Key Terms Protein splicing is the autocatalytic process by which an intein is removed from a protein and the exteins on either side become connected by a standard peptide bond. Extein sequences remain in the mature protein that is produced by processing a precursor via protein splicing. An intein is the part that is removed from a protein that is processed by protein splicing. Key Concepts

• An intein has the ability to catalyze its own removal from a protein in such a way that the flanking exteins are connected.

• Protein splicing is catalyzed by the intein. • Most inteins have two independent activities: protein splicing and a homing endonuclease.

Protein splicing has the same effect as RNA splicing: a sequence that is represented within the gene fails to be represented in the protein. The parts of the protein are named by analogy with RNA splicing: exteins are the sequences that are represented in the mature protein, and inteins are the sequences that are removed. The mechanism of removing the intein is completely different from RNA splicing. Figure 26.24 shows that the gene is translated into a protein precursor that contains the intein, and then the intein is excised from the protein. About 100 examples of protein splicing are known, spread through all classes of organisms. The typical gene whose product undergoes protein splicing has a single intein.

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Figure 26.24 In protein splicing the exteins are connected by removing the intein from the protein.

The first intein was discovered in an archaeal DNA polymerase gene in the form of an intervening sequence in the gene that does not conform to the rules for introns (2327). Then it was demonstrated that the purified protein can splice this sequence out of itself in an autocatalytic reaction (2328). The reaction does not require input of energy and occurs through the series of bond rearrangements shown in Figure 26.25. It is a function of the intein, although its efficiency can be influenced by the exteins.


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Figure 26.25 Bonds are rearranged through a series of transesterifications involving the -OH groups of serine or proline or the -SH group of cysteine until finally the exteins are connected by a peptide bond and the intein is released with a circularized C-terminus.

The first reaction is an attack by an -OH or -SH side chain of the first amino acid in the intein on the peptide bond that connects it to the first extein. This transfers the extein from the amino-terminal group of the intein to an N-O or N-S acyl connection. Then this bond is attacked by the -OH or -SH side chain of the first amino acid in the second extein. The result is to transfer extein1 to the side chain of the amino-terminal acid of extein2. Finally, the C-terminal asparagine of the intein cyclizes, and the terminal NH of extein2 attacks the acyl bond to replace it with a conventional peptide bond. Each of these reactions can occur spontaneously at very low rates, but their occurrence in a coordinate manner rapidly enough to achieve protein splicing requires catalysis by the intein (for review see 2267). Inteins have characteristic features. They are found as in-frame insertions into coding sequences. They can be recognized as such because of the existence of homologous genes that lack the insertion. They have an N-terminal serine or cysteine (to provide the -XH side chain) and a C-terminal asparagine. A typical intein has a sequence of Protein splicing is autocatalytic | SECTION 5.26.12 © 2004. Virtual Text / www.ergito.com

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~150 amino acids at the N-terminal end and ~50 amino acids at the C-terminal end that are involved in catalyzing the protein splicing reaction. The sequence in the center of the intein can have other functions. An extraordinary feature of many inteins is that they have homing endonuclease activity. A homing endonuclease cleaves a target DNA to create a site into which the DNA sequence coding for the intein can be inserted (see Figure 26.11 in Molecular Biology 5.26.5 Some group I introns code for endonucleases that sponsor mobility). The protein splicing and homing endonuclease activities of an intein are independent (2329). We do not really understand the connection between the presence of both these activities in an intein, but two types of model have been suggested. One is to suppose that there was originally some sort of connection between the activities, but that they have since become independent and some inteins have lost the homing endonuclease. The other is to suppose that inteins may have originated as protein splicing units, most of which (for unknown reasons) were subsequently invaded by homing endonucleases. This is consistent with the fact that homing endonucleases appear to have invaded other types of units also, including most notably group I introns. Last updated on 2-6-2002


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Reviews 2267. Paulus, H. (2000). Protein splicing and related forms of protein autoprocessing. Annu. Rev. Biochem. 69, 447-496.

References 2327. Perler, F. B. et al. (1992). Intervening sequences in an Archaea DNA polymerase gene. Proc. Natl. Acad. Sci. USA 89, 5577-5581. 2328. Xu, M. Q., Southworth, M. W., Mersha, F. B., Hornstra, L. J., and Perler, F. B. (1993). in vitro protein splicing of purified precursor and the identification of a branched intermediate. Cell 75, 1371-1377. 2329. Derbyshire, V., Wood, D. W., Wu, W., Dansereau, J. T., Dalgaard, J. Z., and Belfort, M. (1997). Genetic definition of a protein-splicing domain: functional mini-inteins support structure predictions and a model for intein evolution. Proc. Natl. Acad. Sci. USA 94, 11466-11471.



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CATALYTIC RNA

5.26.13 Summary Self-splicing is a property of two groups of introns, which are widely dispersed in lower eukaryotes, prokaryotic systems, and mitochondria. The information necessary for the reaction resides in the intron sequence (although the reaction is actually assisted by proteins in vivo). For both group I and group II introns, the reaction requires formation of a specific secondary/tertiary structure involving short consensus sequences. Group I intron RNA creates a structure in which the substrate sequence is held by the IGS region of the intron, and other conserved sequences generate a guanine nucleotide binding site. It occurs by a transesterification involving a guanosine residue as cofactor. No input of energy is required. The guanosine breaks the bond at the 5 ′ exon-intron junction and becomes linked to the intron; the hydroxyl at the free end of the exon then attacks the 3 ′ exon-intron junction. The intron cyclizes and loses the guanosine and the terminal 15 bases. A series of related reactions can be catalyzed via attacks by the terminal G-OH residue of the intron on internal phosphodiester bonds. By providing appropriate substrates, it has been possible to engineer ribozymes that perform a variety of catalytic reactions, including nucleotidyl transferase activities. Some group I and some group II mitochondrial introns have open reading frames. The proteins coded by group I introns are endonucleases that make double-stranded cleavages in target sites in DNA; the cleavage initiates a gene conversion process in which the sequence of the intron itself is copied into the target site. The proteins coded by group II introns include an endonuclease activity that initiates the transposition process, and a reverse transcriptase that enables an RNA copy of the intron to be copied into the target site. These types of introns probably originated by insertion events. The proteins coded by both groups of introns may include maturase activities that assist splicing of the intron by stabilizing the formation of the secondary/tertiary structure of the active site. Catalytic reactions are undertaken by the RNA component of the RNAase P ribonucleoprotein. Virusoid RNAs can undertake self-cleavage at a "hammerhead" structure. Hammerhead structures can form between a substrate RNA and a ribozyme RNA, allowing cleavage to be directed at highly specific sequences. These reactions support the view that RNA can form specific active sites that have catalytic activity. RNA editing changes the sequence of an RNA after or during its transcription. The changes are required to create a meaningful coding sequence. Substitutions of individual bases occur in mammalian systems; they take the form of deaminations in which C is converted to U, or A is converted to I. A catalytic subunit related to cytidine or adenosine deaminase functions as part of a larger complex that has specificity for a particular target sequence. Additions and deletions (most usually of uridine) occur in trypanosome mitochondria and in paramyxoviruses. Extensive editing reactions occur in trypanosomes in which as many as half of the bases in an mRNA are derived from editing. The editing reaction uses a template consisting of a guide RNA that is complementary to the mRNA sequence. The reaction is catalyzed by an enzyme complex that includes an Summary | SECTION 5.26.13 © 2004. Virtual Text / www.ergito.com

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endonuclease, terminal uridyltransferase, and RNA ligase, using free nucleotides as the source for additions, or releasing cleaved nucleotides following deletion. Protein splicing is an autocatalytic reaction that occurs by bond transfer reactions and input of energy is not required. The intein catalyzes its own splicing out of the flanking exteins. Many inteins have a homing endonuclease activity that is independent of the protein splicing activity. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.5.26.13


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PROTEIN TRAFFICKING

6.27.1 Introduction Key Terms A sorting signal (targeting signal) is the part of a protein that allows it to be incorporated into a nascent transport vesicle. The vesicle moves the protein from one compartment to another. Sorting signals consist of either a short sequence of amino acids or a covalent modification.

A great variety of molecules move out of and into the cell. At one extreme of the size range, proteins may be secreted from the cell into the extracellular fluid or may be internalized from the cell surface. At the other extreme, ions such as K+, Na+, and Ca2+ may be pumped out of or into the cell. In this chapter, we are concerned with the processes by which proteins are physically transported through membranous systems to the plasma membrane or other organelles, or from the cell surface to organelles within the cell. In Molecular Biology 6.28 Signal transduction we discuss the pathways by which an interaction at the surface can trigger internal pathways. Proteins enter the pathway that leads to secretion by co-translational transfer to the membranes of the endoplasmic reticulum (for introduction see Molecular Biology Supplement 32.6 ER and Golgi). They are then transferred to the Golgi apparatus, where they are sorted according to their final intended destination. Figure 27.1 summarizes the routes by which proteins are carried forward or diverted to other organelles. Their destinations are determined by specific sorting signals, which take the form of short sequences of amino acids or covalent modifications that are made to the protein.


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Figure 27.1 Proteins that enter the endoplasmic reticulum are transported to the Golgi and towards the plasma membrane. Specific signals cause proteins to be returned from the Golgi to the ER, to be retained in the Golgi, to be retained in the plasma membrane, or to be transported to endosomes and lysosomes. Proteins may be transported between the plasma membrane and endosomes.

The transport machinery consists of small membranous vesicles. A soluble protein is carried within the lumen of a vesicle, and an integral membrane protein is carried within its membrane. Figure 27.2 illustrates the nature of the budding and fusion events by which the vesicles move between adjacent compartments. A vesicle buds off from a donor surface and then fuses with a target surface. Its proteins are released into the lumen or into the membrane of the target compartment, depending on their nature, and must be loaded into new vesicles for transport to the next compartment. The series of events is repeated at each transition between membrane surfaces, for example, during passage from the ER to the Golgi, or between cisternae of the Golgi stacks.


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Figure 27.2 Vesicles are released when they bud from a donor compartment and are surrounded by coat proteins (left). During fusion, the coated vesicle binds to a target compartment, is uncoated, and fuses with the target membrane, releasing its contents (right).

Once a protein enters a membranous environment, it remains in the membrane until it reaches its final destination. A membrane protein that enters the endoplasmic reticulum is inserted into the membrane with the appropriate orientation (N-terminal lumenal, C-terminal cytosolic for group I proteins, the reverse for group II proteins). The orientation is retained as it moves through the system. The process starts in the same way irrespective of whether the protein is destined to reside in the Golgi, lysosome, or plasma membrane. In each case, it is transported in membrane vesicles along the secretory pathway to the appropriate destination, where some structural feature of the protein is recognized and it is permanently secured (or secreted from the cell). Two important changes occur to a protein in the endoplasmic reticulum: it becomes folded into its proper conformation; and it is modified by glycosylation. A protein is translocated into the ER in unfolded form. Folding occurs as the protein enters the lumen; probably a series of domains each folds independently as the protein passes through the membrane. Folding of a 50 kD protein is complete in 30 hours. It provides the cell with the means of taking up iron. • Receptor and ligand both are degraded. The EGF receptor binds its ligand as a requirement for internalization. Although EGF and its receptor appear to dissociate at low pH, they are both carried on to the lysosome, where they may be degraded, as indicated in Figure 27.33. We do not know how and whether these events are related to the ability of EGF to change the phenotype of a target cell via binding to the receptor. • Receptor and ligand are transported elsewhere. The route illustrated in Figure 27.34 is available in certain polarized cells. A receptor-ligand combination is taken up at one cell surface, transported to the endosome, and then released for transport to the far surface of the cell. This is called transcytosis. By this means, receptors can transport immunoglobulins across epithelial cells.

Figure 27.31 LDL receptor transports apo-B (and apo-E) into endosomes, where receptor and ligand separate. The receptor recycles to the surface, apo-B (or apo-E) continues to the lysosome and is degraded, and cholesterol is released.

Receptors recycle via endocytosis | SECTION 6.27.15 © 2004. Virtual Text / www.ergito.com

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Figure 27.32 Transferrin receptor bound to transferrin carrying iron releases the iron in the endosome; the receptor now bound to apo-transferrin (lacking iron) recycles to the surface, where receptor and ligand dissociate.


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Figure 27.33 EGF receptor carries EGF to the lysosome where both the receptor and ligand are degraded.


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Figure 27.34 Ig receptor transports immunoglobulin across the cell from one surface to the other.

Rapid recycling in general occurs for receptors that bring ligands into the cell, not for those that trigger pathways of signal transduction. Receptors involved in signaling changes from the surface are usually degraded if they are endocytosed.


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Reviews 278. Goldstein, J. L., Brown, M. S., Anderson, R. G., Russell, D. W., and Schneider, W. J. (1985). Receptor-mediated endocytosis: concepts emerging from the LDL receptor system. Annu. Rev. Cell Biol. 1, 1-39.



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PROTEIN TRAFFICKING

6.27.16 Internalization signals are short and contain tyrosine Key Concepts

• Signals for internalization are short (~4 amino acids), are located in the

cytoplasmic tail near the plasma membrane, and usually contain a tyrosine residue that is exposed by the protein conformation.

What features of protein structure are required for endocytosis? Mutations that prevent internalization can be used to identify the relevant sequences in the receptors. In fact, the characterization of an internalization defect provided evidence that entry into coated pits is needed for receptor-mediated endocytosis of LDL. In cells from human patients with such defects in the LDL receptor, the receptor gathers in small clusters over the plasma membrane, and cannot enter coated pits in the manner observed for wild-type cells. The mutations responsible for this type of defect all affect the cytoplasmic domain of the receptor, which functions independently in sponsoring endocytosis. A recombinant protein whose extracellular domain is derived from influenza virus hemagglutinin, which is not usually endocytosed, can be internalized if it is provided with a cytoplasmic domain from an endocytosed receptor. We do not yet have a clear view of all of the sorting signals that mediate endocytosis, but two features are common. The relevant region of the protein comprises a relatively short part of the cytoplasmic tail close to the plasma membrane. And the presence of a tyrosine residue in this region often is necessary. The most common motif is YXX φ (where φ is a hydrophobic amino acid). Removal of the tyrosine prevents internalization; conversely, substituting tyrosine in the relevant region of a protein that is not endocytosed (such as influenza virus hemagglutinin) allows it to be internalized. Tyrosine is also found in another signal for internalization: NPXY (Asn-Pro-X-Tyr). The essential tyrosine can be replaced by other aromatic amino acids, but not by other types of amino acids. The NPXY motif is found in several group I proteins that are internalized, and is usually located close to the plasma membrane. (It was first identified in the LDL receptor.) It could be a general signal for endocytosis. In proteins that are internalized in response to ligand binding, the internalization signal may be generated by a change in conformation as a result of the binding. Do internalized receptors interact directly with proteins on the vesicles that transport them? Clathrin forms an outer polyhedral layer on clathrin-coated vesicles, but other proteins form an inner layer. The adaptins recognize the appropriate sequences in the cytoplasmic domains of receptors that are to be internalized (see Figure 27.12). Figure 27.35 illustrates a model in which, as a coated pit forms, the adaptins bind to the receptor cytoplasmic domain, immobilizing the receptor in the pit. As a result, Internalization signals are short and contain tyrosine | SECTION 6.27.16 © 2004. Virtual Text / www.ergito.com

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the receptor is retained by the coated vesicle when it pinches off from the plasma membrane.

Figure 27.35 The cytoplasmic domain of an internalized receptor interacts with proteins of the inner layer of a coated pit. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.27.16

Internalization signals are short and contain tyrosine | SECTION 6.27.16 © 2004. Virtual Text / www.ergito.com

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PROTEIN TRAFFICKING

6.27.17 Summary Proteins that reside within the reticuloendothelial system or that are secreted from the plasma membrane enter the ER by cotranslational transfer directly from the ribosome. They are transported through the Golgi in the anterograde (forward) direction. Specific signals may cause them to be retained in the ER or a Golgi stack, or directed to other organelles such as endosomes. The default pathway is to be transported to the plasma membrane. Retrograde transport is less well characterized, but proteins that reside in the ER are retrieved from the Golgi by virtue of specific signals; an example is the C-terminal KDEL. Proteins are transported between membranous surfaces as cargoes in membrane-bound coated vesicles. The vesicles form by budding from donor membranes; they unload their cargos by fusing with target membranes. The protein coats are added when the vesicles are formed and must be removed before they can fuse with target membranes. Anterograde transport does not result in any net flow of membrane from the ER to the Golgi and/or plasma membrane, so membrane moving with anterograde transport must be returned to the ER by a retrograde mechanism. Modification of proteins by addition of a preformed oligosaccharide starts in the endoplasmic reticulum. High mannose oligosaccharides are trimmed. Complex oligosaccharides are generated by further modifications that are made during transport through the Golgi, determined by the order in which the protein encounters the enzymes localized in the various Golgi stacks. Proteins are sorted for different destinations in the trans Golgi. The signal for sorting to lysosomes is the presence of mannose-6-phosphate. Different types of vesicles are responsible for transport to and from different membrane systems. The vesicles are distinguished by the nature of their protein coats. COP-I-coated vesicles are responsible for retrograde transport from the Golgi to the ER. COP-I vesicles are coated with coatomer. One of the proteins of coatomer, β-COP, is related to the β-adaptin of clathrin-coated vesicles, suggesting the possibility of a common type of structure between COP-I-coated and clathrin-coated vesicles. COP-II vesicles undertake forward movement from the ER to Golgi. Vesicles that transport proteins along the Golgi stacks have not yet been identified. Vesicles responsible for constitutive (bulk) movement from the Golgi to the plasma membrane also have not been identified. An alternative model for anterograde transport proposes that cis-Golgi cisternae actually become trans-Golgi cisternae, so that there is a continuous process of cisternal maturation from the cis to the trans face. In the pathway for regulated secretion of proteins, proteins are sorted into clathrin-coated vesicles at the Golgi trans face. Some vesicles may fuse into (larger) Summary | SECTION 6.27.17 © 2004. Virtual Text / www.ergito.com

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secretory granules. Vesicles also move to endosomes, which control trafficking to the cell surface. Secretory vesicles are stimulated to unload their cargos at the plasma membrane by extracellular signals. Similar vesicles are used for endocytosis, the pathway by which proteins are internalized from the cell surface. The predominant protein in the outer coat of these vesicles is clathrin. The inner coat contains an adaptor complex, consisting of adaptin subunits, which bind to clathrin and to cargo proteins. There are (at least) three types of adaptor complex, with different specificities. Budding and fusion of all types of vesicles is controlled by a small GTP-binding protein. This is ARF for clathrin and COP-I-coated vesicles, and Sar1P for COP-II-coated vesicles. When activated by GTP, ARF/Sar1p inserts into the membrane and causes coat proteins to assemble. This leads ultimately to budding. Further proteins, such as dynamin, may be required to "pinch off" the budding vesicle from the donor membrane. When ARF/Sar1p is inactivated because GTP is hydrolyzed to GDP, it withdraws from the membrane and the coat proteins either disassemble spontaneously (COP-coated vesicles) or are caused to do so by other proteins (clathrin-coated vesicles). Vesicles initially recognize appropriate target membranes by a tethering reaction in which a tethering complex recognizes a Rab protein on the vesicle and brings the vesicle close to the membrane. Rabs are prenylated monomeric GTP-binding proteins. The fusion reaction is triggered when a v-SNARE on the vesicle pairs specifically with a t-SNARE on the target membrane. Pairing occurs by a coiled-coil interaction in which the SNARE complex lies parallel to the membrane surface. This causes the inner leaflets of the membranes to fuse to form a hemifusion complex; this is followed by fusion of the outer leaflets . The 20S fusion complex includes the soluble ATPase NSF and SNAP, and uses hydrolysis of ATP to release the SNAREs after pairing, which allows them to recycle. Receptors may be internalized either continuously or as the result of binding to an extracellular ligand. Receptor-mediated endocytosis initiates when the receptor moves laterally into a coated pit. The cytoplasmic domain of the receptor has a signal that is recognized by proteins that are presumed to be associated with the coated pit. An exposed tyrosine located near the transmembrane domain is a common signal; it may be part of the sequence NPXY. When a receptor has entered a pit, the clathrin coat pinches off a vesicle, which then migrates to the early endosome. The acid environment of the endosome causes some receptors to release their ligands; the ligand are carried to lysosomes, where they are degraded, and the receptors are recycled back to the plasma membrane by means of coated vesicles. A ligand that does not dissociate may recycle with its receptor. In some cases, the receptor-ligand complex is carried to the lysosome and degraded. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.27.17


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SIGNAL TRANSDUCTION

6.28.1 Introduction Key Terms A ligand is an extracellular molecule that binds to the receptor on the plasma membrane of a cell, thereby effecting a change in the cytoplasm. A receptor is a transmembrane protein, located in the plasma membrane, that binds a ligand in a domain on the extracellular side, and as a result has a change in activity of the cytoplasmic domain. (The same term is sometimes used also for the steroid receptors, which are transcription factors that are activated by binding ligands that are steroids or other small molecules.) A transporter is a type of receptor that moves small molecules across the plasma membrane. It binds the molecules on its extracellular surface, and releases them into the cytoplasm. Internalization is a process through which a ligand-receptor complex is brought into the cell. Endocytosis is the process whereby cells internalize small molecules and particles from their surroundings. There are several forms of endocytosis, all of which involve the formation of a membranous vesicle from the plasma membrane. Signal transduction describes the process by which a receptor interacts with a ligand at the surface of the cell and then transmits a signal to trigger a pathway within the cell. A second messenger is a small molecule that is generated when a signal transduction pathway is activated. The classic second messenger is cyclic AMP, which is generated when adenylate cyclase is activated by a G protein (when the G protein itself was activated by a transmembrane receptor). The ability of a species of kinase to phosphorylate itself is referred to as autophosphorylation. Autophosphorylation does not necessarily occur on the same polypeptide chain as the catalytic site; for example, in a dimer, each subunit may phosphorylate the other. G proteins are guanine nucleotide-binding proteins. Trimeric G proteins are associated with the plasma membrane. When bound by GDP the trimer remains intact and is inert. When the GDP is replaced by GTP, the α subunit is released from the β γ dimer. Either the α monomer or the β γ dimer then activates or represses a target protein. Monomeric G proteins are cytosolic and work on the same principle that the form bound to GDP is inactive, but the form bound to GTP is active.

The plasma membrane separates a cell from the surrounding environment. It is permeable only to small lipid-soluble molecules, such as the steroid hormones, which can diffuse through it into the cytoplasm. It is impermeable to water-soluble material, including ions, small inorganic molecules, and polypeptides or proteins. The response of the cell to hydrophilic material in the environment depends on Introduction | SECTION 6.28.1 © 2004. Virtual Text / www.ergito.com

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interactions that occur on the extracellular side of the plasma membrane. The hydrophilic material binds specifically to the extracellular domain of a protein embedded in the membrane. The extracellular molecule typically is called the ligand, and the plasma membrane protein that binds it is called the receptor. Two fundamental types of response to an external stimulatory molecule that cannot penetrate the membrane are reviewed in Figure 28.1:

Figure 28.1 Overview: information may be transmitted from the exterior to the interior of the cell by movement of a ligand or by signal transduction.

• Material – molecular or macromolecular – is physically transmitted from the outside of the membrane to the inside by transport through a proteinaceous channel in the lipid bilayer. • A signal is transmitted by means of a change in the properties of a membrane protein that activates its cytosolic domain. The physical transfer of material extends from ions to small molecules such as sugars, and to macromolecules such as proteins. Three major transport routes controlled by plasma membrane proteins are reviewed in Figure 28.2:


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Figure 28.2 Three means for transferring material of various sizes into the cell are provided by ion channels, receptor-mediated ligand transport, and receptor internalization.

• Channels control the passage of ions: different channels exist for potassium, sodium, and calcium ions. By opening and closing in response to appropriate signals, the channels establish ionic levels within the cell (a feature of particular significance for cells of the neural network). • One means to import small molecules is for a receptor to transport the molecule from one side of the membrane to the other. Transporters are responsible for the import of small molecules (such as sugars) across the membrane. The target molecule binds to the receptor on the extracellular side, but then is released on the cytoplasmic side. • Ligand-binding may trigger the process of internalization, in which the receptor-ligand combination is brought into the cell by the process of Introduction | SECTION 6.28.1 © 2004. Virtual Text / www.ergito.com

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endocytosis. In due course, the receptor and ligand are separated; the receptor may be returned to the surface for another cycle, or may be degraded. As described in Molecular Biology 6.27.15 Receptors recycle via endocytosis, endocytosis involves the passage of membrane proteins from one surface to another via coated vesicles. The transmission of a signal involves the interaction of an extracellular ligand with a transmembrane protein that has domains on both sides of the membrane. Binding of ligand converts the receptor from an inactive to an active form. The basic principle of this interaction is that ligand binding on the extracellular side activates the receptor domain on the cytoplasmic side. The process is called signal transduction, because a signal has in effect been transduced across the membrane. The amplitude of the cytosolic signal is much greater than the original extracellular signal (the ligand), so signal transduction amplifies the original signal. Figure 28.3 illustrates the principle of signal transduction. A receptor typically is activated by being caused to dimerize when a ligand binds to its extracellular domain. Its cytosolic domain then interacts with a protein at the plasma membrane. This interaction most often takes one of two forms. It increases the quantity of a small molecule (called a second messenger) inside the cell. Or it directly activates a protein whose role is to activate other proteins; one common type of protein that is activated early in such pathways is a monomeric GTP-binding protein.


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Figure 28.3 A signal transduction pathway carries a signal from the cell surface into the cytoplasm and (sometimes) into the nucleus.

However a signal transduction pathway is initiated, a common means of propagating the pathway through the cytosol is to activate a protein kinase, which activates a series of other protein kinases. Ultimately the signal leads to the activation of effectors, which trigger changes in the cell. Some of the effectors act in the cytosol (for example, to affect the cytoskeleton), but some carry the signal into the nucleus, where the ultimate target is the activation of transcription factors that cause new patterns of gene expression. The major signal transduction pathways that we discuss in this chapter are extensively conserved throughout animal evolution, and can be found in most or all animal cells. They are essentially absent from plants, which have evolved different Introduction | SECTION 6.28.1 © 2004. Virtual Text / www.ergito.com

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pathways for signal transduction. Two major types of signal transduction are reviewed in Figure 28.4:

Figure 28.4 A signal may be transduced by activating the kinase activity of the cytoplasmic domain of a transmembrane receptor or by dissociating a G protein into subunits that act on target proteins in the membrane.

• The receptor has a protein kinase activity in its cytosolic domain. The activity of the kinase is activated when ligand binds to the extracellular domain. The kinase phosphorylates its own cytoplasmic domain; this autophosphorylation enables the receptor to associate with and activate a target protein, which in turn acts upon new substrates within the cell. The most common kinase receptors are tyrosine kinases, but there are also some serine/threonine kinase receptors. Such pathways most often lead to activation of a GTP-binding protein that activates a cascade of cytosolic kinases. • The receptor may interact with a trimeric G protein that is associated with the cytosolic face of the membrane (for introduction see Molecular Biology Supplement 32.10 G proteins). G proteins are named for their ability to bind guanine nucleotides. The inactive form of the G protein is a trimer bound to GDP. Receptor activation causes the GDP to be replaced with GTP; as a result, the G protein dissociates into a single subunit carrying GTP and a dimer of the Introduction | SECTION 6.28.1 © 2004. Virtual Text / www.ergito.com

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two other subunits. Either the monomer or the dimer then acts upon a target protein, often also associated with the membrane, which in turn reacts with a target(s) in the cytoplasm. This chain of events often stimulates the production of second messengers, the classic example being the production of cyclic AMP. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.28.1


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SIGNAL TRANSDUCTION

6.28.2 Carriers and channels form water soluble paths through the membrane Key Terms A concentration gradient is a change in the concentration of a molecule or ion from one point to another. The gradient might be gradual (as in a solution that is not homogenous) or abrupt (created by a membrane). An electrical gradient is a change in the amount of charge from one point to another. A change in the concentration of ions from one point to another produces an electrochemical gradient. The term indicates that there is a change in the concentration of both electrical charge and of a chemical species. Passive transport describes movement of molecules along their electrochemical gradient; no energy is required. Active transport is an energy-consuming process that moves molecules against an electrochemical gradient. Energy for the movement is provided by hydrolysis of ATP. A carrier protein moves directly a solute from one side of the plasma membrane to the other. In the process, the protein undergoes a conformational change. A uniporter is a type of carrier protein that moves only one type of solute across the plasma membrane. A symporter is a type of carrier protein that moves two different solutes across the plasma membrane in the same direction. The two solutes can be transported simultaneously or sequentially. An antiporter is a type of carrier protein that simultaneously moves two different types of solutes in opposite directions across the plasma membrane. An ion channel is a transmembrane protein which selectively allows the passage of one type of ion across the membrane. Ion channels are usually oligomers with a central aqueous pore through which the ion passes. A channel which only allows passage of its substrate under certain conditions is referred to as "gated". Gated channels can exist in at least two conformations, one of which is open and the other closed. Ligand-gated channels open or close in response to the binding of a specific molecule. Voltage-gated channels are open or closed depending on the voltage across the membrane. A second messenger gated channel is an ion channel whose activity is controlled by small signaling molecules inside the cell. Key Concepts

• The electric gradient across the plasma membrane (inside is more negative) favors Carriers and channels form water soluble paths through the membrane | SECTION 6.28.2 © 2004. Virtual Text / www.ergito.com

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entry of cations and opposes entry of anions. gradient depends on the ion, typically with low intracellular • The concentration + – + levels of Na and Cl and high levels of K .

• If the overall electrochemical gradient is favorable an ion can enter passively, otherwise it needs to be actively transported against the gradient.

• Carrier proteins that transport solutes across the membrane can be uniporters (one

solute), symporters (two solutes) or antiporters (two solutes in opposite directions).

• Ion channels are water-soluble pores in the membrane that may be gated (controlled) by voltage or by ligands.

The impermeability of the plasma membrane to water-soluble compounds enables different aqueous conditions to be maintained on either side. The ionic environment of the cytosol is quite different from the extracellular ionic milieu. Within the cytoplasm, different organelles offer different ionic environments. A striking example is the maintenance of an acid pH in endosomes and lysosomes, with immediate implications for the functions of the proteins that enter them (see Molecular Biology 6.27.15 Receptors recycle via endocytosis). Another example is the maintenance of a store of Ca2+ in the endoplasmic reticulum. (The nucleus is an exception to the rule that conditions in membrane-bounded organelles usually differ from the cytosol. The nucleoplasm essentially is subjected to the same conditions as the cytosol. This happens because the nuclear pores form relatively large openings in the nuclear envelope, through which ions and other small molecules can diffuse freely.) A notable feature of the cytosolic environment is that there are more free cations (positively charged) (~150 mM) than anions (~10 mM). The reason is that many cellular constituents are negatively charged – for example, nucleic acids have multiple negative charges for every phosphate group in the phosphodiester backbone. The superfluity of cations therefore establishes electrical neutrality by balancing these fixed charges. The intracellular concentrations of Na+ and Cl– are low (~10 mM) while those outside the cell are high (>100 mM); and the situation for K+ is reversed. This creates a concentration gradient across the membrane for each ion. The plasma membrane is electrically charged (due to the different phospholipid compositions of the inner and outer leaflets). There is an electrical gradient in which the inside is negative compared to the outside. This voltage difference favors the entry of cations and opposes the entry of anions. Together the concentration gradient and electrical gradient constitute the electrochemical gradient, which is characteristic for each solute. A solute whose gradient is favorable can enter the cell when a channel opens; the gradient is sufficient to drive passive transport of a solute such as Na+ or Cl– into the cell. But a solute that faces an unfavorable gradient requires active transport in which energy is used to pump it into the cell against the gradient. Carriers and channels form water soluble paths through the membrane | SECTION 6.28.2 © 2004. Virtual Text / www.ergito.com

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The passage of ions (and other small solutes) through the plasma membrane is mediated by resident transmembrane proteins. A common feature of these proteins is their large size and the presence of multiple membrane-spanning regions, features which together argue that they provide a relatively static feature of the membrane. Figure 28.5 illustrates two general means of transport across the membrane:

Figure 28.5 A carrier (porter) transports a solute into the cell by a conformational change that brings the solute-binding site from the exterior to the interior, while an ion channel is controlled by the opening of a gate (which might in principle be located on either side of the membrane).

• A carrier protein binds a solute on one side of the membrane and then experiences a conformational change that transports the solute to the other side of the membrane. By binding the solute on one side and releasing it on the other, the carrier in effect directly transports the solute across the membrane. Several types of carriers are distinguished by the number of solutes that they transport, and the directions in which they transport them. Carriers that transport a single solute across the membrane are called uniporters; carriers that simultaneously or sequentially transport two different solutes are called symporters; and carriers that transport one solute in one direction while transporting a different solute in the opposite direction are called antiporters. Carrier proteins may be Carriers and channels form water soluble paths through the membrane | SECTION 6.28.2 © 2004. Virtual Text / www.ergito.com

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used for passive transport or linked to an energy source to provide active transport. Energy for active transport is provided by hydrolysis of ATP, the classic example being the Na+-K+ pump that functions as an antiporter, pumping sodium out of the cell and potassium into it. Another source of energy is the electrochemical gradient itself; a symporter brings Na+ into the cell together with some other solute, using the favorable gradient of sodium to overcome the unfavorable gradient of the other solute. • An ion channel comprises a water-soluble pore in the membrane. Its activity is controlled by regulation of the opening and closing of the channel. When it is open, ions can diffuse passively, as driven by the electrochemical gradient. Ion channels allow only passive transport. The resting state of an ion channel is closed, and the gates that control channel activity usually open only briefly, in response to a specific signal. Ligand-gated channels are receptors that respond to binding of particular molecules, amongst which the neurotransmitters acetylcholine, glycine, GABA ( γ-amino-butyric acid), and glutamate are prominent examples. Voltage-gated channels respond to electric changes, again a prominent feature of the neural system. Second messenger gated channels provide yet another means for signal transduction, one interesting example comprising channels that respond to activation of G proteins. The structures of both carriers and channels present a paradox. They are transmembrane proteins that have multiple membrane-spanning domains, each consisting of a stretch of amino acids of sufficient hydrophobicity to reside in the lipid bilayer. Yet within these hydrophobic regions must be a highly selective, water-filled path that permits ions to travel through the membrane. One solution to this problem lies in the structure of the transmembrane regions. Instead of comprising unremittingly hydrophobic stretches like those of single membrane-pass proteins, they contain some polar amino acids. They are likely to be organized as illustrated in Figure 28.6 as amphipathic helices in which the hydrophobic face associates with the lipid bilayer, while the polar faces are aligned with one another to create the channel.

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Figure 28.6 A channel may be created by amphipathic helices, which present their hydrophobic faces to the lipid bilayer, while juxtaposing their charged faces away from the bilayer. In this example, the channel is lined with positive charges, which would encourage the passage of anions. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.28.2

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SIGNAL TRANSDUCTION

6.28.3 Ion channels are selective Key Terms Ion selectivity refers to the specificity of an ion channel for a particular type of ion. Key Concepts

• Channels typically consists of several protein subunits with the water-soluble pore at the axis of symmetry.

• Selectivity is determined by the properties of the pore. • The gate acts by a mechanism resembling a ball and chain.

The importance of the interior of the channel is indicated by the ion selectivity. Different channels permit the passages of different ions or groups of ions. The channels are extremely narrow, so ions must be stripped of their associated water molecules in order to pass through. The channel possesses a "filter" at the entrance to the pore that has specificity for its particular ion, presumably based upon its geometry and electrostatic charge. The structures of particular ion channels are beginning to reveal their general features. A common feature is that the constituent proteins are large and have several membrane-spanning regions. A channel probably consists of a "ring" of 4, 5, or 6 subunits, organized in a symmetrical or quasi-symmetrical manner. The water-filled pore is found at the central axis of symmetry. The size of the pore generally increases with the number of subunits in the ring. The subunits are always related in structure, and sometimes are identical. They may consist of separate proteins or of related domains in a single large protein (for review see 296; 297). Voltage-gated sodium channels have a single type of subunit, a protein of 1820 amino acids with a repetitive structure that consists of 4 related domains. Each domain has several membrane-spanning regions. The four domains are probably arranged in the membrane in a pseudo-symmetrical structure. Two smaller subunits are associated with the large protein. Potassium channels have a smaller subunit, equivalent to one of the domains of the sodium channel; four identical subunits associate to create the channel. Six transmembrane domains are identified in the protein subunit by hydrophobicity analysis; they are numbered S1-S6. The S4 domain has an unusual structure for a transmembrane region: it is highly positively charged, with arginine or lysine residues present at every third or fourth position. The S4 motif is found in voltage-gated K+, Na+, and Ca2+ channels, so it seems likely that it is involved with a common property, thought to be channel opening. Some potassium channels have only the S5-S6 membrane-spanning domains, and they appear to be basically shorter versions of the protein. Ion channels are selective | SECTION 6.28.3 © 2004. Virtual Text / www.ergito.com

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Analysis of the shaker potassium channel of the fly has revealed some novel features, illustrated in Figure 28.7. The region that forms the pore has been identified by mutations that alter the response to toxins that inhibit channel function. It occupies the region between transmembrane domains S5 and S6, forming two membrane-spanning stretches that are not organized in the usual hydrophobic α-helical structure. The structure could be a rather extended β-hairpin. The state of the channel (open or closed) is controlled by the N-terminal end, which resembles a ball on a chain. The ball is in effect tethered to the channel by a chain, and plugs it on the cytoplasmic side. The length of the chain controls the rate with which the ball can plug the channel after it has been opened.

Figure 28.7 A potassium channel has a pore consisting of unusual transmembrane regions, with a gate whose mechanism of action resembles a ball and chain.

A major question about potassium channels is how their selectivity is maintained. K+ and Na+ ions are (positively charged) spheres of 1.33 Å and 0.95 Å, respectively. K+ ions are selected over Na+ ions by a margin of 104×, but at the same time, up to 108 ions per second can move through the pore, basically close to the diffusion limit. The salient features of the pore of a potassium channel, based on the crystal structure, are summarized in Figure 28.8, and shown as a cutaway model in Figure 28.9. The pore is ~45 Å long and consists of three regions. It starts inside the cell with a long internal pore, opens out into a central cavity of ~10 Å diameter, and then passes to the extracellular space with a narrow selectivity filter. The lining of the inner pore and central cavity is hydrophobic, providing a relatively inert surface to a diffusing potassium ion. The central cavity is aqueous, and may serve to lower the electrostatic barrier to crossing the membrane (which is at its maximum in the center). The selectivity filter has negative charges and is lined with the polypeptide backbone. When a K+ ion loses its hydrating water on entering the filter, the contacts that it made with the water will be replaced by contacts with the oxygens of the polypeptide carbonyl groups. The size of the pore may be set so that a smaller sodium ion would not be close enough to make these substitute contacts (822).

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Figure 28.8 The pore of a potassium channel consists of three regions.

Figure 28.9 A model of the potassium channel pore shows electrostatic charge (blue = positive, white = neutral, red = negative) and hydrophobicity (= yellow). Photograph kindly provided by Rod MacKinnon (see 822).


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Reviews 296. Unwin, N. (1989). The structure of ion channels in membranes of excitable cells. Neuron 3, 665-676. 297. Miller, C. (1989). Genetic manipulation of ion channels: a new approach to structure and mechanism. Neuron 2, 1195-1205.

References 822. Doyle, D. A., Morais Cabral, J., Pfuetzner, R. A., Kuo, A., Gulbis, J. M., Cohen, S. L., Chait, B. T., and MacKinnon, R. (1998). The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280, 69-77.



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SIGNAL TRANSDUCTION

6.28.4 Neurotransmitters control channel activity Key Concepts

• Neurotransmitter-gated receptors are ion channels that are controlled by neurotransmitters such as acetylcholine, glycine, or GABA.

• The nicotinic acetylcholine receptor is a+5-subunit ion channel that admits several cations but is largely used to control Na uptake by the cell.

Neurotransmitter-gated receptors form a superfamily of related proteins in the 5-member channel class. The nicotinic acetylcholine receptor has been characterized in the most detail, and is a pentamer with the structure α2 β δ γ. As illustrated in Figure 28.10, the bulk of the 5 subunits projects above the plasma membrane into the extracellular space. The openings to the channel narrow from a diameter of ~25 Å until reaching the pore itself. The entrance on the extracellular side is very deep, ~60 Å; the distance on the cellular side is shorter, 20 Å. The pore extends through the 30 Å of the lipid bilayer and is only ~7 Å in diameter.

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Figure 28.10 The acetylcholine receptor consists of a ring of 5 subunits, protruding into the extracellular space, and narrowing to form an ion channel through the membrane.

Ligand binding occurs on the α subunits. Both α subunits must bind an acetylcholine for the gate to open. Where is the gate? Since the channel is really narrow only in the region within the lipid bilayer, the gate seems likely to be located well within the receptor. Structural changes that occur upon opening seem greatest just by the cytoplasmic side of the lipid bilayer, so it is possible that the gate is located at the level of the phospholipid heads on the cytoplasmic boundary. So the acetylcholine receptor, like many other receptors, must transmit information about ligand binding internally, from the extracellular acetylcholine binding site to the near-cytoplasmic gate. How does the gate function? It might consist of an electrostatic repulsion, in which positive groups are extruded into the channel to prevent passage of cations. Or it may take the form of a physical impediment to passage, in which a conformational change brings bulky groups to block the pore. Neurotransmitters control channel activity | SECTION 6.28.4 © 2004. Virtual Text / www.ergito.com

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Ion selectivity may be determined by the walls of the wide entry passage. The walls lining the entrances to the pore have negatively charged groups; each subunit carries ~10 negative charges in its extracellular region. These charge clusters could modify the ionic environment at the entrance to the channel, concentrating the desired ions and diluting ions that are selected against. The structure of the acetylcholine receptor allows passage of Na+, K+, or Ca2+ ions, but because of the prevailing gradients, its main use in practice is to allow the entry of Na+ into the cell. The acetylcholine receptor is an example of a superfamily of receptors gated by neurotransmitters. All appear to have the same general organization, consisting of 5 subunits whose structures are related to one another. All the subunits are about the same size (~50 kD), and each is probably organized in the membrane as a bundle of 4 helices (each helix containing a transmembrane domain). In each case, one of the four transmembrane domains (called M2) has an amphipathic structure and seems likely to be involved in lining the walls of the pore itself. The presence of serine and threonine residues, and some paired acid-basic residues, may assist ion passage. The sequences of subunits of the glycine and GABA receptors are related to the acetylcholine receptor subunits. Some changes in the sequences seem likely to reflect the ion selectivity. So the glycine and GABA receptors have positively charged groups in the entrance walls, consistent with their transport of anions such as Cl–. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.28.4

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SIGNAL TRANSDUCTION

6.28.5 G proteins may activate or inhibit target proteins Key Terms A receptor is a transmembrane protein, located in the plasma membrane, that binds a ligand in a domain on the extracellular side, and as a result has a change in activity of the cytoplasmic domain. (The same term is sometimes used also for the steroid receptors, which are transcription factors that are activated by binding ligands that are steroids or other small molecules.) G proteins are guanine nucleotide-binding proteins. Trimeric G proteins are associated with the plasma membrane. When bound by GDP the trimer remains intact and is inert. When the GDP is replaced by GTP, the α subunit is released from the β γ dimer. Either the α monomer or the β γ dimer then activates or represses a target protein. Monomeric G proteins are cytosolic and work on the same principle that the form bound to GDP is inactive, but the form bound to GTP is active. An effector is the target protein for the activated G protein. A second messenger is a small molecule that is generated when a signal transduction pathway is activated. The classic second messenger is cyclic AMP, which is generated when adenylate cyclase is activated by a G protein (when the G protein itself was activated by a transmembrane receptor). A serpentine receptor has 7 transmembrane segments. Typically it activates a trimeric G protein. Key Concepts

• Ligand binding to a serpentine membrane receptor causes it to activate a G protein. • The G protein is a trimer bound to GDP in its inactive state. • The mechanism of activation is that the receptor causes the GDP bound by the G-protein to be replaced with GTP.

G proteins transduce signals from a variety of receptors to a variety of targets. The components of the general pathway can be described as: • The receptor is a resident membrane protein that is activated by an extracellular signal. • A G protein is converted into active form when an interaction with the activated receptor causes its bound GDP to be replaced with GTP. • An effector is the target protein that is activated (or – less often – inhibited) by the G protein; sometimes it is another membrane-associated protein. G proteins may activate or inhibit target proteins | SECTION 6.28.5 © 2004. Virtual Text / www.ergito.com

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• Second messengers are small molecules that are released as the result of activation of (certain types) of effectors. Another terminology that is sometimes used to describe the relationship of the components of the transduction pathway is to say that the receptor is upstream of the G protein, while the effector is downstream. The effectors linked to different types of G proteins are summarized in Figure 28.11. The important point is that there is a large variety of G proteins, activated by a wide variety of receptors. The activation of an individual G protein may cause it to stimulate or to inhibit a particular effector; and some G proteins act upon multiple effectors (causing the activation in turn of multiple pathways). Two of the classic G proteins are Gs, which stimulates adenylate cyclase (increasing the level of cAMP), and Gt, which stimulates cGMP phosphodiesterase (decreasing the level of cGMP). The cyclic nucleotides are a major class of second messengers; another important group consists of small lipid molecules, such as inositol phosphate or DAG (diacylglycerol; for review see 306).

Figure 28.11 Classes of G proteins are distinguished by their effectors and are activated by a variety of transmembrane receptors.

Although the receptors that couple to G proteins respond to a wide variety of ligands, they have a common type of structure and mode of binding the ligand. They are serpentine receptors, with 7 transmembrane regions, and function as monomers. The greatest conservation of sequence is found in hydrophobic transmembrane regions, which in fact are used to classify the serpentine receptors into individual families (for review see 304; 3438). The binding sites for small hydrophobic ligands lie in the transmembrane domains, so that the ligand becomes bound in the plane of the membrane. The smallest ligands, such as biogenic amines, may be bound by a single transmembrane segment. Larger ligands, such as extended peptides, may have more extensive binding sites in which extracellular domains provide additional points of contact. Large peptide hormones may be bound mainly by the extracellular domains. When the ligand binds to its site, it triggers a conformational change in the receptor that causes it to interact with a G protein. A well characterized (although not typical) G proteins may activate or inhibit target proteins | SECTION 6.28.5 © 2004. Virtual Text / www.ergito.com

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case is that of rhodopsin, which contains a retinal chromophore covalently linked to an amino acid in a transmembrane domain. Exposure to light converts the retinal from the 11-cis to the all trans conformation, which triggers a conformational change in rhodopsin that causes its cytoplasmic domain to associate with the Gt protein (transducin).

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Reviews 304. Strader, D. (1994). Structure and function of G protein-coupled receptors. Annu. Rev. Biochem. 63, 101-132. 306. Divecha, N. and Irvine, R. F. (1995). Phospholipid signaling. Cell 80, 269-278. 3438. Pierce, K. L., Premont, R. T., and Lefkowitz, R. J. (2002). Seven-transmembrane receptors. Nat. Rev. Mol. Cell Biol. 3, 639-650.


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SIGNAL TRANSDUCTION

6.28.6 G proteins function by dissociation of the trimer Key Concepts

• When GDP is replaced by GTP, a trimeric G protein dissociates into an α-GTP subunit and a β γ dimer.

• It is most often the α subunit that activates the next component (the effector) in the pathway.

• Less often the β γ activates the effector.

G proteins are trimers whose function depends on the ability to dissociate into an α monomer and a β γ dimer. The dissociation is triggered by the activation of an associated receptor. In its inactive state, the α subunit of the G protein is bound to GDP. Figure 28.12 shows that the activated receptor causes the GDP to be replaced by GTP. This causes the G protein to dissociate into a free α-GTP subunit and a free β γ dimer. (The basic interactions of G proteins are reviewed in Molecular Biology Supplement 32.10 G proteins). They are found in all classes of eukaryotes.

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Figure 28.12 Activation of Gs causes the α subunit to activate adenylate cyclase.

The interaction between receptor and G protein is catalytic. After a G protein has dissociated from an activated receptor, the receptor binds another (inactive) trimer, and the cycle starts again. So one ligand-receptor complex can activate many G protein molecules in a short period, amplifying the original signal. The most common action for the next stage in the pathway calls for the activated α subunit to interact with the effector. In the case of Gs, the αs subunit activates adenylate cyclase; in the case of Gt, the αt subunit activates cGMP phosphodiesterase. In other cases, however, it is the β γ dimer that interacts with the effector protein. In some cases, both the α subunit and the β γ dimer interact with effectors (for review see 295; 301; 312). Consistent with the idea that it is more often the α subunits that interact with effectors, there are more varieties of α subunits (16 known in mammals) than of β (5) or γ subunits (11). However, irrespective of whether the α or β γ subunits carry the signal, the common feature in all of these reactions is that a G protein usually acts upon an effector enzyme that in turn changes the concentration of some small molecule(s) in the cell. (There are some other pathways in which G proteins behave by activating a kinase.) G proteins function by dissociation of the trimer | SECTION 6.28.6 © 2004. Virtual Text / www.ergito.com

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In either the intact or dissociated state, G proteins are associated with the cytoplasmic face of the plasma membrane. But the individual subunits are quite hydrophilic, and none of them appears to have a transmembrane domain. The β γ dimer has an intrinsic affinity for the membrane because the γ subunit is prenylated. The αi and αo types of subunit are myristoylated, which explains their ability to remain associated with the membrane after release from the β γ dimer. The αs subunit is palmitoylated. Because several receptors can activate the same G proteins, and since (at least in some cases) a given G protein has more than one effector, we must ask how specificity is controlled. The most common model is to suppose that receptors, G proteins, and effectors all are free to diffuse in the plane of the membrane. In this case, the concentrations of the components of the pathway, and their relative affinities for one another, are the important parameters that regulate its activity. We might imagine that an activated α-GTP subunit scurries along the cytoplasmic face of the membrane from receptor to effector. But it is also possible that the membrane constrains the locations of the proteins, possibly in a way that restricts interactions to local areas. Such compartmentation could allow localized responses to occur (for review see 315).


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Reviews 295. Neer, E. J. and Clapham, D. E. (1988). Roles of G protein subunits in transmembrane signaling. Nature 333, 129-134. 301. Clapham, D. E. and Neer, E. J. (1993). New roles of G protein β γ-dimers in transmembrane signaling. Nature 365, 403-406. 312. Neer, E. J. (1995). Heterotrimeric G proteins: organizers of transmembrane signals. Cell 80, 249-257. 315. Sprang, S. R. (1997). G protein mechanisms: insights from structural analysis. Annu. Rev. Biochem. 66, 639-678.



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SIGNAL TRANSDUCTION

6.28.7 Protein kinases are important players in signal transduction Key Terms A protein kinase is a protein which transfers the terminal phosphate group from ATP onto another protein. A protein serine/threonine kinase phosphorylates cytosolic proteins on either their serine or threonine residues. A protein tyrosine kinase is a kinase enzyme whose target is a tyrosine amino acid in a protein. A dual specificity kinase is a protein kinase that can phosphorylate tyrosine or threonine or serine amino acids. RTK is an abbreviation for "receptor tyrosine kinase". These kinases are membrane-bound proteins with large cytoplasmic and extracellular domains. Specific binding of a ligand, such as a growth factor, to the extracellular domain causes the cytoplasmic domain to phosphorylate other proteins on tyrosine residues. Key Concepts

• Protein kinases fall into groups that phosphorylate Ser/Thr or Tyr on target proteins.

• Receptor protein kinases are most often protein tyrosine kinases. • Cytosolic protein kinases are most often protein Ser/Thr kinases.

There are many types of protein kinases involved in signal transduction. They all have the same basic catalytic activity: they add a phosphate group to an amino acid in a target protein. The phosphate is provided by hydrolyzing ATP to ADP. A protein kinase has an ATP-binding site and a catalytic center that can bind to the target amino acid (for review see 293; 294). The phosphorylation of the target protein changes its properties so that it in turn acts to carry the signal transduction pathway to the next stage. Protein kinases can be classified both by the types of amino acids that they phosphorylate in the protein target and by their location in the cell. Three groups of protein kinases are distinguished by the types of amino acid targets: • Protein serine/threonine kinases are responsible for the vast majority of phosphorylation events in the cell. As their name indicates, they phosphorylate either serine or threonine in the target protein. Protein kinases are important players in signal transduction | SECTION 6.28.7 © 2004. Virtual Text / www.ergito.com

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• Protein tyrosine kinases phosphorylate tyrosine in the target protein. • Dual specificity kinases are less common and can phosphorylate target proteins on either tyrosine or serine/threonine. Protein kinases are found in two types of location: • Cytosolic protein kinases are most often Ser/Thr protein kinases. They are responsible for the vast majority of phosphorylation events in the cell. One particularly important class are the cdk (cyclin-dependent kinase) enzymes that control the cell cycle (see Molecular Biology 6.29 Cell cycle and growth regulation). Dual specificity kinases are found in the MAP kinase signal transduction pathway (see Molecular Biology 6.28.16 A MAP kinase pathway is a cascade). The products of some oncogenes, for which src is the paradigm, are protein tyrosine kinases (see Molecular Biology 6.30.16 Src is the prototype for the proto-oncogenic cytoplasmic tyrosine kinases). • Receptor protein kinases are found in the plasma membrane. They have a domain on the exterior of the cell that binds a ligand, and a catalytic domain within the cell that can act on a target protein (for review see 2899; 2266). Most receptors with protein kinase activity are protein tyrosine kinases (abbreviated as RTK for receptor tyrosine kinase), although there are also some receptors of the Ser/Thr kinase class. All kinases have an active site that binds ATP and a short sequence of the target protein that includes the amino acid to be phosphorylated. The sequence bound at the active site usually conforms to a consensus, typically 3-4 amino acids long. Recognition of the target protein also depends on interactions involving other regions of both the kinase and the target. Figure 28.13 illustrates the structure of a dual specificity kinase of the MAP kinase family, based on its crystal structure (2900; 2901). The active site consists of a short loop of the enzyme that forms a deep cleft. The sequence of the catalytic loop is generally conserved. ATP binds at the bottom of the cleft. Adjacent to the catalytic loop, on the surface of the enzyme, is a sequence called the phosphorylation lip; many kinases in this group have amino acids in this sequence whose phosphorylation activates the enzyme activity. The phosphorylation lip contacts the amino acid on the N-terminal side of the amino acid that is phosphorylated.

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Figure 28.13 The active site of a cytosolic kinase is identified by the catalytic loop (green). The target sequence for phosphorylation binds to the active site. Amino acid 0 is phosphorylated. The amino acid on the N-terminal side (indicated by N) is immediately adjacent to the phosphorylation lip, which is a sequence in the kinase that is often itself phosphorylated.

Receptor tyrosine kinases have some common features. The extracellular domain often has characteristic repeating motifs. It contains a ligand-binding site. The transmembrane region is a single short membrane-spanning alpha helix. The catalytic domain is large (~250 amino acids), and often occupies the bulk of the cytoplasmic region. Certain conserved features are characteristic of all kinase catalytic domains. Sometimes the catalytic domain is broken into two parts by an interruption of some other sequence (which may have an important function in selecting the substrate). Figure 28.14 illustrates the features of a receptor tyrosine kinase. Because the receptors are embedded in membranes, we do not yet have crystal structures of intact proteins. However, several extracellular and cytoplasmic domains have been independently crystallized (for review see 2266). The extracellular and cytoplasmic domains of the RTK group both show large variations in size (for review see 2899). The receptors are usually activated by binding a polypeptide ligand, which can be a significant size relative to the extracellular domain (the example in the figure shows the scale for an FGF ligand binding to its receptor; see 2902). The cytoplasmic domain of the RTK is large, and contains many sites involved in signaling, as well as the kinase catalytic domain. Crystal structures have identified the features of the active site (2903; 2904). At the active site, the catalytic loop is adjacent to an activation loop, which contains 2-3 tyrosines. When these tyrosines are phosphorylated, the activation loop swings away from the catalytic loop, freeing it to bind the substrate (for review see 2899).


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Figure 28.14 A receptor tyrosine protein kinase has an extracellular domain that binds a ligand, which is usually a polypeptide, and a cytoplasmic domain that includes a kinase region. ATP binds adjacent to the catalytic site. An important feature is the activation loop, which contains tyrosine residues whose phosphorylation activates the kinase activity. The activation loop containing these tyrosines changes its position when they are phosphoryated.

Phosphorylation usually activates the target protein, but this is not a golden rule – there are some cases in which phosphorylation inhibits the activity of the target. One way to reverse the effects of a phosphorylation event is for a phosphatase (typically a cytosolic phosphatase) to remove the phosphate that was added by a protein kinase (for review see 310).There are phosphatases with specificity for the appropriate amino acids to match each type of kinase. Most phosphatases are cytosolic, although there are some receptor phosphatases. Last updated on 9-6-2002 Protein kinases are important players in signal transduction | SECTION 6.28.7 © 2004. Virtual Text / www.ergito.com

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Reviews 293. Hunter, T. and Cooper, J. A. (1985). Protein-tyrosine kinases. Annu. Rev. Biochem. 54, 897-930. 294. Hunter, T. (1987). A thousand and one protein kinases. Cell 50, 823-829. 310. Hunter, T. (1995). Protein kinases and phosphatases: the Yin and Yang of protein phosphorylation and signaling. Cell 80, 237-248. 2266. Hubbard, S. R. and Till, J. H. (2000). Protein tyrosine kinase structure and function. Annu. Rev. Biochem. 69, 373-398. 2899. Yarden, Y. and Ullrich, A. (1988). Growth factor receptor tyrosine kinases. Annu. Rev. Biochem. 57, 443-478.

References 2900. Zhang, F., Strand, A., Robbins, D., Cobb, M. H., and Goldsmith, E. J. (1994). Atomic structure of the MAP kinase ERK2 at 2.3 A resolution. Nature 367, 704-711. 2901. Canagarajah, B. J., Khokhlatchev, A., Cobb, M. H., and Goldsmith, E. J. (1997). Activation mechanism of the MAP kinase ERK2 by dual phosphorylation. Cell 90, 859-869. 2902. Plotnikov, A. N., Schlessinger, J., Hubbard, S. R., and Mohammadi, M. (1999). Structural basis for FGF receptor dimerization and activation. Cell 98, 641-650. 2903. Hubbard, S. R., Wei, L., Ellis, L., and Hendrickson, W. A. (1994). Crystal structure of the tyrosine kinase domain of the human insulin receptor. Nature 372, 746-754. 2904. Mohammadi, M., Schlessinger, J., and Hubbard, S. R. (1996). Structure of the FGF receptor tyrosine kinase domain reveals a novel autoinhibitory mechanism. Cell 86, 577-587.



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SIGNAL TRANSDUCTION

6.28.8 Growth factor receptors are protein kinases Key Terms A growth factor is a ligand, usually a small polypeptide, that activates a receptor in the plasma membrane to stimulate growth of the target cell. Growth factors were originally isolated as the components of serum that enabled cells to grow in culture. A cytokine is a small polypeptide that affects the growth of particular types of cells. Key Concepts

• Binding of a ligand to the extracellular domain of a growth factor receptor activates the kinase activity of the cytoplasmic domain.

• The receptor may activate a second messenger or may activate a cascade of kinases.

Growth factor receptors take their names from the nature of their ligands, which usually are small polypeptides (casually called growth factors, more properly called cytokines) that stimulate the growth of particular classes of cells. The factors have a variety of effects, including changes in the uptake of small molecules, initiation or stimulation of the cell cycle, and ultimately cell division. The ligands most usually are secreted from one cell to act upon the receptor of another cell. Examples of secreted cytokines are EGF (epidermal growth factor), PDGF (platelet-derived growth factor), and insulin. In some cases, ligands instead take the form of components of the extracellular matrix, or membrane proteins on the surface of another cell (these are sometimes called counter-receptors). The receptors share a general characteristic structure: they are group I integral membrane proteins, spanning the membrane once, with an N-terminal protein domain on the extracellular side of the membrane, and the C-terminal domain on the cytoplasmic side. Most receptors, such as those for EGF or PDGF, consist of single polypeptide chains. An exception is provided by receptors of the insulin family, which are disulfide-bonded dimers (each dimer being a group I protein). The effector pathways that are activated by receptor tyrosine kinases (RTKs) fall into two groups: • An enzymatic activity is activated that leads to the production of a small molecule second messenger. The second messenger may be the immediate product of an enzyme that is activated directly by the receptor, or may be produced later in the pathway. Lipids are common second messengers in these pathways. The enzymes include phospholipases (which cleave lipids from larger substrates) and kinases that phosphorylate lipid substrates. Some common pathways are summarized in Figure 28.15. The second messengers that are Growth factor receptors are protein kinases | SECTION 6.28.8 © 2004. Virtual Text / www.ergito.com

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released in each pathway act in the usual way to activate or inactivate target proteins. • The effector pathway is a cascade that involves a series of interactions between macromolecular components. The most common components of such pathways are protein kinases; each kinase activates the next kinase in the pathway by phosphorylating it, and the ultimate kinases in the pathway typically act on proteins such as transcription factors that may have wide-ranging effects upon the cell phenotype.

Figure 28.15 Effectors for receptor tyrosine kinases include phospholipases and kinases that act on lipids to generate second messengers.

The basic principle underlying the function of all types of effector pathway is that the signal is amplified as it passes from one component of the pathway to the next. When some components have multiple targets, the pathway branches, thus creating further diversity in the response to the original stimulus. When a ligand binds to the extracellular domain of a growth factor receptor, the catalytic activity of the cytoplasmic domain is activated. Phosphorylation of tyrosine is identified as the key event by which the growth factor receptors function because mutants in the tyrosine kinase domain are biologically inactive, although they continue to be able to bind ligand. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.28.8

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SIGNAL TRANSDUCTION

6.28.9 Receptors are activated by dimerization Key Terms The ability of a species of kinase to phosphorylate itself is referred to as autophosphorylation. Autophosphorylation does not necessarily occur on the same polypeptide chain as the catalytic site; for example, in a dimer, each subunit may phosphorylate the other. Key Concepts

• Ligand binding to receptor monomers causes them to dimerize by interactions between the extracellular domains.

• Dimerization is made possible by the ability of membrane proteins to move laterally within the membrane bilayer.

• Dimerization activates the cytoplasmic domains by an autophosphorylation in

which the kinase activity of each monomer phosphorylates the other monomer.

A key question in the concept of how a signal is transduced across a membrane is how binding of the ligand to the extracellular domain activates the catalytic domain in the cytoplasm. The general principle is that a conformational change is induced that affects the overall organization of the receptor. An important factor in this interaction is that membrane proteins have a restricted ability to diffuse laterally (in contrast with the continuous motion of the lipids in the bilayer). This enables their state of aggregation to be controlled by external events. Lateral movement plays a key role in transmitting information from one side of the membrane to the other. Figure 28.16 shows that binding of ligand induces a conformation change in the N-terminal region of a group I receptor that causes the extracellular domains to dimerize. This causes the transmembrane domains to diffuse laterally, bringing the cytoplasmic domains into juxtaposition. The stabilization of contacts between the C-terminal cytosolic domains causes a change in conformation that activates the kinase activity. In some cases, phosphorylation also causes the receptor to interact with proteins present on the cytoplasmic surface of a coated pit, leading to endocytosis of the receptor. An extreme case of lateral diffusion is seen in certain cases of receptor internalization, when receptors of a given type aggregate into a "cap" in response to an extracellular stimulus.

Receptors are activated by dimerization | SECTION 6.28.9 © 2004. Virtual Text / www.ergito.com

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Figure 28.16 The principle underlying signal transduction by a tyrosine kinase receptor is that ligand binding to the extracellular domain triggers dimerization; this causes a conformational change in the cytoplasmic domain that activates the tyrosine kinase catalytic activity. This is a static version of an interactive figure; see http://www.ergito.com/main.jsp?bcs=MBIO.6.28.9 to view properly.

Figure 28.17 shows that dimerization can take several forms (for review see 2898). The most common is that a ligand binds to one or to both monomers to induce them to dimerize (2905). A variation is that a dimeric ligand binds to two monomers to bring them together (2902). In the case of the insulin receptor family, the ligand binds to a dimeric receptor (which is stabilized by extracellular disulfide bridges) to cause an intramolecular change of conformation. The major consequence of dimerization is to allow transmission of a conformational change from the extracellular domain to the cytoplasmic domain without requiring a change in the structure of the transmembrane region (793; for review see 307; 2266).


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Figure 28.17 Binding of ligand to the extracellular domain can induce aggregation in several ways. The common feature is that this causes new contacts to form between the cytoplasmic domains.

Dimerization initiates the signaling pathway by triggering an autophosphorylation in the cytoplasmic domains of the receptor. When the two cytoplasmic domains are brought together in the dimer, each phosphorylates the other. It is necessary for both subunits to have kinase activity for the receptor to be activated; if one subunit is defective in kinase activity, the dimer cannot be activated. Autophosphorylation has two consequences. Phosphorylation of tyrosines in the kinase domain causes the "activation loop" to swing away from the catalytic center, Receptors are activated by dimerization | SECTION 6.28.9 © 2004. Virtual Text / www.ergito.com

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thus activating the ability of the kinase to bind its substrate (see Figure 28.14). Phosphorylation of tyrosines at other regions of the cytoplasmic domain provides the means by which substrate proteins are enabled to bind to the receptor. The existence of these phosphorylated tyrosine(s) in specific signaling motifs causes the cytoplasmic domain to associate with its target proteins (see Molecular Biology 6.28.10 Receptor kinases activate signal transduction pathways; for review see 298; 305).


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Reviews 298. Ullrich, A. and Schlessinger, J. (1990). Signal transduction by receptors with tyrosine kinase activity. Cell 61, 203-212. 305. van der Geer, P., Hunter, T., and Lindberg, R. A. (1994). Receptor protein-tyrosine kinases and their signal transduction pathways. Annu. Rev. Cell Biol. 10, 251-337. 307. Heldin, C.-H. (1995). Dimerization of cell surface receptors in signal transduction. Cell 80, 213-223. 2266. Hubbard, S. R. and Till, J. H. (2000). Protein tyrosine kinase structure and function. Annu. Rev. Biochem. 69, 373-398. 2898. Schlessinger, J. (2000). Cell signaling by receptor tyrosine kinases. Cell 103, 211-225.

References 793. Cunningham, B. C. et al. (1991). Dimerization of the extracellular domain of the human growth hormone receptor by a single hormone molecule. Science 254, 821-825. 2902. Plotnikov, A. N., Schlessinger, J., Hubbard, S. R., and Mohammadi, M. (1999). Structural basis for FGF receptor dimerization and activation. Cell 98, 641-650. 2905. Wiesmann, C., Fuh, G., Christinger, H. W., Eigenbrot, C., Wells, J. A., and de Vos, A. M. (1997). Crystal structure at 1.7 A resolution of VEGF in complex with domain 2 of the Flt-1 receptor. Cell 91, 695-704.



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6.28.10 Receptor kinases activate signal transduction pathways Key Terms Oncogenes are genes whose products have the ability to transform eukaryotic cells so that they grow in a manner analogous to tumor cells. Oncogenes carried by retroviruses have names of the form v-onc. Key Concepts

• Receptor activation causes phosphorylation of Tyr at several short sequence motifs in the cytoplasmic domain.

• Different substrate proteins bind to particular motifs. • The substrate proteins may be docking proteins that bind other proteins, or

signaling proteins that have an enzymatic activities that are activated by associating with the receptor.

Figure 28.18 shows that we can distinguish several types of proteins with which the activated receptor may interact:

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Figure 28.18 Phosphorylation of the intracellular domain of a receptor creates sites that bind cytosolic proteins. Three types of proteins that propagate signaling pathways are adaptors that bind other proteins, enzymes that are activated by associating with the receptor, and substrates, especially kinases, that are activated by being phosphorylated. Other targets can include structural proteins (not shown).

• The protein may be an intermediary that has no catalytic activity of its own, but serves merely to bring other proteins to the receptor. "Docking proteins" or Receptor kinases activate signal transduction pathways | SECTION 6.28.10 © 2004. Virtual Text / www.ergito.com

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"adaptors" bind to an activated receptor, and then other proteins(s) bind to them, and may therefore become substrates for the receptor. By assembling complexes via such intermediaries, receptors can extend their range (for review see 2908). • The protein may be a target that is activated by its association with the receptor, but which is not itself phosphorylated. For example, some enzymes are activated by binding to a receptor, such as PI3 kinase (see Figure 28.15). • If the protein is a substrate for the enzyme, it becomes phosphorylated. If the substrate is itself an enzyme, it may be activated by the phosphorylation (example: c-Src or PLC γ ; see Figure 28.15). Sometimes the substrate is a kinase, and the pathway is continued by a cascade of kinases that successively activate one another. • Some substrates may be end-targets, such as cytoskeletal proteins, whose phosphorylation changes their properties, and causes assembly of a new structure. A receptor tyrosine kinase can initiate a signaling cascade at the membrane. However, in many cases, the activation of the kinase is followed by its internalization, that is, it is removed from the membrane and transported to the interior of the cell by endocytosis of a vesicle carrying a patch of plasma membrane. The relationship between kinase activity and endocytosis is unclear. Phosphorylation at particular residues may be needed for endocytosis; whether the kinase activity as such is needed may differ for various receptors. It is possible that endocytosis of receptor kinases serves principally to clear receptor (and ligand) from the surface following the response to ligand binding (thus terminating the response). However, in some cases, movement of receptors to coated pits followed by internalization could be necessary for them to act on the target proteins. Because growth factor receptors generate signals that lead to cell division, their activation in the wrong circumstances is potentially damaging to an organism, and can lead to uncontrolled growth of cells. Many of the growth factor receptor genes are represented in the oncogenes, a class of mutant genes active in cancers. The mutant genes are derived by changes in cellular genes; often the mutant protein is truncated in either or both of its N-terminal or C-terminal regions. The mutant protein usually displays two properties: the tyrosine kinase has been activated; and there is no longer any response to the usual ligand. As a result, the tyrosine kinase activity of the receptor is either increased or directed against new targets (see Molecular Biology 6.30.15 Growth factor receptor kinases can be mutated to oncogenes). Last updated on 9-10-2002


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Reviews 2908. Pawson, T. and Scott, J. D. (1997). Signaling through scaffold, anchoring, and adaptor proteins. Science 278, 2075-2080.



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6.28.11 Signaling pathways often involve protein-protein interactions Key Terms An SH2 domain (named originally as the Src Homology domain because it was identified in the Src product of the Rous sarcoma virus) is a region of ~100 amino acids that is bound by the SH2-binding domain of the protein upstream in a signal transduction cascade. An SH3 domain is used by some proteins that contain SH2 domains to enable them to bind to the next component downstream in a signal transduction cascade. Key Concepts

• An SH2-binding site has a phospho-Tyrosine residue that is recognized by an SH2 domain.

• A receptor may have several SH2-binding sites, which are recognized by the SH2 domains of different signaling proteins.

• The signaling protein may have an SH3 domain that recognizes the next protein in the pathway.

A common means for propagating a signal transduction pathway is for a protein specifically to recognize the next protein in the pathway by means of a physical interaction (for review see 2908). (This contrasts with the generation of a small molecule [second messenger] that interacts with the next protein in the pathway.) The usual mechanism for a protein-protein interaction in a signal pathway is for a domain in one protein to recognize a rather short motif in a second protein. The salient feature of the target motif may be its sequence or its structure. Phospho-Tyr residues are often components of such motifs, allowing the motif to be made active by phosphorylation or made inactive by dephosphorylation (for review see 3430). Another common feature of target motifs is the amino acid proline, which causes a characteristic turn in a polypeptide chain (for review see 2918). Two motifs found in a variety of cytoplasmic proteins that are involved in signal transduction are used to connect proteins to the components that are upstream and downstream of them in a signaling pathway. The domains are named SH2 and SH3, for Src homology, because they were originally described in the c-Src cytosolic tyrosine kinase (see Molecular Biology 6.30.16 Src is the prototype for the proto-oncogenic cytoplasmic tyrosine kinases; for review see 299; 302). The presence of SH2 and SH3 domains in various proteins is summarized in Figure 28.19. The cytoplasmic tyrosine kinases comprise one group of proteins that have these domains; other prominent members are phospholipase C γ and the regulatory subunit (p85) of PI3 kinase (both targets for activation by receptor tyrosine kinases; see Figure 28.15). The extreme example of a protein with these domains is Signaling pathways often involve protein-protein interactions | SECTION 6.28.11 © 2004. Virtual Text / www.ergito.com

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Grb2/sem5, which consists solely of an SH2 domain flanked by two SH3 domains (see later).

Figure 28.19 Several types of proteins involved in signaling have SH2 and SH3 domains.

Some proteins contain multiple SH2 domains, which increases their affinity for binding to phosphoproteins or confers the ability to bind to different phosphoproteins. A receptor may contain different SH2-binding sites, enabling it to activate a variety of target proteins. Figure 28.20 summarizes the organization of the cytoplasmic domain of the PDGF receptor, which has ~10 distinct SH2-binding sites, each created by a different phosphorylation event. Different pathways may be triggered by the proteins that bind to the various phosphorylated residues (794).

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Figure 28.20 Autophosphorylation of the cytosolic domain of the PDGF receptor creates SH2-binding sites for several proteins. Some sites can bind more than one type of SH2 domain. Some SH2-containing proteins can bind to more than one site. The kinase domain consists of two separated regions (shown in blue), and is activated by the phosphorylation site in it.

A protein that contains an SH2 domain is activated when it binds to an SH2-binding site. The activation may involve the SH2-containing protein directly (when it itself has an enzymatic activity) or may be indirect. The enzymatic activities that are regulated directly are most commonly kinases, phosphatases, or phospholipases. An example of a protein containing an SH2 domain that does not have a catalytic activity is provided by p85, the regulatory subunit of PI3 kinase; when p85 binds to a receptor, it is the associated PI3K catalytic subunit that is activated. Figure 28.21 shows that the SH3 domain provides the effector function by which some SH2-containing proteins bind to a downstream component. The case of the "adaptor" Grb2 strengthens this idea; consisting only of SH2 and SH3 domains, it uses the SH2 domain to contact the component upstream in the pathway, and the SH3 domain to contact the component downstream. SH3 binds the motif PXXP in a Signaling pathways often involve protein-protein interactions | SECTION 6.28.11 © 2004. Virtual Text / www.ergito.com

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sequence-specific manner (see Molecular Biology 6.28.13 Prolines are important determinants in recognition sites). When an activated receptor binds Grb2, the SH3 domain of Grb2 binds to a target protein that contains the PXXP motif (805).

Figure 28.21 SH2 and SH3 domains are used for protein-protein interactions in signal transduction cascades. Typically a phosphorylated receptor recognizes an SH2 target in its substrate, and an SH3 domain in the substrate then recognizes the next protein in the cascade.

SH3 domains often provide connections to small GTP-binding proteins (of which Ras is the paradigm). Another role that has been proposed for SH3 domains (and in particular for the SH3 domain of c-Src) is the ability to interact with proteins of the cytoskeleton, thus triggering changes in cell structure. Last updated on 9-10-2002


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Reviews 299. Koch, C. A. (1991). SH2 and SH3 domains: elements that control interactions of cytoplasmic signaling proteins. Science 252, 668-674. 302. Cohen, G. B., Ren, R., and Baltimore, D. (1995). Molecular binding domains in signal transduction proteins. Cell 80, 237-248. 2908. Pawson, T. and Scott, J. D. (1997). Signaling through scaffold, anchoring, and adaptor proteins. Science 278, 2075-2080. 2918. Kay, B. K., Williamson, M. P., and Sudol, M. (2000). The importance of being proline: the interaction of proline-rich motifs in signaling proteins with their cognate domains. FASEB J. 14, 231-241. 3430. Yaffe, M. B. (2002). Phosphotyrosine-binding domains in signal transduction. Nat. Rev. Mol. Cell Biol. 3, 177-186.

References 794. Fantl, W. J. et al. (1992). Distinct phosphotyrosines on a growth factor receptor bind to specific molecules that mediate different signaling pathways. Cell 69, 413-423. 805. Booker, G. W. et al. (1993). Solution structure and ligand-binding site of the SH3 domain of the p85 α subunit of phosphatidylinositol 3-kinase. Cell 73, 813-822.



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6.28.12 Phosphotyrosine is the critical feature in binding to an SH2 domain Key Terms An SH2 domain (named originally as the Src Homology domain because it was identified in the Src product of the Rous sarcoma virus) is a region of ~100 amino acids that is bound by the SH2-binding domain of the protein upstream in a signal transduction cascade. An SH2-binding site is an area on a protein that interacts with the SH2 domain of another protein. Key Concepts

• An SH2-binding site consists of phospho-Tyrosine and 104 through the pathway. However, the combination of the last three kinases into one complex would presumably restrict amplification at these stages. In mammalian cells, the pathway can be fully activated by very weak signals; for example, the ERK1,2 MAP kinases are fully activated when 7 STATs; each STAT is phosphorylated by a particular set of JAK kinases. The phosphorylation occurs while the JAK is associated with the receptor at the plasma membrane. A pair of JAK kinases associates with an activated receptor, and both may be necessary for the pathway to function. An example is that stimulation by the interferon IFN γ requires both JAK1 and JAK2 (803; 804; for review see 303). STAT phosphorylation leads to the formation of both homodimers and heterodimers. The basis for dimerization is a reciprocal interaction between an SH2 domain in one subunit and a phosphorylated Tyr in the other subunit (810). The STAT dimers translocate to the nucleus, and in some cases associate with other proteins. They bind to specific recognition elements in target genes, whose transcription is activated (for review see 313). Given a multiplicity of related cytokine receptors, JAK kinases, and STAT transcription factors, how is specificity achieved? The question is sharpened by the fact that many receptors can activate the same JAKs, but activate different STATs. Control of specificity lies with formation of a multipartite complex containing the receptor, JAKs, and STATs. The STATs interact directly with the receptor as well as with the JAKs, and an SH2 domain in a particular STAT recognizes a binding site in a particular receptor. So the major control of specificity lies with the STAT. Stimulation of a JAK-STAT pathway is only transient. Its activation may be terminated by the action of a phosphatase. An example is the pathway activated by binding of erythropoietin (red blood cell hormone) to its receptor. This activates JAK2 kinase. Recruitment of another component terminates the reaction; the phosphatase SH-PTP1 binds via its SH2 domain to a phosphotyrosine site in the erythropoietin receptor. This site in the receptor is probably phosphorylated by JAK2. The phosphatase then dephosphorylates JAK2 and terminates the activation of the corresponding STATs. This creates a simple feedback circuit: erythropoietin receptor activates JAK2, JAK2 acts on a site in the receptor, and this site is recognized by the phosphatase that in turn acts on JAK2. This again emphasizes the way in which formation of a multicomponent complex may be used to ensure specificity in controlling the pathway (811).

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Reviews 303. Darnell, J. E., Kerr, I. M., and Stark, G. R. (1994). JAK-STAT pathways and transcriptional activation in response to IFN γ and other extracellular signaling proteins. Science 264, 1415-1421. 313. Schindler, C. and Darnell, J. E. (1995). Transcriptional responses to polypeptide ligands: the JAK-STAT pathway. Annu. Rev. Biochem. 64, 621-651.

References 803. Dale, T. C. et al. (1989). Rapid activation by interferon α of a latent DNA-binding protein present in the cytoplasm of untreated cells. Proc. Natl. Acad. Sci. USA 86, 1203-1207. 804. Velazquez, L. et al. (1992). A protein tyrosine kinase in the interferon α / β signaling pathway. Cell 70, 313-322. 810. Shuai, K. et al. (1994). Interferon activation of the transcription factor STAT91 involves dimerization through SH2-phosphotyrosyl peptide interactions. Cell 76, 821-828. 811. Klingmuller, U. et al. (1995). Specific recruitment of SH-PTP1 to the erythropoietin receptor causes inactivation of JAK2 and termination of proliferative signals. Cell 80, 729-738.


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6.28.21 TGF β signals through Smads Key Concepts

• TGF β activates the heterodimeric type II receptor. • The activated type II receptor phosphorylates the heterodimeric type I receptor. • As part of the tetrameric complex, the type I receptor phosphorylates a cytosolic Smad protein.

• The Smad forms a dimer with a related protein (Smad4) which moves to the nucleus and activates transcription.

Another pathway in which phosphorylation at the membrane triggers migration of a transcription factor to the nucleus is provided by TGF β signaling (for review see 3650). The TGF β family contains many related polypeptide ligands. They bind to receptors that consist of two types of subunits, as illustrated in Figure 28.46. Both subunits have serine/threonine kinase activity. (Actually all serine/threonine receptor kinases are members of the TGF β receptor family.)

Figure 28.46 Activation of TGF β receptors causes phosphorylation of a Smad, which is imported into the nucleus to activate transcription.

The ligand binds to the type II receptor, creating a receptor-ligand combination that has high affinity for the type I receptor. A tetrameric complex is formed in which the TGF β signals through Smads | SECTION 6.28.21 © 2004. Virtual Text / www.ergito.com

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type II receptor phosphorylates the type I receptor. (A variation occurs in a subset of these receptors that bind BMPs – bone morphogenetic proteins – which are members of the TGF β family. In this case, both type I and type II subunits have low affinity for the ligand, but the combination of subunits has high affinity.) Once the active complex has formed, the type I receptor phosphorylates a member of the cytosolic Smads family. Typically a Smad activator is phosphorylated at the motif SSXS at the C-terminus. This causes it to form a dimer with the common partner Smad4. The heterodimer is imported into the nucleus, where it binds to DNA and activates transcription (812; for review see 314; 316). The 9 Smad proteins fall into three functional categories. The pathway-specific activators are Smad2,3 (which mediate TGF β/activin signaling) and Smad1,5 (which activate BMP signaling). Smad4 is a universal partner which can dimerize with all of the pathway-specific Smads. Inhibitory Smads act as competitive inhibitors of the activator Smads, providing another level of complexity to the pathway. Each ligand in the TGF β superfamily activates a particular receptor that signals through a characteristic combination of Smads proteins. Various other proteins bind to the Smads dimers and influence their capacity to act on transcription. Signaling systems of this type are important in early embryonic development, where they are part of the pathways that lead to development of specific tissues (typically bone formation and the development of mesoderm). Also, because TGF β is a powerful growth inhibitor, this pathway is involved in tumor suppression. The TGF β type II receptor is usually inactivated in hereditary nonpolyposis colorectal cancers, and mutations in Smad4 occur in 50% of human pancreatic cancers. One striking feature of the JAK-STAT and TGF β pathways is the simplicity of their organization, compared (for example) with the Ras-MAPK pathway. The specificity of these pathways depends on variation of the components that assemble at the membrane – different combinations of JAK-STATs in the first case, different Smad proteins in the second. One the pathway has been triggered, it functions in a direct linear manner. The component that is phosphorylated at the plasma membrane (STAT in the JAK-STAT pathway, Smad in the TGF β pathway) itself provides the unit that translocates to the nucleus to activate transcription – perhaps the ultimate demonstration of the role of localization

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Reviews 314. Massague, J. (1996). TGF β ; signaling: receptors, transducers, and Mad proteins. Cell 85, 947-950. 316. Massague, J. (1998). TGF β signal transduction. Annu. Rev. Biochem. 67, 753-791. 3650. Attisano, L. and Wrana, J. L. (2002). Signal transduction by the TGF-beta superfamily. Science 296, 1646-1647.

References 812. Macias-Silva, M. et al. (1996). Madr2 is a substrate of the TGF β receptor and its phosphorylation is required for nuclear accumulation and signaling. Cell 87, 1215-1224.


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SIGNAL TRANSDUCTION

6.28.22 Structural subunits can be messengers It is a common need to carry a signal from an event at the cell surface to the nucleus. Activation of a receptor in the plasma membrane can stimulate a signaling pathway in many ways, but a common feature is that some component of the pathway either itself translocates to the nucleus or causes the translocation of another protein. The component that translocates can be distant in the pathway (as in the case of MAP kinases) or a protein that actually itself interacts with the receptor (as in the case of the TGF β or JAK-STAT pathways. A more extreme case is presented by some pathways in which the translocating component may be actually a structural subunit of the complex at the plasma membrane. Figure 28.47 shows a generic model for such a pathway. When a multisubunit complex is activated at the plasma membrane, one of its subunits is released. The subunit translocates to the nucleus, where (directly or indirectly) it activates transcription. A typical pattern is for the translocating subunit to activate a transcription factor that is already in the nucleus.

Structural subunits can be messengers | SECTION 6.28.22 © 2004. Virtual Text / www.ergito.com

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Figure 28.47 Activation of a complex at the plasma membrane triggers release of a subunit that migrates to the nucleus to activate transcription. This is a static version of an interactive figure; see http://www.ergito.com/main.jsp?bcs=MBIO.6.28.22 to view properly.

A pathway that follows this general model signals from the synapse (the connection between neurons in the nervous system). One of the multiprotein complexes that assembles at the synapse has cell surface proteins bound to a group of cytosolic proteins that include a MAGUK (membrane-associated guanylate kinase). The MAGUK has PDZ and SH3 domains (typically involved in protein-protein interactions), a protein kinase-like domain, and a guanylate kinase-like domain – all the hallmarks of a "scaffold" protein involved in assembling the components of the signaling complex. One example is the protein CASK in the nuclei of embryonic neurons. The guanylate kinase domain of CASK interacts with the nuclear transcription factor Tbr (955). This raises the possibility that CASK is a bifunctional protein: one function being in assembly of the cell surface complex, the other being in activating transcription in the nucleus. Last updated on 10-31-2001


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References 955. Diebel, C. E., Proksch, R., Green, C. R., Neilson, P., Walker, M. M., Diebel, C. E., Proksch, R., Green, C. R., Neilson, P., and Walker, M. M. (2000). Magnetite defines a vertebrate magnetoreceptor. Nature 406, 299-302.



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6.28.23 Summary Lipids may cross the plasma membrane, but specific transport mechanisms are required to promote the passage of hydrophilic molecules. Integral proteins of the plasma membrane offer several means for communication between the extracellular milieu and the cytoplasm. They include ion channels, transporters, and receptors. All such proteins reside in the plasma membrane by means of hydrophobic domains. Ions may be transported by carrier proteins, which may utilize passive diffusion or may be connected to energy sources to undertake active diffusion. The detailed mechanism of movement via a carrier is not clear, but is presumed to involve conformational changes in the carrier protein that directly or indirectly allow a substrate to move from one side of the membrane to the other. Ion channels can be used for passive diffusion (driven by the gradient). They may be gated (controlled) by voltage, extracellular ligands, or cytoplasmic second messengers. Channels typically have multiple subunits, each with several transmembrane domains; hydrophilic residues within the transmembrane domains face inward so as to create a hydrophilic path through the membrane. Receptors typically are group I proteins, with a single transmembrane domain, consisting exclusively of uncharged amino acids, connecting the extracellular and cytosolic domains. Many receptors for growth factors are protein tyrosine kinases. Such receptors have a binding site for their ligand in the extracellular domain, and a kinase activity in their cytoplasmic domain. When a ligand binds to the receptor, it causes the extracellular domain to dimerize; most often the product is a homodimer, but there are some cases where heterodimers are formed. The dimerization of the extracellular domains causes the transmembrane domains to diffuse laterally within the membrane, bringing the cytoplasmic domains into contact. This results in an autophosphorylation in which each monomeric subunit phosphorylates the other. The phosphorylation creates a binding site for the SH2 motif of a target protein. Specificity in the SH2-binding site typically is determined by the phosphotyrosine in conjunction with the 4-5 neighboring amino acids on its C-terminal side. The next active component in the pathway may be activated indirectly or directly. Some target proteins are adaptors that are activated by binding to the phosphorylated receptor, and they in turn activate other proteins. An adaptor typically uses its SH2 domain to bind the receptor and uses an SH3 domain to bind the next component in the pathway. Other target proteins are substrates for phosphorylation, and are activated by the acquisition of the phosphate group. One group of effectors consists of enzymes that generate second messengers, most typically phospholipases and kinases that generate or phosphorylate small lipids. Another type of pathway consists of the activation of a kinase cascade, in which a series of kinases successively activate one another, leading ultimately to the phosphorylation and activation of transcription factors in the nucleus. The MAP kinase pathway is the paradigm for this type of response. The connection from receptor tyrosine kinases to the MAP kinase pathway passes Summary | SECTION 6.28.23 © 2004. Virtual Text / www.ergito.com

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through Ras. An adaptor (Grb2 in mammalian cells) is activated by binding to the phosphorylated receptor. Grb2 binds to SOS, and SOS causes GDP to be replaced by GTP on Ras. Ras is anchored to the cytoplasmic face of the membrane. The activated Ras binds the Ser/Thr kinase Raf, thus bringing Raf to the membrane, which causes Raf to be activated, probably because it is phosphorylated by a kinase associated with the membrane. Raf phosphorylates MEK, which is a dual-specificity kinase that phosphorylates ERK MAP kinases on both tyrosine and threonine. ERK MAP kinases activate other kinases; ERK2 MAP kinase also translocates to the nucleus, where it phosphorylates transcription factors that trigger pathways required for cell growth (in mammalian cells) or differentiation (in fly retina, worm vulva, or yeast mating). An alternative connection to the MAP kinase cascade exists from serpentine receptors. Activation of a trimeric G protein causes MEKK to be activated. One component in the pathway between G β γ and MEKK in S. cerevisiae is the kinase STE20. The MEKK (STE11), MEK (STE7), and MAPK (Fus3) form a complex with the scaffold protein STE5 that may be necessary for the kinases to function. The use of scaffolding proteins allows the same kinases to participate in different pathways, but to signal to the downstream components only of the pathway that activates them. The cyclic AMP pathway for activating transcription proceeds by releasing the catalytic subunit of PKA in the cytosol. It diffuses to the nucleus, where it phosphorylates the transcription factor CREB. The activity of this factor is responsible for activating cAMP-inducible genes. The response is down regulated by phosphatases that dephosphorylate CREB and by an inhibitor that exports the C subunit back to the cytosol. JAK-STAT pathways are activated by cytokine receptors. The activated receptor associates with a JAK kinase and activates it. The target for the kinase is a STAT(s); STATs associate with a receptor-JAK kinase complex, are phosphorylated by the JAK kinase, dimerize, translocate to the nucleus, and form a DNA-binding complex that activates transcription at a set of target genes. In an analogous manner, TGF β ligands activate type II/type I receptor systems that phosphorylate Smad proteins, which then are imported into the nucleus to activate transcription. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.28.23


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CELL CYCLE AND GROWTH REGULATION

6.29.1 Introduction Key Terms The cell cycle is the set of stages through which a cell progresses from one division to the next. Interphase is the period between mitotic cell divisions; divided into G1, S, and G2. Mitosis (M phase) is the process by which the cell nucleus divides, resulting in daughter cells that contain the same amount of genetic material as the parent cell. The two cells that result from a cell division are referred to as daughter cells. In budding yeast only the cell derived from the bud is called the daughter cell. G1 is the period of the eukaryotic cell cycle between the last mitosis and the start of DNA replication. S phase is the restricted part of the eukaryotic cell cycle during which synthesis of DNA occurs. G2 phase is the period of the cell cycle separating the replication of a cell's chromosomes (S phase) from the following mitosis (M phase). The spindle guides the movement of chromosomes during cell division. The structure is made up of microtubules.

The act of division is the culmination of a series of events that have occurred since the last time a cell divided. The period between two mitotic divisions defines the somatic cell cycle. The time from the end of one mitosis to the start of the next is called interphase. The period of actual division, corresponding to the visible mitosis, is called M phase. In order to divide, a eukaryotic somatic cell must double its mass and then apportion its components equally between the two daughter cells. Doubling of size is a continuous process, resulting from transcription and translation of the genes that code for the proteins constituting the particular cell phenotype. By contrast, reproduction of the genome occurs only during a specific period of DNA synthesis. Mitosis of a somatic cell generates two identical daughter cells, each bearing a diploid complement of chromosomes. Interphase is divided into periods that are defined by reference to the timing of DNA synthesis, as summarized in Figure 29.1:


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Figure 29.1 Overview: interphase is divided into the G1, S, and G2 periods. One cell cycle is separated from the next by mitosis (M). Cells may withdraw from the cycle into G0 or reenter from it.

• Cells are released from mitosis into G1 phase, when RNAs and proteins are synthesized, but there is no DNA replication. • The initiation of DNA replication marks the transition from G1 phase to the period of S phase. S phase is defined as lasting until all of the DNA has been replicated. During S phase, the total content of DNA increases from the diploid value of 2n to the fully replicated value of 4n. • The period from the end of S phase until mitosis is called G2 phase; during this period, the cell has two complete diploid sets of chromosomes. [S phase was so called as the synthetic period when DNA is replicated, G1 and G2 standing for the two "gaps" in the cell cycle when there is no DNA synthesis (823).] The changes in cellular components are summarized in Figure 29.2. During interphase, there is little visible change in the appearance of the cell. The more or less continuous increase of RNA and protein contrasts with the discrete doubling of DNA. The nucleus increases in size predominantly during S phase, when proteins accumulate to match the production of DNA. Chromatin remains a compact mass in which no change of state is visible.


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Figure 29.2 Synthesis of RNA and proteins occurs continuously, but DNA synthesis occurs only in the discrete period of S phase. The units of mass are arbitrary.

Mitosis segregates one diploid set of chromosomes to each daughter cell. Individual chromosomes become visible only during this period, when the nuclear envelope dissolves, and the cell is reorganized on a spindle. The mechanism for specific segregation of material applies only to chromosomes, and other components are apportioned essentially by the flow of cytoplasm into the two daughter cells. Virtually all synthetic activities come to a halt during mitosis. In a cycling somatic animal cell, this sequence of events is repeated every 18-24 hours. Figure 29.1 shows that G1 phase usually occupies the bulk of this period, varying from ~6 h in a fairly rapidly growing animal cell to ~12 h in a more slowly growing cell. The duration of S phase is determined by the length of time required to replicate all the genome, and a period of 6-8 h is typical. G2 phase is usually the shortest part of interphase, essentially comprising the preparations for mitosis. M phase (mitosis) is a brief interlude in the cell cycle, usually 6 generations. Mouse telomeres are exceptionally long, and range from 10-60 kb. In the absence of telomerase, telomeres shorten at 50-100 bp per cell division. There are ~60 divisions in sperm cell production, and ~25 divisions in oocyte production, which fits with the observed rate of shortening of ~4.8 kb per male mouse generation. This gives an expectation that after about 7 generations, a telomerase-negative mouse will have run down its telomeres to around zero length. By the 6th generation, chromosomal abnormalities become more frequent, and the mice become infertile (due to the inability to produce sperm). The effects of lack of telomerase are first seen in tissues consisting of highly proliferative cells (as might be expected). All of these observations demonstrate the importance of telomerase for continued cell division. However, cells from the telomerase-negative mice can pass through crisis and can be transformed to give tumorigenic cells, so the presence of telomerase is not essential, or at least is not the only means, of supporting an immortal state (although reactivation of telomerase is by far the most common mechanism) (see Molecular Biology 5.19.19 Telomeres are essential for survival) (879). Telomerase-negative mice can develop tumors, but do so at a rate lower than wild-type mice (2852). The effect of telomere loss on formation of a cancer cell is therefore confined to its role in provoking a genetic instability that stimulates tumor initiation. After that, it is in fact inhibitory to cancer formation. There is a curious inconsistency between the results obtained with cultured cells and the survival of telomerase-negative mice. Crisis of mouse cells occurs typically after 10-20 divisions in culture, but we would not expect the telomeres to have reached a limiting length at this point. Mice of the first telomerase-negative generation have passed a greater number of cell divisions without telomerase, and without suffering any ill effects. Mice of the third telomerase-negative generation are to all intents and purposes normal, although their cells have gone through more divisions than would have triggered crisis in culture. Lack of telomerase is clearly associated with inability to continue growth, and reactivation of telomerase is one means by which cells can behave as immortal. It is not clear whether telomerase is the only relevant factor in driving cells into crisis and to what extent other mechanisms might be able to compensate for lack of telomerase. We do not know what pathway is responsible for controlling telomerase production in vivo, and how it is connected to pathways that control cell growth. Last updated on 8-21-2002

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References 878. Meyerson, M. et al. (1997). hEST2, the putative human telomerase catalytic subunit gene, is up-regulated in tumor cells and during immortalization. Cell 90, 785-795. 879. Blasco, M. A. et al. (1997). Telomere shortening and tumor formation by mouse cells lacking telomerase RNA. Cell 91, 25-34. 1380. Kim, N. W., Piatyszek, M. A., Prowse, K. R., Harley, C. B., West, M. D., Ho, P. L., Coviello, G. M., Wright, W. E., Weinrich, S. L., and Shay, J. W. (1994). Specific association of human telomerase activity with immortal cells and cancer. Science 266, 2011-2015. 1390. Wright, W. E., Brasiskyte, D., Piatyszek, M. A., and Shay, J. W. (1996). Experimental elongation of telomeres extends the lifespan of immortal x normal cell hybrids. EMBO J. 15, 1734-1741. 2852. Greenberg, R. A., Chin, L., Femino, A., Lee, K. H., Gottlieb, G. J., Singer, R. H., Greider, C. W., and DePinho, R. A. (1999). Short dysfunctional telomeres impair tumorigenesis in the INK4a(delta2/3) cancer-prone mouse. Cell 97, 515-525.


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ONCOGENES AND CANCER

6.30.25 Immortalization depends on loss of p53 Key Concepts

• Loss of p53 is the crucial step in immortalization.

When cells enter senescence as the result of telomere shortening, p53 is activated, leading to growth arrest or apoptosis. Figure 30.41 shows that the trigger that activates p53 is the loss of the telomere-binding protein TRF2 from the chromosome ends. In effect, TRF2 protects the end of the DNA, but when it is lost, the free 3 ′ overhanging end activates p53 (1998; 3669).

Figure 30.41 TRF2 protects telomeres, but when it is lost, the exposed ends can bind and activate p53.

The loss of p53 is the crucial event that allows the cells to survive and divide. A variety of events can be associated with immortalization, but they converge upon causing either the loss or inactivation of p53 protein. Remember that p53 was discovered as the protein of host cells that binds the T antigen (the transforming protein) of polyomaviruses. A major part of the activity of T antigen is its ability to inactivate p53. The T antigens of different viruses work in different ways, but the consequences of the interaction are especially clear in the case of HPV E6 (the equivalent of T antigen), which targets p53 for degradation. In effect, HPV converts a target cell into a p53– state. p53 provides an important function in immortalization, but may not be sufficient by itself. Established cell lines have usually lost p53 function, which suggests that the role of p53 is connected with the acquisition of ability to support prolonged growth. However, loss of the known functions of p53 is not enough by itself to explain Immortalization depends on loss of p53 | SECTION 6.30.25 © 2004. Virtual Text / www.ergito.com

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An interesting convergence is seen in the properties of the tumor antigens from different DNA tumor viruses, . The antigens always bind to both the cellular tumor suppressor products RB and p53. The two cellular proteins are recognized independently. Either different T antigens of the virus bind separately to RB and to p53, or different domains of the same antigen do so. So adenovirus E1A binds RB, while E1B binds p53; HPV E7 binds RB, while E6 binds p53. SV40 T antigen can bind both RB and p53. The loss of p53 (and/or RB) is a major step in the transforming action of DNA tumor viruses, and explains some significant part of the action of the T antigens. The critical events are inhibition of p53's ability to activate transcription, and loss of RB's ability to bind substrates such as E2F. Loss of the tumor suppressors (especially p53) is the major route in the immortalization pathway. Inability to trigger either growth arrest or apoptosis could lead to continued growth. We do not know whether only one or both of these activities are required for immortalization in vitro. We know that more than the growth arrest pathway is needed for p53's contribution to tumorigenesis, because a p21– mouse shows deficiencies in the G1 checkpoint (as would be expected) but does not develop tumors. The contrast with the increased susceptibility of a p53– mouse to tumors shows that other functions of p53 are involved besides its control of p21. We are now in a position to put together the various events involved in immortalization (for review see 2853). Figure 30.42 summarizes the order in which they occur. Checkpoints normally stop a cell from dividing when its telomeres become too short. Cells then enter replicative senescence and stop growing. If they manage to bypass the checkpoints to enter early crisis, the loss of TRF2 from the telomeres activates p53, which causes growth arrest and/or apoptosis. In the absence of p53 activity, they pass into late crisis, where the dysfunction of the telomeres causes genetic instability as seen in large scale chromosomal rearrangements. To survive this stage, they must activate telomerase or find an alternative means of maintaining the telomeres. A cell that survives through all of these stages will be immortal, but almost certainly will have an altered genetic constitution.

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Figure 30.42 Several changes are required to allow a cell to pass the replicative limit and to become immortalized, including bypassing the checkpoints that respond to short telomeres, losing or preventing the ability of p53 to trigger apoptosis, and reactivating telomerase or finding other means to stabilize telomeres.



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Reviews 2853. Maser, R. S. and DePinho, R. A. (2002). Connecting chromosomes, crisis, and cancer. Science 297, 565-569.

References 877. Donehower, L. A. et al. (1992). Mice deficient for p53 are developmentally normal but susceptible for spontaneous tumors. Nature 356, 215-221. 1998. Karlseder, J., Broccoli, D., Dai, Y., Hardy, S., and de Lange, T. (1999). p53- and ATM-dependent apoptosis induced by telomeres lacking TRF2. Science 283, 1321-1325. 3669. Li, G. Z., Eller, M. S., Firoozabadi, R., and Gilchrest, B. A. (2003). Evidence that exposure of the telomere 3' overhang sequence induces senescence. Proc. Natl. Acad. Sci. USA 100, 527-531.



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6.30.26 Different oncogenes are associated with immortalization and transformation Key Terms Cooperativity in protein binding describes an effect in which binding of the first protein enhances binding of a second protein (or another copy of the same protein). Key Concepts

• A tumor cell has independently acquired changes necessary to immortalize it and to transform it.

• Established cell lines grown in culture usually have been immortalized and need to acquire only transforming properties.

• Primary cells require the actions of different oncogenes to be immortalized and to be transformed.

Most tumors arise as the result of multiple events. Some of these events involve the activation of oncogenes, while others take the form of inactivation of tumor suppressors. The requirement for multiple events reflects the fact that normal cells have multiple mechanisms to regulate their growth and differentiation, and several separate changes may be required to bypass these controls. Indeed, the existence of single genes in which mutations were tumorigenic would no doubt be deleterious to the organism, and has been selected against. Nonetheless, oncogenes and tumor suppressors define genes in which mutations create a predisposition to tumors, that is, they represent one of the necessary events. It is an open question as to whether the oncogenes and tumor suppressor genes identified in available assays are together sufficient to account entirely for the occurrence of cancers, but it is clear that their properties explain at least many of the relevant events. Figure 30.43 gives an overview of the stages of tumor formation. There are two discrete stages, which may loosely be viewed as being concerned with immortalization or with transformation (for review see 346; 350).

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Figure 30.43 Crisis is induced by attempts to divide in the absence of telomerase. Immortalization occurs when p53 activity is lost, as the result of mutation in p53 or in a pathway that acts on it. Transformation requires oncogenes to induce further changes in the growth properties of the cells. For cells that develop solid tumors, angiogenic development is required for provision of nutrients. Metastasis is the result of further changes that allow a cell to migrate to form a colony in a new location.

The immortalization step is to bypass crisis (or its equivalent in the in vivo situation). Crisis is provoked when cells continue to divide in the absence of telomerase (see Molecular Biology 6.30.24 Telomere shortening causes cell senescence). When the telomeres become too short, damage to DNA is caused by attempts at replication, and this triggers the activation of p53. The role of p53 is to cause cell death. If p53 is absent, a cell may survive, although at the expense of a genetic catastrophe in which telomere malfunction leads to chromosome fusions and other rearrangements. Immortalized cells can pass through an unlimited number of cell divisions, but they do not have other tumorigenic properties, such as independence of factors required for growth. The second step, transformation, converts immortalized cells into tumorigenic cells. Whether further changes are involved in creating a cancerous state depends on the nature of the tumor cell. A leukemia cell can multiply freely in the blood. However, a cell type that forms a solid tumor needs to develop a blood supply for the tumor (requiring angiogenic development), and may later pass to the stage of metastasis, when cells are detached from the tumor and migrate to form new tumors at other locations. The minimum requirement to enter the tumorigenic state is therefore the occurrence of successive, independent events that involve different tumor suppressors and/or oncogenes. The need for multiple functions of different types is sometimes described as the requirement for cooperativity.


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The involvement of multiple functions fits with the pattern established by some DNA tumor viruses, in which (at least) two types of functions are needed to transform the usual target cells: • Adenovirus carries the E1A region, which allows primary cells to grow indefinitely in culture, and the E1B region, which causes the morphological changes characteristic of the transformed state. • Polyoma produces three T antigens; large T elicits indefinite growth, middle T is responsible for morphological transformation, and small T is without known function. Large T and middle T together can transform primary cells. • Consistent with the classification of oncogenic functions, adenovirus E1A together with polyoma middle T can transform primary cells. This suggests that one function of each type is needed. In the same way, expression of two or more oncogenes in the cellular transfection assay is usually needed to convert a primary cell (one taken directly from the organism) into a tumor cell. Several cellular oncogenes have been identified by transforming ability in the 3T3 transfection assay; 10-20% of spontaneous human tumors have DNA with detectable transforming activity in this assay. Of course, 3T3 cells have been adapted to indefinite growth in culture over many years, and have passed through some of the changes characteristic of tumor cells (see Great Experiments 9.1 The story of 3T3 cells: A voyage of discovery without an itinerary). The exact nature of these changes is not clear, but generally they can be classified as involving functions concerned with immortalization. Oncogenic activity in this assay therefore depends on the ability to induce further changes in an established cell line. The principal products of 3T3 transfection assays are mutated c-ras genes. They do not have the ability to transform primary cells in vitro, and this supports the implication that their functions are concerned with the act of transforming cells that have previously been immortalized. ras oncogenes clearly provide one major pathway for transforming immortalized cells; we do not know how many other transforming pathways may exist that are independent of ras. Although ras oncogenes alone cannot transform primary fibroblasts, dual transfection with ras and another oncogene can do so. The ability to transform primary cells in conjunction with ras provides a general assay for oncogenes that have an immortalization-like function. This group includes several retroviral oncogenes, v-myc, v-jun, and v-fos. It also includes adenovirus E1A and polyoma large T. Mutant p53 genes have the same effect. In fact, the action of the immortalizing oncogenes is most likely to cause inactivation or loss of p53. However, note that the distinction between immortalizing and transforming proteins is not crystal clear. For example, although E1A is classified as having an immortalizing function, it has (some) of the functions usually attributed to transforming proteins, and loss of p53 confers some properties that are usually considered transforming. One way to investigate the oncogenic potential of individual oncogenes Different oncogenes are associated with immortalization and transformation | SECTION 6.30.26 © 2004. Virtual Text / www.ergito.com

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independently of the constraints that usually are involved in their expression is to create transgenic animals in which the oncogene is placed under control of a tissue-specific promoter. A general pattern is that increased proliferation often occurs in the tissue in which the oncogene is expressed. Oncogenes whose expression have this effect with a variety of tissues include SV40 T antigen, v-ras, and c-myc (588). Increased proliferation (hyperplasia) is often damaging and sometimes fatal to the animal (usually because the proportion of one cell type is increased at the expense of another). However, the expression of a single oncogene does not usually cause malignant transformation (neoplasia), with the production of tumors that kill the animal. Tumors resulting from the introduction of an oncogene (for example, in transgenic mice) are probably due to the occurrence of a second event. The need for two types of event in malignancy is indicated by the difference between transgenic mice that carry either the v-ras or activated c-myc oncogene, and mice that carry both oncogenes. Mice carrying either oncogene develop malignancies at rates of 10% for c-myc and 40% for v-ras; mice carrying both oncogenes develop 100% malignancies over the same period. These results with transgenic mice are even more striking than the comparable results on cooperation between oncogenes in cultured cells (866; 867). In some systems, immortalization may be connected with an inability of the cells to differentiate. Growth and differentiation are often mutually exclusive, because a cell must stop dividing in order to differentiate. An oncoprotein that blocks differentiation may allow a cell to continue proliferating (in a sense resembling the immortalization of cultured cells); continued proliferation in turn may provide an opportunity for other oncogenic mutations to occur. This may explain the occurrence among the oncoproteins of products that usually regulate differentiation. A connection between differentiation and tumorigenesis is shown by avian erythroblastosis virus (AEV). The AEV-H strain carries only v-erbB, but the AEV-E54 strain carries two oncogenes, v-erbB and v-erbA. The major transforming activity of AEV is associated with v-erbB, a truncated form of the EGF receptor, which is equivalent to the single oncogene carried by other tumor retroviruses: it can transform erythroblasts and fibroblasts. The other gene, v-erbA, cannot transform target cells alone, but it increases the transforming efficiency of v-erbB. Expression of v-erbA itself has two phenotypic effects upon target cells: it prevents the spontaneous differentiation (into erythrocytes) of erythroblasts that have been transformed by v-erbB; and it expands the range of conditions under which transformed erythroblasts can propagate. v-erbA may therefore contribute to tumorigenicity by a combination of inhibiting differentiation and stimulating proliferation. In fact, v-erbA has a similar effect in extending the efficacy of transformation by other oncogenes that induce sarcomas, notably v-src, v-fps, and v-ras. Correlations between the activation of oncogenes and the successful growth of tumors are strong in some cases, but by and large the nature of the initiating event remains open. It seems clear that oncogene activity assists tumor growth, but activation could occur (and be selected for) after the initiation event and during early growth of the tumor. We hope that the functions of c-onc genes will provide insights into the regulation of cell growth in normal as well as aberrant cells, so that it will become possible to define the events needed to initiate and establish tumors. Different oncogenes are associated with immortalization and transformation | SECTION 6.30.26 © 2004. Virtual Text / www.ergito.com

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Reviews 346. Hanahan, D. (1988). Dissecting multistep tumorigenesis in transgenic mice. Annu. Rev. Genet. 22, 479-519. 350. Hunter, T. (1991). Cooperation between oncogenes. Cell 64, 249-270.

References 588. Brinster, R. L. et al. (1984). Transgenic mice harboring SV40 T-antigen genes develop characteristic brain tumors. Cell 37, 367-379. 866. Stewart, T. A., Pattengale, P. K., and Leder, P. (1984). Spontaneous mammary adenocarcinomas in transgenic mice that carry and express MTV/myc fusion genes. Cell 38, 627-637. 867. Sinn, E., Muller, W., Pattengale, P., Tepler, I., Wallace, R., and Leder, P. (1987). Coexpression of MMTV/v-Ha-ras and MMTV/c-myc genes in transgenic mice: synergistic action of oncogenes in vitro. Cell 49, 465-4.



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6.30.27 p53 may affect ageing Key Concepts

• Shortening of telomeres below a critical length is associated with reduced longevity.

• Increase of p53 above wild-type levels can decrease tumor formation, but also decreases longevity.

We have very little idea what is responsible for ageing of an animal. The general drift of evolutionary theories of ageing is that natural selection operates only via reproduction, and therefore there is little advantage to the survival of the organism past the stage when it is reproductively active. In other words, there is no selection for longevity beyond the reproductive state. . We do not know how ageing of the organism relates to changes in individual cells, but one possibility is that ageing results from the accumulation of damage at the cellular level. Within this model, one contribution could be inappropriate expression of genes resulting from failure of regulation. It is an open question whether aging of the organism is connected with the senescence of individual cells. Cessation of telomerase activity in adult lineages causes telomeres to shorten as cells divide. When telomere lengths reach zero, cells enter the senescent state. Shortened telomeres can reduce lifespan. Figure 30.44 shows that mice from the fourth or fifth telomerase-negative generations have a slightly reduced lifespan, and mice from the sixth generation have a much reduced lifespan (2248). Whereas normal mice have a 50% survival rate at ~25 months, the figure for the sixth generation mice is ~17 months. Increased cancer incidence accounts for only half of the accelerated deaths. The other mice die from unknown causes (this is typical of ageing), and they prematurely show several of the characteristics of ageing (such as reduced wound healing).

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Figure 30.44 Telomerase-negative mice of the sixth generation (G6) have sharply reduced longevity, and mice of the fourth to fifth generations (G4-5) have slightly reduced longevity.

We know that loss of p53 is one of the mechanisms allowing cultured cells to pass through the crisis provoked by loss of telomeres (see Molecular Biology 6.30.24 Telomere shortening causes cell senescence). What role might p53 loss play in telomerase-negative mice? A major role is seen in the pattern of inheritance. An increased proportion of the progeny are p53–. This is because loss of p53 reduces the apoptosis (death) of germ line cells that is caused by loss of telomerase (2249). Direct attempts to see whether p53 might have any effect on ageing have been unsuccessful because p53– mice die early as the result of accumulating tumors, and mice that over-express p53 cannot be made, probably because of deleterious effects of excess p53 during embryonic development. However, striking results have been obtained from the serendipitous production of a mouse that has a mutant form of p53 (2250). The mutant gene, called the m allele, has lost its first 6 exons, and makes a truncated protein. Heterozygous p53+/m mice have a reduced frequency of tumor formation. Comparison with wild-type and hemizygous p53 mice shows: p53 +/– (one active allele):>80% tumors p53 +/+ (two active alleles): > 45% tumors p53 +/m: 6% tumors. Mice with two active alleles form tumors at about half the frequency of mice with only one active allele, which corresponds to the relative rate at which one allele is likely to be spontaneously inactivated compared to two alleles (see Figure 30.31). The much reduced rate of tumor formation in the p53+/m mice suggests that the m allele has the unexpected effect of increasing p53 activity. This is confirmed by directly measuring some of the known responses to p53 in cells from the p53+/m mice. We might expect the reduction in tumor formation in p53+/m mice to be associated with an increase in longevity, but exactly the reverse is found when they are compared with wild-type mice. Although the p53+/m mice have only 6% tumors and the wild-type mice develop 45% tumors, half of the p53+/m mice have died by 22 months, whereas the wild-type mice survive on average to 27 months. (Mice that are p53+/– or p53–/– die more quickly because of the high accumulation of tumors.) The effect seems to result from the interaction of the m mutant protein with the wild-type p53 may affect ageing | SECTION 6.30.27 © 2004. Virtual Text / www.ergito.com

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References 2248. Rudolph, K. L., Chang, S., Lee, H. W., Blasco, M., Gottlieb, G. J., Greider, C., and DePinho, R. A. (1999). Longevity, stress response, and cancer in aging telomerase-deficient mice. Cell 96, 701-712. 2249. Chin, L., Artandi, S. E., Shen, Q., Tam, A., Lee, S. L., Gottlieb, G. J., Greider, C. W., and DePinho, R. A. (1999). p53 deficiency rescues the adverse effects of telomere loss and cooperates with telomere dysfunction to accelerate carcinogenesis. Cell 97, 527-538. 2250. Tyner, S. D., Venkatachalam, S., Choi, J., Jones, S., Ghebranious, N., Igelmann, H., Lu, X., Soron, G., Cooper, B., Brayton, C., Hee Park, S., Thompson, T., Karsenty, G., Bradley, A., and Donehower, L. A. (2002). p53 mutant mice that display early ageing-associated phenotypes. Nature 415, 45-53.



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6.30.28 Genetic instability is a key event in cancer Key Terms Genetic instability (Genome instability) refers to a state in which there is large increase (×100-fold) in the frequency of changes in the genome as seen by chromosomal rearrangements or other events that affect the genetic content. This is a key occurrence in the generation of cancer cells. Key Concepts

• Tumor cells have rates of genetic change that are increased above the usual rate of somatic mutation.

• Gross chromosomal changes are observed in most types of colorectal cancer. • Chromosome rearrangements can be generated by mutations of checkpoint

pathways and other pathways that act on the genome in a yeast model system.

The inactivation of tumor suppressors and the activation of oncogenes are key events in creating a tumor, but several such events (typically 4-10 in the case of human cancers) are required to generate a fully tumorigenic state. This number of events would not be predicted to occur during the life of a cell or organism if the individual changes occurred at the normal rate of spontaneous mutation. Many cancers are associated with genetic instability that significantly increases the number of events. Genetic instability is revealed by increases in the frequency of genomic changes. These range from reorganizations at the level of the chromosome to individual point mutations. We can get a sense of their relative importance from their occurrence in colon cancers, where the genetic changes have been well characterized. The majority of colorectal tumors show high rates of gross alteration in chromosomes, often involving changes in the number of copies of a gene. This is the most common way to generate the basic changes that fuel tumorigenesis. In the minority of cases (~15%), there are no gross changes, but there are many individual mutations, resulting from a highly increased rate of mutation (see Molecular Biology 6.30.29 Defects in repair systems cause mutations to accumulate in tumors). Either of these types of change in the cell can propagate a tumor; it is rare for both to occur together. Gross chromosomal alterations involve deletion, duplication, or translocation. They may result in changes in the number of copies of a gene. Figure 30.45 illustrates two major causes:

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Figure 30.45 DNA ends induce genome rearrangements. This may happen because of the failure of either telomere maintenance or DNA damage checkpoints.

• loss of telomeres because cells continue to divide in the absence of telomerase; • creation of free double-strand ends that result from an unrepaired break in DNA. The events that occur when a cell passes through crisis are a paradigm for the generation of chromosomal alterations (for review see 2853). Loss of telomeres induces DNA rearrangements (see Molecular Biology 6.30.24 Telomere shortening causes cell senescence). Failure of the normal protective mechanisms allows the damaged cells to survive. Gross rearrangements can also be provoked by failure of protective mechanisms during a normal cell cycle (for review see 2854). Checkpoint pathways respond to DNA damage by halting the cell cycle in its current phase (see Figure 29.21 in Molecular Biology 6.29.13 DNA damage triggers a checkpoint). The checkpoint triggers an effector pathway that repairs the damage, after which the cell cycle is allowed to proceed. Mutations in the S phase checkpoint pathway in S. cerevisiae result in an increase of more than 100× in the rate of genome rearrangements Genetic instability is a key event in cancer | SECTION 6.30.28 © 2004. Virtual Text / www.ergito.com

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(2856; 2857). This happens because the cell cycle is allowed to proceed in spite of the presence of breaks in DNA. Figure 30.46 shows that similar effects are produced by mutations of some recombination-repair pathways or pathways for telomere maintenance. Analogous events could be involved in creating genetic instabilities that lead to cancer.

Figure 30.46 DNA replication errors or loss of telomeres may generate double-strand breaks in DNA. Checkpoints detect the breaks and protect the cell from perpetuating errors. If the checkpoints fail, DNA rearrangements occur.



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Reviews 2853. Maser, R. S. and DePinho, R. A. (2002). Connecting chromosomes, crisis, and cancer. Science 297, 565-569. 2854. Kolodner, R. D., Putnam, C. D., and Myung, K. (2002). Maintenance of genome stability in S. cerevisiae. Science 297, 552-557. 2856. Schar, P. (2001). Spontaneous DNA damage, genome instability, and cancer--when DNA replication escapes control. Cell 104, 329-332.

References 2857. Myung, K. and Kolodner, R. D. (2002). Suppression of genome instability by redundant S-phase checkpoint pathways in S. cerevisiae. Proc. Natl. Acad. Sci. USA 99, 4500-4507.



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6.30.29 Defects in repair systems cause mutations to accumulate in tumors Key Concepts

• Loss of mismatch-repair systems generates a high mutation rate in HNPCC.

All cells have systems to protect themselves against damage from the environment or errors that may occur during replication (see Molecular Biology 4.15 Recombination and repair). The overall mutation rate is the result of the balance between the introduction of mutations and their removal by these systems. One means by which cancer cells increase the rate of mutation is to inactivate some of their repair systems, so that spontaneous mutations accumulate instead of being removed. In effect, a mutation that occurs in a mutator gene causes mutations to accumulate in other genes. (A mutator gene can be any type of gene – such as a DNA polymerase or a repair enzyme – whose function affects the integrity of DNA sequences.) The MutSL system is a particularly important target. This system is responsible for removing mismatches in newly replicated bacterial DNA. Its homologues perform similar functions in eukaryotic cells. During replication of a microsatellite DNA, DNA polymerase may slip backward by one or more of the short repeating units. The additional unit(s) are extruded as a single-stranded region from the duplex. If not removed, they result in an increase in the length of the microsatellite in the next replication cycle (see Figure 4.28). This is averted when homologues of the MutSL system recognize the single-stranded extrusion and replace the newly-synthesized material with a nucleotide sequence that properly matches the template (see Figure 15.47). In the human disease of HNPCC (hereditary nonpolyposis colorectal cancer), new microsatellite sequences are found at a high frequency in tumor cells when their DNA sequences are compared with somatic cells of the same patient (2256; 2257). Figure 30.47 shows an example. This microsatellite has a repeat sequence of AC (reading just one strand of DNA) . The length of the repeat varies from 14-27 copies in the population. Any particular individual shows two repeat lengths, one corresponding to each allele in the diploid cell. In many patients, the repeat length is changed. Most often it is reduced at both alleles, as shown in the example in the figure.

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Figure 30.47 The normal tissue of a patient has two alleles for a microsatellite, each with a different number of repeats of the dinucleotide (AC). In tumor cells, both of these alleles have suffered deletions, one reducing the repeat number from 25 to 23, the other reducing it from 19 to 16. The repeat number of each allele in each situation is in fact probably unique, but some additional bands are generated as an artefact during the amplification procedure used to generate the samples. Data kindly provided by Bert Vogelstein. The bands remaining at the normal position in the tumor samples are due to contamination of the tumor sample with normal tissue. (From 2256).

The idea that this type of change might be the result of loss of the mismatch-repair system was confirmed by showing that mutS and mutL homologues (hMSH2, hMLH1) are mutated in the tumors (2258; 2259). As expected, the tumor cells are deficient in mismatch-repair. Change in the microsatellite sequences is of course only one of the types of mutation that result from the loss of the mismatch-repair system (it is especially easy to diagnose). The case of HNPCC illustrates both the role of multiple mutations in malignancy and the contribution that is made by mutator genes. At least 7 independent genetic events are required to form a fully tumorigenic colorectal cancer. More than 90% of cases have mutations in the mismatch-repair system, and the tumor cells have mutation rates that are elevated by 2-3 orders of magnitude from normal somatic cells (for review see 2253). The high mutation rate is responsible for creating new variants in the tumor that provide the raw material from which cells with more aggressive growth properties will arise. Several human diseases are caused by mutations in the systems that execute checkpoints, including Ataxia telangiectasia (see Molecular Biology 6.29.13 DNA damage triggers a checkpoint), Nijmegan breakage syndrome, and Bloom's syndrome, all of which are characterized by chromosomal rearrangements that are triggered by breaks in DNA. Defects in repair systems cause mutations to accumulate in tumors | SECTION 6.30.29 © 2004. Virtual Text / www.ergito.com

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Reviews 2253. Kinzler, K. W. and Vogelstein, B. (1996). Lessons from hereditary colorectal cancer. Cell 87, 159-170.

References 2256. Aaltonen, L.A., Peltomaki, P., Leach, F.S., Sistonen, P., Pylkkanen, L., Mecklin, J.P., Jarvinen, H., Powell, S.M., Jen, J., Hamilton, S.R., et al. (1993). Clues to the pathogenesis of familial colorectal cancer. Science 260, 812-816. 2257. Ionov, Y., Peinado, M. A., Malkhosyan, S., Shibata, D., and Perucho, M. (1993). Ubiquitous somatic mutations in simple repeated sequences reveal a new mechanism for colonic carcinogenesis. Nature 363, 558-561. 2258. Fishel, R., Lescoe, M. K., Rao, M. R., Copeland, N. G., Jenkins, N. A., Garber, J., Kane, M., and Kolodner, R. (1993). The human mutator gene homolog MSH2 and its association with hereditary nonpolyposis colon cancer. Cell 75, 1027-1038. 2259. Leach, F.S., Nicolaides, N.C., Papadopoulos, N., Liu, B., Jen, J., Parsons, R., Peltomaki, P., Sistonen, P., Aaltonen, L.A., Nystrom-Lahti, M., et al. (1993). Mutations of a mutS homolog in hereditary nonpolyposis colorectal cancer. Cell 75, 1215-1225.



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6.30.30 Summary A tumor cell is distinguished from a normal cell by its immortality, morphological transformation, and (sometimes) ability to metastasize. Oncogenes are identified by genetic changes that represent gain-of-functions associated with the acquisition of these properties. An oncogene may be derived from a proto-oncogene by mutations that affect its function or level of expression. Tumor suppressors are identified by loss-of-function mutations that allow increased cell proliferation. The mutations may either eliminate function of the tumor repressor or create a dominant negative version. DNA tumor viruses carry oncogenes without cellular counterparts. Their oncogenes may work by inhibiting the activities of cellular tumor suppressors. RNA tumor viruses carry v-onc genes that are derived from the mRNA transcripts of cellular (c-onc) genes. Some v-onc oncogenes represent the full length of the c-onc proto-oncogene, but others are truncated at one or both ends. Most are expressed as fusion proteins with a retroviral product. Src is an exception in which the retrovirus (RSV) is replication-competent, and the protein is expressed as an independent entity. Some v-onc genes are qualitatively different from their c-onc counterparts, since the v-onc gene is oncogenic at low levels of protein, while the c-onc gene is not active even at high levels. In such cases, proto-oncogenes are activated efficiently only by changes in the protein coding sequence. Other proto-oncogenes can be activated by large (>10× ) increases in the level of expression; c-myc is an example that can be activated quantitatively by a variety of means, including translocations with the Ig or TCR loci or insertion of retroviruses. c-onc genes may have counterpart v-onc genes in retroviruses, but some proto-oncogenes have been identified only by their association with cellular tumors. The transfection assay detects some activated c-onc sequences by their ability to transform rodent fibroblasts. ras genes are the predominant type identified by this assay. The creation of transgenic mice directly demonstrates the transforming potential of certain oncogenes. Cellular oncoproteins may be derived from several types of genes. The common feature is that each type of gene product is likely to be involved in pathways that regulate growth, and the oncoprotein has lack of regulation or increased activity. Growth factor receptors located in the plasma membrane are represented by truncated versions in v-onc genes. The protein tyrosine kinase activity of the cellular receptor is activated only when ligand binds, but the oncogenic versions have constitutive activity or altered regulation. In the same way, mutation of genes for polypeptide growth factors gives rise to oncogenes, because a receptor becomes inappropriately activated. Some oncoproteins are cytoplasmic tyrosine kinases; their targets are largely Summary | SECTION 6.30.30 © 2004. Virtual Text / www.ergito.com

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unknown. They may be activated in response to the autophosphorylation of tyrosine kinase receptors. The molecular basis for the difference between c-Src and v-Src lies in the phosphorylation states of two tyrosines. Phosphorylation of Tyr-527 in the C-terminal tail of c-Src suppresses phosphorylation of Tyr-416. The phosphorylated Tyr-527 binds to the SH2 domain of Src. However, when the SH2 domain recognizes the phosphopeptide sequence created by autophosphorylation of PDGF receptor; the PDGF receptor displaces the C-terminal region of Src, thus allowing dephosphorylation of Tyr-527, with the consequent phosphorylation of Tyr-416 and activation of the kinase activity. v-Src has lost the repressive C-terminus that includes Tyr-527, and therefore has permanently phosphorylated Tyr-416, and is constitutively active. Ras proteins can bind GTP and are related to the α subunits of G proteins involved in signal transduction across the cell membrane. Oncogenic variants have reduced GTPase activity, and therefore are constitutively active. Activation of Ras is an obligatory step in a signal transduction cascade that is initiated by activation of a tyrosine kinase receptor such as the EGF receptor; the cascade passes to the ERK MAP kinase, which is a serine/threonine kinase, and terminates with the nuclear phosphorylation of transcription factors including Fos. Nuclear oncoproteins may be involved directly in regulating gene expression, and include Jun and Fos, which are part of the AP1 transcription factor. v-ErbA is derived from another transcription factor, the thyroid hormone receptor, and is a dominant negative mutant that prevents the cellular factor from functioning. v-Rel is related to the common factor NF- κB, and influences the set of genes that are activated by transcription factors in this family. Retinoblastoma (RB) arises when both copies of the RB gene are deleted or inactivated. The RB product is a nuclear phosphoprotein whose state of phosphorylation controls entry into S phase. Nonphosphorylated RB sequesters the transcription factor E2F. The RB-E2F complex represses certain target genes. E2F is released when RB is phosphorylated by cyclin/cdk complexes; E2F can then activate genes whose products are needed for S phase. Loss of RB prevents repression by RB-E2F, and means that E2F is constitutively available. The cell cannot be restrained from proceeding through the cycle. Adenovirus E1A and papova virus T antigens bind to nonphosphorylated RB, and thus prevent it from binding to E2F. p53 was originally classified as an oncogene because missense mutations in it are oncogenic. It is now classified as a tumor suppressor because the missense mutants in fact function by inhibiting the activity of wild-type p53. The same phenotype is produced by loss of both wild-type alleles. The level of p53 is usually low, but in response to damage to DNA, p53 activity increases, and triggers either of two pathways, depending upon the stage of the cell cycle and the cell phenotype. Early in the cycle, it provides a checkpoint that prevents further progress; this allows damaged DNA to be repaired before replication. Later in the cycle, it causes apoptosis, so that the cell with damaged DNA dies instead of perpetuating itself. Loss of p53 function is common in established cell lines and may be important in immortalization in vitro. Absence of p53 is common in human tumors and may contribute to the progression of a wide variety of tumors, without specificity for cell type. p53 is activated by binding to damaged DNA, for which it uses a (non Summary | SECTION 6.30.30 © 2004. Virtual Text / www.ergito.com

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sequence-specific) DNA-binding domain. One important target that activates p53 is the single-stranded overhanging end that is generated at a shortened telomere. When it is activated, p53 uses another DNA-binding domain to recognize a palindromic ~10 bp sequence. Genes whose promoters have this sequence and which are activated by p53 include the cdk inhibitor p21 and the protein GADD45 (which is activated by several pathways for response to DNA damage). Activation of these and other genes (involving a transactivation domain that interacts directly with TBP) is probably the means by which p53 causes cell cycle arrest. p53 has a less well characterized ability to repress some genes. Mutant p53 lacks these activities, and therefore allows the perpetuation of cells with damaged DNA. Loss of p53 may be associated with increased amplification of DNA sequences. p53 is bound by viral oncogenes such as SV40 T antigen, whose oncogenic properties result, at least in part, from the ability to block p53 function. It is also bound by the cellular proto-oncogene, Mdm2, which inhibits its activity. p53 and Mdm2 are mutual antagonists. The locus INK4A contains two tumor suppressors that together control both major tumor suppressor pathways. p19ARF inhibits Mdm2, so that p19 in effect turns on p53. p16INK4A inhibits the cdk4/6 kinase, which phosphorylates RB. Deletion of INK4A therefore blocks both tumor suppressor pathways by leading to activation of Mdm2 (inhibiting p53) and activation of cdk4/6 (inhibiting RB). Loss of p53 may be necessary for immortalization, because both the G1 checkpoint and the trigger for apoptosis are inactivated. Telomerase is usually turned off in differentiating cells, which provides a mechanism of tumor suppression by preventing indefinite growth. Reactivation of telomerase is usually necessary to allow continued proliferation of tumor cells. The crisis that is encountered by cultured cells results from shortening of telomeres to the point at which genetic instability is created by the chromosome ends. Loss of p53 is important in passing through crisis, because otherwise p53 is activated by the ends generated by telomere loss. Several independent events are required to convert a normal cell into a cancer cell, typically involving both immortalizing and transforming functions. The required number of events is in the range of 4-10, and would not normally be expected to occur during the life span of a cell. Early events may increase the rate of occurrence of mutational change by damaging the repair or other systems that limit mutational damage. One important target is the MutSL system that is responsible for removing mismatches in replicated DNA. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.30.30


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GRADIENTS, CASCADES, AND SIGNALING PATHWAYS

6.31.1 Introduction Key Terms Axes are straight lines passing through an organism, around which the organism is symmetrically arranged. The anterior-posterior axis is the line running from the head to the tail of an animal. The dorsal-ventral axis is the line running from the back to the belly of an animal.

Development begins with a single fertilized egg, but gives rise to cells that have different developmental fates. The problem of early development is to understand how this asymmetry is introduced: how does a single initial cell give rise within a few cell divisions to progeny cells that have different properties from one another? The means by which asymmetry is generated varies with the type of organism. The egg itself may be homogeneous, with the acquisition of asymmetry depending on the process of the initial division cycles, as in the case of mammals. Or the egg may have an initial asymmetry in the distribution of its cytoplasmic components, which in turn gives rise to further differences as development proceeds, as in the case of Drosophila. Early development is defined by the formation of axes. By whatever means are used to develop asymmetry, the early embryo develops differences along the anterior-posterior axis (head-tail) and along the dorsal-ventral axis (top-bottom). At the stage of interpreting the axial information, a relatively restricted set of signaling pathways is employed, and essentially the same pathways are found in flies and mammals. The paradigm for considering the molecular basis for development is to suppose that each cell type may be characterized by its pattern of gene expression, that is, by the particular gene products that it produces. The principal level for controlling gene expression is at transcription, and components of pathways regulating transcription provide an important class of developmental regulators. We may include a variety of activities within the rubric of transcriptional regulators, which could act to change the structure of a promoter region, to initiate transcription at a promoter, to regulate the activity of an enhancer, or indeed sometimes to repress the action of transcription factors. However, the regulators of transcription most often prove to be DNA-binding proteins that activate transcription at particular promoters or enhancers. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.31.1


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6.31.2 Fly development uses a cascade of transcription factors Key Terms A maternal gene (maternal effect gene) (maternal-effect gene) is usually expressed by the mother during oogenesis. Phenotypes resulting from a maternal-effect mutation depend on the genotype of the mother rather than the genotype of the embryo. Segmentation genes are concerned with controlling the number or polarity of body segments in insects. Homeotic genes are defined by mutations that convert one body part into another; for example, an insect leg may replace an antenna. Key Concepts

• The genes that control the early stages of fly development code for transcription factors.

• At each stage, the factors in one area of the egg control the synthesis of further factors that will define smaller areas.

• Maternal genes are expressed during oogenesis and act in the oocyte. • Three successive groups of segmentation genes are expressed after fertilization to control the number or polarity of segments.

• Homeotic genes control the identity of a segment.

The systematic manner in which the regulators are turned on and off to form circuits that determine body parts has been worked out in detail in D. melanogaster. The basic principle is that a series of events resulting from the initial asymmetry of the egg is translated into the control of gene expression so that specific regions of the egg acquire different properties. The means by which asymmetry is translated into control of gene expression differ for each of four systems that have been characterized in the insect egg. It may involve localization of factors that control transcription or translation within the egg, or localized control of the activities of such factors. But the end result is the same: spatial and temporal regulation of gene expression. Early in development, the identities of parts of the embryo are determined: regions are defined whose descendants will form particular body parts. The genes that regulate this process are identified by loci in which mutations cause a body part to be absent, to be duplicated, or to develop as another body part. Such loci are prime candidates for genes whose function is to provide regulatory "switches." Most of these genes code for regulators of transcription. They act upon one another in a hierarchical manner, but they act also upon other genes whose products are actually Fly development uses a cascade of transcription factors | SECTION 6.31.2 © 2004. Virtual Text / www.ergito.com

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responsible for the formation of pattern. The ultimate targets are genes that code for kinases, cytoskeletal elements, secreted proteins, and transmembrane receptors. The establishment of a specific pattern of transcription in a particular region of the embryo leads to a cascade of control, when regulatory events are connected so that a gene turned on (or off) at one stage itself controls expression of other genes at the next stage. Formally, such a cascade resembles those described previously for bacteriophages or for bacterial sporulation (as discussed in Molecular Biology 3.10 The operon), although it is more complex in the case of eukaryotic development. The common feature of regulatory proteins is that they are transcription factors that regulate the expression of other transcription factors (as well as other target proteins). As in the case of prokaryotic regulation, the basic relationship between the regulator protein and the target gene is that the regulator recognizes a short sequence in the DNA of the promoter (or an enhancer) of a target gene. All of the targets for a particular regulator are identified by their possession of a copy of the appropriate consensus sequence. The development of an adult organism from a fertilized egg follows a predetermined pathway, in which specific genes are turned on and off at particular times. From the perspective of mechanism, we have most information about the control of transcription. However, subsequent stages of gene expression are also targets for regulation. And, of course, the cascade of gene regulation is connected to other types of signaling, including cell-cell interactions that define boundaries between groups of cells. The mechanics of development in terms of cellular events are different in different types of species, but we assume that the principle established with Drosophila will hold in all cases: that a regulatory cascade determines the appropriate pattern of gene expression in cells of the embryo and ultimately of the adult. Indeed, homologous genes in distantly related organisms play related roles in development. The same pathways are found in (for example) flies and mammals, although the consequences of their employment are rather different in terms of the structures that develop. Genes involved in regulating development are identified by mutations that are lethal early in development or that cause the development of abnormal structures. A mutation that affects the development of a particular body part attracts our attention because a single body part is a complex structure, requiring expression of a particular set of many genes. Single mutations that influence the structure of the entire body part therefore identify potential regulator genes that switch or select between developmental pathways. In Drosophila, the body part that is analyzed is the segment, the basic unit that can be seen looking at the adult fly. Mutations fall into (at least) three groups, defined by their effect on the segmental structure: • Maternal genes are expressed during oogenesis by the mother. They may act upon or within the maturing oocyte. • Segmentation genes are expressed after fertilization. Mutations in these genes alter the number or polarity of segments. Three groups of segmentation genes act sequentially to define increasingly smaller regions of the embryo. Fly development uses a cascade of transcription factors | SECTION 6.31.2 © 2004. Virtual Text / www.ergito.com

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• Homeotic genes control the identity of a segment, but do not affect the number, polarity, or size of segments. Mutations in these genes cause one body part to develop the phenotype of another part. The genes in each group act successively to define the properties of increasingly more restricted parts of the embryo. The maternal genes define broad regions in the egg; differences in the distribution of maternal gene products control the expression of segmentation genes; and the homeotic genes determine the identities of individual segments (for review see 355; 362).

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Reviews 355. Mahowald, A. P. and Hardy, P. A. (1985). Genetics of Drosophila embryogenesis. Annu. Rev. Genet. 19, 149-177. 362. Lawrence, P. (1992). . The Making of a Fly.


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6.31.3 A gradient must be converted into discrete compartments Key Terms A denticle is a pigmented, hardened spike of cuticle protruding from the ventral epidermis of a Drosophila embryo. Key Concepts

• During the first 13 division cycles, nuclei divide in a common cytoplasm; cells form only at blastoderm.

• Gradients define the polarity of the egg along both the anterior-posterior and dorsal-ventral axes.

• The gradients consist of RNAs or proteins that are differentially distributed in the common cytoplasm.

• The location of a nucleus in the cytoplasm with regard to the two axes determines the fate of the cells that descend from it.

The basic question of Drosophila development is illustrated in Figure 31.1 in terms of three stages of development: the egg; the larva; and the adult fly.

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Figure 31.1 Gradients in the egg are translated into segments on the anterior-posterior axis and into specialized structures on the dorsal-ventral axis of the larva, and then into the segmented structure of the adult fly.

At the start of development, gradients are established in the egg along the anterior-posterior and dorsal-ventral axes. The anterior end of the egg becomes the head of the adult; the posterior end becomes the tail. The dorsal side is on top (looking down on a larva); the ventral side is underneath. The gradients consist of molecules (proteins or RNAs) that are differentially distributed in the cytoplasm. The gradient responsible for anterior-posterior development is established soon after fertilization; the dorsal-ventral gradient is established a little later. It is only a modest oversimplification to say that the anterior-posterior systems control positional information along the larva, while the dorsal-ventral system regulates tissue differentiation (that is, the specification of distinct embryonic tissues, including mesoderm, neuroectoderm, and dorsal ectoderm).


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Insect development involves two quite different types of structures. The first part of development is concerned with elaborating the larva; then the larva metamorphoses into the fly. This means that the structure of the embryo (the larva) is distinct from the structure of the adult (the fly), in contrast with development of (for example) mammals, where the embryo develops the same body parts that are found in the adult. As the larva develops, it forms some body parts that are exclusively larval (they will not give rise to adult tissues; often they are polyploid), while other body parts are the progenitors that will metamorphose into adult structures (usually they are diploid). In spite of the differences between insect development and vertebrate development, the same general principles appear to govern both processes, and we discover relationships between Drosophila regulators and mammalian regulators. Discrete regions in the embryo correspond to parts of the adult body. They are shown in terms of the superficial organization of the larva in the middle panel of Figure 31.1. Bands of denticles (small hairs) are found in a particular pattern on the surface (cuticle) of the larva. The cuticular pattern has features determined by both the anterior-posterior axis and the dorsal-ventral axis: • Along the anterior-posterior axis, the denticles form discrete bands. Each band corresponds to a segment of the adult fly: in fact, the 11 bands of denticles correspond on a 1:1 basis with the 11 segments of the adult. • Along the dorsal-ventral axis, the denticles that extend from the ventral surface are coarse; those that extend from the dorsal surface are much finer. Although the cuticle represents only the surface body layer, its structure is diagnostic of the overall organization of the embryo in both axes. Much of the analysis of phenotypes of mutants in Drosophila development has therefore been performed in terms of the distortion of the denticle patterns along one axis or the other. The difference in form between the gradients of the egg and the segments of the adult poses some prime questions. How are gradients established in the egg? And how is a continuous gradient converted into discrete differences that define individual cell types? How can a large number of separate compartments develop from a single gradient? The nature of the gradients, and their ability to affect the development of a variety of cell types located throughout the embryo, depend upon some idiosyncratic features of Drosophila development. The early stages are summarized in Figure 31.2.


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Figure 31.2 The early development of the Drosophila egg occurs in a common cytoplasm until the stage of cellular blastoderm.

At fertilization the egg possesses the two parental nuclei and is distinguished at the posterior end by the presence of a region called polar plasm. For the first 9 divisions, the nuclei divide in the common cytoplasm. Material can diffuse in this cytoplasm (although there are probably constraints imposed by cytoskeletal organization). At division 7, some nuclei migrate into the polar plasm, where they become precursors to germ cells. After division 9, nuclei migrate and divide to form a layer at the surface of the egg. Then they divide 4 times, after which membranes surround them to form somatic cells. Up to the point of cellularization, the nuclei effectively reside in a common cytoplasm. At the stage of the cellular blastoderm, the first discrete compartments become evident, and at this time particular regions of the egg are determined to A gradient must be converted into discrete compartments | SECTION 6.31.3 © 2004. Virtual Text / www.ergito.com

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become particular types of adult structures. (Determination is progressive and gradual; over the next few cell divisions, the fates of individual regions of the egg become increasingly restricted.) At the start of this process, nuclei migrate to the surface to form the monolayer of the blastoderm, but they do not do so in any predefined manner. It is therefore the location in which the nuclei find themselves at this stage that determines what types of cells their descendants will become. A nucleus determines its position in the embryo by reference to the anterior-posterior and dorsal-ventral gradients, and behaves accordingly. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.31.3


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6.31.4 Maternal gene products establish gradients in early embryogenesis Key Terms In Drosophila, a female sterile mutation is one in that causes sterility in the female, often because of abnormalities in oogenesis. A morphogen is a factor that induces development of particular cell types in a manner that depends on its concentration. In Drosophila, the anterior system is one of the maternal systems that establishes the polarity of the oocyte. The set of genes in the anterior system play a role in the proper formation of the head and the thorax. In Drosophila, the posterior system is one of the maternal systems that establishes the polarity of the oocyte. The set of genes in the posterior system play a role in the proper formation of the pole plasm and the abdomen. In Drosophila, the terminal system is one of the maternal systems that establishes the polarity of the oocyte. The set of genes in the terminal system play a role in the proper formation of the terminal structures at both ends of the fly. Key Concepts

• Four signaling pathways are initiated outside the egg and each leads to production of a morphogen in the egg.

• The anterior system is responsible for development of head and thorax. • The posterior system is responsible for the segments of the abdomen. • The terminal system is responsible for producing the acron (in the head) and the telson (at the tail).

• The dorsal-ventral system determines development of tissue types (mesoderm, neuroectoderm, ectoderm).

An initial asymmetry is imposed on the Drosophila oocyte during oogenesis. Figure 31.3 illustrates the structure of a follicle in the Drosophila ovary. A single progenitor undergoes four successive mitoses to generate 16 interconnected cells. The connections are known as "cytoplasmic bridges" or "ring canals." Individual cells have 2, 3, or 4 such connections. One of the two cells that has 4 connections undergoes meiosis to become the oocyte; the other 15 cells become "nurse cells." Cytoplasmic material, including protein and RNA, passes from the nurse cells to the oocyte; the accumulation of such material accounts for a considerable part of the volume of the egg. The cytoplasmic connections are made at one end of the oocyte, and this end becomes the anterior end of the egg.

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Figure 31.3 A Drosophila follicle contains an outer surface of follicle cells that surround nurse cells that are in close contact with the oocyte. Nurse cells are connected by cytoplasmic bridges to each other and to the anterior end of the oocyte. Follicle cells are somatic; nurse cells and the oocyte are germline in origin.

Genes that are expressed within the mother fly are important for early development. These maternal genes are identified by female sterile mutations. They do not affect the mother itself, but are required in order to have progeny. Females with such mutations lay eggs that fail to develop into adults; the embryos can be recognized by defects in the cuticular pattern, and they die during development. The common feature in all maternal genes is that they are expressed prior to fertilization (although their products may act either at the time of expression or be stored for later use). The maternal genes are divided into two classes, depending on their site of expression. Genes that are expressed in somatic cells of the mother that affect egg development are called maternal somatic genes. For example, they may act in the follicle cells. Genes that are expressed within the germline are called maternal germline genes. These genes may act either in the nurse cell or the oocyte. Some genes act at both stages. Four groups of genes concerned with the development of particular regions of the embryo can be identified by mutations in maternal genes. The genes in each group can be organized into a pathway that reflects their order of action, by conventional genetic tests (such as comparing the properties of double mutants with the individual mutants) or by biochemical assays (showing which mutants contain components that can bypass the stages that are blocked in other mutants; for review see 355; 362). The components of these pathways are summarized in Figure 31.4, which shows that there is a common principle to their operation. Each pathway is initiated by localized events outside the egg; this results in the localization of a signal within the egg. This signal takes the form of a protein with an asymmetric distribution; this is called a morphogen. Formally, we may define a morphogen as a protein whose local concentration (or activity) causes the surrounding region to take up a particular structure or fate. In each of these systems, the morphogen either is a transcriptional regulator or leads to the activation of a transcription factor in the localized region. Three systems are concerned with the anterior-posterior axis, and one with the dorsal-ventral axis: Maternal gene products establish gradients in early embryogenesis | SECTION 6.31.4 © 2004. Virtual Text / www.ergito.com

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Figure 31.4 Each of the four maternal systems that functions in the egg is initiated outside the egg. The pathway is carried into the egg, where each pathway has a localized product that is the morphogen. This may be a receptor or a regulator of gene expression. The final component is a transcription factor, which acts on zygotic targets that are responsible for the next stage of development.

• The anterior system is responsible for development of the head and thorax. The maternal germline products are required to localize the bicoid product at the anterior end of the egg. In fact, bicoid mRNA is transcribed in nurse cells and transported into the oocyte. Bicoid protein is the morphogen: it functions as a transcriptional regulator, and controls expression of the gene hunchback (and probably also other segmentation and homeotic genes). • The posterior system is responsible for the segments of the abdomen. The nature of the initial asymmetric event is not clear. A large number of products act to cause the localization of the product of nanos, which is the morphogen. This leads to localized repression of expression of hunchback (via control of translation of the mRNA). • The terminal system is responsible for development of the specialized structures at the unsegmented ends of the egg (the acron at the head, and the telson at the tail). As indicated by the dependence on maternal somatic genes, the initial events that create asymmetry occur in the follicle cells. They lead to localized activation of the transmembrane receptor coded by torso; the end product of the pathway has yet to be identified. • The fourth system is responsible for dorsal-ventral development. The pathway is initiated by a signal from a follicle cell on the ventral side of the egg. It is transmitted through the transmembrane receptor coded by Toll. This leads to a gradient of activation of the transcription factor produced by dorsal (by Maternal gene products establish gradients in early embryogenesis | SECTION 6.31.4 © 2004. Virtual Text / www.ergito.com

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controlling its localization within the cell). About 30 maternal genes involved in pattern formation have been identified. All of the components of the four pathways are maternal, so we see that the systems for establishing the initial pattern formation all depend on events that occur prior to fertilization. The two body axes are established independently. Mutations that affect polarity cause posterior regions to develop as anterior structures, or ventral regions to develop in dorsal form. On the anterior-posterior axis, the anterior and posterior systems provide opposing gradients, with sources at the anterior and posterior ends of the embryo, respectively, that control development of the segments of the body. Defects in either system affect the body segments. The terminal and dorsal-ventral systems operate independently of the other systems (for review see 359).


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Reviews 355. Mahowald, A. P. and Hardy, P. A. (1985). Genetics of Drosophila embryogenesis. Annu. Rev. Genet. 19, 149-177. 359. Ingham, P. W. (1988). The molecular genetics of embryonic pattern formation in Drosophila. Nature 335, 25-34. 362. Lawrence, P. (1992). . The Making of a Fly.



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6.31.5 Anterior development uses localized gene regulators Key Terms Ectopic refers to something being out of place. A gene or protein that plays an instructive role in development is one that gives a signal telling the cell what to do. Key Concepts

• The anterior system localizes bicoid mRNA at the anterior end of the egg,

generating a gradient of protein that extends along the anterior 40% of the egg.

• The concentration of bicoid protein determines the types of anterior (head) structures that are produced in each region.

• This system is instructive because it is required for development of the head structures.

Establishing asymmetry in an egg requires that some components – either RNAs or proteins – are localized instead of being diffused evenly through the cytosol. In anterior-posterior development in Drosophila, certain mRNAs are localized at the anterior or posterior end. Figure 31.5 shows that when they are translated, their protein products diffuse away from the ends of the egg, generating a gradient along the anterior-posterior axis.

Anterior development uses localized gene regulators | SECTION 6.31.5 © 2004. Virtual Text / www.ergito.com

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Figure 31.5 Translation of a localized mRNA generates a gradient of protein as the products diffuses away from the site of synthesis.

The existence of localized concentrations of materials needed for development can be tested by the rescue protocol summarized in Figure 31.6. Material is removed from a wild-type embryo and injected into the embryo of a mutant that is defective in early development. If the mutant embryo develops normally, we may conclude that the mutation causes a deficiency of material that is present in the wild-type embryo. This allows us to distinguish components that are necessary for morphogenesis, or that are upstream in the pathway, from the morphogen itself – only the morphogen has the property of localized rescue.


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Figure 31.6 Mutant embryos that cannot develop can be rescued by injecting cytoplasm taken from a wild-type embryo. The donor can be tested for time of appearance and location of the rescuing activity; the recipient can be tested for time at which it is susceptible to rescue and the effects of injecting material at different locations.

The rescue technique identifies bicoid as the morphogen required for anterior development. bicoid mutants do not develop heads; but the defect can be remedied by injecting mutant eggs with cytoplasm taken from the anterior tip of a wild-type embryo. Indeed, anterior structures develop elsewhere in the mutant embryo if wild-type anterior cytoplasm is injected! (This is called ectopic expression.) The extent of the rescue depends on the amount of wild-type cytoplasm injected. And the efficacy of the donor cytoplasm depends on the number of wild-type bicoid genes carried by the donor. These results suggest that the anterior region of a wild-type embryo contains a concentration of some product that depends on the bicoid gene dosage. By purifying the active component in the preparation, it is possible to show that purified bicoid mRNA can substitute for the anterior cytoplasm. This implies that the components on which bicoid acts are ubiquitous, and all that is required to trigger formation of anterior structures is an appropriate concentration of bicoid product. The product of bicoid establishes a gradient with its source (and therefore the highest concentration) at the anterior end of the embryo. The RNA is localized at the anterior tip of the embryo, but it is not translated during oogenesis. Translation begins soon after fertilization. The protein then establishes a gradient along the embryo, as indicated in Figure 31.7 (888). The gradient could be produced by diffusion of the Anterior development uses localized gene regulators | SECTION 6.31.5 © 2004. Virtual Text / www.ergito.com

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protein product from the localized source at the anterior tip. The gradient is established by division 7, and remains stable until after the blastoderm stage.

Figure 31.7 Bicoid protein forms a gradient during D. melanogaster development that extends for ~200 mm along the egg of 500 mm.

What is the consequence of establishing the bicoid gradient? The gradient can be increased or decreased by changing the number of functional gene copies in the mother. The concentration of bicoid protein is correlated with the development of anterior structures. Weakening the gradient causes anterior segments to develop more posterior-like characteristics; strengthening the gradient causes anterior-like structures to extend farther along the embryo. So the bicoid protein behaves as a morphogen that determines anterior-posterior position in the embryo in a concentration-dependent manner. The fate of cells in the anterior part of the embryo is determined by the concentration of bicoid protein. The bicoid product is a sequence-specific DNA-binding protein that regulates transcription by binding to the promoters of its target genes. The immediate effect of bicoid is exercised on other genes that in turn regulate the development of yet further genes. A major target for bicoid is the gene hunchback. Transcription of hunchback is turned on by bicoid in a dose-dependent manner, that is, hunchback is activated above a certain threshold of bicoid protein. The effect of bicoid on hunchback is to produce a band of expression that occupies the anterior part of the embryo (see Figure 31.21; for review see 365). The relationship between bicoid and hunchback establishes the principle that a gradient can provide a spatial on-off switch that affects gene expression. In this way, quantitative differences in the amount of the morphogen (bicoid protein) are transformed into qualitatively different states (cell structures) during embryonic development. bicoid plays an instructive role in anterior development, since it is a positive regulator that is needed for expression of genes that in turn determine the synthesis of anterior structures (for review see 3699). Anterior development uses localized gene regulators | SECTION 6.31.5 © 2004. Virtual Text / www.ergito.com

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Reviews 365. McGinnis, W. and Krumlauf, R. (1992). Homeobox genes and axial patterning. Cell 68, 283-302. 3699. Lawrence, P. A. and Struhl, G. (1996). Morphogens, compartments, and pattern: lessons from Drosophila? Cell 85, 951-961.

References 888. Driever, W. and Nusslein-Volhard, C. (1988). The bicoid protein determines position in the Drosophila embryo in a concentration dependent manner. Cell 54, 95-104.



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6.31.6 Posterior development uses another localized regulator Key Terms A protein that plays a permissive role in development is one that sets up a situation where a certain activity can occur, but does not cause the occurrence itself. Key Concepts

• The posterior system localizes nanos mRNA at the posterior end of the egg,

generating a gradient of nanos protein that extends along the abdominal region.

• This system is permissive because its function is to repress genes whose products would interfere with posterior development.

Posterior development depends on the expression of a large group of genes. Embryos produced by females who are mutant for any one of these genes develop normal head and thoracic segments, but lack the entire abdomen. Some of these genes are concerned with exporting material from the nurse cells to the egg; others are required to transport or to localize the material within the egg. The posterior pathway functions by a series of events in which one product is responsible for localizing the next. Figure 31.8 correlates the order of genes in the genetic pathway with the activities of their products in the embryo. The functions spir and capu are needed for Staufen protein to be localized at the pole. Staufen protein in turn localizes oskar RNA; possibly a complex of Staufen protein and oskar RNA is assembled. These functions are needed to localize Vasa, which is an RNA-binding protein. Its specificity and targets are not known.

Posterior development uses another localized regulator | SECTION 6.31.6 © 2004. Virtual Text / www.ergito.com

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Figure 31.8 The posterior pathway has two branches, responsible for abdominal development and germ cell formation.

If oskar is over-expressed or mislocalized in the embryo, it induces germ cell formation at ectopic sites. It requires only the products of vasa and tudor. This implies that all of the activities that precede oskar in the pathway are needed only to localize oskar RNA. The ability both to form pole cells and to induce abdominal structures is possessed by oskar, in conjunction with vasa and tudor (and of course any components that are ubiquitously expressed in the egg). One effect of oskar function is to localize Vasa protein at the posterior end. The functions of valois and tudor are not known, but it is possible that valois is off the main pathway. Two types of pattern-determining event occur at the posterior pole, and the pathway branches at tudor. The polar plasm contains two morphogens: the posterior determinant (nanos) controls abdominal development; and another signal controls formation of the pole cells, which will give rise to the germline (see Figure 31.2). All of the posterior genes except nanos and pumilio are required for both processes, that is, they are defective in both abdominal development and pole cell formation. nanos and pumilio identify the abdominal branch. We do not know whether there are additional functions representing a separate branch for germ cell formation, or whether the pathway up to tudor is by itself sufficient (901). The posterior system resembles the anterior system in the basic nature of the morphogenetic event: a maternal mRNA is localized at the posterior pole. This is the product of nanos, and provides the morphogen. There are two important differences between the systems. Localization is more complex than in the case of the anterior system, because posterior determinants that originate in the nurse cells must be Posterior development uses another localized regulator | SECTION 6.31.6 © 2004. Virtual Text / www.ergito.com

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transported the full length of the oocyte to the far pole. And nanos protein acts to prevent translation of a transcription factor (hunchback). Its role is said to be permissive, since it functions to repress genes whose products would interfere with posterior development. How do we know that nanos is the morphogen at the end of the pathway? Rescue experiments (along the lines shown previously in Figure 31.6) with the mutants in the posterior group showed that in all but one case the cytoplasm of the nurse cell contained the posterior determinant (although it was absent from the posterior end of the oocyte itself). This indicates that these mutants all act in some subsidiary role, most probably concerned with transporting or localizing the morphogen in the egg. The exception was nanos, whose mutants did not contain any posterior-rescuing activity. Purified nanos RNA can rescue mutants in any of the other posterior genes, indicating that it is the last, or most downstream, component in the pathway. Indeed, injection of nanos RNA into ectopic locations in embryos can induce the formation of abdominal structures, showing that it provides the morphogen. The upper part of Figure 31.9 shows the localization of nanos mRNA at the posterior end of an early embryo. But the localization poses a dilemma: nanos activity is required for development of abdominal segments, that is, for structures occupying approximately the posterior half of the embryo. How does nanos RNA at the pole control abdominal development? The lower part of Figure 31.9 shows that nanos protein diffuses from the site of translation to form a gradient that extends along the abdominal region.


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Figure 31.9 nanos products are localized at the posterior end of a Drosophila embryo. The upper photograph shows the tightly localized RNA in the very early embryo (at the time of the 3rd nuclear division). The lower photograph shows the spreading of nanos protein at the 8th nuclear division. Photographs kindly provided by Ruth Lehmann.

Both bicoid and nanos act on the expression of the hunchback gene. hunchback codes for a repressor of transcription: its presence is needed for formation of anterior structures (in the region of the thorax), and its absence is required for development of posterior structures. It has a complex pattern of expression. It is transcribed during oogenesis to give an mRNA that is uniformly distributed in the egg. After fertilization, the hunchback pattern is changed in two ways. The bicoid gradient activates synthesis of hunchback RNA in the anterior region. And nanos prevents translation of hunchback mRNA in the posterior region; a result of this inhibition is that the mRNA is degraded. The anterior and posterior systems together therefore enhance hunchback levels in the anterior half of the egg, and remove it from the posterior half. The significance of this distribution lies with the genes that hunchback regulates. It represses the genes knirps and (probably) giant, which are needed to form abdominal structures. So the basic role of hunchback is to repress formation of abdominal structures by preventing the expression of knirps and giant in more anterior regions (894).


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References 894. Struhl, G., Johnston, P., and Lawrence, P. A. (1992). Control of Drosophila body pattern by the hunchback morphogen gradient. Cell 69, 237-249. 901. Sprenger, F. and Nusslein-Volhard, C. (1992). Cellular terminal regions of the Drosophila egg. Cell 71, 987-1001.



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6.31.7 How are mRNAs and proteins transported and localized? Key Concepts

• The mRNAs that establish the anterior and posterior systems are transcribed in nurse cells and transported through cytoplasmic bridges into oocytes.

• bicoid mRNA is localized close to the point of entry, but oskar and nanos mRNAs are transported the length of the oocyte to the posterior end.

• Movement is accomplished by a motor attached to microtubules.

Anterior and posterior development both depend on the localization of an mRNA at one end of the egg. How does the mRNA reach the appropriate location and what is responsible for maintaining it there? Similar processes are involved for the anterior-posterior axis and for the dorsal-ventral axis. On the antero-posterior axis, bicoid and oskar mRNAs are localized at opposite ends of the egg. On the dorsal-ventral axis, gurken mRNA is initially localized at the posterior end and then becomes localized on the dorsal side of the anterior end. The principle is that the sites of transcription are distinct from the sites where the mRNAs are localized and translated, and an active transport process is required to localize the mRNAs. bicoid, oskar, and nanos all are transcribed in nurse cells. Figure 31.10 shows that mRNA is transported through the cytoplasmic bridges into the oocyte. Within the oocyte, bicoid mRNA then remains at the anterior end, but oskar mRNA is transported the length of the oocyte to the posterior end. The typical means by which an mRNA is transported to a specific location in a cell involves movement along "tracks", which in principle can be either actin filaments or microtubules. This basically means that the mRNA is attached to the tracks by a motor protein that uses hydrolysis of ATP to drive movement (see Figure 31.10). In the example of the Drosophila egg, microtubules are the tracks used to transport these and other mRNAs (960). In fact, the microtubules form a continuous network that connects the oocyte to the nurse cells through the ring canals (2233).

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Figure 31.10 Some mRNAs are transported into the Drosophila egg as ribonucleoprotein particles. They move to their final sites of localization by association with microtubule.

Genes whose products are needed to transport these mRNAs are identified by mutants in which the mRNAs are not properly localized. The most typical disruption of the pattern is for the mRNAs simply to be distributed throughout the egg. The best characterized of these transport genes are exuperantia (exu) and swallow (swa). Exu protein is part of a large ribonucleoprotein complex (961). This complex assembles in the nurse cell, where it uses microtubule tracks to move to the cytoplasmic bridge (962). Then it passes across the bridge into the oocyte in a way that is independent of microtubules. In the oocyte, it attaches to microtubules to move to its location. The properties of exu and swa mutants show that there are common components for the transport and localization of different mRNAs. We do not yet know what differences exist between the complexes involved in transporting different mRNAs. However, we assume that there must be a component of each complex that is responsible for targeting it to the right location. By following different mRNAs, it seems that in each case the complex is transported to the anterior end of the oocyte, where it aggregates. Then a decision is made on further localization, and the complex is transported to the appropriate site (for review see 2295; 2304). Similar events occur at a later stage of development, in the syncytial blastoderm, when some mRNAs become localized on the apical side of the embryo. The same How are mRNAs and proteins transported and localized? | SECTION 6.31.7 © 2004. Virtual Text / www.ergito.com

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apparatus seems to be involved as in development of the oocyte. Usually it is responsible for apical localization of the products of several pair rule and segmentation genes. However, if the maternal transcripts of gurken, bicoid, or nanos are injected into the syncytial blastoderm, the apparatus localizes them on the apical side of the embryo (2235). This suggests that the RNAs that are localized at early and at later times have the same set of signals to identify themselves as substrates to the localization apparatus. mRNAs that are localized in the syncytial blastoderm are found in particles that are connected to microtubules by the motor dynein (2234). The parallels between the transport systems of the oocyte and the blastoderm suggest that dynein also connects the maternal mRNAs to the microtubules. The proteins Egl (egalitarian) and BicD (Bicaudal-D) associate with localizing transcripts and bind to dynein (2235). In their absence, maternal transcripts do not localize properly in the oocyte. This suggests that an Egl/BicD complex may be the means of connecting the mRNA to the motor. We know that the localization of the bicoid RNA to the anterior end of the oocyte depends upon sequences in the 3 ′ untranslated region (887; 888). This is a common theme, and localization of oskar and nanos mRNAs is controlled in the same way (963). We assume that the 3 ′ sequences provide binding sites for specific protein(s) that are involved in localization. Corresponding sequences in each mRNA will provide binding sites for the proteins that target the RNA to the appropriate sites in the oocyte. However, we are still missing the identification of the crucial protein that binds to the localizing sequence in the mRNA. Localization of RNAs is not sufficient to ensure the pattern of expression. Translation is also controlled. The production of oskar and nanos proteins is controlled by repression of the mRNAs outside of the posterior region. In each case, translation is repressed by a protein that binds to the 3 ′ region (964). In the case of nanos, there is overlap between the elements required for localization and repression (965; 966; 1434). The consequence of this overlap is to make localization and repression mutually exclusive, so that when a nanos mRNA is localized to the posterior end, it cannot be repressed (1433). Last updated on January 26, 2004


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Reviews 2295. Johnstone, O. and Lasko, P. (2001). Translational regulation and RNA localization in Drosophila oocytes and embryos. Annu. Rev. Genet. 35, 365-406. 2304. Palacios, I. M. and Johnston, D. S. (2001). Getting the message across: the intracellular localization of mRNAs in higher eukaryotes. Annu. Rev. Cell Dev. Biol. 17, 569-614.

References 887. Driever, W. and Nusslein-Volhard, C. (1988). A gradient of bicoid protein in Drosophila embryos. Cell 54, 83-93. 888. Driever, W. and Nusslein-Volhard, C. (1988). The bicoid protein determines position in the Drosophila embryo in a concentration dependent manner. Cell 54, 95-104. 960. Pokrywka, N. J. and Stephenson, E. C. (1995). Microtubules are a general component of mRNA localization systems in Drosophila oocytes. Dev. Biol. 167, 363-370. 961. Wilhelm, J. E. et al. (2000). Isolation of a ribonucleoprotein complex involved in mRNA localization in Drosophila oocytes. J. Cell Biol. 148, 427-439. 962. Theurkauf, W. E. and Hazelrigg, T. I. (1998). In vivo analyses of cytoplasmic transport and cytoskeletal organization during Drosophila oogenesis: characterization of a multi-step anterior localization pathway. Development 125, 3655-3666. 963. Gavis, E. R., Curtis, D., and Lehmann, R. (1996). Identification of cis-acting sequences that control nanos localization. Dev. Biol. 176, 36-50. 964. Kim-Ha, J., Kerr, K., and MacDonald, P. M. (1995). Translational regulation of oskar mRNA by bruno, an ovarian RNA-binding protein, is essential. Cell 81, 403-412. 965. Gavis, E. R., Curtis, D., and Lehmann, R. (1996). A conserved 90 nucleotide element mediates translational repression of nanos RNA. Development 122, 2791-2800. 966. Dahanukar, A. and Wharton, R. (1966). The nanos gradient in Drosophila embryos is generated by translation regulation. Genes Dev. 10, 2610-2620. 1433. Bergsten, S. E. and Gavis, E. R. (1999). Role for mRNA localization in translational activation but not spatial restriction of nanos RNA. Development 126, 659-669. 1434. Crucs, S., Chatterjee, S., and Gavis, E. R. (2000). Overlapping but distinct RNA elements control repression and activation of nanos translation. Mol. Cell 5, 457-467. 2233. Theurkauf, W. E., Alberts, B. M., Jan, Y. N., and Jongens, T. A. (1993). A central role for microtubules in the differentiation of Drosophila oocytes. Development 118, 1169-1180. 2234. Wilkie, G. S. and Davis, I. (2001). Drosophila wingless and pair-rule transcripts localize apically by dynein-mediated transport of RNA particles. Cell 105, 209-219. 2235. Bullock, S. L. and Ish-Horowicz, D. (2001). Conserved signals and machinery for RNA transport in Drosophila oogenesis and embryogenesis. Nature 414, 611-616.



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6.31.8 How are gradients propagated? Key Concepts

• Many morphogens form gradients that control differential expression of genes. • A gradient in an egg or in the early Drosophila embryo is propagated by passive diffusion from a localized source.

• A gradient in a cellular tissue may be propagated by passive diffusion in the

intercellular spaces or by an active process in which cells transmit the morphogen to other cells.

• A gradient may also be influenced by degradation of morphogen within cells.

The cytosol of an egg forms a single compartment, as indeed does the syncytium of the early Drosophila embryo. A protein may form a gradient simply by diffusing away from a localized source (see Figure 31.5). In the case of Drosophila, such sources are provided by localized mRNAs at either the anterior end (Figure 31.7) or posterior end (Figure 31.9). Gradients are also important in development of tissues consisting of cells. We know several cases in which a morphogen forms a gradient, and the cells in that gradient respond differently depending upon the local concentration of the morphogen. The differential response of cells can be seen by placing them in tissue culture on a medium that contains a gradient of morphogen. In the tissue, however, there may be more constraints upon the movement of morphogen. For a gradient to form, material must move in intercellular spaces between the cells or must be transported through the cells. Figure 31.11 distinguishes some possible models.

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Figure 31.11 A gradient can form by passive diffusion or by active transport or may be affected by removing the diffusing material.

The simplest situation is for passive diffusion. This appears to be responsible for the gradient of activin (a TGF β homologue) that is secreted by cells in the amphibian embryo and induces formation of the mesoderm tissue layer (see Molecular Biology 6.31.13 TGF β/BMPs are diffusible morphogens). Figure 31.12 shows that the critical experiment is to create a tissue (in vitro) whose continuity is interrupted by a layer of cells that can neither respond to activin nor synthesize it. If the gradient is not stopped by these cells, the activin must be able either to pass freely through them or (more likely) diffuse past them. This turns out to be the case (2169).


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Figure 31.12 Insertion of a layer of cells that cannot interact with activin does not prevent propagation of the gradient.

Formation of the anterior-posterior axis of the Drosophila wing is determined by a gradient of the TGF β homologue, Dpp (2172). Gradient formation may involve both of the mechanisms shown in Figure 31.11 for controlling distribution. The gradient cannot be propagated unless the cells in the issue have an active receptor for TGF β and also the protein dynamin, which is involved in endocytosis (internalization) of Dpp (2171). The basic means of propagating the gradient appears to be transcytosis, in which Dpp is taken up at one face of the cell, transported across the cell, and secreted through the membrane at the other side. Some of the Dpp may be degraded instead of being passed on to the next cell. Figure 31.13 implies that the shape of the gradient may be influenced by the balance between these two processes.

Figure 31.13 A gradient can be propagated by transcytosis, when a morphogen is endocytosed (internalized) at once face of a cell and secreted at the other face. The gradient will be sharpened if some of the morphogen is degraded within the cell.


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References 2169. Gurdon, J. B., Harger, P., Mitchell, A., and Lemaire, P. (1994). Activin signalling and response to a morphogen gradient. Nature 371, 487-492. 2171. Entchev, E. V., Schwabedissen, A., and Gonzalez-Gaitan, M. (2000). Gradient formation of the TGF-beta homolog Dpp. Cell 103, 981-991. 2172. Nellen, D., Burke, R., Struhl, G., and Basler, K. (1996). Direct and long-range action of a DPP morphogen gradient. Cell 85, 357-368.



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6.31.9 Dorsal-ventral development uses localized receptor-ligand interactions Key Concepts

• Gurken mRNA is localized on the dorsal side of the oocyte. • It is translated into a TGF α-like growth factor that interacts with the Torpedo receptor on the adjoining follicle cell.

• The Torpedo receptor triggers a Ras/MAPK pathway that prevents the follicle cell from acquiring a ventral fate.

Dorsal-ventral development displays a complex interplay between the oocyte and follicle cells, involving separate pathways that are required to develop ventral and dorsal structures. The formation of ventral pattern starts with the expression of genes in the oocyte that are needed for proper development of the follicle cells. And then expression of genes in the follicle cells transmits a signal to the oocyte that results in development of ventral structures. Another pathway is responsible for development of dorsal structures in the developing egg. Each of these systems functions by activating a localized ligand-receptor interaction that triggers a signal transduction pathway (886). The localization of gurken mRNA initiates dorsal development, but also plays a role earlier in anterior-posterior patterning. These are key events that define the spatial asymmetry of the egg chamber. (The requirement of gurken for both pathways is the only feature that breaches their independence.) First gurken mRNA is localized on the posterior side of the oocyte. This results in a signal that causes adjacent follicle cells to become posterior. The follicle cells signal back to the oocyte in a process that results in the establishment of a polarized network of microtubules. This is necessary for the localization of the maternal transcripts of bicoid and oskar to opposite poles (see Molecular Biology 6.31.5 Anterior development uses localized gene regulators). Dorsal-ventral polarity is established later when gurken mRNA becomes localized on the dorsal side of the oocyte. Figure 31.14 illustrates the pathway and its consequences. The products of cornichon and brainiac are needed for proper localization of the gurken mRNA or for activation of the protein. Of the group of loci that act earlier, the products of K10 and squid are needed to localize the RNA; and cappuccino and spire mutants have an array of defects that suggest their products have a general role in organizing the cytoskeleton of the oocyte. Accordingly, cappuccino and spire are required also for the earlier localization of gurken mRNA involved in anterior-posterior patterning.

Dorsal-ventral development uses localized receptor-ligand interactions | SECTION 6.31.9 © 2004. Virtual Text / www.ergito.com

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Figure 31.14 Dorsal and ventral identities are first distinguished when grk mRNA is localized on the dorsal side of the oocyte. Synthesis of Grk activates the receptor coded by torpedo, which triggers a MAPK pathway in the follicle cells.

gurken codes for a protein that resembles the growth factor TGF α (2885; 2886). The next locus in the pathway is torpedo, which codes for the Drosophila EGF receptor. It is expressed in the follicle cells. So the pathway moves from oocyte to follicle cells when the ligand (Gurken), possibly in a transmembrane form that exposes the extracellular domain on the oocyte, interacts with the receptor (Torpedo) on the plasma membrane of a follicle cell (for review see 2887). An interesting and general principle emerges from the activation of Torpedo, which is a typical receptor tyrosine kinase. Activation of Torpedo leads to the activation of a Ras signaling pathway, which proceeds through Raf and D-mek (the equivalent of MAPKK), to activate a classic MAP kinase pathway. The ultimate readout of this pathway is not known, but its effect is to prevent activation on the dorsal side of the embryo of the ventral-determining pathway (see Molecular Biology 6.31.10 Ventral development proceeds through Toll). The utilization of this pathway shows that similar pathways may be employed in Dorsal-ventral development uses localized receptor-ligand interactions | SECTION 6.31.9 © 2004. Virtual Text / www.ergito.com

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different circumstances to produce highly specific effects. The trigger to activate the pathway in the oocyte-follicle cell interaction is the specific localization of Gurken. The consequence is a change in the properties of follicle cells that prevents them from acquiring ventral fates. The basic components of the pathway, however, are the same as those employed in signal transduction of proliferation signals in vertebrate systems. The same pathway is employed again in the specific development of retinal cells in Drosophila itself, where another receptor-counter receptor interaction activates the Ras pathway, with specific, but very different effects on cell differentiation. So essentially the same pathway can be employed to interpret an initial signal and produce a response that is predetermined by the cell phenotype. Last updated on 8-30-2002


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Reviews 2887. Wylie, A. A., Murphy, S. K., Orton, T. C., and Jirtle, R. L. (1996). Intercellular signaling and the polarization of body axes during Drosophila oogenesis. Genes Dev. 10, 1711-1723.

References 886. Schupbach, T. (1987). Germ line and soma cooperate during oogenesis to establish the Dorsal-ventral pattern of egg shell and embryo in D. melanogaster. Cell 49, 699-707. 2885. Price, J. V., Clifford, R. J., and Schupbach, T. (1989). The maternal ventralizing locus torpedo is allelic to faint little ball, an embryonic lethal, and encodes the Drosophila EGF receptor homolog. Cell 56, 1085-1092. 2886. Schejter, E. D. and Shilo, B. Z. (1989). The Drosophila EGF receptor homolog (DER) gene is allelic to faint little ball, a locus essential for embryonic development. Cell 56, 1093-1104.



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6.31.10 Ventral development proceeds through Toll Key Concepts

• The follicle cell on the ventral side produces an enzyme that modifies a proteoglycan.

• The proteoglycan triggers a series of proteolytic cleavages in the perivitelline space of the oocyte that activate the spatzle ligand.

• spatzle activates the receptor Toll, which is related to IL1 receptor, and triggers a pathway leading to activation of dorsal, which is related to the vertebrate transcription factor NF- κB.

Development of ventral structures requires a group of 11 maternal genes whose products establish the dorsal-ventral axis between the time of fertilization and cellular blastoderm (see Figure 31.4). Figure 31.15 shows that the dorsal-ventral pattern can be viewed from the side by the phenotype of the cuticle, and can be seen in cross-section to represent the formation of different types of tissues. The dorsal system is necessary for the development of ventral structures including the mesoderm and neurogenic ectoderm. (The system was named for the effects of mutations [to dorsalize], rather than for the role of the gene products [to ventralize].) Mutants in any genes of the dorsal group lack ventral structures, and have dorsal structures on the ventral side, as indicated in the figure. But injecting wild-type cytoplasm into mutant embryos rescues the defect and allows ventral structures to develop (for review see 367).

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Figure 31.15 Wild-type Drosophila embryos have distinct dorsal and ventral structures. Mutations in genes of the dorsal group prevent the appearance of ventral structures, and the ventral side of the embryo is dorsalized. Ventral structures can be restored by injecting cytoplasm containing the Toll gene product.

The ventral-determining pathway also begins in the follicle cell and ends in the oocyte. The pathway is summarized in Figure 31.16. The initial steps are not well defined, and require the expression of three loci in the follicle cells on the ventral side (2888). These loci function before fertilization, but the egg does not receive the signal until after fertilization.


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Figure 31.16 The dorsal-ventral pathway is summarized on the right and shown in detail on the left. It involves interactions between follicle cells and the oocyte. The pathway moves into the oocyte when spatzle binds to Toll and activates the morphogen. The pathway is completed by transporting the transcription factor dorsal into the nucleus.

The three loci that act in the follicle cells are nudel (ndl), windbeutel (wind), and pipe. The roles of ndl and wind are not known, but pipe plays an interesting and novel role. pipe codes for an enzyme whose sequence suggests that it is similar to the enzyme heparan sulfate 2-O-sulfotransferase (HSST) that is involved in the synthesis of a class of proteins called proteoglycans. They are components of the extracellular Ventral development proceeds through Toll | SECTION 6.31.10 © 2004. Virtual Text / www.ergito.com

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matrix. These proteins have covalently attached carbohydrate side-chains called glycosaminoglycans that have a characteristic pattern of attached monosaccharides. HSST is an enzyme that adds sulfate to the 2-O position of certain sugar residues in the monosaccharide. The pipe gene is expressed in follicle cells on the ventral side of the embryo (2883). We assume that, like HSST, it functions within the Golgi apparatus of the follicle cell. Its substrate is not known, but Figure 31.17 shows that it is probably secreted from the follicle cell into the perivitelline space (the outermost layer of the oocyte). Because the proteoglycan is synthesized on the ventral side, this creates an asymmetry at the surface of the egg.

Figure 31.17 Pipe modifies a proteolglycan within the follicle cells. The modified protein is exported to the perivitelline space

The presence of the proteoglycan in some unknown way triggers a series of proteolytic cleavages that occur in the perivitelline space. Several proteases act in succession, ending with the cleavage of the spatzle product (2884). spatzle provides a ligand for a receptor coded by the Toll gene. Toll is the first component of the pathway that functions in the oocyte. Rescue experiments identify Toll as the crucial gene that conveys the signal into the oocyte. Toll– mutants lack any dorsal-ventral gradient, and injection of Toll induces Ventral development proceeds through Toll | SECTION 6.31.10 © 2004. Virtual Text / www.ergito.com

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the formation of dorsal-ventral structures. The other genes of the dorsal group code for products that either regulate or are required for the action of Toll, but they do not establish the primary polarity (896). There is a paradox in the distribution of Toll protein. Toll gene product activity is found in all parts of a donor embryo when cytoplasm is extracted and tested by injection. Yet it induces ventral structures only in the appropriate location in normal development. An initial general distribution of Toll gene product must therefore in some way be converted into a concentration of active product by local events. Toll is a transmembrane protein homologous to the vertebrate interleukin-1 (IL1) receptor. It is located in the plasma membrane of the egg cell, with its ligand-binding domain extending into the perivitelline space (2889). Binding of ligand is sufficient to activate the ventral-determining pathway. The reaction occurs on the ventral side of the perivitelline space. The spatzle ligand either cannot diffuse far from the site where it is generated, or perhaps it binds to Toll very rapidly, with the result that Toll is activated only on the ventral side of the embryo. Loss-of-function mutations in Toll are dorsalized, because the receptor cannot be activated. There are also dominant (TollD) mutations, which confer ventral properties on dorsal regions; these are gain-of-function mutations, which are ventralized because the receptor is constitutively active. Genetic analysis shows that toll acts via tube and pelle. Tube is probably an adaptor protein that recruits the kinase pelle to the activated receptor. The target for the pelle kinase is not proven, but its activation leads to phosphorylation of the product of cactus, which is the final regulator of the transcription factor coded by dorsal. Last updated on 8-30-2002


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Reviews 367. Morisato, D. and Anderson, K. V. (1995). Signaling pathways that establish the dorsal-ventral pattern of the Drosophila embryo. Annu. Rev. Genet. 29, 371-399.

References 896. Anderson, K. V., Bokla, L., and Nusslein-Volhard, C. (1985). Establishment of dorsal-ventral polarity in the Drosophila embryo: the induction of polarity by the Toll gene product. Cell 42, 791-798. 2883. Sen, J., Goltz, J. S., Stevens, L., and Stein, D. (1998). Spatially restricted expression of pipe in the Drosophila egg chamber defines embryonic dorsal-ventral polarity. Cell 95, 471-481. 2884. Morisato, D. and Anderson, K. V. (1994). The spatzle gene encodes a component of the extracellular signaling pathway establishing the dorsal-ventral pattern of the Drosophila embryo. Cell 76, 677-688. 2888. Stein, D., Roth, S., Vogelsang, E., Nusslein-Volhard, C., and Vogelsang, C. (1991). The polarity of the dorsoventral axis in the Drosophila embryo is defined by an extracellular signal. Cell 65, 725-735. 2889. Hashimoto, C., Hudson, K. L., and Anderson, K. V. (1988). The Toll gene of Drosophila, required for dorsal-ventral embryonic polarity, appears to encode a transmembrane protein. Cell 52, 269-279.



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6.31.11 Dorsal protein forms a gradient of nuclear localization Key Concepts

• The activation of dorsal is achieved by releasing it in the cytoplasm so that it can enter the nucleus.

• A gradient of dorsal with regard to nuclear localization is established along the ventral to dorsal axis.

Figure 31.18 shows the parallels between the toll signaling pathway in flies and the IL1 vertebrate pathway (where the biochemistry is well characterized). Activation of the receptor causes a complex to assemble that includes adaptor proteins (several in vertebrates), which bind a kinase. Activation of the vertebrate kinase (IRAK) in turn activates the kinase NIK, which phosphorylates I- κB. It is not clear whether the fly kinase (pelle) acts directly on cactus (the equivalent of I κ-B) or through an intermediate.

Dorsal protein forms a gradient of nuclear localization | SECTION 6.31.11 © 2004. Virtual Text / www.ergito.com

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Figure 31.18 Activation of IL1 receptor triggers formation of a complex containing adaptor(s) and a kinase. The IRAK kinase activates NIK, which phosphorylates I- κ B. This triggers degradation of Iκ B, releasing NF- κ B, which translocates to the nucleus to activate transcription.

At all events, dorsal and cactus form an interacting pair of proteins that are related to the transcription factor NF- κB and its regulator I κB (3662; 3663; 3692). NF- κB consists of two subunits (related in sequence) which are bound by I κB in the cytoplasm. When I κB is phosphorylated, it releases NF- κB, which then moves into the nucleus, where it functions as a transcription factor of genes whose promoters have the κB sequence motif. (An example of the pathway is illustrated in Figure 22.12.) Cactus regulates dorsal in the same way that I κB regulates NF- κB (3693). A cactus-dorsal complex is inert in the cytoplasm, but when cactus is phosphorylated, it releases dorsal protein, which enters the nucleus. The pathway is therefore conserved from receptor to effector, since activation of interleukin-1 receptor has as a principal effect the activation of NF- κ B, and activation of Toll leads to activation of dorsal. A related pathway, triggered by a Toll-like receptor (TLR), is found in the system of innate immunity that is conserved from flies to mammals (see Molecular Biology 5.25.21 Innate immunity utilizes conserved signaling pathways).


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As a result of the activation of Toll, a gradient of dorsal protein in the nucleus is established, from ventral to dorsal side of the embryo. On the ventral side, dorsal protein is released to the nucleus, but on the dorsal side of the embryo it remains in the cytoplasm. A steep gradient is established at the stage of syncytial blastoderm, and becomes sharper during the transition to cellular blastoderm. The proportion of dorsal protein that is in the nucleus correlates with the ventral phenotype that will be displayed by this region. An example of a gradient visualized by staining with antibody against dorsal protein is shown in Figure 31.19. The total amount of dorsal protein in the embryo does not change: the gradient is established solely by a redistribution of the protein between nucleus and cytoplasm (899; 3694; for review see 368).

Figure 31.19 Dorsal protein forms a gradient of nuclear localization from ventral to dorsal side of the embryo. On the ventral side (lower) the protein identifies bright nuclei; on the dorsal side (upper) the nuclei lack protein and show as dark holes in the bright cytoplasm. Photograph kindly provided by Michael Levine.

Dorsal both activates and represses gene expression. It activates the genes twist and snail, which are required for the development of ventral structures. And it represses the genes dpp and zen, which are required for the development of dorsal structures; as a result, these genes are expressed only in the 40% most dorsal of the embryo (see Molecular Biology 6.31.13 TGF β/BMPs are diffusible morphogens). One of the crucial aspects of dorsal-ventral development is the relationship between the different pathways. This is summarized in Figure 31.20. The ability of one system to repress the next is responsible for restricting the localized activities to the appropriate part of the embryo. The initial interaction between gurken and torpedo leads to the repression of spatzle activity on the dorsal side of the embryo. This restricts the activation of dorsal protein to the ventral side of the embryo. Nuclear localization of dorsal protein in turn represses the expression of dpp, so that it forms a gradient diffusing from the dorsal side. In this way, ventral structures are formed in the nuclear gradient of dorsal protein, and dorsal structures are formed in the gradient of dpp protein.


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Figure 31.20 Dorsal-ventral patterning requires the successive actions of three localized systems.

The terminal system is initiated in a way that is similar to the dorsal-ventral system. A transmembrane receptor, coded by the torso gene, is produced by translation of a maternal RNA after fertilization. The receptor is localized throughout the embryo. It is activated at the poles by local production of an extracellular ligand. Torso protein has a kinase activity, which initiates a cascade that leads to local expression of the tailless and huckebein RNAs, which code for factors that regulate transcription. Last updated on 8-30-2002


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Reviews 368. Belvin, M. P. and Anderson, K. V. (1996). A conserved signaling pathway: the Drosophila Toll-Dorsal pathway. Annu. Rev. Cell Dev. Biol. 12, 393-416.

References 899. Roth, S., Stein, D., and Nusslein-Volhard, C. (1989). A gradient of nuclear localization of the dorsal protein determines Dorsal-ventral pattern in the Drosophila embryo. Cell 59, 1189-202. 3662. Kieran, M., Blank, V., Logeat, F., Vandekerckhove, J., Lottspeich, F., Le Bail, O., Urban, M. B., Kourilsky, P., Baeuerle, P. A., and Israel, A. (1990). The DNA binding subunit of NF-kappa B is identical to factor KBF1 and homologous to the rel oncogene product. Cell 62, 1007-1018. 3663. Ghosh, S., Gifford, A. M., Riviere, L. R., Tempst, P., Nolan, G. P., and Baltimore, D. (1990). Cloning of the p50 DNA binding subunit of NF-kappa B: homology to rel and dorsal. Cell 62, 1019-1029. 3692. Ip, Y. T., Kraut, R., Levine, M., and Rushlow, C. A. (1991). The dorsal morphogen is a sequence-specific DNA-binding protein that interacts with a long-range repression element in Drosophila. Cell 64, 439-446. 3693. Kidd, S. (1992). Characterization of the Drosophila cactus locus and analysis of interactions between cactus and dorsal proteins. Cell 71, 623-635. 3694. Rushlow, C. A., Han, K., Manley, J. L., and Levine, M. (1989). The graded distribution of the dorsal morphogen is initiated by selective nuclear transport in Drosophila. Cell 59, 1165-1177.



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6.31.12 Patterning systems have common features The pattern of regulators at each stage of development for each of the systems is summarized in Figure 31.21. Two types of mechanism are used to create the initial asymmetry. For the anterior-posterior axis, an RNA is localized at one end of the egg (bicoid for the anterior system, nanos for the posterior system); localization depends upon the interaction of sequences in the 3 ′ end of the RNA with maternal proteins. In the case of the dorsal-ventral and terminal systems, a receptor protein is specifically activated in a localized manner, as the result of the limited availability of its ligand. All of these interactions depend on RNAs and/or proteins expressed from maternal genes.

Figure 31.21 In each axis-determining system, localized products in the egg cause other maternal RNAs or proteins to be broadly localized at syncytial blastoderm, and zygotic RNAs are transcribed in bands at cellular blastoderm.

The local event leads to the production of a morphogen, which forms a gradient, either quantitatively (bicoid) or by nucleocytoplasmic distribution (dorsal), or is localized in a broad restricted region (nanos). The extent of the region in which the morphogen is active is ~50% across the egg for each of these systems. The morphogens are translated from maternal RNAs, and development is therefore still dependent on maternal genes up to this stage. Establishing anterior-posterior and dorsal-ventral gradients is the first step in determining orientation and spatial organization of the embryo. Under the direction of maternal genes, gradients form across the common cytoplasm and influence the Patterning systems have common features | SECTION 6.31.12 © 2004. Virtual Text / www.ergito.com

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behavior of the nuclei located in it. The next step is the development of discrete regions that will give rise to different body parts. This requires the expression of the zygotic genome, and the loci that now become active are called zygotic genes. Genes involved at this stage are identified by segmentation mutants. The products of the segmentation genes form bands that distinguish individual regions on the anterior-posterior axis. When we consider the results of the anterior and posterior systems together, we see that there are several broad regions (the two regions generated by the anterior system are adjacent to the two regions defined by the posterior system; see Figure 31.27). On the dorsal-ventral axis, there are three rather broad bands that define the regions in which the mesoderm, neuroectoderm, and dorsal ectoderm form (proceeding from the ventral to the dorsal side). Last updated on 8-30-2002 This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.31.12

Patterning systems have common features | SECTION 6.31.12 © 2004. Virtual Text / www.ergito.com

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6.31.13 TGF β/BMPs are diffusible morphogens Key Concepts

• The TGF β/BMP family provides ligands for receptors that activate Smads transcription factors.

• Synthesis of the Dpp member of this family is repressed on the ventral side of the fly embryo.

• It diffuses from a source on the ventral side and induces neural tissues. • A similar pathway functions in vertebrates but is inverted with regard to the dorsal-ventral axis.

The principle of dorsal-ventral development in flies, amphibians, and mammals is the same. On one side of the animal, neural structures (including the CNS) develop. This is the ventral side in flies, and the dorsal side in vertebrates. On the other side, mesenchymal structures develop. This is the dorsal side in flies, and the ventral side in vertebrates. The important point here is that the same relative development is seen from one side of the animal to the other, but its absolute direction is reversed between flies and vertebrates. This must mean that the dorsal-ventral axis was inverted at some point during evolution, causing the CNS to be displaced from the ventral side to the dorsal side. This idea is supported by the fact that the same signaling pathway is initiated on the dorsal side of flies and on the ventral side of vertebrate embryos (for review see 369). Mesenchymal (non-neural) structures are determined by diffusible factors in the TGF β/BMP family. These factors are small polypeptide ligands for receptors that activate the Smads transcription factors (see Figure 28.46). Formation of neural structures requires counteracting activities that also diffuse from a center; they prevent the TGF β/BMP ligands from activating the target receptors. (The names reflect the histories of their discoveries as transforming growth factor β and bone morphogenetic proteins; but in fact the most important role of these polypeptides is as morphogens in development.) The involvement of this pathway in development was first described in Drosophila, where the product of dpp is a member of the TGF β growth factor family. The receptors typically are heterodimers that form transmembrane proteins with serine/threonine kinase activity. The heterodimer consists of a type I component and a type II component. In the dpp pathway, there are two type I members (coded by thick veins and saxophone) and a single type II member (coded by punt). Mutations in tkv and punt have the same phenotype as mutations in dpp, suggesting that the tkv/punt heterodimer is the principal receptor (3695; 3696). The activated receptor phosphorylates the product mad. This is the founding member of the Smad family. The typical pattern of activation in mammalian cells is for the TGF β/BMPs are diffusible morphogens | SECTION 6.31.13 © 2004. Virtual Text / www.ergito.com

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regulated Smad to associate with a general partner to form a heterodimer that is imported into the nucleus, where it activates transcription (see Molecular Biology 6.28.21 TGF β signals through Smads). Because the gene is repressed on the ventral side, Dpp protein is secreted from cells only across the dorsal side of the embryo, as depicted in Figure 31.22. So Dpp is in effect the morphogen that induces synthesis of dorsal structures (902). Several loci influence the production of Dpp, largely by post-translational mechanisms. The net result is to increase Dpp activity on the dorsal side, and to repress it on the ventral side, of the embryo. The concentration of Dpp directly affects the cell phenotype, the most dorsal phenotypes requiring the greatest concentration (2172).

Figure 31.22 The morphogen Dpp forms a gradient originating on the dorsal side of the fly embryo. This prevents the formation of neural structures and induces mesenchymal structures.

The same pathway is involved in inducing the analogous structures in frog or mouse, but it is inverted with regard to the Dorsal-ventral axis. Figure 31.23 shows that Bmp4 is secreted from one side of the egg. It is antagonized by a variety of factors. Neural tissues develop in the (dorsal) regions which Bmp4 is prevented from reaching.

TGF β/BMPs are diffusible morphogens | SECTION 6.31.13 © 2004. Virtual Text / www.ergito.com

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Figure 31.23 Two common pathways are used in early development of Xenopus. The Niewkoop center uses the Wnt pathway to induce the Spemann organizer. The organizer diffuses dorsalizing factors that counteract the effects of the ventralizing BMPs.

The crucial unifying feature is that neural tissues are induced when the activity of Dpp/Bmp4 is antagonized. Typically the Dpp/Bmp diffuses from a source, and different phenotypes may be produced by different concentrations of the morphogen. It is controversial whether the morphogen diffuses extracellularly or whether there may be a relay system that propagates it from cell to cell. Analogous pathways, triggered by different Bmps, are involved in the development of many organs. Figure 31.24 compares the pathways in fly and frog. The basic principle is to control the availability of Dpp/Bmp. An antagonist binds to Dpp/Bmp and prevents it from binding to its receptor. The antagonists are large extracellular proteins. The antagonist is destroyed by a protease, releasing Dpp/Bmp. Neural tissue is formed in regions where Dpp/Bmp actions is prevented, whereas ectodermal tissue is formed in regions where Dpp/Bmp is activated.


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Figure 31.24 The TGF β /Bmp signaling pathway is conserved in evolution. The ligand may be sequestered by an antagonist, which is cleaved by a protease. Ligand binds to a dimeric receptor, causing the phosphorylation of a specific Smad, which together with a Co-Smad translocates to the nucleus to activate gene expression.

The fly pathway is well characterized for dorsal-ventral development. There are two types of mutants. Mutations in sog and tsg identify genes whose products antagonize Dpp, whereas mutations in tolloid suggest that it activates Dpp. Sog fulfills the role of antagonist illustrated in Figure 31.24, and is destroyed by the protease tolloid. Tsg is a sort of co-antagonist, which enhances the effect of Sog (1637; 1638; 1669). The biochemical reactions actually have been better characterized for the corresponding frog proteins (Chordin is related to Sog). Frogs may have several such pathways, with a variety of Bmp ligands that interact in an overlapping manner with a family of receptors. The frog pathway shown in Figure 31.24 is for the ventralizing effects of Bmp4, but the others are similar, although their specific effects on morphogenetic determination are of course different. There can be variation in specificity at each stage of the pathway. The antagonists, ligands, and receptors may be expressed in different places and times, providing specificity with regard to local concentrations of the morphogen, but there may also be partial redundancy. The genes for two proteins (Noggin and Chordin) both must be knocked out in mouse to produce a phenotype. Each receptor has specificity for certain Smads, so that TGF β/BMPs are diffusible morphogens | SECTION 6.31.13 © 2004. Virtual Text / www.ergito.com

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different target genes can be activated in different tissues. Last updated on 4-4-2001


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Reviews 369. De Robertis, E. M. and Sasi, Y. (1996). A common plan for Dorsal-ventral patterning in Bilateria. Nature 380, 37-40.

References 902. Ferguson, E. L. and Anderson, K. V. (1992). decapentaplegic acts as a morphogen to organize dorsal-ventral pattern in the Drosophila embryo. Cell 71, 451-461. 1637. Scott, I. C., Scott, I. C., Scott, I. C., Pappano, W. N., Maas, S. A., Cho, K. W., and Greenspan, D. S. (2001). Homologues of Twisted gastrulation are extracellular cofactors in antagonism of BMP signalling. Nature 410, 475-478. 1638. Ross, J. J., Shimmi, J. J., Vilmos, P., Petryk, A., Kim, H., Gaudenz, K., Hermanson, S., Ekker, S. C., O'Connor, M. B., and Marsh, J. L. (2001). Twisted gastrulation is a conserved extracellular BMP antagonist. Nature 410, 479-483. 1669. Chang, C., Holtzman, D. A., Chau, S., Chickering, T., Woolf, E. A., Holmgren, L. M., Bodorova, J., Gearing, D. P., Holmes, W. E., and Brivanlou, A. H. (2001). Twisted gastrulation can function as a BMP antagonist. Nature 410, 483-487. 2172. Nellen, D., Burke, R., Struhl, G., and Basler, K. (1996). Direct and long-range action of a DPP morphogen gradient. Cell 85, 357-368. 3695. Nellen, D., Affolter, M., and Basler, K. (1994). Receptor serine/threonine kinases implicated in the control of Drosophila body pattern by decapentaplegic. Cell 78, 225-237. 3696. Ruberte, E., Marty, T., Nellen, D., Affolter, M., and Basler, K. (1995). An absolute requirement for both the type II and type I receptors, punt and thick veins, for dpp signaling in vitro. Cell 80, 889-897.



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6.31.14 Cell fate is determined by compartments that form by the blastoderm stage Key Terms Many organisms have a segmented body plan that divides the body into a number of repeating units, called segments, along the anterior-posterior axis. In Drosophila, a parasegment is a unit composed of the rear of one segment and the front of the adjacent segment. In Drosophila the gap genes are a set of genes that help set up the segmentation of the embryo. Gap genes encode transcription factors that are expressed in broad regions of the embryo. Gap genes activate transcription of the pair-rule genes. In Drosophila the pair-rule genes are a set of genes that help set up the segmentation of the embryo. They are expressed in a striped pattern with one stripe in every other future segment. In Drosophila, segment polarity genes are a set of genes that help set up the segmentation of the embryo. They are expressed in a striped pattern with one stripe in every future segment. Each stripe indicates the posterior margin of a segment. Key Concepts

• A compartment of cells is defined at blastoderm and will give rise to a specific set of adult structures.

• Each segment consists of an anterior compartment and posterior compartment. • Segmentation loci are divided into three groups of genes.

By the blastoderm stage, cells have begun to acquire information about the pathways they will follow and the structures they will therefore form. This information derives initially from the maternal regulators, and then it is further refined by the actions of zygotic genes. This makes it possible to draw a "fate map" of the blastoderm embryo to identify each region in terms of the adult segments that will develop from the descendants of the embryonic cells. The concept that is intrinsic in the fate map is that a region identified at blastoderm consists of a "compartment" of cells that will give rise specifically to a particular adult structure. We can consider the development of D. melanogaster in terms of the two types of unit depicted in Figure 31.25: the segment and parasegment.

Cell fate is determined by compartments that form by the blastoderm stage | SECTION 6.31.14 © 2004. Virtual Text / www.ergito.com

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Figure 31.25 Drosophila development proceeds through formation of compartments that define parasegments and segments.

• The segment is a visible morphological structure. The adult fly consists of a series of clearly demarcated segments, and the larva has a series of corresponding segments separated by grooves. We are concerned primarily with the three thoracic (T) and eight abdominal (A) segments, about whose development most is known. The pattern of segmental units is determined by blastoderm, when the main mass of the embryo is divided into a series of alternating anterior (A) and posterior (P) compartments. So a segment consists of an A compartment succeeded by a P compartment; segment A3, for example, consists of compartments A3A and A3P. • Another type of classification originates earlier, when divisions can first be seen at gastrulation. The embryo can be divided into parasegments, each consisting of a P compartment succeeded by an A compartment. Parasegment 8, for Cell fate is determined by compartments that form by the blastoderm stage | SECTION 6.31.14 © 2004. Virtual Text / www.ergito.com

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example, consists of compartments A2P and A3A. In the 5-6 hour embryo, shallow grooves on the surface separate the adjacent parasegments. When segments form at around 9 hours, the grooves deepen and move, so that each segmental boundary represents the center of a parasegment. So the anterior part of the segment is derived from one parasegment, and the posterior part of the segment is derived from the next parasegment. In effect, the segmental units are initially evident as P-A pairs in parasegments, and then are recognized as A-P pairs in segments. How are these compartments defined during embryogenesis? The general nature of segmentation mutants suggests that the functions of segmentation genes are to establish "rules" by which segments form; a mutation changes a rule in such a way as to cause many or all segments to form improperly. The drastic consequences of segment malformation make these mutants embryonic lethals – they die at various stages before metamorphosis into adults (880). Probably ~30 loci are involved in segment formation. Figure 31.26 shows that they can be classified according to the size of the unit that they affect:

Figure 31.26 Segmentation genes affect the number of segments and fall into three groups.

• Gap gene mutants have a group of several adjacent segments deleted from the Cell fate is determined by compartments that form by the blastoderm stage | SECTION 6.31.14 © 2004. Virtual Text / www.ergito.com

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final pattern. Four gap genes are involved in formation of the major body segments, and others are concerned with the head and tail structures. • Pair-rule mutants have corresponding parts of the pattern deleted in every other segment. The afflicted segments may be even-numbered or odd-numbered. There are 8 pair-rule genes. • Segment polarity mutants most often lose part of the P compartment of each segment, and it is replaced by a mirror image duplication of the A compartment. Some mutants cause loss of A compartments or middle segments. There are ~16 segment polarity genes. These groups of genes are expressed at successive periods during development; and they define increasingly restricted regions of the egg, as can be seen from Figure 31.27. The maternal genes establish gradients from the anterior and posterior ends. The maternal gradients either activate or repress the gap genes, which are amongst the earliest to be transcribed following fertilization (following the 11th nuclear division); they divide the embryo into 4 broad regions. The gap genes regulate the pair-rule genes, which are transcribed slightly later; their target regions are restricted to pairs of segments. The pair-rule genes in turn regulate the segment polarity genes, which are expressed during the 13th nuclear division, and by now the target size is the individual segment.


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Figure 31.27 Maternal and segmentation genes act progressively on smaller regions of the embryo.

Many of the maternal genes, the gap genes, and the pair-rule genes are regulators of transcription. Their effects may be either to activate or to repress transcription; in some cases, a given protein may activate some target genes and repress other target genes, depending on its level or the context. The genes in any one class regulate one another as well as regulating the genes of the next class. When we reach the level of segment polarity genes, the nature of the regulatory event changes, and many of the gene products act on communication between cells to maintain borders between compartments, for example, to control the secretion of a protein from one cell to influence its neighbor (for review see 363; 358). The principle that emerges from this analysis is that at each stage a small number of maternal, gap, and pair-rule regulator proteins is used in combinatorial associations to specify the pattern of gene expression in a particular region of the embryo (for review see 3705).


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Reviews 358. Scott, M. P. and Carroll, S. B. (1987). The segmentation and homeotic gene network in early Drosophila development. Cell 51, 689-698. 363. Ingham, P. W. and Martinez-Arias, A. (1992). Boundaries and fields in early embryos. Cell 68, 221-235.

References 880. Nusslein-Vollhard, C. and Wieschaus, E. (1980). Mutations affecting segment number and polarity in Drosophila. Nature 287, 795-801. 3705. Lehmann, R. and Frohnhofer, H. G. (1989). Segmental polarity and identity in the abdomen of Drosophila is controlled by the relative position of gap gene expression. Development 107 Suppl, 21-29.



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6.31.15 Gap genes are controlled by bicoid and by one another Key Concepts

• Gap genes affect a group of segments, and are controlled by bicoid and by interactions among themselves.

• Pair-rule genes are expressed in either even- or odd-numbered segments, and are controlled by the gap genes.

• Segment polarity mutants are controlled by the pair-rule genes, and are expressed in segments where they affect anterior or posterior identification of the compartments.

The gap genes are controlled in two ways: they may respond directly to the bicoid morphogen; and they regulate one another. The four bands shown in Figure 31.27 are created by the levels of the two proteins bicoid and hunchback (3704). The synthesis of hunchback mRNA is activated by bicoid (3702). Hunchback protein forms a gradient in the egg that in turn controls the expression of other genes. The most anterior band in Figure 31.27 consists of hunchback protein. The next band consists of Kruppel protein; transcription of the Kruppel gene is activated by hunchback protein. The next two bands consist of knirps and giant proteins. Transcription of these genes is repressed by hunchback. They are expressed in the posterior part of the embryo because nanos has prevented the expression of hunchback there. Figure 31.28 examines the transition from the 4 band to the 7 striped stage in more detail. The detailed interactions among the gap proteins are determined by examining the pattern of the distribution of other gap proteins in a mutant lacking one particular gap protein. Hunchback plays an especially important role. It is expressed in a broad anterior region, with a gradient of decline in the middle of the embryo. High levels of hunchback repress Kruppel; this determines the anterior boundary of Kruppel expression, which rises just as hunchback falls off, in parasegment 3. But some level of hunchback is needed for Kruppel expression, so when the level of hunchback decreases further, Kruppel is turned off, around parasegment 5. In the same way, expression of giant responds to successive changes in the level of hunchback; and knirps expression requires the absence of hunchback.

Gap genes are controlled by bicoid and by one another | SECTION 6.31.15 © 2004. Virtual Text / www.ergito.com

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Figure 31.28 Expression of the gap genes defines adjacent regions of the embryo. The gap genes control the pair-rule genes, each of which is expressed in 7 stripes.

The control is refined further by interactions among the proteins. The general principle is that one interaction may be required to express a protein in a particular region, and other interactions may be required to repress its expression at the boundaries. The effects are worked out by examining pairwise interactions. For example, overexpression of giant causes the Kruppel band to become much narrower, suggesting that giant contributes to repressing the boundaries of Kruppel. The posterior margins of knirps and giant are determined by the operation of the terminal system. Altogether, these interactions mean that, as we proceed along the egg from anterior to posterior, any particular position can be defined by the levels of the various gap proteins.


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References 3702. Tabata, T., Schwartz, C., Gustavson, E., Ali, Z., and Kornberg, T. B. (1995). Creating a Drosophila wing de novo, the role of engrailed, and the compartment border hypothesis. Development 121, 3359-3369. 3704. Simpson-Brose, M., Treisman, J., and Desplan, C. (1994). Synergy between the hunchback and bicoid morphogens is required for anterior patterning in Drosophila. Cell 78, 855-865.



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6.31.16 Pair-rule genes are regulated by gap genes Key Concepts

• Pair-rule genes are expressed in either even- or odd-numbered segments, and are controlled by the gap genes.

All the gap proteins are regulators of transcription, and in addition to regulating one another, they regulate the expression of the pair-rule genes that function at the next stage. Each pair-rule protein is found in a pattern of 7 "stripes" along the embryo, and Figure 31.28 shows the approximate positions that these stripes will take as the result of expression of the gap genes (3707). (Of course, the parasegments have not developed yet, and are shown just to relate their positions to the protein distribution.) The 7 stripes of a pair-rule gene identify either all the odd-numbered parasegments (like eve) or all the even-numbered parasegments (like ftz). Two of the pair-rule genes, hairy and eve, are called primary pair-rule genes, because they are expressed first, and their pattern of expression influences the expression of the other pair-rule genes. Recall that mutations in pair-rule genes delete half the segments. Figure 31.29 compares the segmentation patterns of wild-type and fushi tarazu (ftz) larvae. The mutant has only half the number of segments, because every other segment is missing.

Figure 31.29 ftz mutants have half the number of segments present in wild-type. Photographs kindly provided by Walter Gehring.

The ftz mRNA is present from early blastoderm to gastrula stages of development. Pair-rule genes are regulated by gap genes | SECTION 6.31.16 © 2004. Virtual Text / www.ergito.com

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Figure 31.30 shows the locations of the transcripts, visualized in situ at blastoderm in wild type. The gene is expressed in 7 stripes, each 3-4 cells wide, running across the embryo. As shown previously in Figure 31.28, the stripes correspond to even-numbered parasegments (4 = T1P/T2A, 6= T3P/A1A, 8 = A2P/A3A, etc.).

Figure 31.30 Transcripts of the ftz gene are localized in stripes corresponding to even numbered parasegments. The expressed regions correspond to the regions that are missing in the ftz mutant of the previous figure. Photograph kindly provided by Walter Gehring.

This pattern suggests a function for the ftz gene: it must be expressed at blastoderm for the structures that will be descended from the even-numbered parasegments to develop. Mutants in which ftz is defective lack these parasegments because the gene product is absent during the period when they must be formed. In other words, expression of ftz is required for survival of the cells in the regions in which it is expressed. The expression of ftz is an example of the general rule that the stripes in which a pair-rule gene is expressed correspond to the regions that are missing from the embryo when the gene is mutated. Compartments are therefore determined by the pattern of expression of segmentation genes. The width of the stripe in which a gene is expressed corresponds to the size of the segmental unit that it affects. Different mechanisms are used to specify the expression patterns of different pair-rule genes; we have the most information about ftz and eve. In the early embryo, ftz is uniformly expressed. If protein synthesis is blocked before the stripes develop, the embryo retains the initial pattern. So the development of stripes depends on the specific degradation of ftz RNA in the regions between the bands and at the anterior and posterior ends of the embryo. Once the stripes have developed, transcription of ftz ceases in the interbands and at the ends of the embryo. The specificity of transcription depends on regions upstream of the ftz promoter, and also on the function of several other segmentation genes. The transcription of ftz responds to other pair-rule genes (and perhaps gap genes) through elements that act on all stripes. The expression pattern of eve is complementary to ftz, but has a different basis: it is controlled separately in each stripe. A detailed reconstruction using subregions of the eve promoter shows that the information for localization in each stripe is coded in a separate part of the promoter; the promoter can be divided into regions that respond to the local levels of gap gene products in particular parasegments. For example, the promoter region that is responsible for eve expression in parasegment 3 has binding Pair-rule genes are regulated by gap genes | SECTION 6.31.16 © 2004. Virtual Text / www.ergito.com

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sites for the gap proteins bicoid, hunchback, giant, and Kruppel. Figure 31.31 shows that this part of the promoter extends for 480 bp. It works in the following way. eve transcription is activated by hunchback and bicoid. The two boundaries are determined because the promoter is repressed by giant on the anterior side and by Kruppel on the posterior side (see also Figure 31.28). Other parts of the promoter respond to the protein levels in other parts of the embryo. So the different stripes of the primary pair-rule gene products are regulated by separate pathways, each of which is susceptible to activation by a particular combination of gap gene products and other regulators.

Figure 31.31 The eve stripe in parasegment 3 is activated by hunchback and bicoid. Repression by giant sets the anterior boundary; repression by Kruppel sets the posterior boundary. Multiple binding sites for these proteins in a 480 bp region of the promoter control expression of the gene.

This illustrates in miniature the principle that combinations of proteins control gene expression in local areas. The general principle is that generally distributed proteins (such as bicoid or hunchback) are needed for activation, whereas the borders are formed by selective repression (by giant and Kruppel in this particular example). We should emphasize that the hierarchy of gene control is not exclusively restricted to interactions between successive stages of control (maternal gap pair-rule). For Pair-rule genes are regulated by gap genes | SECTION 6.31.16 © 2004. Virtual Text / www.ergito.com

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example, the involvement of bicoid protein in regulating eve transcription in parasegment 3 shows that a maternal gene may have a direct effect on a pair-rule gene. The stripes of eve and ftz are fuzzy to begin with, and become sharper as development proceeds, corresponding to more finely defined units. Figure 31.32 shows an example of an embryo simultaneously stained for expression of ftz and eve. Initially there is a series of alternating fuzzy stripes, but the stripes narrow from the posterior margin and sharpen on the anterior side as they intensify during development. This may depend on an autoregulatory loop, in which the expression of the gene is regulated by its own product.

Figure 31.32 Simultaneous staining for ftz (brown) and eve (grey) shows that they are first expressed as broad alternating stripes at the time of blastoderm (upper), but narrow during the next 1 hour of development (lower). Photographs kindly provided by Peter Lawrence.

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References 3707. Lawrence, P. A. and Johnston, P. (1989). Pattern formation in the Drosophila embryo: allocation of cells to parasegments by even-skipped and fushi tarazu. Development 105, 761-767.


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6.31.17 Segment polarity genes are controlled by pair-rule genes Key Concepts

• Segment polarity genes are expressed in segments where they affect anterior or posterior identification of the compartments.

The pair-rule genes control the expression of the segment polarity genes, which are expressed in 14 stripes. Each stripe identifies a segment. The compartmental pattern in which segment polarity genes are expressed is exceedingly precise. Perhaps the ultimate demonstration of precision is provided by the pattern gene engrailed. The function coded by engrailed is needed in all segments and is concerned with the distinction between the A and P compartments. engrailed is expressed in every P compartment, but not in A compartments (3701; 3700; 3702). Mutants in this gene do not distinguish between anterior and posterior compartments of the segments. Antibodies against the protein coded by engrailed react against the nucleus of cells expressing it. The regions in which engrailed is expressed form a pattern of stripes. When the stripes of engrailed protein first become apparent, they are only one cell wide.Figure 31.33 shows the pattern at a stage when each segment has a stripe just 1 cell in width, with the stripe beginning to widen into several cells.

Figure 31.33 Engrailed protein is localized in nuclei and forms stripes as precisely delineated as 1 cell in width. Photograph kindly provided by Patrick O#Farrell.

Actually, the pattern of stripes becomes established over a 30 minute period, moving along the embryo from anterior to posterior. Initially one stripe is apparent; then every other segment has a stripe; and finally the complete pattern has a stripe 3-4 cells wide corresponding to the P compartment of every segment. The expression of engrailed is of particular importance, because it defines the boundaries of the actual compartments from which adult structures will be derived. Segment polarity genes are controlled by pair-rule genes | SECTION 6.31.17 © 2004. Virtual Text / www.ergito.com

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The initial 1-cell-wide stripes of engrailed protein form at the anterior boundaries of both the ftz and eve stripes, and delineate what will become the anterior boundary of every P compartment. Why is engrailed initially transcribed exclusively in this anterior edge, within the broader stripes of ftz and eve expression? This question is a specific example of a more general question: how can a broad stripe be subdivided into more restricted, narrower stripes? We can consider two general types of model: • A combinatorial model supposes that different genes are expressed in overlapping patterns of stripes. A pattern of stripes develops for each of the pair-rule genes. The different pair-rule gene stripes overlap, because they are out of phase with one another. As a result of these patterns, different cells in the cellular blastoderm express different combinations of pair-rule genes. Each compartment is defined by the particular combination of the genes that are expressed, and these combinations determine the responses of the cells at next stage of development. In other words, the segmentation genes are controlled by the pair-rule genes in the same general manner that the pair-rule genes are controlled by the gap genes. • A boundary model supposes that a compartment is defined by the striped pattern of expression, but that interactions involving cell-cell communication at the boundaries cause subdivisions to arise within the compartment. In the case of engrailed, we would suppose that some unique event is triggered by the juxtaposition of cells possessing ftz (or eve) with cells that do not, and this is necessary to trigger engrailed expression. Each of the 14 segments is subdivided further into anterior and posterior compartments by the activities of the segment polarity genes. The actions of the segment polarity genes are the same in every segment. For example, engrailed distinguishes the A and P compartments. engrailed is a transcription factor, but other segment polarity genes have different types of functions. The products of the segment polarity genes include secreted proteins, transmembrane proteins, kinases, cytoskeletal proteins, as well as transcription factors. Cell-cell interactions become important at this stage for defining and maintaining the nature of the compartments (see Molecular Biology 6.31.18 Wingless and engrailed expression alternate in adjacent cells).

Segment polarity genes are controlled by pair-rule genes | SECTION 6.31.17 © 2004. Virtual Text / www.ergito.com

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References 3700. Kornberg, T, Sidén, I, O'Farrell, P, and Simon, M (1985). The engrailed locus of Drosophila: in situ localization of transcripts reveals compartment-specific expression. Cell 40, 45-53. 3701. Kornberg, T. (1981). Engrailed: a gene controlling compartment and segment formation in Drosophila. Proc. Natl. Acad. Sci. USA 78, 1095-1099. 3702. Tabata, T., Schwartz, C., Gustavson, E., Ali, Z., and Kornberg, T. B. (1995). Creating a Drosophila wing de novo, the role of engrailed, and the compartment border hypothesis. Development 121, 3359-3369.


Segment polarity genes are controlled by pair-rule genes | SECTION 6.31.17 © 2004. Virtual Text / www.ergito.com

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6.31.18 Wingless and engrailed expression alternate in adjacent cells Key Concepts

• Wingless and engrailed have a mutually reinforcing interaction between neighboring cells.

• Wingless is secreted at the posterior boundary of a cell. • It activates the Fz (or dFz2) receptor in the adjacent cell, which triggers the translocation of Armadillo to the nucleus.

• Armadillo causes engrailed to be expressed, and engrailed causes secretion of hedgehog at the anterior boundary, where it acts on the neighboring cell to maintain Wingless expression.

The circuit that defines the boundaries between anterior and posterior compartments is based on mutual interactions between segment polarity genes. wingless codes for a protein that is secreted and taken up by the adjacent row of cells. It is initially expressed in a row of cells immediately adjacent to the anterior side of the cells expressing engrailed; so wingless comes to identify the posterior boundary of the preceding parasegment. The initial expression of engrailed in response to ftz and eve is shortly replaced by an autoregulatory loop in which secretion of wingless protein from the adjacent cells is needed for expression of engrailed; and expression of engrailed is needed for expression of wingless. This keeps the boundary sharp. The wingless signaling pathway is one of the most interesting, and has close parallels in all animal development. Like other signaling pathways utilized in development, it is initiated by an extracellular ligand, and results in the expression of a transcription factor, although the interactions between components of the pathway are somewhat unusual. In fly embryonic development at the stage of segmental definition, the cells that define the boundaries of the A and P compartments express wingless (Wg) and engrailed (En) in a reciprocal relationship. Figure 31.34 shows that wingless protein is secreted from a cell at a boundary, and acts upon the cell on its posterior side. The wingless signaling pathway causes the engrailed gene to be expressed. Engrailed causes the production of hedgehog (Hh) protein, which in turn is secreted. Hedgehog acts on the cell on its anterior side to maintain wingless expression. Wg is also required for patterning of adult eyes, legs, and wings (hence its name) (898).

Wingless and engrailed expression alternate in adjacent cells | SECTION 6.31.18 © 2004. Virtual Text / www.ergito.com

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Figure 31.34 Reciprocal interactions maintain Wg and Hh signaling between adjacent cells. Wg activates a receptor, which activates a pathway leading to translocation of Arm to the nucleus. This activates engrailed, which leads to expression of Hedgehog protein, which is secreted to act on the neighboring cell, where it maintains Wg expression.

The identification of the receptor for Wg on the posterior cell has actually been very difficult. Wg interacts with frizzled in vitro, but mutational analysis suggests that the related protein, DFz2 (Drosophila frizzled-2) is the receptor. It is possible that these may play redundant roles. Another protein that is required for reception/signaling is the product of arrow, which is a single-pass membrane protein and is classified as a coreceptor (2894). The frizzled family members are 7-membrane pass proteins, with the appearance of classical receptors (although the major pathway does not appear to involve G proteins) (903; 2896). Last updated on 9-3-2002


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References 898. Rijsewijk, F. M. (1987). The Drosophila homolog of the mouse mammary oncogene int-1 is identical to the segment polarity gene wingless. Cell 50, 649-657. 903. Bhanot, P. et al. (1995). A new member of the frizzled family from Drosophila functions as a wingless receptor. Nature 382, 225-230. 2894. Wehrli, M., Dougan, S. T., Caldwell, K., O'Keefe, L., Schwartz, S., Vaizel-Ohayon, D., Schejter, E., Tomlinson, A., and DiNardo, S. (2000). arrow encodes an LDL-receptor-related protein essential for Wingless signalling. Nature 407, 527-530. 2896. Muller, H., Samanta, R., and Wieschaus, E. (1999). Wingless signaling in the Drosophila embryo: zygotic requirements and the role of the frizzled genes. Development 126, 577-586.



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6.31.19 The wingless/wnt pathway signals to the nucleus Key Concepts

• Wingless (in Drosophila) and Wnt (in vertebrates) activate a receptor that blocks the action of a cytosolic Ser/Thr kinase.

• The kinase phosphorylates Armadillo/ β-catenin that is localized in cytosolic complexes.

• In the absence of phosphorylation, Armadillo/ β-catenin is stabilized, and translocates to the nucleus where it activates transcription.

• A separate pool of Armadillo/ β-catenin is present in complexes at the cell surface, but is not a target for the pathway.

• The function of the cancer-causing gene APC is to destabilize β-catenin, and colon cancers caused by mutation in APC have elevated levels of β-catenin.

The interaction between Wingless and its receptor activates a signaling pathway. The effector of the pathway is a protein called Arm (Armadillo) in Drosophila and β-catenin in vertebrate cells. A cytoplasmic pool of the effector protein is constitutively degraded, and the role of the signaling pathway is to block the degradation. When this happens, Arm/ β-catenin is transported to the nucleus, where it activates transcription of a set of target genes. Mutants in other genes that have segment polarity defects similar to wg mutants identify the other components of the pathway. They signal positively to execute the pathway. Mutations that have the opposite phenotype, and that block the pathway, identify proteins that are required to degrade Arm/ β-catenin. Figure 31.35 shows the results of ordering the genes genetically, and defining the biochemical interactions between their products.

The wingless/wnt pathway signals to the nucleus | SECTION 6.31.19 © 2004. Virtual Text / www.ergito.com

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Figure 31.35 Wg secretion is assisted by porc. Wg activates the Dfz2 receptor, which inhibits Zw3 kinase. Active Zw3 causes turnover of Arm. Inhibition of Zw3 stabilizes Arm, allowing it to translocate to the nucleus. In the nucleus, Arm partners Pan, and activates target genes (including engrailed). A similar pathway is found in vertebrate cells (components named in blue).

The fly and vertebrate transduction pathways have homologous components. The components in the order of their function in the pathway are: • Dsh (coded by Dishevelled) is a phosphoprotein that responds to the interaction of wingless/Wnt with the frizzled receptor. • Dsh signals to a Ser/Thr kinase, called Zw3 in Drosophila, but called GSK3 in vertebrates (named for its historical identification as glycogen synthase kinase, but in fact a homologue of Zw3). Zw3/GSK3 is constitutively active, unless and until it is inactivated by Dsh. The wingless/wnt pathway signals to the nucleus | SECTION 6.31.19 © 2004. Virtual Text / www.ergito.com

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Reviews 2897. Zhurinsky, J., Shtutman, M., and Ben-Ze'ev, A. (2000). Plakoglobin and beta-catenin: protein interactions, regulation and biological roles. J. Cell Sci. 113 , 3127-3139.

References 904. Molenaar, M. et al. (1996). XTcf-3 transcription factor mediates β -catenin-induced axis formation in Xenopus embryos. Cell 86, 391-399. 905. Brunner, E. (1997). pangolin encodes a Lef-1 homologue that acts downstream of Armadillo to transduce the Wingless signal in Drosophila. Nature 385, 829-833. 2167. Graham, T. A., Weaver, C., Mao, F., Kimelman, D., and Xu, W. (2000). Crystal structure of a beta-catenin/Tcf complex. Cell 103, 885-896. 4518. Xing, Y., Clements, W. K., Kimelman, D., and Xu, W. (2003). Crystal structure of a beta-catenin/axin complex suggests a mechanism for the beta-catenin destruction complex. Genes Dev. 17, 2753-2764.



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6.31.20 Complex loci are extremely large and involved in regulation Key Terms Homeotic genes are defined by mutations that convert one body part into another; for example, an insect leg may replace an antenna. A complex locus (of D. melanogaster) has genetic properties inconsistent with the function of a gene representing a single protein. Complex loci are usually very large (>100 kb) at the molecular level. Key Concepts

• Complex loci were identified by interallelic interactions that did not fit the usual complementation behavior.

• They are extremely large and may include multiple protein-coding units as well as cis-acting regulatory sites.

• The order of mutations from upstream to downstream corresponds to the order of the body parts that are affected from anterior to posterior.

• ANT-C includes several genes that affect head segments, whereas BX-C has only three genes and many regulatory sites.

• A segment is distinguished from the preceding (anterior-side) segment by expression of an additional protein-coding unit.

• Loss of a protein-coding unit causes a homeotic transformation in which a segment has the identity of the segment on its anterior-side.

Segment polarity genes control the anterior-posterior pattern within each segment. Homeotic genes impose the program that determines the unique differentiation of each segment. Most homeotic genes are expressed in a spatially restricted manner that corresponds to parasegments. Homeotic genes interact in complicated interlocking patterns. Many homeotic genes code for transcription factors that act upon other homeotic genes as well as upon other target loci. As a result, a mutation in one homeotic gene influences the expression of other homeotic genes. The consequence is that the final appearance of a mutant depends not only on the loss of one homeotic gene function, but also on how other homeotic genes change their spatial patterns in response to the loss. Homeotic genes act during embryogenesis. Their expression depends on the prior expression of the segmentation genes; we might regard the homeotic genes as integrating the pattern of signals established by the segmentation genes. Homeotic mutants "transform" part of a segment or an entire segment into another type of segment; they may cause one segment of the abdomen to develop as another, legs to Complex loci are extremely large and involved in regulation | SECTION 6.31.20 © 2004. Virtual Text / www.ergito.com

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develop in place of antennae, or wings to develop in place of eyes. Note that homeotic genes do not create patterns de novo; they modify cell fates that are determined by genes such as the segment polarity genes, by switching the set of genes that functions in a particular place. Indeed, the segment polarity genes are active at about the same time as the peak of expression of the homeotic genes. The genetic properties of some homeotic mutations are unusual and led to the identification of complex loci (for review see 357; 2881; 2882). A conventional gene – even an interrupted one – is identified at the level of the genetic map by a cluster of noncomplementing mutations. In the case of a large gene, the mutations might map into individual clusters corresponding to the exons. A hallmark of a complex locus is that, in addition to rather well-spaced groups of mutations, extending over a relatively large map distance, there are complex patterns of complementation, in which some pairwise combinations complement but others do not. The individual mutations may have different and complex morphological effects on the phenotype. These relationships are caused by the existence of an array of regulatory elements. Many of the bizarre results that are obtained in complementation assays turn out to result from mutations in promoters or enhancers that affect expression in one cell type but not another. We now recognize that complex loci do not have any novel features of genetic organization, apart from the fact that they have many regulatory elements that control expression in different parts of the embryo. Two of the complex loci are involved in regulating development of the adult insect body. The ANT-C and BX-C complex loci together provide a continuum of functions that specify the identities of all of the segmented units of the fly. Each of these complexes contains several homeotic genes. The two separate complexes may have evolved from a split in a single ancestral complex, as suggested by the evolution of the corresponding genes in other species. In the beetle Tribolium, the ANT-C and BX-C complexes are found together at a single chromosomal location. The individual genes may have been derived from duplications and mutations of an original ancestral gene. And in mammals, there are arrays of related genes whose individual members are related sequentially to the genes of the ANT-C and BX-C complexes (see Molecular Biology 6.31.22 The homeobox is a common coding motif in homeotic genes). The homeotic genes clustered at the ANT-C and BX-C complexes show a relationship between genetic order and the position in which they are expressed in the body of the fly. Proceeding from left to right, each homeotic gene in the complex acts upon a more posterior region of the fly. The basic principle is that formation of a compartment requires the gene product(s) expressed in the previous compartment, plus a new function coded by the next gene along the cluster. So loss-of-function mutations usually cause one compartment to have the phenotype of the corresponding compartment on its anterior side. The individual genes code for a set of transcription factors that have related DNA-binding domains (see next section). The identities of the most anterior parts of the fly (parasegments 1-4) are specified by ANT-C, which contains several homeotic genes, including labial (lab), proboscipedia (pb), Deformed (Dfd), Sex combs reduced (Scr), and Antennapedia (Antp). The homeotic genes lie in a cluster over a region of ~350 kb, but several other genes are interspersed; most of these genes are regulators that function at different stages of development (356). Complex loci are extremely large and involved in regulation | SECTION 6.31.20 © 2004. Virtual Text / www.ergito.com

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Figure 31.36 correlates the organization of ANT-C with its effects upon body parts. Adjacent genes are expressed in successively more posterior parts of the embryo, ranging from the leftmost gene labial (the most anterior acting, which affects the head) to the rightmost gene Antp (the most posterior acting, which affects segments T2–T3).

Figure 31.36 The homeotic genes of the ANT-C complex confer identity on the most anterior segments of the fly. The genes vary in size, and are interspersed with other genes. The antp gene is very large and has alternative forms of expression. Complex loci are extremely large and involved in regulation | SECTION 6.31.20 © 2004. Virtual Text / www.ergito.com

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The Antp gene gave its name to the complex, and among the mutations in it are alleles that change antennae into second legs, or second and third legs into first legs. Antp usually functions in the thorax; it is needed both to promote formation of segments T2-T3 and to suppress formation of head structures. Loss of function therefore causes T2-T3 to resemble the more anterior structure of T1; gain of function, for example, by overexpression in the head, causes the anterior region to develop structures of the thorax. (The molecular action of Antp is to prevent the action of genes hth and exd that promote formation of antennal structures. Hth causes exd to be imported into the nucleus, where it switches on the genes that make the antenna.) Figure 31.36 summarizes the organization of the gene. It has 8 exons, separated by very large introns, and altogether spanning ~103 kb. The single open reading frame begins only in exon 5, and apparently gives rise to a protein of 43 kD. The discrepancy between the length of the locus and the size of the protein means that only 1% of its DNA codes for protein (882). Transcription starts at either of two promoters, located ~70 kb apart! One promoter is located upstream of exon 1, the other upstream of exon 3. Use of the first promoter is associated with omission of exon 3. The transcripts generated from either promoter end either within or after exon 8. All the transcripts appear to code for the same protein. Each promoter has its own tissue-specific expression pattern (3708). We do not know if there is any significance to the difference in the structure of the two types of transcript. The other genes of the ANT-C complex are expressed in the head and first thoracic segment. In the most anterior compartments, lab, pb, Dfd, have unique patterns of expression, so that deletions of segmental regions can result from loss-of-function mutations. An exception to the left-right/anterior-posterior order of action is that loss of Scr allows the overlapping Antp to function, that is, the direction of transformation is opposite from usual.

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Reviews 356. Regulski, M., Harding, K., Kostriken, R., Karch, F., Levine, M., and McGinnis, W. (1985). Homeo box genes of the Antennapedia and bithorax complexes of Drosophila. Cell 43, 71-80. 357. Scott, M. P. (1987). Complex loci of Drosophila. Annu. Rev. Biochem. 56, 195-227. 2881. Montgomery, G. (2002). E. B. Lewis and the bithorax complex. Part I. Genetics 160, 1265-1272.

References 882. Scott, M. P. et al. (1983). The molecular organization of the Antennapedia locus of Drosophila. Cell 35, 763-766. 2882. Duncan, I. and Montgomery, G. (2002). E. B. Lewis and the Bithorax Complex. Part ii. from cis-trans test to the genetic control of development. Genetics 161, 1-10. 3708. Jorgensen, E. M. and Garber, R. L. (1987). Function and misfunction of the two promoters of the Drosophila Antennapedia gene. Genes Dev. 1, 544-555.


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6.31.21 The bithorax complex has trans-acting genes and cis-acting regulators Key Terms The bithorax complex is a group of homeotic genes which are responsible for the diversification of the different segments of the fly. Key Concepts

• bithorax controls body structures from T2 to A8. • The ultrabithorax domain has the Ubx transcription unit. • The infraabdominal domain has the AbdA and AbdB transcription units. • The order of units on the genetic map coincides with the order of body parts. • Expression of additional units specifies more posterior body parts.

The classic complex homeotic locus is BX-C, the bithorax complex, characterized by several groups of homeotic mutations that affect development of the thorax, causing major morphological changes in the abdomen. When the whole complex is deleted, the insect dies late in embryonic development. Within the complex, however, are mutations that are viable, but which change the phenotype of certain segments. An extreme case of homeotic transformation is shown in Figure 31.37, in which a triple mutation converts T3A (which carries the halteres [truncated wings]) into the tissue type of the T2 (which carries the wings). This creates a fly with four wings instead of the usual two (881).

Figure 31.37 A four-winged fly is produced by a triple mutation in abx, bx, and pbx at the BX-C complex. Photograph kindly provided by Ed Lewis.

The genetic map of BX-C is correlated with the body structures that it controls in the fly in Figure 31.38. The body structures extend from T2 to A8. The BX-C complex is therefore concerned with the development of the major part of the body of the fly. The bithorax complex has trans-acting genes and cis-acting regulators | SECTION 6.31.21 © 2004. Virtual Text / www.ergito.com

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Like ANT-C, a crucial feature of this complex is also that mutations affecting particular segments lie in the same order on the genetic map as the corresponding segments in the body of the fly. Proceeding from left to right along the genetic map, mutations affect segments in the fly that become successively more posterior (895; for review see 3709).

Figure 31.38 The bithorax (BX-C) locus has 3 coding units. A series of regulatory mutations affects successive segments of the fly. The sites of the regulatory mutations show the regions within which deletions, insertions, and translocations confer a given phenotype.

A difference between ANT-C and BX-C is that ANT-C functions largely or exclusively via its protein-coding loci, but BX-C displays a complex pattern of cis-acting interactions in addition to the effects of mutations in protein-coding The bithorax complex has trans-acting genes and cis-acting regulators | SECTION 6.31.21 © 2004. Virtual Text / www.ergito.com

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regions. The BX-C occupies 315 kb, of which only 1.4% codes for protein. The individual mutations fall into two classes: • Three transcription units (Ubx, abdA, AbdB) produce mRNAs that code for proteins. The transcription units are large (>75 kb for Ubx, and >20 kb each for abdA and AbdB). Each contains several large introns. (The bxd and iab4 regions produce RNAs that do not code for proteins; again, the transcription units are large, and the RNA products are spliced. Their functions are unknown.) • There are cis-acting mutations at intervals throughout the entire cluster. They control expression of the transcription units. Cis-acting mutations of any particular type may occur in a large region. The locations shown on the map are only approximate, and the boundaries within which mutations of each type may occur are not well defined. As a historical note, the complex was originally defined in terms of two "domains." Mutations in the Ultrabithorax domain were characterized first; they have the thoracic segments T2P-T3P and the abdominal compartment A1A as their targets (this corresponds to parasegments 5-6). These mutations lie either in the Ubx transcription unit or in the cis-acting sites that control it. The mutations within the ultrabithorax domain are named for their phenotypes. The bx and bxd types are identified by a series of mutations, in each case dispersed over ~10 kb. The abx and pbx mutations are caused by deletions, which vary from 1-10 kb. Mutations in the Infraabdominal domain were found later; they have the abdominal segments A1P-A8P (parasegments 7-14) as targets. These mutations lie either in the AbdA,B transcription units, or in the cis-acting sites that control them. Within the infraabdominal domain, cis-acting mutations are named systematically as iab2-9. These mutations affect individual compartments, or sometimes adjacent sets of compartments, as shown at the top of the figure (885). Proceeding from left to right along the cluster, transcripts are found in increasingly posterior parts of the embryo, as shown at the bottom of the figure. The patterns overlap. Ubx has an anterior boundary of expression in compartment T2P (parasegment 5), abdA is expressed from compartment A1P (parasegment 7), and AbdB is expressed in compartments posterior to compartment A4P (parasegment 10). Transcription of Ubx has been studied in the most detail. The Ubx transcription unit is ~75 kb, and has alternative splicing patterns that give rise to several short RNAs. A transient 4.7 kb RNA appears first, and then is replaced by RNAs of 3.2 and 4.3 kb. A feature common to both the latter two RNAs is their inclusion of sequences from both ends of the primary transcript. Of course, there may be other RNAs that have not yet been identified. We do not yet have a good idea of whether there are significant differences in the coding functions of these RNAs. The first and last exons are quite lengthy, but the interior exons are rather small. Small exons from within the long transcription unit may enter mRNA products by means of alternative splicing patterns. So far, however, we do not know of any functional differences in the Ubx proteins produced by the various modes of expression. The bithorax complex has trans-acting genes and cis-acting regulators | SECTION 6.31.21 © 2004. Virtual Text / www.ergito.com

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Ubx proteins are found in the compartments that correspond to the sites of transcription, that is in T2P-A1 and at lower levels in A2-A8. So the Ubx unit codes for a set of related proteins that are concentrated in the compartments affected by mutations in the Ultrabithorax domain. Ubx proteins are located in the nucleus, and they fall into the general type of transcriptional regulators whose DNA-binding region consists of a homeodomain. We can understand the general function of the BX-C complex by considering the effects of loss-of-function mutations. If the entire complex is deleted, the larva cannot develop the individual types of segments. In terms of parasegments (which are probably the affected units), all the parasegments differentiate in the same way as parasegment 4; the embryo has 10 repetitions of the repeating structure T1P/T2A all along its length, in place of the usual compartments between parasegments 5 and 14. In effect, the absence of BX-C functions allows Antp to be expressed throughout the abdomen, so that all the segments take on the characteristic of a segment determined by Antp; BX-C functions are needed to add more posterior-type information. Each of the transcription units affects successive segments, according to its pattern of expression. So if Ubx alone is present, the larva has parasegment 4 (T1P/T2A), parasegment 5 (T2P/T3A), and then 8 copies of parasegment 6 (T3P/A1A). This suggests that the expression of Ubx is needed for the compartments anterior to A1A. Ubx is also expressed in the more posterior segments, but in the wild type, abdA and AbdB are also present. If they are removed, the expression of Ubx alone in all the posterior segments has the same effect that it usually has in parasegment 6 (T3P/A1A). The addition of abdA to Ubx adds the wild-type pattern to parasegments 7, 8, and 9. In other words, Ubx plus abdA can specify up to compartments A3P/A4A, and in the absence of AbdB, this continues to be the default pattern for all the more posterior compartments. The addition of AbdB is needed to specify parasegments 10-14. The general model for the function of the ANT-C and BX-C complexes is to suppose that additional functions are added to define successive segments proceeding in the posterior direction. It functions by reliance on a combinatorial pattern in which the addition of successive gene products confers new specificities. This explains the rule that a loss-of-function mutation in one of the genes of the ANT-C/BX-C complexes generally allows the gene on the more anterior side of the mutated gene to determine phenotype, that is, loss-of-function results in homeotic transformation of posterior regions into more anterior phenotypes (884). Expression of Ubx in a more anterior segment than usual should have the opposite effect to a loss-of-function; the segment develops a more anterior phenotype. When this is tested by arranging for Ubx to be expressed in the head, the anterior segments are converted to the phenotype of parasegment 6. So lack of expression of Ubx causes a homeotic transformation in which posterior segments acquire more anterior phenotypes; and overexpression of Ubx causes a homeotic transformation in which anterior segments acquire more posterior phenotypes. This type of relationship is true generally for the cluster as a whole, and explains the properties of cis-acting mutations as well as those in the transcription units. These regulatory mutations cause loss of the protein in part of its domain of expression or The bithorax complex has trans-acting genes and cis-acting regulators | SECTION 6.31.21 © 2004. Virtual Text / www.ergito.com

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cause additional expression in new domains. So they may have either loss-of-function or gain-of-function phenotypes (or sometimes both). The most common is loss-of-function in an individual compartment. For example, bx specifically controls expression of Ubx in compartment T3A; a bx mutation loses expression of Ubx in that compartment, which is therefore transformed to the more anterior type of T2A. This example is typical of the general rule for individual cis-acting mutations in the complex; each converts a target compartment so that it develops as though it were located at the corresponding position in the previous segment. The order of the cis-acting sites of mutation on the chromosomes reflects the order of the compartments in which they function. So the expression of Ubx in parasegments 4, 5, 6 is controlled sequentially by abx (affects parasegment 5), bx (affects T3A), etc. The presence of only 3 genes within the BX-C complex poses two major questions. First, how do the combinations of 3 proteins specify the identity of 10 parasegments? One possibility is that there are quantitative differences in the various regions, allowing for the same sort of varying responses in target genes that we described previously for the combinatorial functioning of the segmentation genes. Second, how do the proteins function in different tissue types? The pattern of expression described above refers generally to the epidermis; the development of other tissues is controlled in a way that is parallel, but not identical. For example, although Ubx is expressed in all posterior segments up to A8 in the epidermis, in mesoderm, it is repressed posterior of segment A7. The posterior boundary reflects repression by abdA, since in abdA mutants, Ubx expression extends posterior in the mesoderm. Why are loci involved in regulating development of the adult insect from the embryonic larva different from genes coding for the everyday proteins of the organism? Is their enormous length necessary to generate the alternative products? Could it be connected with some timing mechanism, determined by how long it takes to transcribe the unit? At a typical rate of transcription, it would take ~100 minutes to transcribe Antp, which is a significant proportion of the 22 hour duration of D. melanogaster embryogenesis. Proceeding from anterior to posterior along the embryo, we encounter the changing patterns of expression of the genes of the ANT-C and BC-C loci. What controls their transcription? As in the case of the segmentation loci, the homeotic loci are controlled partially by the genes that were expressed at the previous stage of development, and partially by interactions among themselves. For example, the expression of Ubx is changed by mutations in bicoid, hunchback, or Kruppel. The anterior boundary of expression respects the parasegment border defined by ftz and eve. The general principle is that all of these regulatory genes function by controlling transcription, either by activating it or by repressing it, and that the gene products may exert specific effects by both qualitative and quantitative combinations.

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Reviews 3709. Lewis, E. B. (1985). Regulation of the genes of the bithorax complex in Drosophila.Drosophila. Cold Spring Harb Symp Quant Biol 50, 155-164.

References 881. Lewis, E. B. (1978). A gene complex controlling segmentation in Drosophila. Nature 276, 565-570. 884. Beachy, P. A., Helfand, S. L., and Hogness, D. S. (1985). Segmental distribution of bithorax complex proteins during Drosophila development. Nature 313, 545-551. 885. Karch, F., Weiffenbach, B., Peifer, M., Bender, W., Duncan, I., Celniker, S., Crosby, M., and Lewis, E. B. (1985). The abdominal region of the bithorax complex. Cell 43, 81-96. 895. Martin, C. H. et al. (1995). Complete sequence of the bithorax complex of Drosophila. Proc. Natl. Acad. Sci. USA 92, 8398-8402.


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6.31.22 The homeobox is a common coding motif in homeotic genes Key Terms The homeobox describes the conserved sequence that is part of the coding region of D. melanogaster homeotic genes; it is also found in amphibian and mammalian genes expressed in early embryonic development. Paralogs are highly similar proteins that are coded by the same genome. Key Concepts

• The homeobox codes for a 60 amino acid protein domain that is a DNA-binding motif.

• Many Drosophila homeotic and segmentation genes code for transcription factors that use a homeodomain to bind DNA.

• The homeodomain does not fully specify the target DNA site, which is influenced by protein-protein combinatorial interactions.

• Homeodomains are found in important regulators of development in a wide range of organisms.

• A vertebrate Hox cluster contains several genes that have homeoboxes, related to the genes in the ANT-C and BX-C fly loci.

• The Hox genes play roles in vertebrate development that are analogous to the roles of the fly genes.

The three groups of genes that control D. melanogaster development – maternal genes, segmentation genes, and homeotic genes – regulate one another and (presumably) target genes that code for structural proteins. Interactions between the regulator genes have been defined by analyses that show defects in expression of one gene in mutants of another. However, we have identified rather few of the structural targets on which these groups of genes act to cause differentiation of individual body parts. Consistent with the idea that the segmentation genes code for proteins that regulate transcription, the genes of 3 gap loci (hb, Kr, kni) contain zinc finger motifs. As first identified in the transcription factors TFIIIA and Sp1 (see Figure 22.13), these motifs are responsible for making contacts with DNA. The products of other loci in the gap class also have DNA-binding motifs; giant encodes a protein with a basic zipper motif, and tailless encodes a protein that resembles the steroid receptors. This suggests that the general function of gap genes is to function as transcriptional regulators. Conserved motifs are found in many of the homeotic and segmentation genes. The The homeobox is a common coding motif in homeotic genes | SECTION 6.31.22 © 2004. Virtual Text / www.ergito.com

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most common of the conserved motifs is the homeobox, a 180 bp region located near the 3 ′ end in several segmentation and homeotic genes. There are ~40 genes in Drosophila that contain a homeobox, and almost all are known to be involved in developmental regulation. (The homeobox was first identified by its predominance in the homeotic genes, from which it took its name.) The protein sequence coded by the homeobox is called the homeodomain; it is a DNA-binding motif in transcription factors (see Figure 22.24 in Molecular Biology 5.22.14 Homeodomains bind related targets in DNA) (883). The fly homeodomains fall into several groups. A major group in Drosophila consists of the homeotic genes in the BX-C/ANT-C complexes; they are called the Antennapedia group. Their homeodomains are 70-80% conserved, and usually occur at the C-terminal end of the protein (see Figure 22.22). A distinct homeodomain sequence is found in the related genes engrailed and invected; it has only 45% sequence conservation with the Antennapedia group (see Figure 22.23). Other types of homeodomain sequences are represented in 2-4 genes each (for review see 360). Many of the Drosophila genes that contain homeoboxes are organized into clusters. Three of the homeotic genes in the BX-C cluster have homeoboxes, the ANT-C complex contains a group of 5 homeotic genes with homeoboxes, and 4 other genes at ANT-C also contain homeoboxes. The homeotic genes at BX-C and ANT-C are sometimes described under the general heading of HOM-C genes (for review see 356). What is the basic function of the HOM-C genes in determining identity on the anterior-posterior axis? We assume that homeodomains with different amino acid sequences recognize different target sequences in DNA. Experiments in which regions have been swapped between different proteins suggest that a major part of the specificity of these proteins rests with the homeodomain. However, the ability to bind to a particular DNA target site may not account entirely for their properties. For example, some of these proteins can either activate or repress transcription in response to the context, that is, their actions depend on the set of other proteins that are bound, not just on recognition of the DNA-binding site (for review see 365). The similarities between the homeodomains of the more closely related members of the group suggest that they could recognize overlapping patterns of target sites. This would open the way for combinatorial effects that could be based on quantitative as well as qualitative differences, that is, there could be competition between proteins with related homeodomains for the same sites. In some cases, different homeoproteins recognize the same target sites on DNA, which poses a puzzle with regard to defining their specificity of action; we assume that there are subtle differences in DNA-binding yet to be discovered, or there are other interactions, such as protein-protein interactions, that play a role. The homeobox motif is extensively represented in evolution. A striking extension of the significance of homeoboxes is provided by the discovery that a DNA probe representing the homeobox hybridizes with the genomes of many eukaryotes. Genes containing homeoboxes have been characterized in detail in frog, mouse, and human DNA. The frog and mammalian genes are expressed in early embryogenesis, which strengthens the parallel with the fly genes, and suggests the possibility that genes containing homeoboxes are involved in regulation of embryogenesis in a variety of species. The homeobox is a common coding motif in homeotic genes | SECTION 6.31.22 © 2004. Virtual Text / www.ergito.com

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Genes in mammals (and possibly all animals) that are related to the HOM-C group have a striking property: like those of the BX-C/ANT-C complexes, they are organized in clusters. The individual mammalian genes are called Hox genes. A cluster of Hox genes may extend 20-100 kb and contain up to 10 genes. Four Hox clusters of genes containing homeoboxes have been characterized in the mouse and human genomes. Their organization is compared with the two large fly clusters in Figure 31.39 (889).

Figure 31.39 Mouse and human genomes each contain 4 clusters of genes that have homeoboxes. The order of genes reflects the regions in which they are expressed on the anterior-posterior axis. The Hox genes are aligned with the fly genes according to homology, which is strong for groups 1, 2, 4, and 9. The genes are named according to the group and the cluster, e.g., HoxA1 is the most anterior gene in the HoxA group. All Hox genes are present in both man and mice except for some mouse genes missing from cluster C.

By comparing the sequences of the homeoboxes (and sometimes other short regions), the mammalian genes can be placed into groups that correspond with the fly genes. This is shown by vertical alignment in the figure. For example, HoxA4 and HoxB4 are best related to Dfd. When these relationships are defined for the cluster as a whole, it appears that within each cluster we can recognize a series of genes that are related to the genes in the ANT-C and BX-C clusters. Groups 1-9 in the mammalian loci are defined as corresponding to the genes of the ANT-C and BX-C loci organized end to end in anterior-posterior orientation. Groups 10-13 appear to have arisen by tandem duplications and divergence of group 9 (the AbdB homologue). The corresponding loci in each cluster are sometimes called paralogs (for example, HoxA4 and HoxB4 are paralogous). This situation could have arisen if the fly and mammalian loci diverged at a point when there was only a single complex, containing all of the genes that define anterior-posterior polarity. The organism Amphioxus, which corresponds to a line of evolution parallel to the vertebrates, has a single Hox cluster containing one member of each paralogous group; this appears to be a direct representative of the original cluster. During evolution, the Drosophila genes broke into two separate clusters, The homeobox is a common coding motif in homeotic genes | SECTION 6.31.22 © 2004. Virtual Text / www.ergito.com

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while the entire group of mammalian genes became duplicated, some individual members being lost from each complex after the duplication (891). The parallel between the mouse and fly genes extends to their pattern of spatial expression. The genes within a Hox cluster are expressed in the embryo in a manner that matches their organization in the genome. Progressing from the left toward the right end of the cluster drawn in Figure 31.39, genes are expressed in the embryo in locations progressively more restricted to the posterior end. The patterns of expression for fly and mouse are compared schematically in Figure 31.40. The domain of expression extends strongly to the posterior boundary shown in the figure, and then tails off into more posterior segments (for review see 361; 364).

Figure 31.40 A comparison of ANT-C/BX-C and HoxB expression patterns shows that the individual gene products share a progressive localization of expression towards the more posterior of the animal proceeding along the gene cluster from left to right. Expression patterns show the regions of transcription in the fly epidermis at 10 hours, and in the central nervous system of the mouse embryo at 12 days.

These results raise the extraordinary possibility that the clusters of genes not only share a common evolution, but also have maintained a common general function in which genome organization is related to spatial expression in fly and mouse, and The homeobox is a common coding motif in homeotic genes | SECTION 6.31.22 © 2004. Virtual Text / www.ergito.com

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there is some correspondence between the homologous genes. The idea of such a relationship is strengthened by the observation that ectopic expression of mouse HoxD4 or HoxB6 in Drosophila cause homeotic transformations virtually identical to those caused by homeotic expression of Dfd or Antp, respectively! Since the homology between these mouse and fly proteins rests almost exclusively with their homeodomains, this reinforces the view that these domains determine specificity. There are some differences in the apparent behavior of the vertebrate Hox clusters and the ANT-C/BX-C fly clusters: • The Hox genes are small, and there is a greater number of protein-coding units. The mouse HoxB cluster is ~120 kb and contains 9 genes. The connection between genomic position and embryonic expression is analogous to that in Drosophila, but describes only the genes themselves; we have no information about cis-acting sites. Of course, this may be a consequence of the much greater difficulty in generating mutations in vertebrates. However, our present information identifies the control of Hox genes only by promoters and enhancers in the region upstream of the startpoint. It remains to be seen whether there is any counterpart to the very extensive and complex regulatory regions of the Drosophila homeotic genes. • In Drosophila, each gene is unique; but in vertebrates, the duplication of the clusters enables multiple genes (paralogs) to have the same or very similar patterns of expression. If the paralogs have redundant or partially redundant functions, so that the absence of one product may be at least partially substituted by the corresponding protein of another cluster, the effects of mutations will be minimized. Disruptions of Hox genes in mice often generate recessive lethals. In the examples of HoxA1 and HoxA3 various structures of the head and thorax are absent. Not all of the structures that usually express the mutant genes are missing, suggesting that there is indeed some functional redundancy, that is, other Hox genes of group 1 or group 3 can substitute in some but not all other tissues for the absence of the HoxA gene. Homeotic transformations are less common with mutants in mice than in Drosophila, but sometimes occur. Loss of HoxC8, for example, causes some skeletal segments to show more anterior phenotypes. It remains to be seen whether this is a general rule. Ectopic expression of Hox genes has been used successfully to demonstrate that gain-of-function can transform the identity of a segment towards the identity usually conferred by the gene. The most common type of effect in Drosophila is to transform a segment into a phenotype that is usually more posterior; in effect, the expression of the homeotic gene has added additional information that confers a more posterior identity on the segment. Similar effects are observed in some cases in the mouse. However, the pattern is not completely consistent (892). Taken together, these results make it clear that the Hox genes resemble their counterparts in Drosophila in determining patterning along the anterior-posterior axis. It may be the case that there is a combinatorial code of Hox gene expression, or there may be differences in degree of functional redundancy between paralogs, but we cannot yet provide a systematic model for their role in determining pattern. The homeobox is a common coding motif in homeotic genes | SECTION 6.31.22 © 2004. Virtual Text / www.ergito.com

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The most striking feature of organization of the Hox loci still defies explanation: why has the organization of the cluster, in which genomic position correlates with embryonic expression, been maintained in evolution? The obvious explanation is that there is some overall control of gene expression to ensure that it proceeds through the cluster, with the result that a gene could be properly expressed only when it is within the cluster. But this does not appear to be true, at least for those individual cases in which genes have been removed from the cluster. Analysis of promoter regions suggests that a Hox gene may be controlled by a series of promoter or enhancer elements that together ensure its overall pattern of expression. Usually these elements are in the region upstream of the startpoint. For example, HoxB4 expression can be reconstructed as the sum of the properties of a series of such elements, tested by introducing appropriate constructs to make transgenic mice. But then why should there have been evolutionary pressure to retain genes in an ordered cluster? One possibility is that an enhancer for one gene might be embedded within another gene, in such a way that, even if an individual gene could function when translocated elsewhere, its removal would impede the expression of other gene(s). An indication that there may be something special about the organization of the region is given by the fact that it has an unusually high density of conserved noncoding sequences and an unusually low density of insertions such as transposons (1442). This suggests the existence of large scale regulatory elements that we have not yet identified. Last updated on 2-16-2001

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Reviews 356. Regulski, M., Harding, K., Kostriken, R., Karch, F., Levine, M., and McGinnis, W. (1985). Homeo box genes of the Antennapedia and bithorax complexes of Drosophila. Cell 43, 71-80. 360. Scott, M. P (1989). The structure and function of the homeodomain. Biochim. Biophys. Acta 989, 25-48. 361. Hunt, P. and Krumlauf, R. (1992). Hox codes and positional specification in vertebrate embryonic axes. Annu. Rev. Cell Biol. 8, 227-256. 364. Krumlauf, R. (1994). Hox genes in vertebrate development. Cell 78, 191-201. 365. McGinnis, W. and Krumlauf, R. (1992). Homeobox genes and axial patterning. Cell 68, 283-302.

References 883. McGinnis, W. et al. (1984). A homologous protein-coding sequence in Drosophila homeotic genes and its conservation in other metazoans. Cell 37, 403-408. 889. Graham, A., Papalopulu, N., and Krumlauf, R. (1989). The murine and Drosophila homeobox gene complexes have common features of organization and expression. Cell 57, 367-378. 891. Garcia-Fernandez, J. and Holland, P. W. H. (1994). Archetypal organization of the amphioxus Hox gene cluster. Nature 370, 563-566. 892. Malicki, J., Schughart, K., and McGinnis, W. (1990). Mouse hox-22 specifies thoracic segmental identity in Drosophila embryos and larvae. Cell 63, 961-967. 1442. Sachidanandam, R. et al. (2001). A map of human genome sequence variation containing 1.42 million single nucleotide polymorphisms. The International SNP Map Working Group. Nature 409, 928-933.


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6.31.23 Summary The development of segments in Drosophila occurs by the actions of segmentation genes that delineate successively smaller regions of the embryo. Asymmetry in the distribution of maternal gene products is established by interactions between the oocyte and surrounding cells. This leads to the expression of the gap genes, in 4 broad regions of the embryo. The gap genes in turn control the pair-rule genes, each of which is distributed in 7 stripes; and the pair-rule genes define the pattern of expression of the segment polarity genes, which delineate individual compartments. At each stage of expression, the relevant genes are controlled both by the products of genes that were expressed at the previous stage, and by interactions among themselves. The segmentation genes act upon the homeotic genes, which determine the identities of the individual compartments. Each of the 4 maternal systems consists of a cascade which generates a locally distributed or locally active morphogen. The morphogen either is a transcription factor or causes the activation of a transcription factor. The transcription factor is the last component in each pathway. The major anterior-posterior axis is determined by two systems: the anterior system establishes a gradient of bicoid from the anterior pole; and the posterior system produces nanos protein in the posterior half of the egg. These systems function to define a gradient of hunchback protein from the anterior end, with broad bands of knirps and giant in the posterior half. The terminal system acts to produce localized events at both termini. The dorsal-ventral system produces a gradient of nuclear localization of dorsal protein on the ventral side, which represses expression of dpp and zen; this leads to the ventral activation of twist and snail, and the dorsal-side activation of dpp and zen. Each system is initiated by localization of a morphogen in the egg as a result of its interaction with the surrounding cells. For the anterior and posterior systems, this takes the form of localizing an RNA; bicoid mRNA is transported into and localized at the anterior end, and nanos mRNA is transported to the posterior end. For the dorsal-ventral system, the Toll receptor is located ubiquitously on the oocyte membrane, but the spatzle ligand is activated ventrally and therefore triggers the pathway on the ventral side. The pathway resembles the mammalian IL-1 signal transduction pathway and culminates in the phosphorylation of cactus, which regulates the dorsal transcription factor. On the dorsal side of the embryo, the morphogen Dpp is released; it is a member of the TGF β family that diffuses to interact with its receptor. A ligand-receptor interaction involving related members of the TGF β/receptor families is also employed in a comparable role in vertebrate development. The early embryo consists of a syncytium, in which nuclei are exposed to common cytoplasm. It is this feature that allows all 4 maternal systems to control the function of a nucleus according to the coordinates of its position on the anterior-posterior and dorsal-ventral axes. At cellular blastoderm, zygotic RNAs are transcribed, and the developing embryo becomes dependent upon its own genes. Cells form at the Summary | SECTION 6.31.23 © 2004. Virtual Text / www.ergito.com

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blastoderm stage, after which successive interactions involve a cascade of transcriptional regulators. Three gap genes are zinc-finger proteins, and one is a basic zipper protein. Their concentrations control expression of the pair-rule genes, which are also transcription factors. In particular, the expression of eve and ftz controls the boundaries of compartments, functioning in every other segment. The segment polarity genes represent the first step in the developmental cascade that involves functions other than transcription factors. Interactions between the segmentation gene products define unique combinations of gene expression for each segment. The segment polarity genes include proteins involved in cell-cell interactions as well as transcription factors. The basic circuitry that determines the anterior and posterior polarities of compartments is maintained by an autoregulatory interaction between the cells at the boundary. An anterior compartment secretes wingless protein, which acts upon the cell on the posterior side. This causes engrailed to be expressed in the posterior cell, which in turn causes secretion of hedgehog on the anterior side. Hedgehog causes the anterior cell to express wingless. Homeotic genes impose the program that determines the unique differentiation of each segment. The complex loci ANT-C and BX-C each contain a cluster of functions, whose spatial expression on the anterior-posterior axis reflects genetic position in the cluster. Each cluster contains one exceedingly large transcription unit as well as other, shorter units. Many of the transcription units (including the largest genes, Ubx and Antp) have patterns of alternative splicing, but no significance has been attributed to this yet. Proceeding from left to right in each cluster, genes are expressed in more posterior tissues. The genes are expressed in overlapping patterns in such a way that addition of a function confers new features of posterior identity; thus loss of a function results in a homeotic transformation from posterior to more anterior phenotype. The genes are controlled in a complex manner by a series of regulatory sites that extend over large regions; mutations in these sites are cis-acting, and may cause either loss-of-function or gain-of-function. The cis-acting mutations tend to act on successive segments of the fly, by controlling expression of the homeotic proteins. The genes of the ANT-C and BX-C loci, and many segmentation genes (including the maternal gene bicoid and most of the pair-rule genes) contain a conserved motif, the homeobox. Homeoboxes are also found in genes of other eukaryotes, including worms, frogs, and mammals. In each case, these genes are expressed during early embryogenesis. In mammals, the Hox genes (which specify homeodomains in the Antennapedia class) are organized in clusters. There are 4 Hox clusters in both man and mouse. These clusters can be aligned with the ANT-C/BX-C clusters in such a way as to recognize homologies between genes at corresponding positions. Proceeding towards the right in a Hox cluster, a gene is expressed more towards the posterior of the embryo. The Hox genes have roles in conferring identity on segments of the brain and skeleton (and other tissues). The analogous clusters represent regulators of embryogenesis in mammals and flies. Hox clusters may be a characteristic of all animals. Drosophila genes containing homeoboxes form an intricate regulatory network, in which one gene may activate or repress another. The relationship between the sequence of the homeodomain, the DNA target it recognizes, and the regulatory Summary | SECTION 6.31.23 © 2004. Virtual Text / www.ergito.com

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consequences, remains to be fully elucidated. Specificity in target choice appears to reside largely in the homeodomain; we have yet to explain the abilities of a particular homeoprotein to activate or to repress gene transcription at its various targets. The general principle is that segmentation and homeotic genes act in a transcriptional cascade, in which a series of hierarchical interactions between the regulatory proteins is succeeded by the activation of structural genes coding for body parts. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.6.31.23


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SUPPLEMENTS

7.32.1 DNA reassociation kinetics Key Terms A Cot curve is a plot of the extent of renaturation of DNA against time. Complexity is the total length of different sequences of DNA present in a given preparation. The fast component of a reassociation reaction is the first to renature and contains highly repetitive DNA. Intermediate component(s) of a reassociation reaction are those reacting between the fast (satellite DNA) and slow (nonrepetitive DNA) components; contain moderately repetitive DNA. The slow component of a reassociation reaction is the last to reassociate; usually consists of nonrepetitive DNA. Nonrepetitive DNA shows reassociation kinetics expected of unique sequences. Repetitive DNA behaves in a reassociation reaction as though many (related or identical) sequences are present in a component, allowing any pair of complementary sequences to reassociate. The repetition frequency is the (integral) number of copies of a given sequence present in the haploid genome; equals 1 for nonrepetitive DNA, >2 for repetitive DNA. Highly repetitive DNA (Simple sequence DNA) is the first component to reassociate and is equated with satellite DNA. The stringency of a hybridization describes describes the effect of conditions on the degree of complementarity that is required for reaction. At the most stringent conditions, only exact complements can hybridize. As the stringency is lowered, an increasing number of mismatches can be tolerated between the two strands that are hybridizing. A tracer is a radioactively labeled nucleic acid component included in a reassociation reaction in amounts too small to influence the progress of reaction.

The general nature of the eukaryotic genome can be assessed by the kinetics of reassociation of denatured DNA. Reassociation between complementary sequences of DNA occurs by base pairing. This reverses the process of denaturation by which they were separated (see Figure 1.17). The kinetics of the reassociation reaction reflect the variety of sequences that are present; so the reaction can be used to quantitate genes and their RNA products. Figure S 1 describes the reaction. Renaturation of DNA depends on random collision of the complementary strands, and follows second-order kinetics. The reaction for any particular DNA can be characterized by conditions required for half-completion. This is the product of C0× t½ and is called the Cot½;. It is inversely proportional to the rate constant. Since the Cot½ is the product of the concentration and time required DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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to proceed halfway, a greater Cot½ implies a slower reaction.

Figure S 1 A DNA reassociation reaction is described by the Cot½.

The reassociation of DNA usually is followed in the form of a Cot curve, which plots the fraction of DNA that has reassociated (1 –C/C0) against the log of the Cot. Figure S 2 gives Cot curves for several simple genomes. The form of each curve is similar, with renaturation occurring over an ~100-fold range of Cot values between the points of 10% reaction and 90% reaction. But the Cot½ for each curve is different.

Figure S 2 Rate of reassociation is inversely proportional to the length of the reassociating DNA.

The genomes in Figure S 2 represent a series of DNAs. Each is unique in sequence, DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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and they become progressively longer. The Cot½ is directly related to the amount of DNA in the genome. This reflects a situation in which, as the genome becomes more complex, there are fewer copies of any particular sequence within a given mass of DNA. For example, if the C0 of DNA is 12 pg, it will contain 3000 copies of each sequence in a bacterial genome whose size is 0.004 pg, but will contain only 4 copies of each sequence present in a eukaryotic genome of size 3 pg. So the same absolute concentration of DNA measured in moles of nucleotides per liter (the C0) will provide a concentration of each eukaryotic sequence that is 3000/4 = 750× less than that of each bacterial sequence. Since the rate of reassociation depends on the concentration of complementary sequences, for the eukaryotic sequences to be present at the same relative concentration as the bacterial sequences, it is necessary to have 750× more DNA (or to incubate the same amount of DNA for 750 times longer). So the Cot½ of the eukaryotic reaction is 750× the Cot½ of the bacterial reaction. The Cot½ of a reaction therefore indicates the total length of different sequences that are present. This is described as the complexity, usually given in base pairs. The Cot½ for the renaturation of the DNA of any genome (or part of a genome) is proportional to its complexity. The complexity of any DNA can be determined by comparing its Cot½ with that of a standard DNA of known complexity. Usually E. coli DNA is used as a standard. Assuming that the E. coli genome of 4.2 × 106 bp consists of unique sequences:

When the DNA of a eukaryotic genome is characterized by reassociation kinetics, usually the reaction occurs over a range of Cot values spanning up to eight orders of magnitude. This is much broader than the 100-fold range expected from the examples of Figure S 2. The reason is that each of these curves follows the equation that describes the kinetics of reassociation for a single component. A eukaryotic genome actually includes several such components, each reassociating with its own characteristic kinetics. The Cot curve reveals a crucial difference between bacterial and eukaryotic genomes: bacterial genomes essentially consist of a single kinetic component, but eukaryotic genomes are much more complex. Figure S 3 shows the reassociation of a (hypothetical) eukaryotic genome, starting at a Cot of 10–4 and terminating at a Cot of 104. The reaction falls into three distinct phases, outlined by the shaded boxes. Each of these phases represents a different kinetic component of the genome:

DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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Figure S 3 The reassociation kinetics of eukaryotic DNA show three types of component (indicated by the shaded areas). The arrows identify the Cot½ values for each component.

• The fast component is the first fraction to reassociate. In this case, it represents 25% of the total DNA, renaturing between Cot values of 10–4 and ~2 × 10–2, with a Cot½ value of 0.0013. • The next fraction is called the intermediate component. This represents 30% of the DNA. It renatures between Cot values of ~0.2 and 100, with a Cot½ value of 1.9. • The slow component is the last fraction to renature. This is 45% of the total DNA; it extends over a Cot range from ~100 to ~10,000, with a Cot½ of 630. To calculate the complexities of these fractions, each must be treated as an independent kinetic component whose reassociation is compared with a standard DNA. The slow component represents 45% of the total DNA, so its concentration in the reassociation reaction is 0.45 of the measured C0 (which refers to the total amount of DNA present). The Cot½ applying to the slow fraction alone is 0.45 × 630 = 283. Suppose that under these conditions, E. coli DNA reassociates with a Cot½ of 4.0. This corresponds to a complexity for the slow fraction of 3.0 × 108 bp (= 4.2 × 106× 283 /4). Treating the other components in a similar way shows that the intermediate component has a complexity of 6 × 105 bp, and the fast component has a complexity of only 340 bp. This provides a quantitative basis for our statement that, the faster a component reassociates, the lower is its complexity. DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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Reversing the argument, suppose we took three DNA preparations, each containing a unique sequence of the appropriate length (340 bp, 6 × 105 bp, and 3 × 108 bp, respectively) and mixed them in the proportions of mass 25:30:45. Each would renature as though it were a single component. Together the mixture would display the same kinetics as those determined for the whole genome of Figure S 3. The complexity of the slow component corresponds with its physical size. Suppose that the genome reassociating in Figure S 3 has a haploid DNA content of 7.0 × 108 bp, determined by chemical analysis. Then 45% of it is 3.15 × 108 bp, which is the same (within experimental error) as the value of 3.0 × 108 bp measured by the kinetics of reassociation. The complexity of the slow component corresponds to its physical length. The slow component comprises sequences that are unique in the genome: on denaturation, each single-stranded sequence is able to renature only with the corresponding complementary sequence. This part of the genome is the sole component of prokaryotic DNA and is usually a major component in eukaryotes. It is called nonrepetitive DNA. What is the nature of the components that renature more rapidly than the nonrepetitive (slow) DNA? In the example of Figure S 2, the intermediate component occupies 30% of the genome. Its chemical complexity is 0.3 × 7 × 108 = 2.1 × 108 bp. But its kinetic complexity is only 6 × 105 bp. The unique length of DNA that corresponds to the Cot½ for reassociation is much shorter than the total length of the DNA chemically occupied by this component in the genome. In other words, the intermediate component behaves as though consisting of a sequence of 6 × 105 bp that is present in 350 copies in every genome (because 350 × 6 × 105 = 2.1 × 108). Following denaturation, the single strands generated from any one of these copies are able to renature with their complements from any one of the 350 copies. This effectively raises the concentration of reacting sequences in the reassociation reaction, explaining why the component renatures at a lower Cot½. Sequences that are present in more than one copy in each genome are called repetitive DNA. The number of copies present per genome is called the repetition frequency (f). Repetitive DNA is often classed into two general types, corresponding approximately to the intermediate and fast components of Figure S 3: • Moderately repetitive DNA occupies the intermediate fraction, usually reassociating in a range between a Cot of 10–2 and that of nonrepetitive DNA. • Highly repetitive DNA occupies the fast fraction, reassociating before a Cot of 10–2 is reached. The behavior of a repetitive DNA component represents only an average that is useful for describing its sequences. The relevant parameters do not necessarily represent the properties of any particular sequence. DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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Molecular Biology The moderately repetitive component of Figure S 3 includes a total length of 6 × 105 bp of DNA, repeated ~350× per genome. But this does not correspond to a single, identifiable, continuous length of DNA. Instead, it is made up of a variety of individual sequences, each much shorter, whose total length together comes to 6 × 105 bp. These individual sequences are dispersed about the genome. Their average repetition is 350, but some will be present in more copies than this and some in fewer. When a eukaryotic genome is analyzed by reassociation kinetics, the individual sequence components are rarely so well separated as shown in Figure S 3. In fact, they often overlap extensively, so that in reality there is probably a continuum of repetitive components, reassociating over a range from >10× to >20,000× that of the nonrepetitive component. The different components of eukaryotic DNA can be isolated in the form of the DNA that becomes double-stranded after renaturation to a particular Cot value. The properties of renatured nonrepetitive and repetitive DNA differ significantly. Nonrepetitive DNA forms duplex material that behaves very much like the original preparation of DNA before its denaturation. When denatured again, the duplex molecules melt sharply at a Tm only slightly below that of the original native DNA. This shows that strand reassociation has been accurate: each unique sequence has annealed with its exact complement. Different behavior is shown by renatured repetitive DNA. The reassociated double strands tend to melt gradually over rather a wide temperature range, as shown in Figure S 4. This means that they do not consist of exactly paired molecules. Instead, they must contain appreciable mispairing. The more mispairing in a particular molecule, the fewer hydrogen bonds need be broken to melt it, and thus the lower the T m.


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Figure S 4 The denaturation of reassociated nonrepetitive DNA takes place over a narrow temperature range close to that of native DNA, but reassociated repetitive DNA melts over a wide temperature range.

The breadth of the melting curve shows that renatured repetitive DNA contains a spectrum of sequences, ranging from those that have been formed by reassociation between sequences that are only partially complementary, to those formed by reassociation between sequences that are very nearly or even exactly complementary. How can this happen? Repetitive DNA components consist of families of sequences that are not exactly the same, but are related. The members of each family consist of a set of nucleotide sequences that are sufficiently similar to renature with one another. The differences between the individual members are the result of base substitutions, insertions, and deletions, all creating points within the related sequences at which the complementary strands cannot base pair. The proportion of these changes establishes the relationship between any two sequences. When two closely related members of the family renature, they form a duplex with high Tm.. When two more distantly related members associate, they form a duplex with a lower Tm. Overall, we see the broad range represented in the figure. The ability of related but not identical complementary sequences to recognize each other can be controlled by the stringency of the conditions imposed for reassociation. A higher stringency is imposed by (for example) an increase in temperature, which requires a greater degree of complementarity to allow base pairing. So by performing the hybridization reaction at high temperatures, reassociation is restricted to rather closely related members of a family; at lower temperatures, more distantly related members may anneal. The measured size of a repetitive family is arbitrary, since it is determined by the hybridization conditions. DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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Moderately repetitive DNA is dispersed throughout the genome, usually in the form of relatively short individual sequences. It is responsible for the high degree of secondary structure formation in pre-mRNA, when (inverted) repeats in the introns pair to form duplex regions. Highly repetitive DNA often forms discrete clusters (see Molecular Biology 1.4 Clusters and repeats). Neither class represents protein. The genome sequence components represented in mRNA can be determined by using the RNA as a tracer in a reassociation experiment. A very small amount of radioactively labeled RNA (or cDNA) is included together with a much larger amount of cellular DNA. The tracer RNA (or cDNA) participates in the reaction as though it were just another member of the sequence component from which it was transcribed. The Cot values at which the labeled RNA hybridizes identify the repetition frequencies of the corresponding genomic sequences. Figure S 5 shows a typical result for a population of mRNAs. A small proportion of the RNA, generally 10% or less, hybridizes with a Cot½ corresponding to moderately repetitive sequences. The major component hybridizes with nonrepetitive DNA.

Figure S 5 The hybridization of an mRNA tracer preparation in a reassociation curve shows that most mRNA sequences are derived from nonrepetitive DNA, the remainder from moderately repetitive DNA, and none from highly repetitive DNA.

Reassociation analysis can be also used to measure the complexity of an RNA population. One method is to hybridize nonrepetitive DNA with an excess of RNA; the proportion of the DNA that is bound at saturation identifies the complexity of the RNA population. Another method is to follow the kinetics of hybridization between a excess of an RNA population and a DNA copy prepared from it. This is exactly analogous to reassociation analysis of genomic DNA. The reaction is described in terms of the Rot½(where R0 is the starting concentrationof RNA). In looking at the DNA that hybridizes with mRNA, we are basically examining the exons in the genome. The conclusion therefore is that most exons are present at low repetition frequency – depending on the stringency of hybridization, they may be DNA reassociation kinetics | SECTION 7.32.1 © 2004. Virtual Text / www.ergito.com

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unique or present in a small number of copies. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.7.32.1


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SUPPLEMENTS

7.32.2 Mendel's laws and dominance Key Terms An allele is one of several alternative forms of a gene occupying a given locus on a chromosome. An individual is said to be homozygous when it has identical alleles of a given gene. An individual is said to be heterozygous when it has different alleles of a given gene on each of its homologous chromosomes. Complete dominance is the state in which the phenotype is the same when the dominant allele is homozygous or heterozygous. A dominant allele determines the phenotype displayed in a heterozygote with another (recessive) allele. A recessive allele is obscured in the phenotype of a heterozygote by the dominant allele, often due to inactivity or absence of the product of the recessive allele. Incomplete dominance is a state in which the heterozygote has a phenotype in between that of each of the homozygotes. Two alleles are said to be codominant when they are each equally evident in the phenotype of the heterozygote. Mendel's law of independent assortment states that the assortment of one gene does not influence the assortment of another. A parental genotype is one that is identical to the genotype of one of the contributing parents. Recombinant progeny have a different genotype from that of either parent.

The essential attributes of the gene were defined by Mendel more than a century ago. As he concluded in his analysis of pea genetics in 1865: "The law of combination of different characters, which governs the development of the hybrids, finds therefore its explanation in the principle enunciated, that the hybrids produce egg cells and pollen cells, which in equal numbers, represent all constant forms which result from combinations of the characters brought together in fertilization." Summarized in his two laws, the gene was recognized as a "particulate factor" that passes unchanged from parent to progeny. A gene may exist in alternative forms that determine the expression of some particular characteristic. For example, the color of a flower may be red or white. The forms of the gene are called alleles. Mendel's first law describes the segregation of alleles: alleles have no permanent effect on one another when present in the same plant, but segregate unchanged by passing into different gametes. When an organism has two identical alleles of a gene, it is said to be homozygous (or true-breeding) for the trait conveyed by that gene. If the alleles are different, the organism is heterozygous (or hybrid). The phenotype of a homozygote directly Mendel's laws and dominance | SECTION 7.32.2 © 2004. Virtual Text / www.ergito.com

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reflects the genotype of the (single type of) allele, but the phenotype of a heterozygote depends on the relationship between the types of alleles that are present. Mendel's first law recognizes that the genotype of a heterozygote includes both alleles, irrespective of the phenotype that is displayed. When a homozygote for one allele is crossed with a homozygote for another allele, all the progeny in the first (F1) generation are heterozygotes with the same phenotype. But when the heterozygotes are crossed with one another to generate a second (F2) generation, the genotypes of the original parents reappear. The critical point is that the alleles must consist of discrete physical entities that contribute independently (or fail to contribute) to the phenotype. Figure S 6 shows how the results of such crosses differ according to the type of relationship between alleles:

Figure S 6 Mendel#s first law: alleles segregate each generation.

• The case analyzed by Mendel corresponds to complete dominance, and is shown in the first column of results. When one allele is dominant and the other is recessive, the phenotype of a heterozygote is determined by the dominant allele. The recessive allele makes no contribution. The single dominant allele produces the same phenotype that is seen in a wild-type homozygote. The appearance of the heterozygote is indistinguishable from that of the homozygous Mendel's laws and dominance | SECTION 7.32.2 © 2004. Virtual Text / www.ergito.com

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dominant parent. The F1 resembles the dominant parent. Complete dominance generates the classic 3:1 ratio of dominant to recessive phenotypes in the F2. • Some alleles exhibit incomplete dominance (or partial dominance), as shown in the middle column. The phenotype of the heterozygote is intermediate between that of the two homozygotes. In the snapdragon, for example, a cross between red and white generates heterozygotes with pink flowers. However, the same rule is observed that the first hybrid (F1) generation is uniform in phenotype; and the same ratios are generated in the second (F2) generation, except that three phenotypes can be distinguished instead of two (AA is red, 2 Aa are pink, aa is white). This type of situation arises through quantitative effects; the single red allele in the heterozygote produces half as much pigment as the two red alleles in a homozygote. • Alleles are said to be codominant when they contribute equally to the phenotype, as shown in the final column. In human blood groups, the AA and BB combinations are homozygous, and AB is a codominant heterozygote in which the A and B groups are equally expressed. The result is that each genotypic class produces a different phenotype. The F1 has the properties of both parents, and the F2 has the phenotypes A: 2 AB: B. Mendel's second law summarizes the independent assortment of different genes. When a homozygote that is dominant for two different characters is crossed with a homozygote that is recessive for both characters, as before the F1 consists of plants whose phenotype is the same as the dominant parent. But in the next (F2) generation, two general classes of progeny are found: • One class consists of the two parental genotypes. • The other class consists of new phenotypes, representing plants with the dominant feature of one parent and the recessive feature of the other. These are called recombinant types; and they occur in both possible (reciprocal) combinations. Figure S 7 shows that the ratios of the four phenotypes comprising the F2 can be explained by supposing that gamete formation involves an entirely random association between one of the two alleles for the first character and one of the two alleles for the second character. All four possible types of gamete are formed in equal proportion; and then they associate at random to form the zygotes of the next generation. Once again, the phenotypes conceal a greater variety of genotypes.

Mendel's laws and dominance | SECTION 7.32.2 © 2004. Virtual Text / www.ergito.com

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Figure S 7 Mendel#s second law: different genes assort independently in genetic crosses.

The law of independent assortment establishes the principle that the behavior of any pair (or greater number) of genes can be predicted overall by the rules of mathematical combination. The assortment of one gene does not influence the assortment of another. Implicit in this concept is the view that assortment is a matter of statistical probability and not an exact result. The ratio of progeny types will approximate increasingly closely to the predicted proportions as the number of crosses is increased. Appreciation of Mendel's discoveries was inhibited by the lack of any known physical basis for the postulated factors (genes). When the chromosomal theory of inheritance was subsequently proposed, however, it was realized that the behavior of chromosomes at meiosis and fertilization corresponds precisely with the properties of Mendel's particulate units of inheritance. There is an exact parallel between the behavior of chromosomes and Mendel's units of inheritance: • Genes occur in allelic pairs. One member of each pair is contributed by each parent; so the diploid set of chromosomes results from the contribution of a haploid set by each parent.


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• The assortment of nonallelic genes into gametes is independent of (parental) origin; correspondingly, nonhomologous chromosomes undergo independent segregation at meiosis. The critical proviso is that each gamete obtains a complete haploid set, and this condition is fulfilled whether viewed in terms of Mendel's factors or chromosomes. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.7.32.2


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SUPPLEMENTS

7.32.3 Linkage and mapping Key Terms A recombinant genotype is one that consists of a new combination of genes produced by crossing over. A parental genotype is one that is identical to the genotype of one of the contributing parents. Linkage describes the tendency of genes to be inherited together as a result of their location on the same chromosome; measured by percent recombination between loci. A linkage map is a map showing the linear order of genes on a chromosome and the relative distances between them in recombinational units. An allele is one of several alternative forms of a gene occupying a given locus on a chromosome. A backcross describes a genetic cross in which a hybrid strain is crossed to one of its two parental strains. Crossing-over describes the reciprocal exchange of material between chromosomes that occurs during prophase I of meiosis and is responsible for genetic recombination. A chiasma (pl. chiasmata) is a site at which two homologous chromosomes appear to have exchanged material during meiosis. Breakage and reunion describes the mode of genetic recombination, in which two DNA duplex molecules are broken at corresponding points and then rejoined crosswise (involving formation of a length of heteroduplex DNA around the site of joining). Map distance is measured as cM (centimorgans) = percent recombination (sometimes subject to adjustments). A map unit is the distance between two genes that recombine with a frequency of 1%. A locus is the position on a chromosome at which the gene for a particular trait resides; a locus may be occupied by any one of the alleles for the gene. A linkage group includes all loci that can be connected (directly or indirectly) by linkage relationships; equivalent to a chromosome. A marker is an identifiable and inheritable difference that can be mapped to a location on a chromosome. A genetic marker is an allele that is identified with its genetic trait. A molecular marker is a DNA sequence difference that can be identified by molecular methods. Phage T4 is a virus that infects E. coli causing lysis of the bacterium. Rapid lysis (r) mutants display a change in the pattern of lysis of E. coli at the end of an infection by a T-even phage.

Linkage and mapping | SECTION 7.32.3 © 2004. Virtual Text / www.ergito.com

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Mendel's laws predict that genes carried on different chromosomes will segregate independently (for additional description see Molecular Biology Supplement 32.2 Mendel's laws and dominance). However, genes that are on the same chromosome show linked inheritance. The basic observation is that genes on different chromosomes recombine at random from one generation to the next, whereas genes that are linked show a reduction in recombination, that is, they tend to stay together. The results of a genetic cross are analyzed by determining determining the proportions of recombinant genotypes (where an allele of one parent is found with an allele of the other parent) and parental genotypes (which have the same combination of alleles as either parent). Genes on different chromosomes segregate independently, as predicted by Mendel, to give 50% parental and 50% recombinant progeny. Genes on the same chromosome behave differently, because they are present on the same (very long) molecule of DNA. Instead of generating the proportions depicted by independent assortment, the proportion of parental genotypes is greater than expected, because there is a reduction in the formation of recombinant genotypes. The propensity of some characters to remain associated instead of assorting independently is called linkage. Linkage is measured by the per cent recombination between two loci (in formal terms a map distance of 1 centimorgan = 1% recombination). When pairwise combinations of loci on the same chromosome are tested in genetic crosses, loci close to one another are linked, as defined by a map distance 50 cM apart are connected because they show linkage to a locus between them. This genetic map corresponds to the physical existence of the chromosome. A crucial concept in the construction of a genetic map is that the distance between genes does not depend on the particular alleles that are used, but only on the genetic loci. The locus defines the position occupied on the chromosome by the gene representing a particular trait. The various alternative forms of a gene – that is, the alleles used in mapping – all reside at the same location on its particular chromosome. So genetic mapping is concerned with identifying the positions of genetic loci, which are fixed and lie in a linear order. In a mapping experiment, the same result is obtained irrespective of the particular combination of alleles. Figure S 8 shows how a backcross to a recessive homozygote is used to measure linkage. The alleles of the recessive parent make no contribution to the phenotype of the progeny. As a result, the backcross essentially makes it possible to examine directly the genotype of the organism being investigated. In each cross the progeny show an increase in the proportion of parental types (70%) and a decrease in the proportion of recombinant types (30%), compared with the 50% of each type that is expected from independent assortment. The linkage between A and B is measured as 30%.


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Figure S 8 Linkage can be measured by a backcross with a double recessive homozygote.

The smaller the proportion of recombinants in the progeny, the tighter the linkage. A crucial characteristic is that the same proportion of recombinants is obtained irrespective of the arrangement of parental alleles (AB/ab or Ab/aB). And in each case, both of the (reciprocal) recombinant types are present in the same proportions.. Morgan proposed that genetic linkage is the "simple mechanical result of the location of the (genes) in the chromosomes." He suggested that the production of recombinant classes can be equated with the process of crossing-over that is visible during meiosis. Early in meiosis, at the stage when all four copies of each chromosome are organized in a bivalent, pairwise exchanges of material occur between the closely associated (synapsed) chromatids. The visible result of a crossing-over event is called a chiasma, and is illustrated diagrammatically in Figure S 9. A chiasma represents a site at which two of the chromatids in a bivalent have been broken at corresponding points. The broken ends have been rejoined crosswise, generating new chromatids. Each new chromatid consists of material derived from one chromatid on one side of the junction point, with material from the other chromatid on the opposite side. The two recombinant chromatids have reciprocal structures. The event is described as a breakage and reunion.


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Figure S 9 Chiasma formation is responsible for generating recombinants.

If the likelihood that a chiasma will form between two points on a chromosome depends on their distance apart, genes located near each other will tend to remain together. As the distance decreases, the probability of crossing-over between them will decrease. If crossing-over is responsible for recombination, the closer genes lie to one another, the more tightly they will be linked. Reversing the argument, genetic linkage can be taken to be a measure of physical distance. The extent of recombination between two genes on the same chromosome can be used as a map distance to measure their relative locations. The formula to measure genetic distance is:

Map units are defined as 1 unit (or centiMorgan, abbreviated cM) equals 1% crossover. For short distances (. The locus defines the position occupied on the chromosome by the gene representing a particular trait. The various alternative forms of a gene – that is, the alleles used in mapping – all reside at the same location on its particular chromosome. So genetic mapping is concerned with identifying the positions of genetic loci, which are fixed and lie in a linear order. In a mapping experiment, the same result is obtained irrespective of the particular combination of alleles (see Figure S 8, where in either combination, there are 70% parental and 30% recombinant types). Linkage is not displayed between all pairs of genes located on a single chromosome. The maximum recombination between two loci is the 50% corresponding to the independent segregation predicted by Mendel's second law. (Although there is a high probability that recombination will occur between two genes lying far apart on a chromosome, each individual recombination event involves only two of the four associated chromatids, so there is a limit of 50% recombination between the genes.) In spite of their presence on the same chromosome, genes that are far apart therefore assort independently. But although they show no direct linkage, each can be linked to genes that lie between them. This allows the genetic map to be extended beyond the limit of 50% recombination that can be measured directly between any pair of genes. A genetic map is usually based on measurements involving genes that are fairly close together (and is subject to corrections from the simple percent recombination). A linkage group includes all those genes that can be connected either directly or indirectly by linkage relationships. Genes lying close together show direct linkage; those >50 cM apart assort independently. As linkage relationships are extended, the genes of any organism fall into a discrete number of linkage groups. Each gene identified in the organism can be placed into one of the linkage groups. Genes in one linkage group always show independent assortment with regard to genes located in other linkage groups. The number of linkage groups is the same as the (haploid) number of chromosomes. The relative lengths of the linkage groups are similar to the relative sizes of the chromosomes. Mendel's concept of the gene as a discrete particulate factor can therefore be extended into the concept that the chromosome constitutes a linkage group, divided into many genes, whose physical arrangement underlies their genetic behavior. We sometimes use the term genetic marker to describe a gene of interest, for example, one being used in a mapping experiment or identifying a particular region. Thus a chromosome may be said to carry a particular set of markers, that is, alleles. On the genetic maps of higher organisms established during the first half of this century, the genes are arranged like beads on a string. They occur in a fixed order, and genetic recombination involves transfer of corresponding portions of the string between homologous chromosomes. The gene is to all intents and purposes a mysterious object (the bead), whose relationship to its surroundings (the string) is unclear. The resolution of the recombination map of a higher eukaryote is restricted by the Linkage and mapping | SECTION 7.32.3 © 2004. Virtual Text / www.ergito.com

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small number of progeny that can be obtained from each mating. Recombination occurs so infrequently between nearby points that it is rarely observed between different mutations in the same gene. This forces the questions: does recombination occur within a gene; and can its frequency at these close quarters be used to arrange sites of mutation in a linear order? To answer these questions by conventional genetic means requires a microbial system in which a very large number of progeny can be obtained from each genetic cross. A suitable system is provided by phage T4, a virus that infects the bacterium E. coli. Infection of a single bacterium leads to the production of ~100 progeny phages in less than 30 minutes. The constitution of an individual locus was investigated by Benzer in a series of intensive studies of the rII genes of the phage, which are responsible for a change in the pattern of bacterial killing known as rapid lysis. When two different rII mutant phages are used to infect a bacterium simultaneously, the conditions can be arranged so that progeny phages will be produced only if recombination has occurred between the two mutations to generate a wild-type recombinant. The frequency of recombination depends on the distance between sites, just as in the eukaryotic chromosome. The selective power of this technique in distinguishing recombinants of the desired type allows even the rarest recombination events to be quantitated, so that the map distance between any pair of mutations can be measured. About 2400 mutations fall into 304 different mutant sites. (When two mutations fail to recombine, they are assumed to represent independent and spontaneous occurrences at the same genetic site.) The mutations can be arranged into a linear order, showing that the gene itself has the same linear construction as the array of genes on a chromosome. So the genetic map is linear within as well as between loci: it consists of an unbroken sequence within which the genes reside. This conclusion has of course now been extended in molecular terms to all known genetic systems. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.7.32.3


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7.32.4 Protein folding Key Terms A cofactor is a small inorganic component (often a metal ion) that is required for the proper structure or function of an enzyme. Chaperones are a class of proteins which bind to incompletely folded or assembled proteins in order to assist their folding or prevent them from aggregating. A domain of a protein is a discrete continuous part of the amino acid sequence that can be equated with a particular function.

We can consider two principles that might control the folding of a protein into the correct higher-order structure. • Folding is an intrinsic feature of the primary sequence. In this case, the final structure must always be the most stable thermodynamically and can be generated at any time after synthesis of the polypeptide chain is complete. • The correct structure can be generated only during the synthesis of the polypeptide. Then it becomes possible that an intrinsically less stable structure could prevail because the protein becomes "trapped" in it during synthesis. The relationship between higher-order structures and the primary structure may be revealed when a protein is denatured by heating or by chemical treatments that disrupt protein conformation. Most denaturing events involve the breakage of hydrogen and other noncovalent bonds. An exception is the disruption of S–S bridges that results from treatment with reducing agents. However, all of these changes affect the conformation; the primary sequence of amino acids in the polypeptide chain remains unaltered. In some cases, the higher-order structure follows ineluctably from the primary sequence. The enzyme ribonuclease is the classic example (1115). After the protein has been denatured, its active conformation can be regained by reversing the denaturing procedure. All the information necessary to form the secondary structure resides in the primary sequence. Thus the production of active ribonuclease is an inevitable event whenever the intact primary chain is placed in the appropriate conditions. In other cases, proteins can be irreversibly denatured. Thus under certain (nonphysiological) conditions, a protein may have alternative stable conformations. In some cases the correct conformation probably can be attained only during synthesis of the protein. The conformation could depend on specific interactions between regions of the protein that can occur only in the absence of other regions (that is, those that have not yet been synthesized). This is probably the more common situation. Protein folding | SECTION 7.32.4 © 2004. Virtual Text / www.ergito.com

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In some instances, a cofactor that is part of the active protein (such as the iron-binding heme group of the cytochromes) must be present in order for the polypeptide chain to take up its proper conformation. In the case of multimeric proteins, it may be necessary for one subunit to be present in order for another to acquire the proper conformation. Protein folding is usually rapid in vivo, occurring within seconds or less. It begins even before a protein has been completely synthesized. Probably it involves a sequential folding mechanism, in which the reaction passes through discrete (although highly transient) intermediates. The process is initiated by the collapse of hydrophobic side chains into the "core" of the protein; this occurs within milliseconds. Units of secondary structure, largely α-helices and β-sheets, form on the same time scale. The transition from this structure to the final tertiary structure is slower. The process appears to be cooperative, so that formation of one region of secondary structure enhances formation of the next region, and so on (for review see 2389) The acquisition of structure when a protein is synthesized is not a spontaneous process, but may require assistance. More precisely, we should say that spontaneous folding is a slow reaction, which under normal cellular conditions is a rate-limiting step. The rate is significantly increased by several types of additional functions. These are summarized in Figure S 10. They fall into two groups: enzymes that catalyze specific isomerization steps; and factors that act stoichiometrically to influence folding directly.

Figure S 10 Both catalytic and stoichiometric functions are required to assist protein folding

The formation of disulfide bonds is shown in Figure S 11. The animation shows that formation of a disulfide bond may have a major effect on the conformation of the protein. It is influenced by both environment and specific accessory proteins. Disulfide bonds are rare in cytoplasmic proteins, but common in exported proteins. This is related to a difference in the thiol/disulfide redox state between internal and external conditions. It may help to prevent bond formation in a bacterium, but helps to drive it in the periplasm (the layer surrounding the bacterial cell).

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Figure S 11 Formation of a disulfide bridge between the sulfhydryl groups of two cysteines may connect different parts of a polypeptide chain.

Disulfide bond formation can occur spontaneously in vitro, but the rate is slow. It has a t½>15 min, compared with the ability to form disulfide bonds correctly within a few seconds in vivo. The process is catalyzed in vivo by an enzyme, protein disulfide isomerase (PDI). This is a curious protein, which participates in a variety of functions concerned with protein-modification, in addition to its sponsorship of disulfide bridge formation. It is not entirely clear whether it simply helps the initial formation of disulfide bonds or whether it also catalyzes rearrangement of disulfide bonds that have formed incorrectly (for review see 3443) Proline has a major effect upon protein structure because of the restrictions imposed by its ring structure. Proline introduces a bend in a polypeptide chain, because the nitrogen atom is restrained by the ring structure. The existence (and interconversion) of two stereochemical forms of the peptidyl-proline link is an important feature of protein structure. The direction of the bend is determined by whether the proline is in the cis or trans configuration, as shown in Figure S 12.


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Figure S 12 The configuration of proline has an important effect on protein conformation.

Proteins containing proline fold slowly because the peptidyl-proline link does not necessarily form in correct stereochemical conformation. The enzyme peptidyl-prolyl isomerase (PPI) catalyzes the cis-trans conversion, and by this means significantly accelerates the folding reaction. Enzymes with PPI activity fall into two major groups, named for their abilities to bind certain drugs: cyclophilin PPI binds the drug cyclosporin A, and FKBP PPI binds the drug FK506. Members of the cyclophilin class are better characterized, and they vary in their specificity of action from those that appear to be generic (able to act on any protein) to those that appear to work only with specific proteins. This makes the point that, although control of proline isomerization is a general feature of many proteins, it can also be used to control specifically the maturation of an individual protein. Proteins that act stoichiometrically on the folding of other proteins are called molecular chaperones. A chaperone forms a complex with a protein during folding, but is required only during assembly, and is not part of the mature structure. The major role of a chaperone is to prevent the formation of incorrectly folded structures, in which the substrate protein might otherwise become trapped during folding (see Figure 8.8 in Molecular Biology 2.8.4 Chaperones may be required for protein folding). Protein folding is an intricate process. The primary sequence of a protein is a crucial Protein folding | SECTION 7.32.4 © 2004. Virtual Text / www.ergito.com

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determinant of its higher-order structures. Sometimes it is the sole determinant, but in most cases additional interactions are involved in acquiring the final conformation. In each case, however, if the primary sequence is synthesized within the appropriate environment, it will acquire the proper higher-order structures. Although higher-order structure follows from primary sequence, the same general tertiary structure can be determined by different primary sequences. For example, the globin (red blood cell) proteins of different species vary substantially in sequence, but have the same general tertiary structure. An important concept is that a protein may consist of domains. A domain is a (relatively) independent region of the protein. In some cases its conformation can be acquired independently by the relevant fragment of the polypeptide chain. Some globular proteins consist of discrete domains connected by "clefts." Sometimes a substrate binds to the cleft between domains. A domain may represent a functional unit that is identified with a particular activity of the protein, for example, its ability to perform a certain catalytic activity, to bind a certain ligand, or to interact specifically with other types of domains. The lengths of recognized domains vary from 30–300 amino acid residues. Certain types of domains may be found in proteins with particular locations; for example, on the exterior of the cell. A domain may represent an evolutionary unit. It may have arisen as a functional polypeptide or region of a polypeptide and later have associated with other domains to generate a new protein with additional abilities. The occurrence of closely related domains in different proteins is common; Figure S 13 compares the use of domains in two blood cell proteins.

Figure S 13 Overlapping arrangements of discrete domains are found in proteins.



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Reviews 2389. Fersht, A. R. and Daggett, V. (2002). Protein folding and unfolding at atomic resolution. Cell 108, 573-582. 3443. Sevier, C. S. and Kaiser, C. A. (2002). Formation and transfer of disulphide bonds in living cells. Nat. Rev. Mol. Cell Biol. 3, 836-847.

References 1115. Anfinsen, C. B. (1973). Principles that govern the folding of protein chains. Science 181, 223-230.



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7.32.5 Membranes and membrane proteins Key Terms Amphipathic structures have two surfaces, one hydrophilic and one hydrophobic. Lipids are amphipathic; and some protein regions may form amphipathic helices, with one charged face and one neutral face. A saturated fatty acid only has single carbon-carbon bonds in its backbone. An unsaturated fatty acid has some double carbon-carbon bonds in its backbone. A phospholipid is a lipid that has a positively charged head that is linked by a phosphate group to the fatty acid tails. A glycolipid has a head consisting of an oligosaccharide, linked to a fatty acid tail. A sterol is a compound containing a planar steroid ring. A lipid bilayer is a structure formed by phospholipids in an aqueous solution. The structure consists of two sheets of phospholipids, in which the hydrophilic phosphate groups face the aqueous solution and the hydrophobic tails face each other. Fluidity is a property of membranes; it indicates the ability of lipids to move laterally within their particular monolayer. A transmembrane protein (Integral membrane protein) extends across a lipid bilayer. A hydrophobic region (typically consisting of a stretch of 20-25 hydrophobic and/or uncharged aminoa acids) or regions of the protein resides in the membrane. Hydrophilic regions are exposed on one or both sides of the membrane. The transmembrane region (transmembrane domain) is the part of a protein that spans the membrane bilayer. It is hydrophobic and in many cases contains approximately 20 amino acids that form an α -helix. It is also called the transmembrane domain. A hydropathy plot is a measure of the hydrophobicity of a protein region and therefore of the likelihood that it will reside in a membrane. The side of the plasma membrane, or of the membrane of an organelle, which faces the cytoplasm is its cytoplasmic face. The extracellular matrix (ECM) is a relatively rigid layer of insoluble glycoproteins that fill the spaces between cells in multicellular organisms. These glycoproteins connect to plasma membrane proteins.

The characteristic properties of membranes result from their high contents of lipids. A crucial feature of lipids creates the membranous environment: they are amphipathic. One end of the molecule consists of a polar "head," while the other end consists of a hydrophobic "tail." The major bulk of a lipid is provided by its hydrophobic tails, which differ in overall Membranes and membrane proteins | SECTION 7.32.5 © 2004. Virtual Text / www.ergito.com

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length and in the nature of the carbon-carbon bonds. One type of fatty acid tail is saturated: all the carbon-carbon links are single bonds. The other type of tail is unsaturated: one or more carbon-carbon links consist of double bonds. Because rotation is restricted around the double bond, the unsaturated tail has a bend, while the saturated tail can extend freely. Fatty acid tails are usually ~20 residues long. Distinguished by their polar heads, membranes contain the three principal types of lipids illustrated in Figure S 14:

Figure S 14 A lipid has a polar head and a hydrophobic tail.

• In phospholipids the head has a positively charged group linked via a negatively charged phosphate group to the rest of the molecule. The example of Figure S 14 has a head consisting of choline-phosphate-glycerol, attached to two hydrophobic tails. Lipids based on glycerol have one saturated and one unsaturated fatty acid tail. • Glycolipids are characterized by the presence of oligosaccharide. The chain of sugars typically consists of 1-15 residues. In animal cells, the connection between the saccharide head and the fatty acid tail is sphingosine (a long amino alcohol). Lipids based on sphingosine have a fatty acid chain in addition to the long hydrocarbon chain of sphingosine itself. In plants and bacteria, glycerol connects the head and tail. • Sterols contain a steroid ring. They lend rigidity to a membrane because the steroid ring is planar. Cholesterol, a prominent component of animal cell Membranes and membrane proteins | SECTION 7.32.5 © 2004. Virtual Text / www.ergito.com

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membranes, has a polar hydroxyl group at the terminus. In an aqueous environment, a lipid is happy to have its polar head exposed, but tries to bury its hydrophobic tail away from the water. Figure S 15 illustrates how this is accomplished in the cell. Two lipid monolayers are juxtaposed to form a lipid bilayer, a sheet in which the polar heads of the lipids face out toward the aqueous environment on either side, while the hydrophobic tails face in to create a hydrophobic environment.

Figure S 15 A lipid bilayer forms in an aqueous environment when the the polar heads are immersed in the water and the hydrophobic tails of the lipids segregate away from water.

Although a membrane consists of a specific type of structure, the lipid bilayer, there is variety in the constitution of different membranes. Overall lipid compositions of membranes vary considerably, with regard to both the ratio of protein to lipid and the types of lipids. These differences mean that different membranes have different biophysical properties. One of the important properties of a membrane is the ability of the constituent lipids to move within it. Lipid molecules rarely move from one monolayer to the other in a bilayer, but frequently move laterally to exchange places with their neighbors within the monolayer. The property of movement is called fluidity; and a membrane is often regarded as a "two dimensional fluid" (1023). The more readily the tails of adjacent lipids can pack together, the more crystalline the membrane structure can become, and the less fluid. The major determinants of membrane fluidity are therefore the types and lengths of the lipid tails. The proportion of saturated versus unsaturated residues in the tails has a major effect on fluidity; unsaturated chains are more difficult to pack, and therefore give a more fluid structure. A protein that resides in a membrane is called a transmembrane protein. The structure of such a protein is illustrated in Figure S 16. A typical transmembrane protein resides in the membrane by means of a specific region called the transmembrane domain. This consists of a stretch of ~21 amino acids with Membranes and membrane proteins | SECTION 7.32.5 © 2004. Virtual Text / www.ergito.com

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sufficient hydrophobicity to be comfortable in the environment of the lipid bilayer. It forms an α-helix that just spans the membrane. The transmembrane domain is flanked by regions that protrude into the interior of the cell or out into the surrounding environment. These regions are generally hydrophilic, like any protein that resides in the cytosol. A protein may have several transmembrane domains, in which case the parts of the polypeptide chain connecting them can be viewed as "looping out" into the cytoplasm or the exterior.

Figure S 16 A transmembrane protein crosses the lipid bilayer. The hydrophobic transmembrane region spans the bilayer, and hydrophilic regions are exposed on either side.

Within the lipid bilayer, water is effectively excluded. As a result, the regions of the proteins located within the bilayer are not subjected to the aqueous environment of the cytosol. The lipid "solvent" does not form hydrogen bonds with the protein, and therefore solvates neither the groups of the peptide backbone nor polar side chains. Hydrogen bonding occurs solely between groups within the protein itself, and functions to form α-helices and (to a lesser degree) ß-sheets. This allows the protein to acquire a different conformation from what would be reached in an aqueous environment. Indeed, it is possible that membrane proteins can attain their natural conformation only in the hydrophobic environment. The hydrophobicity of a sequence of amino acids can be used to predict (although not perfectly) whether it is likely to reside in a membrane. A hydropathy plot shows the sequence of a protein in terms of the hydrophobicity of overlapping segments. There are various means of measuring hydrophobicity, but whichever scale is used, it is conventional to assess hydrophobicity as a positive score and hydrophilicity as a negative score. Most of the scales utilize the energy required in kcal/mol to transfer from a hydrophobic to a hydrophilic phase. In the example of Figure S 17, the hydrophobicity is calculated for each position in the protein by summing the scores of the individual amino acids in the next 21 positions. A region with a positive score is therefore a candidate to provide a transmembrane domain that resides in a membrane.

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Figure S 17 A hydropathy plot identifies potential membrane-spanning regions as the most hydrophobic sequences of a protein.

Proteins may also be associated with a membrane by means of covalent linkage to a fatty acid that is incorporated into the lipid bilayer. Figure S 18 depicts four forms of such association. In each case, a fatty acid or lipid is attached to an amino acid near to or at one terminus of the protein, with the result that the entire polypeptide chain resides on one side of the membrane, but is attached to it.

Figure S 18 Proteins may be associated with one face of a membrane by acyl linkages to fatty acids.

Prenylation is used to attach proteins to both the plasma membrane and internal membranes. Two types of prenyl groups have been identified: farnesyl is a 15 carbon Membranes and membrane proteins | SECTION 7.32.5 © 2004. Virtual Text / www.ergito.com

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isoprenoid (shown in the figure), and geranylgeranyl is a 20 carbon chain. They are added to cysteine residues by a thioester linkage; the cysteine is always located at the fourth position from the C terminus, as part of the sequence CAAX, where A represents aliphatic amino acids and X is methionine or serine for farnesylation, and leucine for geranylgeranylation. The usefulness of the prenyl groups for assisting attachment to the membrane is obvious, but we do not yet know how specificity is conferred with regard to the choice of membrane. Two fatty acids are used to anchor proteins on the cytoplasmic side of the plasma membrane. Palmitic acid, a 16 carbon-chain saturated fatty acid, is linked through a sulfide bond to a cysteine residue located close to the terminus (usually the C-terminus, but sometimes the N-terminus). Myristic acid, a 14 carbon-chain saturated fatty acid, is linked to the amino group of N-terminal glycine. Myristoylated proteins are often, but not always, associated with a membrane. The more complex structure of a glycosyl-phosphatidyl-inositol (GPI) anchor is linked to the carboxyl group of the C-terminal amino acid of protein exposed on the extracellular side of the membrane. Addition of the GPI anchor actually involves cleavage of the original polypeptide chain near the C-terminus, generating a new C-terminus that is linked to the anchor. Enzymes exist that can cleave the GPI anchor from the protein, releasing the protein into the extracellular medium. The membranes bounding different cellular compartments are different not only in their overall composition, but also in the particular proteins that reside within them. The proteins in each type of membrane serve, for example, to control transport into or out of the particular compartment, and are therefore designed to recognize the particular molecules or macromolecules that travel this route. Each membrane has two "faces," as indicated in Figure S 19. In all membranes, the cytoplasmic face is defined as the surface that contacts the general cytosol. The noncytoplasmic face is given various names, depending on the membrane. On a plasma membrane, it provides the outside surface of the cell. In a membrane within the cell, it provides a limit for an interior compartment, comprising the surface that separates the lumen of the compartment from the cytosol.


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Figure S 19 The asymmetry of the membrane bilayer distinguishes the cytoplasmic face from the exterior face.

A major feature of the lipid bilayer is an asymmetry created by biochemical differences between the two faces of the membrane. Three components of the plasma membrane are unevenly distributed: • Different lipids are concentrated in the cytoplasmic and extracellular monolayers. This affects both the polar heads and the hydrophobic tails. Lipids on the cytoplasmic face are more highly charged and tend to be unsaturated. How is the difference between the monolayers established? Lipids are synthesized within the cell, and initially inserted into the cytoplasmic surface of the membrane. A specific protein, a "flippase," may be responsible for transporting a lipid from one monolayer to the other. The existence of specific flippases for different lipids could be responsible for creating some of the asymmetries in lipid distribution between the bilayers. • Proteins are oriented so that different sequences (or even entire proteins) are present on each face. The location of a protein is determined by its sequence, which contains signals that cause it to be inserted in the membrane in a particular orientation. This is discussed in detail in Molecular Biology 2.8.15 Anchor sequences determine protein orientation. • Carbohydrate groups (on glycolipids or glycoproteins) are found exclusively on the extracellular face. The consequence of this organization for the plasma membrane is that the exterior of the cell has a surface rich in oligosaccharides. The plasma membrane circumscribes a cell. It marks the boundary between the cellular milieu inside and the environment outside. In the case of a unicellular organism, the surroundings constitute the environment in which the organism lives. In a multicellular organism, the environment for any one cell is created by other cells. In some cases, the plasma membrane is extended by the presence of additional glycoproteins, connected to those actually included in the membrane. This type of arrangement may form a cell coat. In some cases, the cell coat is extended into an extracellular matrix, rich in glycoproteins, and providing a thicker layer at the cell surface. Membranes and membrane proteins | SECTION 7.32.5 © 2004. Virtual Text / www.ergito.com

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The role of the plasma membrane extends beyond providing a mere barrier to the outside. It controls ingress and egress for molecules both small and large. Within the membrane reside specific transport systems that pump ions in or out, that allow proteins to be secreted from the cell into the environment (see Molecular Biology 6.27 Protein trafficking), and that recognize molecules outside and as a result transmit messages to the interior (see Molecular Biology 6.28 Signal transduction). Plasma membranes of animal cells contain relatively large amounts of cholesterol, which increases mechanical stability because of the steroid rings near its polar head. (Plant cells lack cholesterol, but have other sterols instead.) Plasma membranes also are the only membranes to contain significant amounts of glycolipids. A plasma membrane contains about equal masses of lipid and protein. Internal membranes, such as those surrounding mitochondria, have a greater proportion of protein. The mass of an individual protein molecule is much larger than any lipid, so there are 10–100× more lipid molecules than protein molecules. We might view the basic structure of a membrane as consisting of a lipid bilayer that provides a residence for relatively large protein molecules. Protein components of the membrane can move laterally within the lipid bilayer, although they diffuse much more slowly than the lipid components. As the result of a stimulus, proteins can be "internalized," when they are removed to the interior of the cell. Other proteins are secreted from the interior of the cell to the exterior by passing through the membrane. The lipid bilayer itself, and the proteins associated with it, therefore comprises a dynamic structure.


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References 1023. Singer, S. J. and Nicolson, G. L. (1972). The fluid mosaic model of the structure of cell membranes. Science 175, 720-731.



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7.32.6 ER and Golgi Key Terms The endoplasmic reticulum (ER) is an organelle involved in the synthesis of lipids, membrane proteins, and secretory proteins. It is a single compartment that extends from the outer layer of the nuclear envelope into the cytoplasm. It has subdomains, such as the rough ER and smooth ER. The lumen describes the interior of a compartment bounded by a membrane, usually the endoplasmic reticulum or the Golgi apparatus. Rough endoplasmic reticulum (rough ER) refers to the region of the endoplasmic reticulum to which ribosomes are bound. It is the site of synthesis of membrane proteins and secretory proteins. Smooth ER consists of a regions of endoplasmic reticulum devoid of ribosomes. The ribosome is a large assembly of RNA and proteins that synthesizes proteins under direction from an mRNA template. Bacterial ribosomes sediment at 70S, eukaryotic ribosomes at 80S. A ribosome can be dissociated into two subunits. The Golgi apparatus is an organelle that receives newly-synthesized proteins from the endoplasmic reticulum and processes them for subsequent delivery to other destinations. It is composed of several flattened membrane disks arranged in a stack. The cisternae of the Golgi apparatus are the successive stacks, each bounded by a membrane, that make up individual compartments. The cis face of the Golgi is the side juxtaposed to the nucleus. The trans face of the Golgi is juxtaposed to the plasma membrane. Lipid trafficking is the movement of lipids among the various membranes of a eukaryotic cell. An endosome is an organelle that functions to sort endocytosed molecules and molecules delivered from the trans-Golgi network and deliver them to other compartments, such as lysosomes. It consists of membrane-bounded tubules and vesicles. A lysosome is an organelle that contains hydrolytic enzymes and has an acidic lumen (pH as low as 4.5). Its primary function is the degradation of endocytosed material. The peroxisome is an organelle in the cytoplasm enclosed by a single membrane. It contains oxidizing enzymes.

Membranes occupy a major part of the eukaryotic cell. In addition to surrounding individual organelles, large sheets of membranes are a prominent feature of many cells (especially those involved in secreting proteins). The electron micrograph in Figure 8.19 shows an extensive sheet of membranes that extends from the nucleus. This is the endoplasmic reticulum, a highly convoluted sheet of membranes ER and Golgi | SECTION 7.32.6 © 2004. Virtual Text / www.ergito.com

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representing 30–60% of total membrane. The interior space comprises the lumen. Visualized in the electron micrograph, the endoplasmic reticulum can be divided into two types: rough ER and smooth ER. They are part of the same membrane sheet. The characteristic appearance of the rough ER results from the presence of ribosomes on its cytoplasmic surface. The ribosomes are small particles concerned with the synthesis of proteins. Their presence is an indication that proteins are being synthesized at the cytoplasmic surface of the endoplasmic reticulum, which then processes them for assignment to various cell compartments. Between the endoplasmic reticulum and the plasma membrane lies the Golgi apparatus. The electron micrograph in Figure S 20 shows the Golgi as a series of individual membrane sheets, tightly packed together.

Figure S 20 The Golgi apparatus consists of a series of individual membrane stacks. Photograph kindly provided by Alain Rambourg.

The relationship between the ER and Golgi is depicted diagrammatically in Figure S 21. The ER is shown as a sheet made from the folding of a single lipid bilayer that extends from the outer membrane surrounding the nucleus. The Golgi consists of a "stack" of flat cisternae, like a pile of discs. Each cisterna consists of a closed structure bounded by a single continuous membrane. A stack usually consists of 90-95% of tRNAs. Sometimes the exceptions are individual; sometimes they fall into groups representing some peculiarity of a particular cell. The length of the D loop varies by up to 4 residues. The extra nucleotides relative to the most common structure are denoted 17:1 (lying between 17 and 18) and 20:1 and 20:2 (lying between 20 and 21). However, in the smallest D loops, residue 17 as well as these three is absent. tRNA sequences | SECTION 7.32.8 © 2004. Virtual Text / www.ergito.com

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The most variable feature of tRNA is the so-called extra arm. Depending on the nature of the extra arm, tRNAs can be divided into two classes. Class 1 tRNAs have a small extra arm, consisting of only 3-5 bases. They represent ~75% of all tRNAs. Class 2 tRNAs have a large extra arm – it may even be the longest in the tRNA – with 13–21 bases, and ~5 base pairs in the stem. The additional bases are numbered from 47:1 through 47:18. The functional significance of the extra arm is unknown. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.7.32.8

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7.32.9 Complementation Key Terms Interallelic complementation (intragenic complementation) describes the change in the properties of a heteromultimeric protein brought about by the interaction of subunits coded by two different mutant alleles; the mixed protein may be more or less active than the protein consisting of subunits only of one or the other type. Negative complementation occurs when interallelic complementation allows a mutant subunit to suppress the activity of a wild-type subunit in a multimeric protein. A dominant negative mutation results in a mutant gene product that prevents the function of the wild-type gene product, causing loss or reduction of gene activity in cells containing both the mutant and wild-type alleles. The effect may result from the titration of another factor that interacts with the gene product or by an inhibiting interaction of the mutant subunit on the multimer.

Complementation was originally developed as a test to determine whether two mutations lie in different genes. It consists of comparing the phenotypes when both mutations are the same piece of DNA (called the cis configuration) and when they are on different pieces of DNA (called the trans configuration). Figure S 27 shows that, if the mutations are in the same gene, the trans configuration has a mutant phenotype. Both copies of the gene are mutated (each copy has one of the mutations). The cis configuration has wild phenotype, however, because one copy of the gene has both mutations, and the other has no mutations.

Figure S 27 When mutations are in the same gene, each allele has a different mutation, so only mutant protein is produced in the trans configuration.

Figure S 28 shows that, if the mutations are in different genes, the configuration does not matter. There is always one mutant copy of each gene and one wild-type copy of each gene. Complementation | SECTION 7.32.9 © 2004. Virtual Text / www.ergito.com

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Figure S 28 When mutations are in different genes, one allele has two mutations, but the other has none, and so produces wild-type protein.

The practical form of the test, therefore, is to use the cis configuration as a control (it is always wild-type) and to determine whether the trans configuration is mutant (mutations are in the same gene) or wild-type (mutations are in different genes). The complementation test applies to genes whose products function as independent proteins. When the gene product is a subunit of a multimeric protein, interactions between the subunits can either allow complementation between alleles or cause one allele to suppress the effect of the other. An exception to the rule that only different genes can complement is sometimes found when a gene represents a polypeptide that is the subunit of a homomultimeric protein. In the wild-type cell, the active protein consists of several identical subunits. In a cell containing two mutant alleles, however, their products can mix to form multimeric proteins that contain both types of subunit. Figure S 29 shows that if the two mutations compensate, the mixed-subunit protein is active, even though the proteins consisting solely of either type of mutant subunit are inactive. This effect is called interallelic complementation.

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Figure S 29 Interallelic complementation occurs when different mutations in the subunits of a multimeric protein can compensate to make an active protein even though each separate subunit can only form inactive multimers.

In the reverse type of interaction, a defective subunit produced by one allele inhibits the active subunits produced by another allele. This is called negative complementation. Figure S 30 shows that in effect the "bad" subunit poisons the multimeric protein so that the "good" subunits cannot function. An allele that is able to prevent other alleles from functioning is called a dominant negative.


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Figure S 30 Negative complementation occurs when a mutant subunit prevents the wild-type subunit from functioning.

Dominant negatives can be constructed by targeting mutagenesis at an active site in the protein. Figure S 31 shows that one common use for this technique is to delete the DNA-binding site from a protein that is a subunit in a DNA-binding factor. When the gene for the mutant protein is introduced into the cell, the defective subunit overwhelms the normal subunits, and prevents them from forming multimers with sufficient activity to bind DNA. The same principle can be applied to any situation in which a protein forms a subunit of a multimeric protein. The subunits do not have to be identical. In Figure S 31, one subunit could bind DNA, while the other might interact with other proteins. The function of the second protein would be prevented by a dominant-negative subunit without a DNA-binding site.


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Figure S 31 A dominant negative mutant can be constructed by removing the DNA-binding site from a subunit of a DNA-binding protein.

The same technique can be used to target any active site, for example, the kinase site of a multimeric protein kinase enzyme. It is enormously useful because it can be used in circumstances where it is impossible to mutate the endogenous protein. This allows the function of the protein to be tested directly by introducing the mutant allele. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.7.32.9


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7.32.10 G proteins Key Terms A serpentine receptor has 7 transmembrane segments. Typically it activates a trimeric G protein. A second messenger is a small molecule that is generated when a signal transduction pathway is activated. The classic second messenger is cyclic AMP, which is generated when adenylate cyclase is activated by a G protein (when the G protein itself was activated by a transmembrane receptor).

There are two types of G proteins. The name reflects the ability to bind a guanine nucleotide. The guanine nucleotide can alternate between GDP and GTP, and controls the activity of the protein. Both types of G protein work on the same principle that the GDP-bound form is inactive, and the GTP-bound form is active. Trimeric G proteins are associated with the cytosolic face of the membrane. They are involved in the initial stages of signal transduction. They are activated by transmembrane receptors, most typically by serpentine receptors (7-membrane pass proteins). The three subunits are called α, β, and γ. The α subunit binds the guanine nucleotide. The inactive form of the G protein is bound to GDP. In this form, the G protein is constitutively associated with a membrane receptor. When the receptor is activated (usually by binding ligand) it causes GDP to be displaced from the G protein. Because the concentration of GTP in the cytosol is much greater than that of GDP, the vacant nucleotide binding site is filled with GTP. Figure S 32 shows how trimeric G proteins are activated. Binding of GTP causes the G protein to dissociate into a free α subunit and free β γ dimer. Depending on the individual G protein, it can be either the α subunit or the β γ dimer that transmits the signal to the next stage in the pathway. Whichever is the active component (and sometimes both are active) may either activate or repress the activity of a target protein.

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Figure S 32 When a receptor is activated by hormone binding, it causes GTP to replace GDP on a G α subunit. The G α subunit dissociates from the β γ dimer, and activates an effector such as adenylate cyclase. This is a static version of an interactive figure; see http://www.ergito.com/main.jsp?bcs=MBIO.7.32.10 to view properly.

The target protein often is also associated with the membrane. This chain of events often stimulates the production of second messengers. In one classic example, when the protein Gs is activated, the α subunit then activates adenylate cyclase, which generates cyclic AMP.


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Molecular Biology How long the G protein remains active is controlled by the α subunit. All α subunits are GTPases. When the GTP is hydrolyzed to GDP, the α subunit reassociates with the β γ dimer to reconstitute the trimeric G protein. By removing the individual subunits, the hydrolysis of GTP terminates the physiological response. Each α subunit hydrolyzes its GTP in vitro at a characteristic (slow) rate, typically with a half-life ~15 secs. But some of the physiological reactions are much shorter lived. For example, in the classic system of vision, a light response terminates in ~100 msec. The rate of GTP hydrolysis can be accelerated in vivo by interaction with another component of the system. This type of interaction was originally discovered for monomeric G proteins (see below), where the relevant component is a called a GAP. A common type of protein with GAP function for the α subunits of trimeric G proteins is the RGS (G protein signaling) class (for review see 2276). An RGS acts indirectly by affecting teh conformation of the α subunit so that it becomes a more effective GTPase. Monomeric G proteins are cytosolic and are often used as binary switches in signalling or other pathways. They work on the same principle as the α subunit of a trimeric G protein. A monomeric G protein is a GTPase that hydrolyzes its bound GTP. This converts it from an active state to an inactive state. Figure S 33 shows that three types of ancillary proteins influence the balance between the GDP- and GTP-bound forms of a monomeric G protein.

Figure S 33 Monomeric G proteins are active when bound to GTP and inactive when bound to GDP. Their activity is controlled by other proteins. This is a static version of an interactive figure; see http://www.ergito.com/main.jsp?bcs=MBIO.7.32.10 to view properly.

• A GAP (GTPase activating protein) stimulates the GTPase activity. This is needed for a fast reaction time, because the intrinsic rate of GTP hydrolysis is slow. Thus GAP activity inactivates the G protein. Different GAPs have G proteins | SECTION 7.32.10 © 2004. Virtual Text / www.ergito.com

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specificities for different GTP-binding proteins; they are typically named as Protein-GAP, where Protein is the monomeric G-protein on which they act. • A GEF (guanine nucleotide exchange factor) displaces the GDP bound to an inactive G protein. The principle of replacement is the same as for the trimeric α subunit. Release of the GDP creates an empty site. The concentration of GTP in the cytosol is greater than that of GDP, so the site is then filled with GTP. This activates the protein. GEFs have the same sort of specificity as GAPs, and similarly are named in the form Protein-GEF (for review see 3217). • A GDI (guanine nucleotide dissociation inhibitor) can block the displacement reaction. This maintains the G protein in the inactive state. Examples of specific monomeric G proteins are EF-Tu (see Molecular Biology 2.6.10 Elongation factor Tu loads aminoacyl-tRNA into the A site), Ran (see Molecular Biology 2.8.28 Transport receptors carry cargo proteins through the pore), ARF and Rab (see Molecular Biology 6.27.7 Vesicles can bud and fuse with membranes), and Ras and the Rho family (see Molecular Biology 6.28.15 The activation of Ras is controlled by GTP). Last updated on 1-22-2002


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Reviews 2276. Ross, E. M. and Wilkie, T. M. (2000). GTPase-activating proteins for heterotrimeric G proteins: regulators of G protein signaling (RGS) and RGS-like proteins. Annu. Rev. Biochem. 69, 795-827. 3217. Schmidt, A. and Hall, A. (2002). Guanine nucleotide exchange factors for Rho GTPases: turning on the switch. Genes Dev. 16, 1587-1609.



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SUPPLEMENTS

7.32.11 Restriction mapping Key Terms End labeling describes the addition of a radioactively labeled group to one end (5 ′ or 3 ′ ) of a DNA strand.

The principle of restriction mapping is to break a piece of DNA into several fragments that overlap, and then to place these fragments in order by making use of the overlap. One way of generating overlapping fragments is to use enzymes that cleave at different target sites. Another is to use partial cleavage, so that an enzyme attacks any individual target site with less then 100% efficiency. Figure S 34 shows an example of cleavage by different enzymes. A DNA molecule of length 5000 bp is incubated separately with two restriction enzymes, A and B. After cleavage the DNA is electrophoresed. The sizes of the individual fragments generated by enzyme A (left) or enzyme B (right) are determined by comparison with the positions of fragments of known size, such as the control shown in the center. This demonstrates that enzyme A has cut the substrate DNA into four fragments (lengths 2100, 1400, 1000, and 500 bp), while enzyme B has generated three fragments (lengths 2500, 1300, and 1200 bp).

Restriction mapping | SECTION 7.32.11 © 2004. Virtual Text / www.ergito.com

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Figure S 34 DNA can be cleaved by restriction enzymes into fragments that can be separated by gel electrophoresis.

The patterns of cutting by the two enzymes can be related by several means. Figure S 35 illustrates the principle of analysis by double digestion. In this technique, the DNA is cleaved simultaneously with two enzymes as well as with either one by itself. The most decisive way to use this technique is to extract each fragment produced in the individual digests with either enzyme A or enzyme B and then to cleave it with the other enzyme. The products of cleavage are analyzed again by electrophoresis.


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Figure S 35 Double digests define the cleavage positions of one enzyme with regard to the other.

We can use these data to construct a map of the original 5000 bp molecule of DNA, as illustrated by the stages of Figure S 36.


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Figure S 36 A restriction map can be constructed by relating the A-fragments and B-fragments through the overlaps seen with double digest fragments.

Each gel in Figure S 35 is labeled according to the fragment that was isolated from the gel in Figure S 34. A-2100 identifies the fragment of 2100 bp produced by degrading the original DNA molecule with enzyme A. When this fragment is retrieved and subjected to enzyme B, it is cut into fragments of 1900 and 200 bp. So one of the cuts made by enzyme B lies 200 bp from the nearest site cut by enzyme A on one side, and is 1900 bp from the site cut by enzyme A on the other side. This situation is described by the top map in Figure S 36. A related pattern of cuts is seen when we examine the susceptibility of fragment B-2500 to enzyme A. It is cut into fragments of 1900 and 600 bp. So the 1900 bp fragment is generated by double cuts, with an A site at one end and a B site at the other end. It can be released from either of the single-cut fragments (A-2100 or B-2500) that contain it. These single-cut fragments must therefore overlap in the region of the 1900 bp of the common fragment that can be generated from them. This is described in the second map of Figure S 36, which extends our map to the right to add a cleavage site for enzyme B. The key to restriction mapping is the use of overlapping fragments. Because of the overlap of A-2100 and B-2500 in the central region of 1900 bp, we can relate the A site 200 bp to the left of the 1900 bp region with the B site 600 bp to the right. In the Restriction mapping | SECTION 7.32.11 © 2004. Virtual Text / www.ergito.com

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same way, we can now extend the map farther on either side. The 200 bp fragment at the left is also produced by cutting B-1200 with enzyme A, so the next B site must lie 1000 bp to the left. The 600 bp fragment at the right is also produced by cutting A-1400 with enzyme B, so the next A site must lie 800 bp to the right. This gives the third map in Figure S 36. We can now complete the map by identifying the source of the two fragments at each end. At the left end, the 1000 bp fragment arises from B-1200 or in the form of A-1000, which is not cut by enzyme B. So A-1000 lies at the end of the map. Proceeding from the left end of the complete 5000 bp region, it is 1000 bp to the first A site and 1200 bp to the first B site. (This is why a B cut is not shown at the left end of the map above, although formally we treated the end as a B-cutting site in the analysis.) At the right end of the map, the 800 bp double-cut fragment is generated by cutting B-1300 with enzyme A, so we must add a fragment of 500 bp to the right. This is the terminal fragment, as seen by its presence as A-500 in the single-cut A digest. So our completed map takes the form of the bottom map in Figure S 36. The actual construction of a restriction map usually requires recourse to several enzymes, so it becomes necessary to resolve quite a complex pattern of the overlapping fragments generated by the various enzymes. Several other techniques are used in conjunction with comparison of fragments, including end labeling, in which the ends of the DNA molecule are labeled with a radioactive phosphate (certain enzymes can add phosphate moieties specifically to 5 ′ or to 3 ′ ends). Figure S 37 shows that this allows the fragments containing the ends to be identified directly by their radioactive label. So in the fragment A preparation, A-1000 and A-500 would be placed immediately at opposite ends of the map; similarly, fragments B-1200 and B-1300 would be identified as ends.

Figure S 37 When restriction fragments are identified by their possession of a labeled end, each fragment directly shows the distance of a cutting site from the end. Successive fragments increase in length by the distance between adjacent restriction sites.

A complex set of overlapping fragments can be generated directly by a single enzyme by using conditions of partial cleavage, as illustrated in Figure S 38. Of course, for mapping purposes then it is necessary to distinguish different fragments Restriction mapping | SECTION 7.32.11 © 2004. Virtual Text / www.ergito.com

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that have the same size, and to determine overlaps. However, this is a useful technique when we want essentially to introduce random breaks into a large DNA (for example a whole genome) in order to obtain cloned fragments (see Molecular Biology Supplement 32.12 Genome mapping).

Figure S 38 Partial cleavage by a restriction endonuclease generates a series of overlapping fragments. This content is available online at http://www.ergito.com/main.jsp?bcs=MBIO.7.32.11


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SUPPLEMENTS

7.32.12 Genome mapping Key Terms A library is a set of cloned fragments together representing the entire genome (genomic library) or all the expressed genes (cDNA library). A yeast artificial chromosome (YAC) is a synthetic DNA molecule that contains an origin for replication, a centromere to support segregation, and telomeres to seal the ends. It is used as a means to propagate whatever genes it carries in yeast cells. A bacterial artificial chromosome (BAC) is a synthetic DNA molecule that contains the sequences needed for replication and segregation in bacteria. This is used in genomic cloning to amplify sequences typically 100-200 kb long. They are usually derived from derived from the naturally-occurring F factor episome. A contig is a continuous stretch of genomic DNA generated by assembling cloned fragments by means of their overlaps. Shotgun cloning analyzes an entire genome in the form of randomly generated fragments. An expressed sequence tag (EST) is a short sequence of DNA taken from a cDNA copy of an mRNA. The EST is complementary to the mRNA and can be used to identify genes corresponding to the mRNA.

The principle of genome mapping is the same as restriction mapping (see Molecular Biology Supplement 32.11 Restriction mapping): to break a large DNA molecule into smaller molecules that are assembled in order by the principle of overlaps between their ends. Of course, there is a difference of scale when we start with a whole genome, and special tricks have to be used to assemble the fragments. The starting point is usually the generation of a library of cloned fragments that represent the whole genome. This can be achieved by cleaving DNA at random, for example, by using partial cleavage conditions with a restriction endonuclease of low specificity (see Figure S 38). We can calculate statistically how many clone are needed for the whole genome to be represented in fragments of a given size (1441). Over the past 20 years, it has been possible to move from cloning relatively small fragments of DNA in individual plasmid or phage vectors to using synthetic chromosomes, either YACs (yeast artificial chromosomes) or BACs (bacterial artificial chromosomes). The basic principle is that the artificial chromosome consists of a long length of DNA (100-200 kb)which is able to propagate in the host cell because it has the features required for replication, segregation, and stability. Cloned fragments are joined together by the principle of overlap, in which their ends are compared. When the end of one fragment is identical to the end of another fragment, we may conclude that the two fragments represent overlapping stretches of the genome. A series of fragments may be joined together into a contig, which is effectively a continuous stretch of the genome generated from overlapping Genome mapping | SECTION 7.32.12 © 2004. Virtual Text / www.ergito.com

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fragments. Figure S 39 shows an example of a chromosome "walk", in which successive fragments were joined together by identifying overlapping clones from a library. A subfragment from one end of the first clone is used to isolate clones that extend farther along the chromosome. These clones in turn are used to isolate the next set. In each cycle, a new clone is selected because its restriction map coincides at one end with the end of the previous clone, but at the other end has new material. It is possible to walk for hundreds of kb, typically at a rate of >100 kb per month. Chromosome walking allows large contiguous regions of the chromosome to be represented in a library of clones.

Figure S 39 Chromosome walking is accomplished by successive hybridizations between overlapping genomic clones.

Two approaches have been taken to large-scale mapping of genomes. One is to break the genome first into a series of large contigs, as illustrated in Figure S 40. With the contigs in hand, each contig is then sequenced, and we know that these sequences together will add up to the entire genome. This approach was used by one of the groups that sequenced the human genome (1440). In this case, the raw material was a total of ~29,000 BACs that were assembled into ~1250 contigs. The contigs were mapped to chromosomal locations and merged.

Genome mapping | SECTION 7.32.12 © 2004. Virtual Text / www.ergito.com

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Figure S 40 A genome may be organized into a set of contigs before it is sequenced.

Another approach is to use shotgun cloning, as illustrated in Figure S 41. Here the entire genome is broken into random fragments that are sequenced, and then the sequences are joined by the usual principle of identifying overlaps. This requires very sophisticated computer processing to identify all the overlaps. This was also used to analyze the human genome (1439).


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Figure S 41 The principle of shotgun cloning is to break a DNA at random into fragments that are sequenced. Overlapping fragments are identified by sequence comparisons and connected into a continuous sequence.

Analysis of fragments generated by shotgun cloning is usually involved at some stage. Identifying the overlaps becomes easier the smaller the size of the starting fragment. So when a contig is analyzed by shotgun cloning, the process is simpler than when a genome is analyzed, by virtue of the reduced complexity of the DNA sequences. Joining sequences on the basis of overlaps would be relatively straightforward if the genome consisted only of unique (nonrepetitive) sequences. The major problem in practice is caused by the presence of repetitive sequences, where several stretches of DNA may be so similar that it is impossible to distinguish among them. Figure S 42 illustrates the difficulty. In its simplest form, it is difficult to line up two overlapping fragments because we do not know how many copies of the repeating unit there are. Figure S 43 shows the more complex condition in which the same sequences occur in multiple locations in the genome, we do not know which of any two overlapping fragments should be joined.


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Molecular Biology

Figure S 42 Fragments that overlap in a repeated sequence may have multiple possible alignments.

Figure S 43 Dispersed duplicated sequences may have multiple possible alignments.

In the same way that genomic libraries can be prepared from genomicDNA, cDNA libraries can be prepared by reverse transcribing populations of mRNA. This is an important tool in identifying and mapping expressed genes. In principle, each RNA molecule provides the template for synthesizing a single-stranded cDNA that is Genome mapping | SECTION 7.32.12 © 2004. Virtual Text / www.ergito.com

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complementary to it. The cDNA is then converted to a double-stranded form and cloned in the usual way. Any individual RNA should be represented in the cloned population in proportion to its abundance in the original RNA population (although the actual proportions are distorted by the idosyncracies of reverse transcription). In its simplest form, a cDNA library consists of a population of cloned DNA molecules that represent all of the expressed sequences in the source cell type. However, using cDNA clones as such turns out to be an efficient strategy for identifying and mapping only the shorter genes. More sophisticated variations of the procedure have been introduced to make it possible to automate data collection. Expressed sequence tags (EST) are obtained by sequencing the extremities of cDNA clones, so typically they correspond to the 5 ′ leader and 3 ′ trailer. They can be used to provide large data sets that allow expressed genes to be identified (2220). The principle that an EST is located on the genome by its sequence identity with a cloned sequence of genomic DNA. The most recent development is the use of a strategy qhere sequences are produced from within the transcript, instead of from the ends. This is called ORF EST (meaning that the EST represents an open reading frame). This provides a better data set, with less confusion from overlapping or repeated sequences, and allows large libraries to be made that accurately represent the expresed set of genes (2219). Last updated on 12-13-2001


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References 1439. Venter, J. C. et al. (2001). The sequence of the human genome. Science 291, 1304-1350. 1440. International Human Genome Sequencing Consortium. (2001). Initial sequencing and analysis of the human genome. Nature 409, 860-921. 1441. Maniatis, T., Hardison, R. C., Lacy, E., Lauer, J., O'Connell, C., Quon, D., Sim, G. K., and Efstratiadis, A. (1978). The isolation of structural genes from libraries of eucaryotic DNA. Cell 15, 687-701. 2219. Carmargo, A. A. et al. (2001). The contribution of 700,000 ORF sequence tags to the definition of the human transcriptome. Proc. Natl. Acad. Sci. USA 98, 12103-12108. 2220. Adams, M D. et al. (1991). Complementary DNA sequencing: expressed sequence tags and human genome project. Science 252, 1651-1656.



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SUPPLEMENTS

7.32.13 Two-component signal transduction Two-component signaling systems were discovered in bacteria, where they provide the most common form of signaling pathway that responds to extracellular events (2307). There are ~30 such pathways in E. coli. Figure S 44 illustrates a generic system. They are also found in plant cells (a typical plant may have 10-15 such pathways), occasionally in yeast (~5 pathways), but are relatively rare in animal cells (there are none in mammals), where kinase cascades involving Ser/Thr and Tyr kinases are more common.

Two-component signal transduction | SECTION 7.32.13 © 2004. Virtual Text / www.ergito.com

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Figure S 44 A two component system consists of a sensor that is an autophosphorylating histidine kinase and an effector that can catalyze transfer of phosphate from the sensor to itself.

The sensor protein is a histidine kinase that is located in the bacterial membrane. It can be activated by binding a ligand that is in the extracellular medium. Activation causes the kinase to autophosphorylate (that is, to phosphorylate itself). The reaction transfers the γ phosphate from ATP on to a histidine residue in the kinase. The sensor interacts with an effector protein (also called a "response regulator"). The effector catalyzes transfer of the phosphate group from the histidine on the sensor to an aspartic acid residue in its own regulatory domain. This activates the effector. It is later deactivated by dephosphorylation. Two-component signal transduction | SECTION 7.32.13 © 2004. Virtual Text / www.ergito.com

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The chemistry of phosphorylation by a histidine kinase is different from that of the eukaryotic serine/threonine or tyrosine kinases. Figure S 45 shows that the phosphate is transferred on to a nitrogen atom in the histidine ring, creating a high-energy phosphoramidate bond. This type of high energy bond is used for phosphoryl transfer in many proteins. When the phosphate is transferred to aspartic acid, it generates a high energy acyl bond. By contrast, the bonds formed with serine, threonine, or tyrosine are between phosphate and hydroxyl group, generating a low-energy ester bond.

Figure S 45 Phosphorylation of histidine or aspartic acid occurs through a high energy bond, but phosphorylation of serine, threonine, or tyrosine occurs as a low energy ester.

More elaborate versions of these systems are called phosphorelays. In such cases, the Two-component signal transduction | SECTION 7.32.13 © 2004. Virtual Text / www.ergito.com

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phosphate group is transferred several times. In a typical case, the pathway starts just like a two-component system, but the second protein in turn transfers the phosphate group to another, and the process may continue (for review see 2308; 2263). The high energy bonds tend to be short-lived, which makes the response rapid and transient. The usual end target of a two-component pathway is the regulation of gene transcription, as summarized inFigure S 46. In the typical bacterial pathway, the effector is the terminal component. It has two domains. The regulatory domain catalyzes transfer on to itself of the phosphate from the sensor histidine kinase (for review see 2263). When it is phosphorylated, it activates the effector domain, which most commonly binds DNA to activate or to repress transcription .

Figure S 46 The effector protein of a two-component system has two domains. When the regulator domain is phosphorylated, the effector domain binds to DNA. This may activate transcription (as shown) or may repress it.

Several basic responses to the environment are mediated by two-component systems, in E. coli including the response to osmotic pressure, redox control, and chemotaxis. A two-component system is used by Agrobacterium when it infects a plant cell (see Molecular Biology 4.18.14 T-DNA carries genes required for infection). Sporulation Two-component signal transduction | SECTION 7.32.13 © 2004. Virtual Text / www.ergito.com

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of B. subtilis is initiated by a phosphorelay system Biology 3.9.19 Sporulation is controlled by sigma factors).

(see

Molecular



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Reviews 2263. Stock, A. M., Robinson, V. L., and Goudreau, P. N. (2000). Two-component signal transduction. Annu. Rev. Biochem. 69, 183-215. 2308. Parkinson, J. S. (1993). Signal transduction schemes of bacteria. Cell 73, 857-871.

References 2307. Nixon, B. T., Ronson, C. W., and Ausubel, F. M. (1986). Two-component regulatory systems responsive to environmental stimuli share strongly conserved domains with the nitrogen assimilation regulatory genes ntrB and ntrC. Proc. Natl. Acad. Sci. USA 83, 7850-7854.



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Genes VIII

Genes VIII

VIII

VIII

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Murder Genes

Pandora's Genes

Good Genes

Genes IX

Immunoglobulin Genes

Essential genes

Genes VII

Red Genes, Blue Genes: Exposing Political Irrationality

Good Genes

Pandora's Genes

Deadly Genes

HOX Genes

Good Genes

Good Genes

Genes IX

Murder Genes

Deadly Genes

Immunoglobulin Genes

Henry VIII

Генрих VIII

Blue Genes

Pandora's Genes

Good Genes

Enrico VIII

Volume VIII

Писмо VIII

Genes VIII

Genes VIII

VIII

VIII

Murder Genes

Murder Genes

Pandora's Genes

Good Genes

Genes IX

Immunoglobulin Genes

Essential genes

Genes VII

Red Genes, Blue Genes: Exposing Political Irrationality

Good Genes

Pandora's Genes

Deadly Genes

HOX Genes

Good Genes

Good Genes

Genes IX

Murder Genes

Deadly Genes

Immunoglobulin Genes

Henry VIII

Генрих VIII

Blue Genes

Pandora's Genes

Good Genes

Enrico VIII

Volume VIII

Писмо VIII

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