Current Protocols in Chemical Biology 2011 (Volume 3)

CURRENT PROTOCOLS in Chemical Biology cp Current Protocols in Chemical Biology Online ISBN: 9780470559277 DOI: 10.1...

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CURRENT PROTOCOLS

in Chemical Biology

cp

Current Protocols in Chemical Biology

Online ISBN: 9780470559277 DOI: 10.1002/9780470559277 Editors & Contributors

EDITORIAL BOARD Adam P. Arkin University of California, Berkeley Berkeley, California Lara Mahal New York University New York, New York Floyd Romesberg The Scripps Research Institute La Jolla, California Kavita Shah Purdue University West Lafayette, Indiana Caroline Shamu Harvard Medical School Boston, Massachusetts Craig Thomas NIH Chemical Genomics Center Rockville, Maryland ASSOCIATE EDITORS Michael Burkart University of California, San Diego San Diego, California John Ellman Yale University New Haven, Connecticut Howard Hang The Rockefeller University New York, New York Hans Luecke National Institute of Diabetes and Digestive and Kidney Diseases, NIH Bethesda, Maryland Andreas Marx Universität Konstanz Konstanz, Germany

Michael Rape University of California, Berkeley Berkeley, California Carsten Schultz EMBL Heidelberg Heidelberg, Germany Oliver Seitz Universität zu Berlin Berlin, Germany Katherine L. Seley-Radtke University of Maryland Baltimore, Maryland Nicky Tolliday Broad Institute of Harvard and MIT Cambridge, Massachusetts Gregory A. Weiss University of California, Irvine Irvine, California CONTRIBUTORS Jasmina J. Allen Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Ruben T. Almaraz The Johns Hopkins University Baltimore, Maryland Rogerio Alves de Almeida University of Manchester Manchester, United Kingdom Alma L. Burlingame Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Christopher T. Campbell National Cancer Institute Frederick, Maryland Jennifer Campbell Harvard Medical School Boston, Massachusetts Yong Chi Fred Hutchinson Cancer Research Center Seattle, Washington Bruce E. Clurman Fred Hutchinson Cancer Research Center Seattle, Washington Benjamin F. Cravatt The Scripps Research Institute La Jolla, California Richard D. Cummings Emory University Atlanta, Georgia Arvin C. Dar

Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Jian Du The Johns Hopkins University Baltimore, Maryland Jeremy R. Duvall The Broad Institute of MIT and Harvard Cambridge, Massachusetts Meng Fang University of Georgia Athens, Georgia Matthew Francis University of California, Berkeley Berkeley, California Jeffrey C. Gildersleeve National Cancer Institute Frederick, Maryland Christian Gloeckner University of Konstanz Konstanz, Germany Jay T. Groves Howard Hughes Medical Institute University of California, San Francisco San Francisco, California and National University of Singapore Singapore And Lawrence Berkeley National Laboratory Berkeley, California Howard C. Hang The Rockefeller University New York, New York Rami N. Hannoush Genentech South San Francisco, California Jamie Heimburg-Molinaro Emory University Atlanta, Georgia Nicholas T. Hertz Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Michal Hocek Academy of Sciences of the Czech Republic Prague, Czech Republic Gregory R. Hoffman Harvard Medical School Boston, Massachusetts Eun Ryoung Jang University of Kentucky Lexington, Kentucky Sean Johnston Harvard Medical School Boston, Massachusetts

Hargun S. Khanna The Johns Hopkins University Baltimore, Maryland Kyung Bo Kim University of Kentucky Lexington, Kentucky Ramon Kranaster University of Konstanz Konstanz, Germany Robert D. Kuchta University of Colorado Boulder, Colorado Maya T. Kunkel University of California at San Diego La Jolla, California Wooin Lee University of Kentucky Lexington, Kentucky Jae-Min Lim University of Georgia Athens, Georgia Wan-Chen Lin Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Hana Macickova-Cahová Academy of Sciences of the Czech Republic Prague, Czech Republic Andrew L. MacKinnon University of California San Francisco San Francisco, California Lisa A. Marcaurelle The Broad Institute of MIT and Harvard Cambridge, Massachusetts Gerard Marriott University of California, Berkeley Berkeley, California Andreas Marx University of Konstanz Konstanz, Germany Nathan J. Moerke Harvard Medical School Boston, Massachusetts Nuzhat Motlekar University of Pennsylvania Philadelphia, Pennsylvania Andrew D. Napper University of Pennsylvania Philadelphia, Pennsylvania and University of Manchester Manchester, United Kingdom and Nemours Center for Childhood Cancer Research Wilmington, Delaware

Alexandra C. Newton University of California at San Diego La Jolla, California Takeaki Ozawa The University of Tokyo and Japan Science and Technology Agency Tokyo, Japan Graham D. Pavitt University of Manchester Manchester, United Kingdom Chutima Petchprayoon University of California, Berkeley Berkeley, California Stewart Rudnicki Harvard Medical School Boston, Massachusetts Kevan M. Shokat Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Sharmila Sivendran University of Pennsylvania Philadelphia, Pennsylvania and GlaxoSmithKline Collegeville, Pennsylvania David F. Smith Emory University Atlanta, Georgia Xuezheng Song Emory University Atlanta, Georgia Anna E. Speers The Scripps Research Institute La Jolla, California Elaine Tan The Johns Hopkins University Baltimore, Maryland Jack Taunton University of California San Francisco San Francisco, California Nicola Tolliday The Broad Institute of MIT and Harvard Cambridge, Massachusetts Sara Triffo Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Yoshio Umezawa Musashino University Tokyo, Japan Milan Vrábel Academy of Sciences of the Czech Republic Prague, Czech Republic

Anita Vrcic The Broad Institute of MIT and Harvard Cambridge, Massachusetts Beatrice T. Wang Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Lance Wells University of Georgia Athens, Georgia Leah S. Witus University of California, Berkeley Berkeley, California Kevin J. Yarema The Johns Hopkins University Baltimore, Maryland Jacob S. Yount The Rockefeller University New York, New York Cheng-Han Yu National University of Singapore Singapore Chao Zhang Howard Hughes Medical Institute University of California, San Francisco San Francisco, California Mingzi M. Zhang The Rockefeller University New York, New York Yalong Zhang National Cancer Institute Frederick, Maryland

Parallel High-Throughput Automated Assays to Measure Cell Growth and Beta-Galactosidase Reporter Gene Expression in the Yeast Saccharomyces cerevisiae Andrew D. Napper,1,3 Nuzhat Motlekar,1 Rogerio Alves de Almeida,2 and Graham D. Pavitt2 1 Penn Center for Molecular Discovery, Institute for Medicine and Engineering, and Department of Chemical and Biomolecular Engineering, University of Pennsylvania, Philadelphia, Pennsylvania 2 Faculty of Life Sciences, University of Manchester, Manchester, United Kingdom 3 Nemours Center for Childhood Cancer Research, Wilmington, Delaware

ABSTRACT Parallel high-throughput automated assays are described for the measurement of cell growth and β-galactosidase reporter gene expression from a single culture of the yeast S. cerevisiae. The dual assay measures the effect of test compounds on expression of a specific gene of interest linked to the β-galactosidase reporter gene, and simultaneously tests for compound toxicity and other effects on cell growth. Examples of assay development and validation results are used to illustrate how this protocol may be used to screen two yeast cell lines in parallel. Yeast cells are grown overnight in V-bottom polypropylene 384-well plates, after which portions of the cell suspension are transferred to clear and to white flat-bottom 384-well plates for measurement of cell growth and reporter gene expression, respectively. Cell growth is determined by measurement of absorbance at 595 nm, and β-galactosidase expression is quantified by Beta-Glo, a commercially available luminescent β-galactosidase substrate. Curr. Protoc. Chem. Biol. C 2011 by John Wiley & Sons, Inc. 3:1-14 Keywords: cell growth r yeast r reporter gene r luciferase r cell-based assay r model organism r mutant gene r β-galactosidase r luminescence

INTRODUCTION Here, we describe HTS assays to assess reporter gene activity in the yeast Saccharomyces cerevisiae. Yeasts have been used extensively to study the effect of specific genes and genetic changes on cellular phenotypes. These studies have provided valuable insight into human disease processes and novel approaches for therapeutic intervention due to the similarity of yeast cellular control mechanisms to those in mammalian cells and the relative ease with which yeast genes can be manipulated. The optimization of these assays is exemplified by their validation for a high-throughput screen designed to discover chemical modulators of stress-response pathways in yeast (Alves de Almeida et al., 2008; Motlekar et al., 2009). Both cell growth and expression of a chromosomally integrated GCN4-dependent reporter gene fusion to β-galactosidase were measured in two yeast strains, wild-type Saccharomyces cerevisiae and an eIF2B mutant. GCN4 is a central regulator of general amino acid control stress response (GAAC), many effects of which are mediated through the translation factor eIF2B. Mutant and wildtype yeast strains were screened in parallel to identify compounds acting specifically on either strain: wild type–specific compounds affecting GAAC through functional eIF2B, Current Protocols in Chemical Biology 3: 1-14, January 2011 Published online January 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470559277.ch100119 C 2011 John Wiley & Sons, Inc. Copyright

HighThroughput Assays of Yeast Cell Growth

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and mutant-specific compounds that restore normal function to mutant eIF2B. The gene expression assay described here was optimized for high-throughput screening (HTS) by adaptation of a commercially available coupled β-galactosidase–firefly luciferase system for use with yeast cells (Alves de Almeida et al., 2008; Motlekar et al., 2009). In this assay β-galactosidase present in the yeast cells reacts with 6-O-β-galactopyranosyl-luciferin substrate to generate D-luciferin. This, in turn, reacts with ATP and firefly luciferase (provided in the commercially available assay reagent kit) in the presence of atmospheric oxygen to form oxyluciferin and emit light. In the discussion that follows the protocols, technical challenges encountered in miniaturizing to a robot-compatible 384-well format are described. The progression of protocols is as follows:

r Basic Protocol 1: Preparation of frozen yeast stocks (matched pair, e.g., wild type r r r r r r

BASIC PROTOCOL 1

and mutant gene), optimization of growth conditions, and determination of the effect of DMSO on cell growth Basic Protocol 2: Yeast end-point growth assay: absorbance at 595 nm Basic Protocol 3: Time-course luminescence measurement of β-galactosidase reporter expression Basic Protocol 4: End-point luminescence measurement of β-galactosidase expression Basic Protocol 5: Single concentration compound screening Basic Protocol 6: Data analysis and hit selection Basic Protocol 7: Dose-response testing, curve fitting, IC50 determination, and hit confirmation

PREPARATION OF FROZEN YEAST STOCKS, OPTIMIZATION OF CELL GROWTH CONDITIONS, AND DETERMINATION OF THE EFFECT OF DMSO ON YEAST CELL GROWTH Yeast cell stocks are prepared in sufficient quantity for assay validation, HTS, and anticipated follow-up experiments. These stocks are divided into single-use aliquots to ensure a consistent source of cells throughout all subsequent experiments. Experiments to determine a cell growth time course and the effect of DMSO on cell growth are carried out prior to configuring the assay for HTS.

Materials Matched pair of yeast strains YPD plates (contain YPD medium with an addition of 20 g/liter agar; supplied by Formedium) YPD medium (10 g/liter yeast extract, 20 g/liter bacto peptone, and 20 g/liter glucose; available from Sigma) Freezing medium: YPD medium with 15% (v/v) glycerol Dry ice 0% to 1.6% (v/v) final dimethyl sulfoxide (DMSO)

High-Throughput Assays of Yeast Cell Growth

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30◦ C incubator 250-ml Erlenmeyer flasks Shaking incubator Freezer vials for yeast stocks: 0.2-ml snap-cap PCR tubes (from any supplier of standard laboratory accessories) Polypropylene V-bottom 384-well plates (Greiner Bio-One, cat. no. 781280) Pipetting workstation equipped with 384-tip pipetting head (e.g., JANUS from Perkin Elmer or equivalent; Rudnicki and Johnston, 2009) Reagent dispenser (e.g., Multidrop-384 reagent dispenser from Thermo Scientific; Rudnicki and Johnston, 2009) Plate reader capable of reading absorbance and luminescence in 384-well plates (e.g., EnVision multimode plate reader from Perkin Elmer) Current Protocols in Chemical Biology

Clear 384-well plates (Corning, cat. no. 3702) Breathe-Easy sealing films (Sigma, cat. no. Z380059) Prepare the frozen yeast stocks 1. Remove the strains from the –80◦ C freezer, streak onto YPD plates, and incubate the plates for 3 days at 30◦ C. For the study used as an example here, wild-type yeast strain GP4213 (MATα leu2-3 leu2-112 ura3-52::HIS4–lacZ ino1 gcd6 gcn2::hisG pAV1265[GCD6 CEN6 LEU2]) and an isogenic gcd6 mutant strain GP4198 (MATα leu2-3 leu2-112 ura3-52::HIS4–lacZ ino1 gcd6 gcn2::hisG pAV1744[gcd6-R284H CEN6 LEU2]) (Alves de Almeida et al., 2008) were used. Store at –80◦ C as single-use frozen stocks in YPD containing 15% (v/v) glycerol. Generation of yeast strains is outside the scope of this protocol. It is assumed that the reader has already obtained the desired strains.

2. Prepare cell stocks of the two yeast strains by inoculating a single colony into 50 ml of YPD medium at room temperature in a 250-ml Erlenmeyer flask and growing the cultures overnight at 30◦ C with vigorous shaking (200 rpm). 3. Measure absorbance at 595 nm (A595) after the overnight growth. The cell suspension should give an absorbance of at least 4 to 6 before proceeding to step 4. (Absorbance at 595 nm is a measure of cell density: increased cell density increases turbidity of the cell suspension, and turbidity may be measured by absorbance at 595 nm.) Note that dilution of the culture might be necessary to obtain a reading in the linear range of the plate reader. See Critical Parameters for a more detailed explanation of the use of absorbance to measure cell density.

4. Centrifuge the cell culture at 5 min 3000 × g, 4◦ C, decant the supernatant, and resuspend the pelleted cells in freezing medium to generate a cell suspension with A595 = 5.0. 5. Divide the cells into aliquots in freezer vials for single use. Freeze on dry ice and store up to 1 year at –80◦ C. A volume of 0.5 ml contains sufficient cell stock for screening of twenty plates, taking into account dead volume and wastage of the liquid dispenser.

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Absorbance at 595 nm

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Figure 1 Dilution of a suspension of yeast cell stock used to determine the linear range of absorbance determination at 595 nm (A595 ). Previously published in Motlekar (2009). Current Protocols in Chemical Biology


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Determine linear range of absorbance values as a measure of cell density 6. Thaw the yeast cell stock frozen at –80◦ C (from step 5) and serially dilute in a clear 384-well plate in YPD medium to obtain 50 μl per well of dilutions ranging from 1:5 to 1:500. Measure A595 in the plate reader. 7. Graph A595 data versus dilution to determine the linear range of absorbance values useful for monitoring yeast growth (Fig. 1). See Critical Parameters for more detailed notes on determining the linear range of absorbance values useful for monitoring yeast growth.

Determine optimal time to incubate yeast suspensions prior to recording growth 8. Select the optimal time for end-point reading in the growth assay by determining a growth time course over 24 hr as follows: a. For each time point fill a clear polystyrene 384-well plate with 25 μl per well of YPD medium. b. Dilute the thawed wild-type and mutant yeast cells 1:500 in YPD medium. Inoculate 25 μl of each of the diluted stocks to specific wells as follows: Add 25 μl diluted wild-type cells to wells A-P in columns 2 and 3-12, add 25 μl diluted mutant cells to wells A-P in columns 13-22 and 24, and add 25 μl of YPD medium in place of yeast cells to wells designated as blanks (wells A-P in columns 1 and 23). c. Seal the plates with Breathe-Easy sealing films and incubate the yeast cultures at 30◦ C without shaking for 4, 8, 16, or 24 hr. d. After incubation, remove the Breathe-Easy membranes and measure A595 in the plate reader. The optimal incubation time prior to recording growth will be one in which the strain is still growing in exponential phase and the cell density is in the linear range of absorbance values (Fig. 2). See Critical Parameters for more details.

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Figure 2 Time course of yeast growth. Growth time course of wild-type (square) and mutant (triangle) yeast determined by measurement of A595 at each time point. Blanks consisting of medium alone (diamond) were used to monitor A595 in the absence of cell growth. Previously published in Motlekar (2009).

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Figure 3 Effect of DMSO on yeast growth. Cell growth after 16 hr was determined by measurement of A595 of a suspension of wild-type cells (light gray), mutant cells (black), or medium blank (dark gray). Error bars represent mean ± standard deviation of 16 replicate wells. Previously published in Motlekar (2009).

Determine the effect of DMSO on yeast cell growth 9. Grow both wild-type and mutant yeast cells for 16 hr in the presence of 0% to 1.6% (v/v) final DMSO concentration. Perform the growth assay as described in step 8a-c, except mix DMSO with the medium in the assay plate prior to the addition of yeast cells. Include wells without added DMSO as controls. To calculate the percent growth inhibition caused by addition of DMSO, see Basic Protocol 6. Figure 3 shows the effect of DMSO on the growth of wild-type and mutant cells. See Critical Parameters for more details on the effects of DMSO on the growth of S. cerevisiae.

YEAST END-POINT GROWTH ASSAY: ABSORBANCE AT 595 NM If growth of two yeast cell lines (e.g., mutant and wild type) is to be tested in parallel, a control column of the other cell line should be included in each test plate, as shown in Figure 4. Only negative control wells (yeast grown in the absence of test compounds) are included in the plate layout described here. Positive control wells (containing compounds known to affect the growth of the yeast strains in the assay) could be included if desired in columns 2 or 24.

BASIC PROTOCOL 2

Materials YPD medium (10 g/liter yeast extract, 20 g/liter bacto peptone, and 20 g/liter glucose; available from Sigma) Test compounds in DMSO Yeast strains, wild type and mutant Z buffer (see recipe) Polypropylene V-bottom 384-well plates (Greiner Bio-One, cat. no. 781280) Pipetting workstation equipped with pintool consisting of 384 pins with nominal transfer volume of 100 nl (e.g., JANUS MDT from Perkin Elmer or equivalent; Rudnicki and Johnston, 2009) Pipetting workstation equipped with 384-tip pipetting head (e.g., JANUS from Perkin Elmer or equivalent; Rudnicki and Johnston, 2009)


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Figure 4

Assay plate formats showing (A) the wild-type assay plate and the (B) mutant assay plate.

Vortex Breathe-Easy sealing films (Sigma, cat. no. Z380059) 30◦ C incubator White 384-well plates (Corning Life Sciences, cat. no. 3652) Clear 384-well plates (Corning Life Sciences, cat. no. 3702) Plate reader capable of reading absorbance and luminescence in 384-well plates (e.g., EnVision multimode plate reader from Perkin Elmer) 1. Fill the V-bottom polypropylene 384-well plates with 25 μl of YPD medium using a reagent dispenser, and add an additional 25 μl of YPD medium to column 1 (for blanks). See Figure 4 for plate layout. 2. Add test compounds by pintool transfer to give a final concentration not to exceed 25 μM of each compound in 0.25% DMSO. Reserve wells in columns 1, 2, 23, and 24 for controls and blanks containing 0.25% DMSO but no compound.


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In the example assay here, a 384-pin pintool was used to transfer 120 nl of a 10 mM solution in DMSO into the medium-containing V-bottom growth plates, giving a final concentration of 24 μM of each compound in 0.24% DMSO in columns 3-22 once yeast cells are added. Columns 1, 2, 23, and 24 of the library plates contain DMSO in this case, so 0.24% DMSO (final concentration), but no compound, is transferred to wells in those columns.

3. Thaw frozen stocks of wild-type and mutant yeast cells and dilute 1:500 in YPD medium. Mix by vortexing to ensure even resuspension. Current Protocols in Chemical Biology

4a. Wild-type yeast test plates: Add 25 μl of diluted wild-type stock to all columns, except 1 and 23, in V-bottom growth plates using a reagent dispenser. Add 25 μl of diluted mutant stock to column 23. 4b. Mutant strain test plates: Add 25 μl of diluted mutant stock to all columns, except 1 and 23, in V-bottom growth plates using a reagent dispenser. Add 25 μl of diluted wild-type stock to column 23. 5. Seal plates with Breathe-Easy sealing film and incubate the yeast cultures for 16 hr at 30◦ C. This incubation time was selected based on earlier determinations of growth curves in Basic Protocol 1. The optimal time may vary between different strains and needs to be determined empirically in each case.

6. Mix the cell suspension by repeated aspirate-dispense cycles using 30-μl disposable tips on the 384-well pipetting head in the pipetting workstation. Transfer 5 μl to a white 384-well plate containing 20 μl Z buffer for use in Basic Protocol 3. Transfer 30 μl from each well of the portion that remains into wells in a clear 384-well flat-bottom plate, taking care to avoid formation of bubbles in the destination plate. Record A595 in the clear flat-bottom plate using the plate reader. 7. Calculate percent inhibition of growth from the A595 data, as described in Basic Protocol 6. β-GALACTOSIDASE REPORTER EXPRESSION TIME COURSE IN YEAST This protocol measures expression of a gene of interest linked to the expression of the reporter gene β-galactosidase. Enzymatic conversion of the Beta-Glo reagent by β-galactosidase results in long-lived luminescence. To obtain consistent results, the light output should be allowed to reach a steady state prior to recording luminescence. The time to reach a steady state is determined from a luminescence time course.

BASIC PROTOCOL 3

Materials Yeast cell suspension prepared in Basic Protocol 2, step 6 Z buffer Beta-Glo assay system (Promega, cat. no. E4780) White, flat-bottomed 384-well plates Reagent dispenser (e.g., Multidrop-384 reagent dispenser from Thermo Scientific; Rudnicki and Johnston, 2009) Pipetting workstation equipped with 384-tip pipetting head (e.g., JANUS from Perkin Elmer or equivalent; Rudnicki and Johnston, 2009) Plate reader capable of reading absorbance and luminescence in 384-well plates (e.g., EnVision multimode plate reader from Perkin Elmer) 1. Add 20 μl Z buffer to each well of a white, flat-bottomed 384-well plates using a reagent dispenser. 2. Add 5 μl of yeast cell suspension by tip transfer from the V-bottom yeast growth plates using a 384-well pipetting head (see Basic Protocol 2, step 6). 3. Mix the cells and Z buffer by pipetting, using the pipets on the pipetting head on the workstation from step 2, and incubate for 20 min at room temperature to permeabilize the cells. 4. Add 25 μl of Beta-Glo reagent using a reagent dispenser. 5. Measure luminescence at 2-min intervals for 4 hr on the Envision plate reader. For data workup, see Basic Protocol 6. Figure 5 depicts sample data from a luminescence time course. Current Protocols in Chemical Biology


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800

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Figure 5 Time course of luminescence resulting from GCN4-dependent expression of the βgalactosidase reporter gene. Wild-type (circle) and eIF2B mutant (diamond) yeast strains were grown overnight and mixed with Beta-Glo reagent. Error bars represent mean ± standard deviation of 16 replicate wells. In this experiment, there was no significant difference in growth rates between the mutant and wild-type strain and the data shown have not been corrected for cell growth. Previously published in Motlekar (2009).

BASIC PROTOCOL 4

LUMINESCENCE END-POINT β-GALACTOSIDASE EXPRESSION ASSAY

BASIC PROTOCOL 5

SINGLE-CONCENTRATION COMPOUND SCREENING

BASIC PROTOCOL 6

DATA ANALYSIS AND HIT SELECTION

This protocol is identical to Basic Protocol 3, except in step 5, luminescence is read once after light output has reached steady state. Using the strain and protocol described here, that time is 90 min after Beta-Glo reagent is added.

Add test compounds to yeast growth plates as described in Basic Protocol 2. Measure the cell growth (Basic Protocol 2) and β-galactosidase reporter gene expression at a single timepoint (Basic Protocol 4).

Yeast cell growth data and β-galactosidase reporter gene expression data obtained in Basic Protocol 5 is analyzed as follows to determine the percent inhibition of growth and percent inhibition of luminescence. Test compounds are selected as hits if these values exceed a defined threshold. 1. Calculate percent inhibition of growth or percent inhibition of luminescence for each test compound using the signal in absorbance units (OD) or luminescence units (LU) readout for each well, and the mean of the plate’s negative controls (columns 2 and 24), and the mean of the plate’s blanks (column 1; see plate maps in Figure 4):

% Inhibition =


100 × [1 - (signal-blank mean) ] ( negative control mean-blank mean) Equation 1

The reference yeast strain (column 23) is not used in the calculation.

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2. Correct percent inhibition of luminescence for cell growth:

Corrected % inhibition =

100 × [1 – (100 − % reporter gene inhibition)] (100 − % inhibition of growth) Equation 2

3. Equations 1 and 2 may also be used if the goal of the compound screening is to identify activators of growth or luminescence. A negative value of percent inhibition indicates activation, in which growth or luminescence in the presence of test compound is increased relative to the negative control mean. These negative percent inhibition values may be converted to percent activation as follows: Percent activation = −1 × percent inhibition Equation 3

4. Select hits based on percent inhibition (or activation) exceeding a defined threshold. For the hits to be statistically significant (i.e., not just due to random scatter in the data), the threshold should be below (or above for activators) the mean of the plate’s negative controls by a minimum of three standard deviations of the controls. The standard deviation of the plate controls in the yeast growth and luminescence assays typically ranges from 10% to 15%; thus, the minimal hit threshold would be set at 30% to 45% inhibition. For assays with a high hit rate (>1%), the threshold may be set as high as 80% inhibition to keep the number of hits to a manageable number for follow-up testing.

DOSE-RESPONSE TESTING, CURVE FITTING, IC50 DETERMINATION, AND HIT CONFIRMATION

BASIC PROTOCOL 7

This protocol is identical to single compound screening (Basic Protocol 5), except that compounds are tested at multiple concentrations obtained by serial dilution. A suitable layout is sixteen two-fold dilutions of each compound, giving an assay concentration range of 100 μM to 3 nM. In this case, compounds may be serially diluted vertically down each plate such that columns 3 to 22 each contain the dilutions of one compound. Following dose-response testing, the yeast growth and luminescence data are fit to a dose-response curve to calculate IC50 (for inhibitors) or EC50 (for activators).

Materials Compound stocks (10 mM in DMSO) Dimethyl sulfoxide (DMSO) 384-well V-bottom polypropylene plates Reagent dispenser (e.g., Multidrop-384 reagent dispenser from Thermo Scientific; Rudnicki and Johnston, 2009) Pipetting workstation equipped with pintool consisting of 384 pins with nominal transfer volume of 100 nl (e.g., JANUS MDT from Perkin Elmer or equivalent; Rudnicki and Johnston, 2009) Microsoft Excel: for calculation of percent inhibition or activation (if screening database is not available) GraphPad Prism or equivalent: for graphing data and curve fitting for IC50 and EC50 calculation (if screening database is not available) OpenHTS (CeuticalSoft), ActivityBase (IDBS), or equivalent screening database: for calculation of percent inhibition or activation, comparison of data sets and selection of hits



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Perform serial dilution in DMSO 1. Add 20 μl of each compound stock (10 mM in DMSO) arranged twenty per plate in wells A3 to A22 of 384-well V-bottom polypropylene plates. 2. Add 20 μl DMSO to wells A1, A2, A23, and A24 3. Add 10 μl DMSO per well to the entire plate except row A using a reagent dispenser. 4. Serial dilute the compounds two-fold by transferring 10 μl row-by-row from row A to row P using a single row of disposable tips, then remove and discard 10 μl from row P. The resulting dose-response plates contain sixteen two-fold dilutions of each compound, ranging from 10 mM to 305 nM, arranged one compound per column in columns 3 to 22. Pintool transfer into growth plates (Basic Protocol 2) gives a final range of 100 μM to 3 nM.

5. Proceed with the screening assay (Basic Protocol 5).

Fit the data to determine IC50 or EC50 6. Calculate percent growth and β-galactosidase activity for each dilution of each compound from A595 and luminescence, respectively, and the means of the plate controls and plate blanks (see plate maps in Fig. 4): % Activity = 100 × [(signal-blank mean)/(control mean-blank mean)] Equation 4

7. If there is a dose-dependent effect on growth (as revealed by a decrease in percent growth as the test compound concentration increases), correct the luminescence assay percent activity values at each compound dose for cell growth using the following equation: Corrected % activity = % activity/% growth (relative to growth controls) Equation 5

8. Fit percent activity data to a dose-response curve using nonlinear regression. A screening database or a curve-fitting and graphing program, such as GraphFit Prism, will perform a four-parameter logistic fit and calculate IC50 or EC50 as appropriate.

REAGENTS AND SOLUTIONS Z buffer 82 mM disodium hydrogen phosphate 9 mM sodium dihydrogen phosphate 0.1% (w/v) SDS 1 mM DTT Protease inhibitor tablets (Roche Applied Science, cat. no. 1873580) The sodium phosphate/SDS buffer is stable at room temperature for at least 1 year, but DTT and protease inhibitors should be added fresh on each day of use. DTT is stored frozen at −20◦ C as a 1 M stock (1000×).

COMMENTARY Background Information High-Throughput Assays of Yeast Cell Growth

Reporter gene fusions have been widely employed in biological research for decades. They provide a simple experimental means to study the control of expression of genes.

Ease and sensitivity of measurement, and determination of whether the host system naturally expresses the reporter gene are important considerations that govern the usefulness and choice of reporter gene fusion.

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Expression of commonly employed reporter genes may be detected by specific substrates that produce colored chemical products or emit light, easily detected with a spectrophotometer or luminometer (Sambrook and Russell, 2001). The Escherichia coli lacZ gene is one such reporter. The β-galactosidase enzyme was first studied at least 60 years ago (Lederberg, 1950) and is encoded by lacZ. β-galactosidase naturally cleaves lactose into glucose and galactose. The development of colorimetric substrates that mimic the natural lactose substrate, including O-nitrophenylβ-D-galactopyranoside (ONPG), greatly enhanced its subsequent widespread utilization. ONPG is colorless, but when cleaved it releases O-nitrophenol, which is yellow and absorbs at A420 . Thus, in the presence of excess ONPG, the yellow color develops at a rate proportional to the amount of LacZ protein present in a cell lysate (Miller, 1972). In recent years, parallel development of instrumentation, data analysis, and substrates with enhanced sensitivity has made it possible for reporter gene studies to be performed in a variety of high-throughput screening (HTS) formats. Luminescent reagents have been especially useful in facilitating low-microliter volume “mix-and-read” assays in 384- and 1536-well plates amenable to high-throughput robotic liquid handling (Inglese et al., 2007).

sure cell density after an overnight incubation, provided the yeast cells remain in an exponential growth phase. To ensure optimal growth conditions in Basic Protocol 2, yeast growth is first studied and optimized in Basic Protocol 1. Yeast growth is determined by measurement of absorbance at 595 nm (A595 ). Points to note: i. In this assay, absorbance is an indirect measure of turbidity. Turbidity of the yeast suspension increases in proportion to cell number during growth. ii. Increase in A595 is due to light scattering by the turbid suspension and not absorbance. Measurement at a long wavelength (595 nm or above) ensures that there is no absorbance due to the yeast cells or most test compounds. iii. At low cell density, there is a direct linear relationship between A595 and yeast cell number determined by turbidity of the suspension. Above a certain cell density, the relationship between absorbance and cell number deviates from linearity, and A595 understates the true cell number (Warringer and Blomberg, 2003). iv. To ensure that A595 provides a true measure of cell number, it is necessary to obtain a standard curve of A595 against yeast cell density to determine the linear range of the growth assay (see iii above). v. DMSO is known to affect yeast cells (Murata et al., 2003), so sensitivity of yeast cell growth to DMSO should be assessed.

Critical Parameters and Troubleshooting

Yeast end-point growth assay: Absorbance at 595 nm In Basic Protocol 2, yeast cells are grown overnight, after which growth is measured and a portion of the cells are removed for testing in the β-galactosidase reporter gene assay (Basic Protocol 4). Points to note: i. Incubation time for the growth assay should be selected based on optimization of growth conditions (Basic Protocol 1, steps 6 to 8). A 16-hr incubation time was selected for the yeast cell lines described here. ii. IMPORTANT: Overnight growth in Vbottom polypropylene plates instead of flatbottom plates was necessary for consistent mixing of the suspension with pipet tips and precise cell transfer by automated liquid handling for the β-galactosidase reporter gene assay (also see Luminescence end-point βgalactosidase expression assay). iii. The concentration of the test compounds should be no more than 25 μM to avoid nonspecific toxicity. Likewise, DMSO concentration should be below the level at which effects on growth were observed (Basic Protocol 1, step 9).

The yeast growth (Basic Protocols 1 and 2) and β-galactosidase expression assays (Basic Protocols 3 and 4) require careful and extensive optimization. Given the multitude of parameters that can affect data quality and reproducibility, careful analysis of the results is required. These issues are discussed here. Preparation of frozen yeast stocks Yeast strains are grown once to generate cell suspension sufficient for all assay development, screening, and hit confirmation. Advantages include: Consistency: A portion of the same cell suspension is used for each experiment. Ease of assay setup: Avoids the need to grow yeast strains before each experiment. Cell suspension samples are stored in 0.2ml PCR tubes, as these allow for rapid freezing and thawing. Optimization of growth conditions To allow sufficient time to discern test compound effects, the yeast end-point growth assay (Basic Protocol 2) was designed to mea-



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Luminescence end-point β-galactosidase expression assay Yeast cells are grown overnight in V-bottom polypropylene plates, after which a portion of the cells is removed for testing in the βgalactosidase reporter gene assay (Basic Protocol 4). Points to note: i. IMPORTANT: Mixing and transferring of the cell suspensions from the yeast growth plate to the β-galactosidase assay plate must be carefully optimized. After overnight growth, the yeast cells needed to be resuspended to ensure reproducible transfer of a small volume of cells for the luminescence assay. Mixing in a flat-bottom plate tends to concentrate the cells to the rims of the well; therefore, cells should be grown in V-bottom polypropylene plates. Efficient mixing necessitates slow aspiration of the cell suspension with each pipet tip a fraction of a millimeter above the well bottom. ii. Luminescence should be read after light output has reached a steady state.


Data analysis and hit selection The selection of “hits” following compound screening is a critical step. It is important that the data be analyzed in such a way that compounds with the desired activity are selected. Points to note: i. Luminescence values obtained in Basic Protocol 4 will be a true measure of inhibition (or activation) of β-galactosidase gene expression only if the data are corrected to take account of each compound’s effect on cell growth in Basic Protocol 2. In the absence of this growth correction (Basic Protocol 6, step 2), luminescence values are decreased or increased in proportion to the cell density following overnight yeast growth. ii. If the primary screening data are highly scattered (standard deviation of plate controls is >15%), the correction described in (i) above may not be reliable. In this case, the primary screening should be performed in duplicate, or hits should be selected based on uncorrected percent inhibition (or activation) of luminescence, and the correction made upon doseresponse testing (Basic Protocol 7). iii. If two cell lines are screened in parallel (Basic Protocol 2), compounds selectively active against one of the cell lines may be identified after primary screening, provided the data quality is acceptable (standard deviation of plate controls is 2fold higher luminescence. This difference provided an assay signal window sufficient for parallel screening of both wild-type and eIF2B mutant yeast to identify test compounds that restore the function of mutant eIF2B (Motlekar et al., 2009).

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Figure 6 Reporter gene assay validation plate. eIF2B mutant control (square), wild-type control (triangle), blank (black circle), and test (diamond) wells to which DMSO has been added to a final concentration of 0.24% were assayed for β-galactosidase expression. Previously published in Motlekar (2009).

Basic Protocol 4: Luminescence end-point β-galactosidase expression assay Figure 6 depicts results from a reporter gene assay validation plate. The cell suspension to which DMSO had been added by pintool (no test compounds) was grown overnight in a V-bottom 384-well plate, after which 5 μl was transferred to a white 384-well plate and mixed with Z buffer and Beta-Glo. Luminescence results shown provide a measure of well-to-well variability and confirm the 2-fold difference between wild-type and eIF2B mutant expression of the β-galactosidase reporter gene. Depending on the hit threshold chosen for screening (Basic Protocol 6, step 4), several outliers in Figure 6 might be selected as hits (known as false positives since the deviation from the plate control mean is not due to compound activity). Luminescence values from wells 120, 240, and 255, in particular, deviate significantly from the remainder of the data. If outliers such as these are observed, a repeat of the experiment will reveal whether there is a problem associated with a specific pin in the pintool or a specific channel in the pipetting head. If the outliers are not replicated at the same well locations between experiments, then they may be attributed to random data scatter. If further refinement of the liquid handling steps fails to reduce the occurrence of such outliers, screening may nonetheless

proceed. False-positive outliers will be eliminated during dose-response testing (Basic Protocol 7).

Time Considerations Preparation of frozen yeast stocks (Basic Protocol 1): Total growth time is 4 days, with 1 to 2 hr of hands-on time on days 0, 3, and 4. Yeast stocks are stable at −80◦ C for at least one year, so this procedure may be required only once. Optimization of growth conditions (Basic Protocol 1): Total growth time is 1 day, with 2 to 3 hr of hands-on time on days 0 and 1. Provided the experiment works according to the protocol, this optimization will only need to be performed once. End-point growth assay: absorbance at 595 nm (Basic Protocol 2): Total growth time is 16 hr, with 2 to 3 hr of hands-on time on days 0 and 1. β-galactosidase reporter expression time course (Basic Protocol 3): Total growth time is 16 hr, followed by a 3-hr luminescence time. Allow 1 to 2 hr before and 2 to 4 hr after for set up and data analysis, respectively. Luminescence end-point β-galactosidase expression assay (Basic Protocol 4): Total growth time is 16 hr, followed by a 90-min luminescence assay. Allow 1 to 2 hr before and 2 to 4 hr after for set up and data collection, respectively.



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Single concentration compound screening (Basic Protocol 5): Total growth time is 16 hr, followed by a 90-min luminescence assay. Up to twenty plates may be nested for batchwise testing. For a twenty-plate screening run, allow 4 to 5 hr before starting yeast growth, and 8 hr the following day for luminescence assay set up and data collection. Data analysis and hit selection (Basic Protocol 6): Allow 1 day per week during screening. Dose-response testing, curve fitting, IC50 determination, and hit confirmation (Basic Protocol 7): Allow 1 day to select hits and serially dilute. Testing time similar to single-concentration compound screening above. Allow 1 day for data analysis of 100 to 200 compounds.

Acknowledgements Development and validation of the assays reported here was supported by the NIH Molecular Libraries Screening Center Network (Grant U54HG003915-02) and grant ELA 2005-008C5 from the European Leukodystrophy Association.

Literature Cited Alves de Almeida, R., Burgess, D., Shema, R., Motlekar, N., Napper, A.D., Diamond, S.L., and Pavitt, G.D. 2008. A Saccharomyces cerevisiae cell-based quantitative beta-galactosidase assay compatible with robotic handling and highthroughput screening. Yeast 25:71-76.

Inglese, J., Johnson, R.L., Simeonov, A., Xia, M., Zheng, W., Austin, C.P., and Auld, D.S. 2007. High-throughput screening assays for the identification of chemical probes. Nat. Chem. Biol. 3:466-479. Lederberg, J. 1950. The beta-D-galactosidase of Escherichia coli, strain K-12. J. Bacteriol. 60:381392. Miller, J.H. 1972. Experiments in Molecular Genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Motlekar, N., de Almeida, R.A., Pavitt, G.D., Diamond, S.L., and Napper, A.D. 2009. Discovery of chemical modulators of a conserved translational control pathway by parallel screening in yeast. Assay Drug Dev. Technol. 7:479494. Murata, Y., Watanabe, T., Sato, M., Momose, Y., Nakahara, T., Oka, S., and Iwahashi, H. 2003. Dimethyl sulfoxide exposure facilitates phospholipid biosynthesis and cellular membrane proliferation in yeast cells. J. Biol. Chem. 278:33185-33193. Rudnicki, S. and Johnston, S. 2009. Overview of liquid handling Instrumentation for highthroughput screening applications. Curr. Protoc. Chem. Biol. 1:43-54. Sambrook, J., and Russell, D.W. 2001. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Warringer, J. and Blomberg, A. 2003. Automated screening in environmental arrays allows analysis of quantitative phenotypic profiles in Saccharomyces cerevisiae. Yeast 20:53-67.


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Development of Chemical Probes for Biochemical Detection and Cellular Imaging of Myristoylated and Palmitoylated Proteins Rami N. Hannoush1 1

Genentech, South San Francisco, California

ABSTRACT Fatty acylation of proteins regulates their spatial localization and activity in living cells. Methods to monitor fatty acylation are invaluable for studying its role in regulating protein dynamics. The protocols in this unit describe a procedure that involves metabolic labeling with ω-alkynyl fatty acids for detecting and cellular imaging of fatty-acylated proteins, namely myristoylated and palmitoylated proteins. Curr. Protoc. Chem. Biol. C 2011 by John Wiley & Sons, Inc. 3:15-26 Keywords: myristoylation r palmitoylation r alkynyl fatty acids r chemical probes r metabolic labeling r click chemistry r fatty acylation

INTRODUCTION This unit describes a general procedure for the biochemical detection and cellular imaging of myristoylated and palmitoylated proteins using ω-alkynyl fatty acid probes and click chemistry. Fatty acylation of cellular proteins regulates their spatial localization and activity in living cells. On a molecular level, fatty acylation involves the enzymecatalyzed addition of 14-carbon (myristoylation) or 16-carbon (palmitoylation) fatty acid chains to cellular proteins via amide (N-myristoylation) or thioester (S-palmitoylation) linkages, respectively. A wide variety of mammalian proteins are fatty acylated, and these include a subpopulation of kinases, GTPases, heterotrimeric G proteins, cytokines, and phosphatases (Resh, 2006; Linder and Deschenes, 2007). Methods to monitor the status of protein myristoylation or palmitoylation are invaluable for studying cellular fatty acylation and its role in regulating protein behavior (Hannoush and Sun, 2010). The method described herein utilizes fatty acid analogues modified at their termini with alkyne groups (Hannoush and Arenas-Ramirez, 2009). These probes are added to cultured cells and are incorporated into cellular proteins (Fig. 1). Proteins modified with these probes are selectively conjugated to biotin-azide or rhodamine-azide that reacts with the alkyne group via a Cu(I)-catalyzed [3+2] Huisgen cycloaddition reaction (or click reaction) (Rostovtsev et al., 2002; Tornoe et al., 2002; Wang et al., 2003). The labeled proteins are then separated by SDS-PAGE and detected by either immunoblotting or in-gel fluorescence (Basic Protocol 1 and Fig. 2). Alternatively, the user can image fattyacylated proteins by fluorescence microscopy (Fig. 1). This entails fixing cells that have been metabolically labeled with the fatty acid probes and processing them with click chemistry (Basic Protocol 2 and Fig. 3). The labeling procedure has a number of attractive features. It is nonradioactive, highly sensitive, and applicable across a wide range of cellular systems. It also requires short detection time. Furthermore, the alkynyl fatty acid reagents are portable, tunable in concentration, and can be conveniently stored in a freezer for immediate use. All of

Current Protocols in Chemical Biology 3: 15-26, February 2011 Published online February 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470559277.ch100143 C 2011 John Wiley & Sons, Inc. Copyright

Probes for Imaging Myristoylated, Palmitoylated Proteins

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A

O

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n=9 n = 10 n = 12 n = 14

OH

Symbol

Chemical name

Detection

Alk-C13 Alk-C14 Alk-C16 Alk-C18

12-Tridecynoic acid 13-Tetradecynoic acid 15-Hexadecynoic acid 17-Octadecynoic acid

myristoylation myristoylation palmitoylation palmitoylation

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fix and permeabilize cells

N N

CuSO4, TCEP, TBTA

O OH

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n

Detection: immunoblot or in-gel fluorescence

N3 cellular imaging CuSO4, TCEP, TBTA

Figure 1 Schematic depicting the method for monitoring myristoylated and palmitoylated cellular proteins. (A) ω-Alkynyl fatty acid probes used for detecting protein myristoylation and palmitoylation. (B) ω-Alkynyl fatty acids added to growth medium are metabolically incorporated into cellular fatty-acylated proteins. The cells are then either lysed and the proteome detected by immunoblotting or fixed for imaging by fluorescence microscopy. In both cases, click chemistry is used to chemoselectively conjugate a biotin-azide or rhodamineazide to fatty-acylated proteins. Alk-C13 and Alk-C14 are probes for detecting protein myristoylation, while Alk-C16 and Alk-C18 detect protein palmitoylation.

this makes this technique well suited for characterizing and studying fatty acylation of cellular proteins of interest in various contexts. A key application of the alkynyl fatty acid probes, when combined with fluorescence microscopy, is imaging the global subcellular distribution of myristoylated and palmitoylated proteins in different cell lines (Hannoush and Arenas-Ramirez, 2009). This provides important information about the dynamics and behavior of fatty-acylated proteins in living cells.

STRATEGIC PLANNING


Researchers first need to determine the cell type to be used for studying fatty acylation. For detecting protein myristoylation, Alk-C13 or Alk-C14 should be used (Fig. 1). Alternatively, Alk-C16 and Alk-C18 are appropriate probes for detecting protein palmitoylation (Fig. 1). In general, these probes work across a wide range of mammalian cell lines and their uptake may vary depending on the particular cell type used. In every case, the researcher should determine the optimal concentration of the probe for obtaining the maximum signal-to-noise ratio. Furthermore, one needs to check for palmitoylation or myristoylation by verifying the type of linkage via which the individual probe is incorporated into cellular proteins (thioester versus amide linkage). This is done by measuring sensitivity to hydroxylamine (in Basic Protocol 1), a reagent that cleaves thioester- but not amide-linked acyl chains.

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BIOCHEMICAL DETECTION OF MYRISTOYLATED AND PALMITOYLATED CELLULAR PROTEINS BY IMMUNOBLOTTING OR IN-GEL FLUORESCENCE

BASIC PROTOCOL 1

This protocol enables researchers to monitor biochemically the myristoylation and palmitoylation of cellular proteins. It involves metabolic labeling of cells with ω-alkynyl fatty acids and click chemistry followed by immunoblotting or in-gel fluorescence. The expected outcome will reveal the status of fatty acylation of cellular proteins.

Materials Cell line of interest: Examples include: Raw 264.7 macrophages (ATCC #CCL-2278) MDCK epithelial cells (ATCC #CCL-34) PC-3 cells (ATCC #CRL-1435) Mouse L-cells (ATCC #CRL-2648) HeLa cells (ATCC #CCL-2) Jurkat T cells (ATCC #TIB-152) COS-7 cells (ATCC #CRL-1651) Appropriate cell culture growth medium (e.g., DMEM, F-12K, RPMI) Fatty acid stock solution (see recipe) Bovine serum albumin (BSA; fatty acid-free, Sigma-Aldrich) Dimethyl sulfoxide (DMSO) Ethanol Phosphate-buffered saline (PBS; see recipe) Lysis buffer (see recipe) BCA protein assay kit (Thermo Scientific) Biotin-azide or rhodamine-azide (see recipe) Tris (2-carboxyethyl)phosphine hydrochloride (TCEP; Sigma-Aldrich) Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA; Sigma-Aldrich) Copper sulfate (CuSO4 ; Sigma-Aldrich) Acetone, ice-cold Novex Tris glycine SDS sample buffer (2×) (Invitrogen) NuPAGE sample reducing agent (10×) (Invitrogen) Tris-glycine gels, precast (Invitrogen) PBS-T (see recipe) Nonfat dried milk (any grocery store) Streptavidin-horseradish peroxidase (Invitrogen) Hydroxylamine solution, 50% in water (NH2 OH; Sigma-Aldrich) Restore western blot stripping buffer (Thermo Scientific) Anti-β-tubulin HRP antibody (Invitrogen) 37◦ C, 5% CO2 humidified incubator 6-well plates Sonicator (Branson) Cell scraper (25 cm, Starstedt) Centrifuge (Eppendorf) Nanocep centrifugal ultrafiltration devices (Pall Corporation) Vortexer (VWR) Thermomixer heating block (Eppendorf) ECL immunoblotting detection kit (GE Healthcare) Amersham Hyperfilm ECL (GE Healthcare) Desktop scanner ImageJ or AdobePhotoshop Typhoon scanner



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Additional reagents and equipment for transferring the proteins onto a nitrocellulose membrane (Gallagher et al., 2008) Culture the cells 1. Grow the preferred cells in suitable growth medium in a 5% CO2 humidified chamber at 37◦ C for at least 24 hr. Examples of cell lines in which this method has been validated include: Raw 264.7 macrophages (culture in high-glucose DMEM supplemented with 10% FBS and 2 mM GlutaMax); PC-3 cells (culture in F-12K medium supplemented with 10% FBS); Jurkat (culture in RPMI supplemented with 10% FBS); MDCK, L-cells, HeLa, and COS-7 (culture in DMEM supplemented with 10% FBS). Typically, ∼8 × 105 cells per sample are needed for an experiment.

Seed the cells 2. Seed the cells with growth medium onto 6-well plates at a density of 8 × 105 cells/well. 3. Incubate for 24 hr in a 5% CO2 humidified chamber at 37◦ C before treating with fatty acids.

Prepare the fatty acid probe-containing medium 4. Dissolve the fatty acid stock solution in serum-free growth medium supplemented with 5% BSA (fatty acid-free) to a final concentration of 100 μM just prior to the experiment. As a control, dissolve DMSO to 100 μM. Perform all dissolutions in a tissue culture hood to keep the medium sterile. 5. Sonicate the solutions in closed plastic tubes in a water bath for 15 min at room temperature and then incubate for 15 min at room temperature. Wipe the tube with ethanol to minimize contamination before transferring it back to the tissue culture hood.

Treat the cells with fatty acid probes 6. Aspirate the growth medium from the seeded cells that have been growing for 24 hr. 7. Wash the cells once with 2 ml PBS. 8. Add 2 ml of fatty acid probe-containing medium per well. 9. Incubate for 24 hr at 37◦ C, 5% CO2 .

Prepare the cell lysates 10. Remove the medium and wash the cells three times, each time with 1 ml cold PBS. 11. Add 400 μl lysis buffer, scrape the cells with a cell scraper, collect the lysate in a 1.5-ml sterile plastic tube, and rock at speed 2 for 1 hr at 4◦ C. 12. Centrifuge cell lysates 10 min at 16,000 × g, 4◦ C, and collect the supernatant. 13. Concentrate the supernatant by centrifuging 20 min at 16,000 × g, 4◦ C, with Nanocep centrifugal ultrafiltration devices. This will yield a final proteome concentration of ∼2 mg/ml, and this may vary depending on cell type and confluency.


14. Measure the protein concentration by BCA protein assay kit following the manufacturer’s protocol. At this point, cell lysates can be snap-frozen in liquid nitrogen and stored for up to 4 months at −80◦ C without detectable changes in their integrity.

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Label the cell lysates with biotin-azide or rhodamine-azide These volumes are based on a 25-μl reaction scale. 15. Add 23 μl of cell lysate at ∼2 mg/ml into a 1.5-ml plastic tube. Protein concentration in the range 1 to 2 mg/ml would work as long as 40 to 50 μg of total cell lysate is included in the reaction.

16. Add 0.5 μl of 5 mM biotin-azide to a final concentration of 0.1 mM. 17. Add 0.5 μl of 50 mM TCEP to a final concentration of 1 mM. 18. Add 0.5 μl of 10 mM TBTA to a final concentration of 0.2 mM. 19. Vortex for 5 sec at a high setting. 20. Add 0.5 μl of 50 mM CuSO4 to a final concentration of 1 mM. 21. Vortex for 5 sec at a high setting. 22. Incubate the click reaction cocktail (steps 15 to 21) for 1 hr at room temperature in dark. The order of addition of the reagents to the protein extract is important and must be performed as described above. TCEP is a reducing agent, TBTA acts as a stabilizing ligand for Cu(I), and Cu(II)SO4 is a catalyst for the click reaction.

Precipitate the proteins and prepare the samples for gel loading 23. Add 250 μl of ice-cold acetone. 24. Vortex for 5 sec (at a high setting) and incubate for 2 hr or overnight at –20◦ C. Recovery is about the same for the two incubation periods; the overnight incubation is for convenience.

25. Centrifuge 10 min at 16,000 × g, 4◦ C. There should be a visible pellet.

26. Aspirate the supernatant. 27. Air dry the tube for 5 to 10 min. 28. Resuspend the pellet in 16 μl lysis buffer. 29. Add 15.5 μl of 2× SDS sample buffer and 3.5 μl of 10× sample reducing agent. 30. Heat the sample for 5 min at 95◦ C in a heating block. 31. Load 35 μl/lane on a 10-well gel (∼30 to 40 μg protein/lane). See Critical Parameters for suggestions regarding controls.

Detect signal by immunoblotting 32. Resolve biotin-labeled protein lysates by SDS-PAGE using 4% to 20% Tris-glycine or 4% to 12% Bis-Tris gels. 33. Transfer proteins onto a nitrocellulose membrane (e.g., Gallagher et al., 2008). 34. Block the membrane with 5 ml PBS-T and 5 ml 5% nonfat dried milk for 2 hr at room temperature or overnight at 4◦ C. 35. Wash the membrane three times with PBS-T, 5 min each time. 36. Incubate the membrane with streptavidin-linked horseradish peroxidase (1:5000 in PBS-T) for 1 hr at room temperature.



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37. Wash the membrane three times, each time with 5 ml PBS-T, 10 min each. 38. Develop signal using ECL detection kit according to manufacturer’s protocol. Capture signal on film with 1- to 2-min exposure time.

Check for protein incorporation of fatty acid probes via amide or thioester linkages This protocol involves the use of hydroxylamine and is carried out right after protein transfer (step 33 above). 39. Incubate membranes for 65 to 72 hr at room temperature with 5 ml PBS-T and 5 ml 5% NH2 OH. 40. Block the membrane with 5 ml 5% nonfat dried milk in PBS-T for 2 hr at room temperature or overnight at 4◦ C. 41. Analyze by streptavidin blot as described above (steps 35 to 38).

Demonstrate equal levels of protein loading This can be carried out either after step 38 or 41.There are several ways to check for levels of protein loading which include: Coomassie staining (total protein levels), anti-actin, anti-tubulin, etc. Below is a procedure for detecting tubulin. 42. Incubate the streptavidin blots with Restore western blot stripping buffer for 15 min at room temperature. 43. Repeat steps 34 to 35. 44. Incubate the membrane with anti-β-tubulin HRP antibody (1:10,000 in PBS-T) for 1 hr at room temperature. 45. Repeat steps 37 to 38.

B SO

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-tubulin

Figure 2 Biochemical detection of fatty-acylated proteins. Cell lysates were prepared from MDCK cells treated with different ω-alkynyl fatty acids (100 μM) for 24 hr. The proteome was subjected to a click reaction with biotin-azide and then separated by SDS-PAGE. The membrane was soaked in PBS-T buffer (A) or in hydroxylamine (B), and the signal was detected by streptavidin-linked horseradish peroxidase (see Basic Protocol 1). Lanes: 1, Alk-C10 (n = 6); 2, Alk-C11 (n = 7); 3, Alk-C13 (n = 9); 4, Alk-C14 (n = 10); 5, Alk-C16 (n = 12); 6, Alk-C18 (n = 14). In (A), asterisks indicate specific bands labeled by the various probes. Arrows indicate nonspecific bands labeled in both nontreated (DMSO) and treated lanes. Alk-C14 yields more labeled bands than Alk-C13.

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Obtain images 46. Scan films using a desktop scanner and import images into ImageJ or AdobePhotoshop. See Figure 2 for typical staining with Alk-C13, Alk-C14, Alk-C16, Alk-C18, and other fatty acid probes.

Detect signal by in-gel fluorescence In cases where fluorescence scanners are available, use of rhodamine-azide is recommended as it results in minimal background and high signal-to-noise ratio relative to biotin-azide. 47. Follow steps 15 to 32 above, except replace biotin-azide with rhodamine-azide in step 16. Minimize exposure of the gel to light while carrying out the protocol. 48. Scan gel on a Typhoon scanner using the Cy3/rhodamine fluorescence channel. 49. Export and process the image in the Typhoon scanner image-processing software.

CELLULAR IMAGING OF MYRISTOYLATED AND PLAMITOYLATED PROTEINS

BASIC PROTOCOL 2

This protocol enables imaging of the global cellular myristoylated and palmitoylated proteome by fluorescence microscopy. It allows the user to capture a view of the spatial localization and dynamics of myristoylated and palmitoylated proteins under different conditions and cellular states.

Materials Phosphate-buffered saline (PBS; see recipe) Methanol (Sigma-Aldrich), prechilled PBS/0.1% (v/v) Triton X-100 Rhodamine-azide (see recipe) Tris (2-carboxyethyl)phosphine hydrochloride (TCEP; see recipe) Copper sulfate (CuSO4 ; see recipe) Hoechst 33342 (Invitrogen, cat. no. H21492) 12-well tissue culture plates Glass coverslips, washed in ethanol and left to dry for 10 min under UV light in a tissue culture hood before use 37◦ C, 5% CO2 incubator Microscope slides Fluorescence microscope and a 40× or 63× objective Standard image analysis software: e.g., Slidebook 5.0 (Intelligent Imaging Innovation) or ImageJ (NIH) Culture the cells 1. Perform step 1 in Basic Protocol 1. Seed the cells 2. Seed the cells onto 12-well plates (4 × 105 cells/well) containing ethanol-washed and dried glass coverslips and incubate for 24 hr before treatment in a 37◦ C, 5% CO2 humidified incubator. Prepare the fatty acid probe-containing medium 3. Perform steps 4 and 5 in Basic Protocol 1. Treat the cells with fatty acid probes 4. Aspirate the growth medium from wells containing seeded cells.



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5. Wash the cells gently once with 1 ml PBS. 6. Add 1 ml of fatty acid probe–containing medium per well and incubate for 4 hr at 37◦ C, 5% CO2 .

Fix and permeabilize the cells 7. Wash the cells four times, each time with 1 ml PBS to remove excess probe. 8. Fix the cells with 1 ml prechilled methanol at −20◦ C for 10 min. 9. Aspirate the methanol. 10. Permeabilize the cells with 1 ml PBS/0.1% (v/v) Triton X-100 for 5 min at room temperature. 11. Wash the cells extensively six times, each time with 1 ml PBS.

Label the cells with rhodamine-azide 12. Remove the coverslips from the wells and place them in a humidified chamber for subsequent steps. 13. Prepare 50 μl of click reaction cocktail as follows: a. Add 47 μl of PBS into a 1.5-ml plastic tube. b. Add 1 μl of 5 mM rhodamine-azide to a final concentration of 0.1 mM. c. Add 1 μl of 50 mM TCEP to a final concentration of 1 mM. d. Add 1 μl of 50 mM CuSO4 to a final concentration of 1 mM.

nuclei

fatty-acid labeling

merge

DMSO

Alk-C14

Alk-C16

Alk-C18

10 m


Figure 3 Cellular imaging of myristoylated and palmitoylated proteins. Prostate cancer cells were treated with fatty acid probes (100 μM) for 3 hr. After fixation, the cells were processed with rhodamine-azide and imaged by epifluorescence microscopy using a 40× objective (see Basic Protocol 2). Images show the perinuclear and punctuate distribution of cellular proteins labeled with Alk-C14, Alk-C16, and Alk-C18.

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14. Add the click reaction cocktail (step 13 above) to the coverslips and incubate for 1 hr in the dark at room temperature in the humidified chamber. 15. Rinse the cells extensively six times, each time with 100 μl PBS. 16. Incubate the cells with 50 μl of Hoechst 33342 (1:10,000 in PBS) for 10 min at room temperature. 17. Wash the cells three times, each time with 100 μl PBS. 18. Mount the coverslips onto the microscope glass slides.

Capture images by fluorescence microscopy 19. Capture images by using a fluorescence microscope and a 40× or 63× objective. We recommend an inverted Zeiss AX10 microscope equipped with a CoolSnap CCD camera (Roper Scientific), a Piezo stage for obtaining optical Z-sections, and standard excitation filters for Hoechst/DAPI (377 nm) and rhodamine (568 nm).

20. Acquire 50 to 70 optical Z-sections per image with 0.3-μm spacing.

Analyze the images 21. Export and process the images in a standard image analysis software, such as Slidebook 5.0 (Intelligent Imaging Innovation) or ImageJ (NIH). See Figure 3 for typical staining with Alk-C14, Alk-C16, and Alk-C18. REAGENTS AND SOLUTIONS Use Milli-Q purified water or equivalent in all recipes and protocol steps.

Biotin-azide or rhodamine-azide Reagents can be obtained from our laboratory, synthesized as described earlier (Lewis et al., 2004; Speers and Cravatt, 2004; Hsu et al., 2007; Hannoush and Arenas-Ramirez, 2009), or purchased from Invitrogen. Dissolve powder in dimethyl sulofoxide (DMSO) to a concentration of 5 mM. Divide into 50-μl aliquots and store up to 1 year at −20◦ C. Protect from light.

CuSO4 Dissolve powder in phosphate-buffered saline (PBS; see recipe) to a concentration of 50 mM just prior to use in the click reaction (see Critical Parameters). Prepare fresh. Fatty acid stock solution Dissolve lyophilized fatty acid powder [obtained from our laboratory or synthesized as described earlier in Hannoush and Arenas-Ramirez (2009)] in dimethyl sulfoxide (DMSO) to a concentration of 50 mM. Divide the solution into 50-μl aliquots and store indefinitely at −80◦ C. Alk-C13 and Alk-C18 can be obtained from Otava and Sigma-Aldrich, respectively.

Lysis buffer 100 mM sodium phosphate, pH 7.5 150 mM NaCl 1% (v/v) nonidet P-40 Store up to 3 months at 4◦ C Immediately before the experiment, add fresh protease and phosphatase inhibitor cocktail (Pierce)



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PBS-T Phosphate-buffered saline (PBS; see recipe) 0.1% (v/v) Tween-20 Store up to 6 months at room temperature Phosphate-buffered saline (PBS) 8 g NaCl 0.2 g KCl 1.13 g Na2 HPO4 (sodium phosphate, dibasic anhydrous) 0.2 g KH2 PO4 (potassium phosphate, monobasic) 1 liter of H2 O Adjust pH to 7.2 ± 0.1 with 6 N HCl Store up to 6 months at room temperature Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA) Dissolve powder in DMSO/t-butanol (1:4, v:v) solution to a concentration of 10 mM. Divide the solution into 100-μl aliquots and store up to 1 year at −20◦ C.

Tris (2-carboxyethyl)phosphine hydrochloride (TCEP) Dissolve powder in water to a concentration of 50 mM just prior to use in the click reaction (see Critical Parameters).

COMMENTARY Background Information


A number of techniques have been employed to monitor cellular fatty acylation (Hannoush and Sun, 2010). Radioactivity has been the standard method for detecting myristoylation and palmitoylation of cellular proteins. Typically, [3 H]- or [125 I]-labeled myristic and palmitic acids are added to cells for metabolic incorporation into cellular proteins (Schlesinger et al., 1980; Peseckis et al., 1993). The signal is developed by autoradiography and requires lengthy film exposures. Because of the hazards and costs associated with radioactivity, this method is less than ideal for use. Recently, the acyl-biotin exchange method was introduced for detecting protein palmitoylation (Drisdel and Green, 2004). This method relies on blocking the free sulfhydryl groups of palmitoylated proteins, followed by cleavage of the palmitate moiety and subsequent tagging of the unmasked cysteine with biotin for affinity capture and enrichment. While this method is a breakthrough, it has limitations primarily associated with false positives (Roth et al., 2006), and it only detects S-palmitoylation but not myristoylation. For a detailed description of the various methods and their limitations, the reader is referred to a comprehensive review on this topic (Hannoush and Sun, 2010). Recently, nonradioactive probes based on alkyne fatty acids have been developed for

metabolic labeling of myristoylated and palmitoylated proteins (Hannoush and ArenasRamirez, 2009). We reasoned that appending an alkyne group to the terminal end of a fatty acid would not interfere with the hydrophobic nature of the fatty acid and the mechanism by which it inserts into lipid membranes. The alkynyl fatty acid probes are portable and can be stored indefinitely in the freezer for immediate use. They are metabolically incorporated into fatty-acylated proteins in human and mouse cell lines. The tagged proteins are then conjugated to biotin-azide or rhodamine azide via a click reaction and detected by immunoblotting or in-gel fluorescence, respectively. Additionally, these probes enable cellular imaging of the global subcellular distribution of fatty-acylated proteins and shed light on the dynamics and turnover of such proteins (Hannoush and Arenas-Ramirez, 2009). Based on hydroxylamine sensitivity, Alk-C14 is a probe for protein myristoylation while Alk-C16 and Alk-C18 are probes for protein palmitoylation. Other emerging applications of these probes include in vitro labeling of recombinant proteins, monitoring turnover of palmitoylated proteins, enrichment of trace proteins, proteomics studies, and immunoprecipitation of specific proteins of interest for profiling their fatty acylation status (Heal et al., 2008; Martin and Cravatt, 2009; Hannoush and Sun, 2010; Yap et al., 2010). However,

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this method has some limitations. The toxicity of the copper reagent makes it inappropriate for monitoring fatty acylation by live-cell imaging. Furthermore, the presence of certain endogenous proteins that chelate copper may interfere with the signal of the assay, and this varies depending on the particular cellular system being studied.

Critical Parameters Reagent preparation and concentrations Avoid multiple freeze-thaw cycles of the fatty acid probe stocks as this may affect reagent quality. Also, copper sulfate and TCEP reagents should be freshly prepared just prior to use in each experiment. Because the optimal concentration of ωalkynyl fatty acid needed to obtain the best signal-to-noise ratio may vary depending on the particular cell type used, it is recommended to first test the individual probe at various increasing concentrations (recommended starting range is 1 to 200 μM) and for different times of incubation to determine the optimal conditions for their cellular system. Experimental controls Two protein gels should be run simultaneously for each set of samples. One gel is run and analyzed as described in steps 31 to 38 (Basic Protocol 1), while the other gel is treated with hydroxylamine and analyzed as described in steps 39 to 41 (Basic Protocol 1). This allows the user to determine the type of linkage via which fatty acid probes incorporate into cellular proteins. Sometimes, detection by streptavidin-HRP results in varying degrees of background staining, which stems from streptavidin labeling of endogenously biotinylated proteins. Therefore, care should be taken to decipher nonspecifically labeled bands on the gel, and it is critical that the researcher include negative controls, such as those lacking copper sulfate or the fatty acid probes. Furthermore, for imaging by fluorescence microscopy, it is critical to ensure that the signal observed is not due to nonspecific incorporation of the fatty acid probe into biological membranes. Therefore, optimizing the time for methanol fixation and the permeabilization step for the particular cell type used is crucial to avoid nonspecific incorporation into lipid bilayers. The intensity of staining will decrease with longer detergent extraction times if there is nonspecific incorporation of the probes.

Troubleshooting While the protocols described herein detail a robust method for detecting protein myristoylation and palmitoylation, in very few cases a signal may not be observed and below are recommended troubleshooting procedures. Reagent quality and concentration Fatty acid probes were not used at the appropriate concentrations to obtain the best signal-to-noise ratio. Also, the click chemistry reaction works best only when the reagents are freshly prepared just prior to the experiment. Signal optimization Sometimes, staining is not homogeneous throughout the membrane blot. This may be due to uneven transfer of proteins from the gel. In this case, ensuring that the appropriate transfer procedure is used is critical. Also, if the intensity of bands on the membrane is high, then reducing exposure times or using a more diluted stock of streptavidin-HRP can resolve this issue. Another source of reduction or loss in signal is proteins that may interfere with the click reaction, such as those that may chelate copper. These proteins may exist in a particular cellular system, and steps need to be taken to eliminate them if feasible.

Anticipated Results Basic Protocol 1 will generate gel scans of myristoylated and palmitoylated proteins (Fig. 2). This enables researchers to analyze the myristoylation or palmitoylation status of specific proteins of interest when combined with standard immunoprecipitation protocols on cells that have been metabolically labeled with Alk-C14, Alk-C16, or Alk-C18. In Basic Protocol 2, a cellular view of the distribution of myristoylated and palmitoylated proteins will be obtained (Fig. 3). The labeling method described here is very robust and the experimental results are typically consistent and reproducible across multiple runs.

Time Considerations Once the cells are seeded in plates and all the necessary reagents are in hand, the procedures described above can be completed within 2 to 3 days.

Acknowledgement I would like to thank Natalia Arenas, JingLucy Sun, and other members of the Early Discovery Biochemistry Department who contributed directly or indirectly to this project.



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Literature Cited Drisdel, R.C. and Green, W.N. 2004. Labeling and quantifying sites of protein palmitoylation. Biotechniques 36:276-285. Gallagher, S., Winston, S.E., Fuller, S.A., and Hurrell, J.G. 2008. Immunoblotting and Immunodetection. Curr. Protoc. Mol. Biol. 83:10.8.110.8.28. Hannoush, R.N. and Arenas-Ramirez, N. 2009. Imaging the lipidome: Omega-alkynyl fatty acids for detection and cellular visualization of lipid-modified proteins. ACS Chem. Biol. 4:581587. Hannoush, R.N. and Sun, J. 2010. The chemical toolbox for monitoring protein fatty acylation and prenylation. Nat. Chem. Biol. 6:498-506. Heal, W.P., Wickramasinghe, S.R., Bowyer, P.W., Holder, A.A., Smith, D.F., Leatherbarrow, R.J., and Tate, E.W. 2008. Site-specific N-terminal labelling of proteins in vitro and in vivo using Nmyristoyl transferase and bioorthogonal ligation chemistry. Chem. Commun. (Camb) 4:480-482. Hsu, T.L., Hanson, S.R., Kishikawa, K., Wang, S.K., Sawa, M., and Wong, C.H. 2007. Alkynyl sugar analogs for the labeling and visualization of glycoconjugates in cells. Proc. Natl. Acad. Sci. U.S.A. 104:2614-2619. Lewis, W.G., Magallon, F.G., Fokin, V.V., and Finn, M.G. 2004. Discovery and characterization of catalysts for azide−alkyne cycloaddition by fluorescence quenching. J. Am. Chem. Soc. 126:9152-9153. Linder, M.E. and Deschenes, R.J. 2007. Palmitoylation: Policing protein stability and traffic. Nat. Rev. Mol. Cell Biol. 8:74-84. Martin, B.R. and Cravatt, B.F. 2009. Large-scale profiling of protein palmitoylation in mammalian cells. Nat. Methods 6:135-138. Peseckis, S.M., Deichaite, I., and Resh, M.D. 1993. Iodinated fatty acids as probes for myristate pro-

cessing and function. Incorporation into pp60vsrc. J. Biol. Chem. 268:5107-5114. Resh, M.D. 2006. Trafficking and signaling by fatty-acylated and prenylated proteins. Nat. Chem. Biol. 2:584-590. Rostovtsev, V.V., Green, L.G., Fokin, V.V., and Sharpless, K.B. 2002. A stepwise Huisgen cycloaddition process: Copper(I)-catalyzed regioselective “ligation” of azides and terminal alkynes. Angew. Chem. Int. Ed. Engl. 41:25962599. Roth, A.F., Wan, J., Green, W.N., Yates, J.R., and Davis, N.G. 2006. Proteomic identification of palmitoylated proteins. Methods 40:135142. Schlesinger, M.J., Magee, A.I., and Schmidt, M.F.G. 1980. Fatty acid acylation of proteins in cultured cells. J. Biol. Chem. 255:1002110024. Speers, A.E. and Cravatt, B.F. 2004. Profiling enzyme activities in vivo using click chemistry methods. Chem. Biol. 11:535-546. Tornoe, C.W., Christensen, C., and Meldal, M. 2002. Peptidotriazoles on solid phase: [1,2,3]Triazoles by regiospecific copper(i)-catalyzed 1,3-dipolar cycloadditions of terminal alkynes to azides. J. Org. Chem. 67:3057-3064. Wang, Q., Chan, T.R., Hilgraf, R., Fokin, V.V., Sharpless, K.B., and Finn, M.G. 2003. Bioconjugation by copper(I)-catalyzed azide-alkyne [3 + 2] cycloaddition. J. Am. Chem. Soc. 125:3192-3193. Yap, M.C., Kostiuk, M.A., Martin, D.D.O., Perinpanayagam, M.A., Hak, P.C., Siddam, A., Majjigapu, J.R., Rajaiah, G., Keller, B.O., Prescher, J.A., Wu, P., Bertozzi, C.R., Falck, J.R., and Berthiaume, L.G. 2010. Rapid and selective detection of fatty acylated proteins using omega-alkynyl-fatty acids and click chemistry. J. Lipid Res. 51:1566-1580.


26 Volume 3


Imaging of Endogenous RNA Using Genetically Encoded Probes Takeaki Ozawa1 and Yoshio Umezawa2 1

Department of Chemistry, Graduate School of Science, The University of Tokyo, Tokyo, and Japan Science and Technology Agency, Tokyo, Japan 2 Research Institute of Pharmaceutical Sciences, Musashino University, Tokyo, Japan

ABSTRACT Imaging of RNAs in single cells revealed their localized transcription and specific function. Such information cannot be obtained from bulk measurements. This unit contains a protocol of an imaging method capable of visualizing endogenous RNAs bound to genetically encoded fluorescent probes in single living cells. The protocol includes methods of design and construction of the probes, their characterization, and imaging a target RNA in living cells. The methods for RNA imaging are generally applicable to many kinds of RNAs and may allow for elucidating novel functions of localized RNAs and C 2011 understanding their dynamics in living cells. Curr. Protoc. Chem. Biol. 3:27-37 by John Wiley & Sons, Inc. Keywords: RNA r imaging r GFP r fluorescence r molecular beacon

INTRODUCTION Because genome sequences of many species have been documented completely over the past decade, elucidating the respective functions of gene products, such as RNAs, is extremely important. Numerous data related to RNAs and their functions are now available. Nevertheless, the information typically represents an average of a population of cells, which sometimes obscures the actual characteristics and behavior of RNAs. Consequently, the continued development of new tools and methods to probe RNAs is vital for additional elucidation of their localization and dynamics. This challenge necessitates single-cell-based analyses using fluorescent probes and microscopes. Fluorescence imaging technologies using green fluorescent protein (GFP) and spectral variants of GFP have been widely used to visualize molecules with high spatial and temporal resolution in live cells. Among the technologies, the split-GFP complementation and reconstitution methodologies have been most commonly applied toward elucidation of protein-protein interactions in living cells (Ozawa et al., 2000, 2003; Hu and Kerppola, 2003), wherein reassembly of conditional GFP fragments is facilitated by the direct interactions of two proteins. Based on this concept of GFP fragments reassembly, we have recently developed genetically encoded fluorescent probes to visualize a target mRNA in live cells (Ozawa et al., 2007). The RNA probes consist of split fragments of enhanced GFP (EGFP) or yellow fluorescent protein (Venus) (Nagai et al., 2002), each of which is connected with a sequence-specific RNA binding domain of PUMILIO (Pumilio homology domain; PUM-HD) (Fig. 1A). PUMILIO was discovered in D. melanogaster as a protein that regulates mRNA expression by binding to specific sequences in the 3’untranslated region (UTR) of target mRNAs. The PUMILIO family of proteins has been identified in many species including C. elegans, S. cerevisiae, and H. sapiens. The crystal structure of H. sapiens PUM-HD (HsPUM1-HD) in complex with its RNA revealed that it comprises eight sequence repeats (named from R1 to R8), which recognize a consensus sequence 5 -UGUAUAUA-3 (Wang et al., 2002). Each RNA base is recognized by three

Current Protocols in Chemical Biology 3: 27-37, February 2011 Published online February 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470559277.ch100152 C 2011 John Wiley & Sons, Inc. Copyright

Imaging RNA Using Genetically Encoded Probes

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A C-terminal R8 mRNA

R7

N-terminal

R6 R5 R1

R4 R2

R3

B PUM mutant 2 (mPUM2)

PUM mutant 1 (mPUM1)

reconstituted EGFP (fluorescence)

mRNA

C-EGFP

N-EGFP ND6 mRNA (186–210) : 5 - AA UGAUGGUU GUCUU UGGAUAUA CU -3 binding proteins: mPUM1 mPUM2

C PUM1 repeat wild PUM-HD

RNA sequence

R8 Q1126 Y1123 N1122

R7 E1083 N1080 S1079

R6 Q1047 Y1044 N1043

R5 Q1031 R1008 C1007

R4 Q975 H972

R3 Q939 R936 C935

R2 Q903 Y900 N899

R1 Q867 R864 S863

5 -U1

G2

U3

A4

U5

A6

U7

A8-3

Figure 1 (A) Structure of the human PUM-HD complexed with RNA. The helical repeats are shown alternately blue and yellow, which are labeled as repeat 1(R1) to repeat 8(R8). Each repeat recognizes a specific base of RNA. (B) Basic principle of the RNA probes. Two RNA-binding domains of PUM are engineered to recognize specific sequences on a target mRNA (mPUM1 and mPUM2). In the presence of the target mRNA, mPUM1 and mPUM2 bind to their target sequences, bringing together the N- and C-terminal fragments of EGFP, resulting in functional reconstitution of the fluorescent protein. (C) RNA sequences of PUM-HD for RNA. The amino acids that interact with RNA bases are shown. The amino acids in square frames are necessary for stacking between upper and lower RNA bases. The other amino acids are for hydrogen bonds or van der Waals interactions.


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conserved amino acids at specific positions in the second of three α helices in each repeat of HsPUM1-HD. Two amino acid side chains make hydrogen bonds or van der Waals interactions like the Watson-Crick edge of an RNA base, and the third amino acid side chain forms a stacking interaction with aromatic rings of the RNA bases. The recognition sequences can be tailored based on an eight-base sequence of target RNAs (Cheong and Hall, 2006); the construct is determined simply by changing the two residues in a repeat that interact with the Watson-Crick edge of the base. There is no need to change the third side chain used for stacking interactions. However, contributions of the 8-base repeat to the binding affinity are not equivalent (Wang et al., 2002). Although there is no reported systematic analysis of the affinity of PUM-HD-RNA complex, see Cheong and Hall (2006) for additional information on the design of PUM-HD mutants. The applicability of PUM-HD to technology was first demonstrated for imaging mitochondrial mRNA, specifically NADH dehydrogenase 6 (ND6) mRNA. The HsPUM1-HD (hereafter named PUM1) was designed to match the RNA sequence of ND6, which was connected with EGFP fragments (Fig. 1B). Upon interacting with RNA, PUM1 brings the fragments of EGFP into an orientation and proximity in which they are sufficiently close for the fragments to associate and fluoresce. Monitoring the fluorescence signals allows spatial and temporal analysis of mRNA localization in individual living cells. The basic concept has recently been applied for imaging of viral RNA genomes in plants (Tilsner et al., 2009). PUM1, fused to either the N-terminal or the C-terminal halves of split mCitrine, was engineered to recognize two closely adjacent eight-nucleotide sequences in the genomic RNA of tobacco mosaic virus (TMV). They demonstrated that most RNAs were sequestered in characteristic inclusion bodies known as viral replication complexes (VRCs). The PUM1-based approach can be adopted for various RNAs as structural understanding of the PUM-HD–RNA interaction increases in the future. Described herein is the basic experimental protocol of PUM1-based imaging of mitochondrial mRNA.

STRATEGIC PLANNING Design of Mutant PUM for Specific RNA Recognition The specificity and selectivity of engineered PUM1 to RNA sequences have been examined in vitro (Wang et al., 2002). PUM1 comprises eight tandem repeats (Fig. 1A). The RNA runs antiparallel to the protein such that nucleotides from U1 to A8 (5 U1 G2 U3 A4 U5 A6 U7 A8 -3 of PUM1) are recognized individually by the repeats from R8 to R1, respectively. Each repeat recognizes a single RNA base through three conserved side chains of amino acids; two chains form hydrogen bonds or van der Waals interactions with the Watson-Crick edge of an RNA base, and a third side chain stacks with the same base and/or the preceding base (Fig. 1C). Construction of a mutant PUM1 requires only a change of the two residues in a repeat; there is no need to change the third side chain used for stacking interactions. The two amino acid residues that recognize an RNA base are named B1 and B5 from the N-terminal end (e.g., B5 Q867 and B1 R864 interact with A8). When two amino acids in a repeat of PUM1 are mutated to match an RNA base (X), the following combination (X: B1, B5) is recommended: (U: Asn, Gln), (G: Ser, Glu) and (A: Cys or Ser, Gln). No information is currently available on how to mutate B1 and B5 to match the RNA base T. The contributions of the eight repeats to RNA binding affinity are not equivalent. The most important factor is the consensus sequences of PUM1 beginning with 5 -U1 G2 U3 -3 , which recognize R8-R6. The specificity is sensitive to mutations and it is therefore recommended that the consensus sequence not be changed, if possible. The R5 recognizes a purine base (A or G). The repeats from R4 to R1 bind to the sequence, 5 -U5A6U7A8-3 .



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Recognition of each repeat to the base is also sensitive to mutations. In addition to the general rules, many empirical data have been reported (Cheong and Hall, 2006; Miller et al., 2008; Lu et al., 2009). The empirical data will be useful for the design of PUM1 mutants. BASIC PROTOCOL 1

CHARACTERIZATION OF RNA PROBES Before acquiring images of a target RNA with the RNA probes, it is important to examine whether the mutated PUM binds directly to the RNA in the cells of interest. For this purpose, it is recommended to use immunoprecipitation (Bonifacino et al., 2001) and reverse transcription PCR (RT-PCR; Beverly, 2001). In addition, it is recommended to examine the binding affinity of the engineered PUM to a target sequence using an electrophoretic mobility shift assay (gel-shift assay; Buratowski and Chodosh, 2001) to confirm the specificity of the RNA sequence recognition. Representative methods of immunoprecipitation and RT-PCR are described below. The experimental procedure is illustrated in Figure 2.

cultured cells transfect plasmids: GN-mPUM1, VN-mPUM1, mPUM2-GC, mPUM2-VC incubate for 48 hr cells expressing the probes lyse the cells with lysis buffer A centrifuge 3 min at 15,000

precipitate (discarded)

g

supernatant add anti-flag antibody or anti-GFP antibody incubate for 1 hr at 4°C on a rotator mix ProteinSepharose 4FF beads incubate for 1 hr at 4°C on a rotator wash the beads four times with lysis buffer B add 25 to 50 l of 2

loading buffer

boil at 95° to 100°C for 5 min centrifuge 3 min at 15,000

precipitate (discarded)

g

supernatant (including RNA) convert the extracted RNA into cDNA supernatant (including cDNA) run PCR using the cDNA as a template


PCR products (the size analysis)

Figure 2

Flow chart of RNA-probe characterization.

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BamHI

Not I

Xho I

EcoRI

GN-mPUM1 MTS

BamHI

EGFP-N (1 – 157 a.a.)

Not I

mPUM1

Xho I

FLAG

EcoRI

VN-mPUM1 MTS

Venus-N (1 – 154 a.a.)

XbaI

Not I

mPUM1

Xho I

FLAG

XbaI

mPUM2-GC MTSX3

XbaI

mPUM2

Not I

EGFP-C (158 – 238 a.a.)

Xho I

XbaI

mPUM2-VC MTSX3

BamHI

mPUM2

Venus-C (155 – 238 a.a.)

Not I

Xho I

MTS-DsRed-Ex MTS

DsRed-Express (full length)

Figure 3 Constructs of the plasmids. FLAG, FLAG epitope; MTS, matrix-targeting signal derived from subunit VIII of cytochrome C oxidase. The cDNA is inserted into an expression vector.

Materials HeLa cells DMEM with 10% FBS Plasmids: GN-mPUM1, VN-mPUM1, mPUM2-GC, mPUM2-VC, and MTS-DsRed-Ex, cloned into a mammalian expression vector, pcDNA3.1 (+) (Invitrogen) (Fig. 3) Lipofectamine 2000 (Invitrogen) Lysis buffer A (see recipe) Mouse monoclonal anti-Flag antibody (Sigma) or mouse monoclonal anti-GFP antibody (Roche) Protein Sepharose 4FF beads (GE healthcare) Lysis buffer B (see recipe) 2× loading buffer (see recipe) cDNA synthesis kit (Invitrogen) Primers: ND6F (5 -ATGATGTATGCTTTGTTTCT-3 ) and ND6R (5 -CCTATTCCCCCGAGCAATCT-3 ) for mitochondrial ND6 mRNA, and ND1F (5 -ATACCCATGGCCAACCTCCT-3 ) and ND1R (5 -TTAGGTTTGAGGGGGAATGC-3 ) for controls 6-well plates 37◦ C cell culture incubator 1.5-ml tubes Rotator UV transilluminator Additional reagents and equipment for agarose gel electrophoresis (Voytas, 2001)



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NOTE: The plasmids and more information on their cDNA sequences and enzyme sites may be obtained upon request from the authors’ laboratory (e-mail: [email protected]). 1. Seed 1.0 × 106 HeLa cells in 6-well plates and culture the cells in DMEM with 10% FBS at 37◦ C for 24 hr. 2. When the cells are 90% confluent, transfect the cells with plasmids GN-mPUM1, VN-mPUM1, mPUM2-GC, or mPUM2-VC using Lipofectamine 2000 according to the manufacturer’s instructions. 3. Incubate the cells 48 hr at 37◦ C. 4. Lyse the cells by adding 1 ml of ice-cold lysis buffer A. Shake the plates gently for 3 min on ice and then collect the lysates in a 1.5-ml tube. 5. Centrifuge the cells 3 min at 15,000 × g, 4◦ C, and collect the supernatant. 6. Add anti-flag antibody (1/1000 v/v) or anti-GFP antibody (1/2000 v/v) into the solution and incubate for 1 hr at 4◦ C with a rotator. 7. To absorb the immunoprecipitates, add 10 μl of protein Sepharose 4FF beads to the solution, and then incubate for 1 hr at 4◦ C. 8. Wash the beads four times, each time with 1 ml lysis buffer B at 4◦ C. 9. To denature the protein and mRNA and separate them from the beads, add 25 to 50 μl of 2× loading buffer and boil 5 min at 95◦ C. 10. Centrifuge the solution 3 min at 15,000 × g, 4◦ C, and collect the supernatant. 11. Convert the RNAs in the supernatant into cDNA using a cDNA synthesis kit (Invitrogen) according to the manufacturer’s protocol. 12. Perform PCR using the 1 μl of cDNA as a template. Select a pair of the following primers, ND6F (5 -ATGATGTATGCTTTGTTTCT-3 ) and ND6R (5 -CCTATTCCCCCGAGCAATCT-3 ) for mitochondrial ND6 mRNA, and ND1F (5 ATACCCATGGCCAACCTCCT-3 ) and ND1R (5 -TTAGGTTTGAGGGGGAATGC-3 ) as a control experiment. In the control experiments, run PCR using cDNAs prepared from total RNA in the HeLa cells as a template. Use the following PCR conditions: 30 cycles: 30 sec 30 sec 4 min

98◦ C (denaturation) 58◦ C (annealing) 72◦ C (extension).

13. Load 10 μl of the PCR products and molecular weight marker (x) onto a 2% agarose gel, which includes ethidium bromide. Electrophorese for 10 min at 100 V (Voytas, 2001). 14. Transfer the gel onto a UV transilluminator. Measure the size of the amplified DNA at 525 bp and 957 bp, which corresponds to the ND6 mRNA and ND1 mRNA, respectively. Confirm that amplified DNAs at both 525 bp and 957 bp are obtained from the total RNAs of HeLa cells, whereas only amplified DNA at 525 bp is obtained from the immunoprecipitated RNAs. BASIC PROTOCOL 2

IMAGING ENDOGENOUS RNA USING GENETICALLY ENGINEERED FLUORESCENT PROBES Described below is a procedure for imaging mitochondrial RNA using HeLa cells. The procedure can be adapted for many other cell types, including plant cells, by replacing the transfection method with a conventional one.

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A

B

Mito Tracker

C

ND6 mRNA

D

DAPI

E

ND6 mRNA Mito Tracker

ND6 mRNA DAPI

Figure 4 Fluorescence images of HeLa cells expressing GN-mPUM1 and mPUM2-GC stained with MitoTracker and DAPI: (A) Localization of mitochondria, (B) reconstituted EGFP, and (C) mtDNA. Panels D and E show their merged images. Bar, 5 μm. The insets are enlarged images of the boxed region of (A) (bar, 1.2 μm). White arrows indicate colocalization of mtDNA and ND6 mRNA.

Materials HeLa cells DMEM with 10% FBS Plasmids: GN-mPUM1, mPUM2-GC, mPUM2-VC, and MTS-DsRed-Ex, cloned into a mammalian expression vector, pcDNA3.1 (+) (Invitrogen) (Fig. 3) Lipofectamine 2000 (Invitrogen) DAPI (Invitrogen) MitoTracker (Invitrogen) HBSS (Sigma) containing 5% FBS (Invitrogen) 10-cm culture dishes 37◦ C cell culture incubator 3.5-cm glass-bottom dish Inverted fluorescence microscope, IX71 (Olympus), equipped with 100×, 1.40-NA oil objective, a 100-W mercury arc lamp for illumination and 50-W xenon lamp for bleaching with a double lamp-house system EM-CCD camera (iXon, ANDOR Technology) to acquire cell images MetaMorph software



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NOTE: The plasmids and more information on their cDNA sequences and enzyme sites may be obtained upon request from the authors’ laboratory (e-mail: [email protected]). 1. Seed HeLa cells (1 × 106 cells) onto a 10-cm dish 24 hr before transfection, and incubate the cells in DMEM with 5% FBS at 37◦ C. 2. Mix three plasmids to yield a 2.5-μl solution of 1 μl GN-mPUM1, 1 μl mPUM2-GC, and 0.1 μl MTS-DsRed-Ex. 3. Transfect the three-plasmid mixture into the HeLa cells using Lipofectamine 2000 according to the manufacturer’s instructions. 4. Incubate the cells for 12 hr at 37◦ C and then seed onto a 3.5-cm glass-bottom dish. Incubate the cells in DMEM with 5% FBS for an additional 12 hr at 37◦ C. 5. To visualize DNAs and mitochondria, add 10 μM DAPI and 1 μM MitoTracker in the culture medium. Incubate the cells 5 min at 37◦ C. 6. Aspirate the medium and replace with 1 ml HBSS containing 5% FBS. 7. Place the glass-bottom dish containing the HeLa cells on an inverted fluorescence microscope equipped with a mercury arc lamp and the 100×, 1.40 numerical aperture (NA) oil objective. Adjust the excitation filter, emission filter, and a dichroic mirror as needed. Set the cooling temperature of EM-CCD camera at –50◦ C. 8. Acquire images with the CCD camera and analyze using MetaMorph software. Typical results are shown in Figure 4.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Loading buffer, 2× 10% 2-mercaptoethanol 4% (w/v) SDS 250 mM Tris·Cl, pH 6.8 Store up to 6 months at 4◦ C Lysis buffer A 100 mM NaCl 1mM EDTA 10 mM NaF 2 mM sodium orthovanadate 1 mM phenylmethylsulfonyl fluoride (PMSF; add PMSF right before use) 10 μg/ml pepstatin 10 μg/ml leupeptin 10 μg/ml aprotinin 0.1% Triton X-100 50 mM Tris·Cl, pH 7.4 Store up to 1 month at 4◦ C Lysis buffer B


10% (w/v) SDS 250 mM Tris·Cl, pH 6.8 Store up to 6 months at 4◦ C

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COMMENTARY Background Information The RNA imaging method using PUMbased probes described here has many advantages over other methods. The PUM-based probes are genetically encoded; therefore, the cDNA encoding the probes can be introduced into living cells or inserted into chromosomal genes without damage. In addition, the principle is based on complementation of fluorescent protein fragments, which are able to emit fluorescence only in the presence of a target RNA. High-contrast fluorescence images of localized RNA can be acquired because of low background fluorescence. In addition, the method accommodates probes designed to match the sequence of target mRNAs by simple amino acid mutations. The probes currently most commonly used are nucleic acid probes, termed molecular beacons (Tyagi and Kramer, 1996). The molecular beacons are hairpin-shaped DNAs with an internally quenched fluorophore whose fluorescence is restored when they bind to a target nucleic acid sequence. The specificity for a target RNA is defined by the complementary sequence of the nucleotides of the probes, typically using ∼15 nucleotides. Therefore, the specificity is relatively high and the design of the molecule is quite simple. However, microinjection techniques are often required for inserting the probes into living cells, which causes significant damage to the cells. It is also difficult to target the beacons to a distinct intracellular organelle. Another method for RNA imaging is the use of GFP-tagged MS2 protein (Bertrand et al., 1998). A drawback of this approach is that the expressed probes emit luminescence. Because of the high background fluorescence, it is often difficult to distinguish GFP-tagged protein bound to mRNA from unbound tagged protein. In addition, the approach requires an injection of target artificial RNAs, indicating that it is impossible to visualize endogenous RNAs in living cells. From this comparison of methods, it can be concluded that the PUMbased probes are generally and widely applicable for imaging RNAs in living cells (Tyagi, 2007).

Critical Parameters and Troubleshooting Validation of imaging data is important for confirming RNA localization in living cells. Structures of fluorescent proteins reconstituted from complementary fragments are very stable

and their fragments cannot be dissociated spontaneously. The reconstituted proteins remain fluorescent even after degradation of RNA so the fluorescent proteins themselves do not indicate precise location of target RNA. To overcome this issue, it is recommend to use VN-mPUM1 and mPUM2-VC, which encode split fragments of the Venus protein. Venus emits fluorescence as strong as EGFP and is suitable for bleaching experiments. In order to visualize only the Venus that makes a complex with RNA, the fluorescence recovery after photobleaching (FRAP) technique is used. The fluorescence of Venus is bleached by irradiation with light at 480 nm, and then fluorescence images are acquired every few minutes. Fluorescence of Venus is recovered gradually by reconstitution of the Venus fragments complexed with target RNA. If the time for data acquisition is faster than the rate of RNA decomposition, the localization of reconstituted Venus indicates accurate localization of RNA in living cells. In one study, a signal sequence of subunit VIII of cytochrome c oxidase (MSVLTPLLLRGLTGSARRLPVPRAKIHSL) was used to target probes into mitochondria (Hegeman et al., 1995). GN–mPUM1 localized exclusively to the mitochondrial matrix, whereas three repeats of MTS were required for targeting mPUM2-GC; use of only one or two MTS sequences failed to localize mPUM2-GC to the mitochondrial matrix. Short linker amino acids (AAA) were used to connect the signal sequences to the probes. Other signal sequences may be used for targeting the probes to the nucleus or chloroplasts (Emanuelsson and von Heijne, 2001). When the signal sequence is used, it is then necessary to verify localization of the probes inside the cells. The length of the linker between the split fragments of EGFP and mutant PUM-HD is variable. There is no data available on the effect of the linker length on efficiency of fluorescence recovery. When an expected fluorescence intensity is not obtained, probes with different linker lengths should be constructed and FRAP behavior examined. In the design of the probes, the exact distance between the two probes may not be critical for efficient complementation of fluorescent proteins. In the case of mitochondrial ND6 mRNA, the RNA sequence that bridges mPUM1 and mPUM2 is five bases (GUCUU). For imaging plant virus, seven bases (AGUUUUU) were used for bridging the mutant



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PUM-HDs. Because a single strand of RNA is flexible, it is likely that the RNA sequence between the two PUM-HD recognition sequences does not affect the binding of RNA to PUM-HD.

Anticipated Results Localization of ND6 mRNA in mitochondria is confirmed by staining mitochondria with MitoTracker Red (CMXRos) and mtDNA with DAPI (Fig. 4). The fluorescence intensity of reconstituted EGFP (Fig. 4B) is very weak owing to the low amounts of mtRNA; therefore, the exposure time of the CCD camera may be set to >300 msec. In contrast, each image of mitochondria (Fig. 4A) and mtDNA (Fig. 4C) required 95% free of contaminating structures. This can be accomplished by starting the process with a purified, structurally defined glycan or

40,000 20,000 0

blood group H antibody (10 ␮g/ml) a2

a2

b4 a3 b3

b4 a3

20,000 blood group B antibody (10 ␮g/ml) 15,000 RFU

RFU

60,000

b4

5,000 0

1 4 7 10 13 16 19 22 25 Glycan number

30,000 25,000

a3 b4 a3

SLex antibody 2 (50 ␮g/ml)

15,000 10,000

4,000 0

1 4 7 10 13 16 19 22 25 Glycan number

20,000

12,000

RFU

RFU

20,000 SLex antibody 1 (50 ␮g/ml) 16,000

8,000

a3 a2 b4 a3

10,000

a3 b4 a3

5,000 1 4 7 10 13 16 19 22 25 Glycan number open-ring Glc

Gal

GlcNAc

0

1 4 7 10 13 16 19 22 25 Glycan number closed-ring Fuc

Neu5Ac

Figure 3 Comparison of the binding of four different antibodies to open-ring and closed-ring AEAB conjugates on a glycan array. The arrays were printed using a piezo printer (Perkin-Elmer) with open- and closed-ring AEAB derivatives at 300 μM on NHS-derivatized slides. Antibodies were applied to the glycan array at the concentrations indicated in the figure, and detected with appropriate fluorescently labeled secondary antibodies (Song et al., 2009b). The x axis represents different glycans on the array by number, and the y axis represents the relative fluorescence units (RFU) detected on the microarray.

Derivatizing Glycans on the Microscale


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oligosaccharides are not compatible with either carbograph SPE or PGC HPLC, presumably due to extremely strong retention. The capacity of carbograph SPE columns is ∼100 mg carbon/1 mg glycan.

Anticipated Results The purified closed-ring and open-ring conjugates can be treated as any other synthetic material with a primary alkyl amine group. Their reactivity toward both epoxy- and NHSderivatized surfaces is high. Although the printing, assay, and data analysis of glycan microarray is not within the scope of this article, Figure 3 shows several histograms that demonstrate the comparison of four antibodies binding to a glycan microarray printed with 26 different open-ring glycan-AEAB derivatives and 26 corresponding closed-ring glycanAEAB derivatives (Song et al., 2009b). The structures of the bound glycans are indicated using symbols in each panel. In the case of the human anti-H and anti-B blood group antibodies, the binding is apparently directed more toward the nonreducing end of the oligosaccharide, as binding is observed with both the natural closed-ring and open-ring structure. However, in the case of the two anti-sialylLewis x (SLex) antibodies, the open-ring GlcNAc of the AEAB derivative destroys the glycan epitope. Thus, if the binding motif of a GBP is at the nonreducing end of the glycan, which is generally the case, both types of conjugates show similar binding. However, if the binding motif is close to or includes the reducing end, the open-ring AEAB conjugates may not be appropriate for binding assays.

Time Considerations With the available reagents, the procedure for preparing 1 to 20 glycans can be completed in 3 to 4 days. This includes scheduling the freeze-drying overnight between steps. The procedure can be shortened significantly if the drying steps are concluded more quickly. In addition, multiple parallel syntheses can be carried out at the same time, which will greatly increase the efficiency. Testing the glycan array with GBPs, although not covered in this article, takes 3 to 4 hr, depending on the number of samples and number of steps for detecting the GBPs (Smith et al., 2010). Derivatizing Glycans on the Microscale

Literature Cited Alvarez, R.A. and Blixt, O. 2006. Identification of ligand specificities for glycan-binding pro-

teins using glycan arrays. Methods Enzymol. 415:292-310. Bigge, J.C., Patel, T.P., Bruce, J.A., Goulding, P.N., Charles, S.M., and Parekh, R.B. 1995. Nonselective and efficient fluorescent labeling of glycans using 2-amino benzamide and anthranilic acid. Anal. Biochem. 230:229-238. Blixt, O., Head, S., Mondala, T., Scanlan, C., Huflejt, M.E., Alvarez, R., Bryan, M.C., Fazio, F., Calarese, D., Stevens, J., Razi, N., Stevens, D.J., Skehel, J.J., van Die, I., Burton, D.R., Wilson, I.A., Cummings, R., Bovin, N., Wong, C.H., and Paulson, J.C. 2004. Printed covalent glycan array for ligand profiling of diverse glycan binding proteins. Proc. Natl. Acad. Sci. U.S.A. 101:17033-17038. Cummings, R.D. 2009. The repertoire of glycan determinants in the human glycome. Mol. Biosyst. 5:1087-1104. de Paz, J.L. and Seeberger, P.H. 2006. Recent advances in carbohydrate microarrays. QSAR Combin. Sci. 25:1027-1032. Dubois, M., Gilles, K., Hamilton, J.K., Rebers, P.A., and Smith, F. 1951. A colorimetric method for the determination of sugars. Nature 168:167. Feizi, T. and Chai, W. 2004. Oligosaccharide microarrays to decipher the glyco code. Nat. Rev. Mol. Cell Biol. 5:582-588. Horlacher, T. and Seeberger, P.H. 2008. Carbohydrate arrays as tools for research and diagnostics. Chem. Soc. Rev. 37:1414-1422. Luyai, A., Lasanajak, Y., Smith, D.F., Cummings, R.D., and Song, X. 2009. Facile preparation of fluorescent neoglycoproteins using pnitrophenyl anthranilate as a heterobifunctional linker. Bioconjug. Chem. 20:1618-1624. Manger, I.D., Rademacher, T.W., and Dwek, R.A. 1992. 1-N-glycyl beta-oligosaccharide derivatives as stable intermediates for the formation of glycoconjugate probes. Biochemistry 31:1072410732. Paulson, J.C., Blixt, O., and Collins, B.E. 2006. Sweet spots in functional glycomics. Nat. Chem. Biol. 2:238-248. Smith, D.F., Song, X., and Cummings, R.D. 2010. Use of glycan microarrays to explore specificity of glycan-binding proteins. Methods Enzymol. 480:417-444. Song, X., Lasanajak, Y., Rivera-Marrero, C., Luyai, A., Willard, M., Smith, D.F., and Cummings, R.D. 2009a. Generation of a natural glycan microarray using 9-fluorenylmethyl chloroformate (FmocCl) as a cleavable fluorescent tag. Anal. Biochem. 395:151-160. Song, X., Lasanajak, Y., Xia, B., Smith, D.F., and Cummings, R.D. 2009b. Fluorescent glycosylamides produced by microscale derivatization of free glycans for natural glycan microarrays. ACS Chem. Biol. 4:741-750. Song, X., Xia, B., Stowell, S.R., Lasanajak, Y., Smith, D.F., and Cummings, R.D. 2009c. Novel fluorescent glycan microarray strategy reveals ligands for galectins. Chem. Biol. 16:36-47.

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Song, X., Lasanajak, Y., Xia, B., HeimburgMolinaro, J., Rhea, J.M., Ju, H., Zhao, C., Molinaro, R.J., Cummings, R.D., and Smith, D.F. 2011. Shotgun glycomics: A microarray strategy for functional glycomics. Nat. Methods 8:85-90. Stevens, J., Blixt, O., Paulson, J.C., and Wilson, I.A. 2006. Glycan microarray technologies: Tools to survey host specificity of influenza viruses. Nat. Rev. Microbiol. 4:857-864. Stowell, S.R., Arthur, C.M., Dias-Baruffi, M., Rodrigues, L.C., Gourdine, J.P., HeimburgMolinaro, J., Ju, T., Molinaro, R.J., RiveraMarrero, C., Xia, B., Smith, D.F., and Cummings, R.D. 2010. Innate immune lectins kill bacteria expressing blood group antigen. Nat. Med. 16:295-301. Xia, B., Kawar, Z.S., Ju, T., Alvarez, R.A., Sachdev, G.P., and Cummings, R.D. 2005. Versatile fluorescent derivatization of glycans for glycomic analysis. Nat. Methods 2:845-850.

Derivatizing Glycans on the Microscale


Volume 3

Visualization and Identification of Fatty Acylated Proteins Using Chemical Reporters Jacob S. Yount,1 Mingzi M. Zhang,1 and Howard C. Hang1 1

The Rockefeller University, New York, New York

ABSTRACT Protein fatty acylation, the covalent addition of a lipid chain at specific amino acids, changes the inherent hydrophobicity of a protein, often targeting it to cellular membrane compartments and regulating protein activity, stability, and interactions. Fatty acylation can be analyzed using chemical reporters that mimic natural lipids and contain bioorthogonal chemical handles, allowing them to be reacted with detection tags such as fluorophores or affinity tags. Our laboratory has used alkynyl-chemical reporters of protein myristoylation, S-palmitoylation, prenylation, and acetylation to provide robust, nonradioactive methods for examining the acylation states of full cellular proteomes and individual proteins of interest by (1) metabolically incorporating these chemical reporters into proteins in living cells; (2) selectively reacting the labeled proteins in cell lysates with azido-rhodamine via click chemistry, and globally visualizing them with fluorescence gel scanning; (3) analyzing protein acylation on individual candidate proteins using immunoprecipitation, click chemistry, and fluorescence gel scanning; and (4) identifying novel fatty acylated proteins by reacting chemical reporter–labeled proteins with azido-biotin via click chemistry and selective retrieval using streptavidin beads. This is particularly valuable for the examining protein S-palmitoylation, which does not occur in readily predicted consensus amino acid motifs. Curr. Protoc. Chem. C 2011 by John Wiley & Sons, Inc. Biol. 3:65-79 Keywords: fatty acylation r S-palmitoylation r click chemistry

INTRODUCTION Protein fatty acylation is the covalent addition of a lipid chain at specific amino acids. This modification changes the inherent hydrophobicity of a protein, often targeting it to cellular membrane compartments (Linder and Deschenes, 2007). Acylation may also regulate protein activity, stability, and protein-protein interactions (Linder and Deschenes, 2007). Fatty acylation can be analyzed using chemical reporters that mimic natural lipids and contain bioorthogonal chemical handles that allow them to be reacted with secondary detection tags such as fluorophores or affinity tags (Charron et al., 2009b). For example, our laboratory has successfully utilized alkynyl-chemical reporters of protein myristoylation (Charron et al., 2009b), S-palmitoylation (Charron et al., 2009b; Yount et al., 2010; Zhang et al., 2010; Fig. 1), prenylation (Fig. 1; Charron et al., 2010), and acetylation (Yang et al., 2010a). Basic Protocol 1 describes metabolic incorporation of these chemical reporters onto proteins in living cells. Basic Protocol 2 describes the global visualization of reporter-labeled proteins by selectively reacting alkyne-containing chemical reporter–labeled proteins in cell lysates with azido-rhodamine via click chemistry and fluorescence gel scanning. Basic Protocol 3 describes analysis of protein acylation on individual candidate proteins using immunoprecipitation, click chemistry, and fluorescence gel scanning. Finally, Basic Protocol 4 allows identification of novel fatty acylated proteins by reacting chemical reporter–labeled proteins with azido-biotin via click chemistry and selective retrieval using streptavidin beads. This may be particularly Current Protocols in Chemical Biology 3: 65-79, May 2011 Published online May 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470559277.ch100225 C 2011 John Wiley & Sons, Inc. Copyright

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A

O O

S

H N

N H

COOH

H2N

N H

O

N-myristoylation

B

H2N

COOH

N H

O

S-palmitoylation

H N

COOH

O

S-prenylation

O

O OH

O

OH

alk-12

C

S

H N

alk-16

OH

alk-FOH

O N3

5 N

O S

N O

Et2N

HN O Cl

NEt2 ⫺

O N H

O

O

O

N H

NH

N

N

N3

HO

O

azido-azo-biotin

azido-rhodamine

Figure 1 Chemical tools for studying protein fatty acylation. (A) Chemical structures for protein myristoylation of Nterminal glycines, palmitoylation of cysteine residues, and prenylation of C-terminal CaaX motifs (cysteine, two aliphatic amino acids, and a variable amino acid). (B) Alkyne chemical reporters used for studying protein myristoylation (alk-12), palmitoylation (alk-16), and prenylation (alk-FOH). (C) Secondary detection reagents for visualization (azido-rhodamine) or affinity enrichment (azido-azo-biotin) of alkyne chemical reporter–labeled proteins.

valuable for the examination of S-palmitoylomes in different cell types or activation states, as these modifications do not occur on readily predicted consensus amino acid motifs. Overall, these techniques provide robust, nonradioactive methods for examining the acylation states of full cellular proteomes and individual proteins of interest (Fig. 2). BASIC PROTOCOL 1

METABOLIC INCORPORATION OF CHEMICAL REPORTERS OF PROTEIN FATTY ACYLATION IN LIVING CELLS Cellular growth medium is supplemented with alkyne-bearing chemical reporters of fatty acylation, allowing cellular uptake and metabolic incorporation of the reporters onto proteins at sites of acylation. NOTE: The volume of cell culture that should be grown depends upon the type of cell used, amount of protein of interest produced by the cells, and type of analysis being performed. These parameters should be determined before beginning labeling studies. The numbers of HeLa cells required for the analyses described in Basic Protocols 2, 3, and 4 are given in those protocols.

Materials


Cultured cells of interest, growing in appropriate complete cell culture medium Serum-free cell culture medium appropriate for growing the cell type of interest, supplemented with 2% (v/v) charcoal/dextran-stripped fetal bovine serum (FBS; e.g., Invitrogen), 37◦ C DMSO (USB grade) 50 mM alkyne-fatty acid chemical reporter stock in DMSO, e.g., Alk-12 (Charron et al., 2009b; for studying myristoylation) Alk-16 (Sigma-Aldrich cat no. O8382; also see Charron et al., 2009b; for studying palmitoylation) Alk-FOH (Charron et al., 2010; for studying prenylation)

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visualize alk-labeled protein (Basic Protocol 3)

alk-16: ⫺

⫹

⫺

⫹

rho -N3 fluorescence immunoblot

immunoprecipitation

click chemistry alkyne-labeled protein

alk-16: ⫺

⫹

⫺

⫹

visualize alk-labeled proteome (Basic Protocol 2) rho -N3

cell

chemical reporter labeling (Basic Protocol 1)

click chemistry alkyne-labeled cell lysate

fluorescence coomassie

alk-16: ⫺

⫹

biotin -N3

1) click chemistry

biotin N

N

2) SM removal 3) Streptavidin beads

N

Na2S2O4 elution

identify alk-labeled proteins (Basic Protocol 4)

coomassie

proteomics

Figure 2 Workflow in protocols used for studying protein fatty acylation. Basic Protocol 1 describes labeling of proteins in cells with alkyne chemical reporters of fatty acylation. Basic Protocols 2 to 4 utilize the labeled cells for visualization of labeled proteomes by bioorthogonal ligation with azido-rhodamine (rho, Basic Protocol 2), visualization of individual candidate proteins using immunoprecipitation and bioorthogonal ligation with azido-rhodamine (Basic Protocol 3), and/or affinity enrichment of labeled proteins by bioorthogonal ligation with azido-azo-biotin (biotin) and selective elution from streptavidin beads (Basic Protocol 4). Data shown for Basic Protocols 2 and 4 is from DC2.4 cells labeled with 50 μM alk-16 for 1 hr. Data shown for Basic Protocol 3 is from HeLa cells transfected with HA-tagged IFITM3 protein and labeled with 50 μM alk-16 for 1 hr, followed by immunoprecipitation, click chemistry, and immunoblotting with anti-HA antibodies. Abbrevation: SM, small molecule.

Phosphate-buffered saline (PBS; see recipe), ice-cold Liquid nitrogen or dry-ice/ethanol bath 37◦ C, 5% CO2 cell culture incubator Cell scrapers Refrigerated centrifuge 1. Replace the serum-containing complete cell culture medium with 37◦ C, serum-free cell culture medium, supplemented with 2% charcoal/dextran-filtered FBS, and either DMSO as a solvent control or 20 to 100 μM (final concentration) alkyne-fatty acid chemical reporter. We label our HeLa cells when they are confluent. For other studies, cells may need to be labeled at other stages in the cell cycle when the proteins of interest are being produced. Medium at 37◦ C allows solubilization of fatty acid chemical reporters. Charcoal/dextran-filtered FBS removes lipids present in serum, allowing cellular uptake of lipid chemical reporters without competition from serum lipids.



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Fatty acid chemical reporters are typically effective and show minimal toxicity when used at concentrations of 20 to100 μM. A concentration of 20 μM can be used as a starting point for most experiments and can be increased if enhanced signal strength is desired.

2. Incubate the cells with chemical reporters 1 hr at 37◦ C. Longer incubation times may be necessary to label protein sites with minimal turnover and to label sites that are modified upon de novo protein synthesis. This should be titrated for each protein of interest, although a 1-hr labeling time is sufficient for most proteins.

3. Harvest the cells by scraping them into suspension and centrifuging 3 min at 1000 × g, 4◦ C. Use of trypsin to harvest the cells may degrade cell surface proteins.

4. Remove the supernatant and wash the cells by suspending them in a large volume of ice-cold PBS (e.g., the volume of the container with the cell pellet). Centrifuge 3 min at 1000 × g, 4◦ C. This removes the serum proteins.

5. Remove the supernatant, and repeat the wash. 6. Remove the supernatant, freeze the washed cell pellets in liquid nitrogen or in a dry ice/ethanol bath, and store up to 6 months at −80◦ C, or continue to Basic Protocol 2, 3, or 4. Cell pellets have been stored for up to 6 months without apparent loss of signal. BASIC PROTOCOL 2

GLOBAL FLUORESCENT PROFILING OF FATTY ACYLATED PROTEINS IN WHOLE-CELL LYSATES Chemical reporters mimic natural lipids but contain bioorthogonal groups allowing a selective chemical reaction for appending detection tags to the labeled proteins. Proteins labeled with alkyne-bearing chemical reporters of fatty acylation are reacted with the azide-bearing detection tag azido-rhodamine via click chemistry, allowing fluorescent visualization of protein bands after SDS-PAGE.

Materials


Chemical reporter–labeled cells (Basic Protocol 1), e.g., ∼1 × 106 HeLa cells/pellet 4% (w/v) sodium dodecyl sulfate (SDS) with EDTA-free protease inhibitors (see recipe) 250 U/μl Benzonase (Sigma-Aldrich, ultrapure) BCA assay reagents (Pierce Protein Research) 5 mM azido-rhodamine(Charron et al., 2009b) in dimethyl sulfoxide (DMSO) or tetramethylrhodamine-5-carbonyl azide (Invitrogen; see Martin and Cravatt, 2009) in DMSO 50 mM tris(2-carboxyethyl)phosphine (TCEP): prepare fresh 2 mM tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl] amine (TBTA) in 1:4 (v/v) DMSO/butanol 50 mM CuSO4 : prepare fresh Methanol, ice-cold Chloroform, ice-cold Water, ice-cold 4× loading buffer (see recipe) 2-mercaptoethanol (2-ME) 18-well 4% to 20% Tris·Cl protein gels (Bio-Rad)

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Fluorescent protein molecular weight ladder (15 to 250 kDa; Bio-Rad), diluted as necessary Fluorescence gel destaining solution: 40% (v/v) methanol/50% (v/v) acetic acid/10% (v/v) water Coomassie blue staining reagents (Pierce Protein Research) 1.5-ml microcentrifuge tubes Bath sonicator (optional) 95◦ C heating block Fluorescence gel scanner with a 532-nm excitation and 580-nm detection filters and 30-nm band-pass (e.g., Typhoon 9400, Amersham Biosciences) Additional reagents and equipment for quantifying protein concentration using BCA (Olson and Markwell, 2007), carrying out SDS-PAGE (Gallagher, 2006), and staining proteins (e.g., see Sasse and Gallagher, 2009) Lyse cells 1. Lyse chemical reporter–labeled cells (e.g., ∼1 × 106 HeLa cells) by adding 50 μl of 4% SDS buffer with EDTA-free protease inhibitors and 1 μl (250 U) of Benzonase to the cell pellet and vortexing. Lysing cells with SDS-containing buffer maximally solubilizes the proteins. Protease inhibitors containing EDTA are not compatible with Benzonase or click chemistry reactions.

2. Quantify the protein concentrations using a standard BCA assay (Olson and Markwell, 2007). Protein concentrations obtained for cell lines are generally 1 to 10 mg/ml, depending on the cell type.

3. Dispense aliquots of equal amounts of protein (∼50 μg) for each sample into 1.5-ml microcentrifuge tubes, and bring the volumes to 44.5 μl with 4% SDS buffer with EDTA-free protease inhibitors. Generally, 50 μg total protein is adequate for visualization of fatty acylation with chemical reporters. Using less protein makes precipitation more difficult in later steps.

Ligate labeled proteins to azido-rhodamine 4. Prepare a click chemistry master mix, by combining the following volumes of reagents per sample: 1 μl of 5 mM azido-rhodamine in DMSO 1 μl of 50 mM TCEP 2.5 μl of 2 mM TBTA in 1:4 (v/v) DMSO/butanol 1 μl of 50 mM CuSO4 . We synthesize the azido-rhodamine used in this step (synthesis described in Charron et al., 2009b). Tetramethyl rhodamine-5-carbonyl azide has also been used in click chemistry reactions (Martin and Cravatt, 2009) and is commercially available. Solutions of TCEP and CuSO4 should be prepared fresh for each experiment. A master mix ensures that equal amounts of each reagent are added to each sample.

5. Add 5.5 μl of the click chemistry master mix to each protein sample for a final volume of 50 μl, and vortex to mix. 6. Incubate 1 hr at room temperature. Visualization of Fatty Acylated Proteins


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Remove unreacted molecules 7. Perform a chloroform/methanol precipitation of protein to remove unreacted azidorhodamine by adding the following ice-cold reagents to each tube: 200 μl methanol 75 μl chloroform 150 μl water. Vortex, and centrifuge 15 min at 20,000 × g, at 4◦ C. Precipitation reagents should be ice-cold.

8. Remove and discard the upper aqueous phase, leaving the lower organic phase and the white layer of protein between the two layers. The upper aqueous layer should be clear, and the lower organic layer contains the pink-colored rhodamine. A white layer of protein should have formed between the two layers.

9. Add 1 ml of ice-cold methanol to each sample and mix gently, causing the protein pellet to sink to the bottom of the tube. Centrifuge 10 min at 20,000 × g, 4◦ C. The protein pellet should be a light pink color.

10. Remove all of the liquid by pipetting, being careful not to disturb the pellet. Wash the protein pellet by adding 1 ml of ice-cold methanol and inverting the tube. Centrifuge 10 min at 20,000 × g, 4◦ C. 11. Carefully remove the methanol leaving behind the protein pellet. Allow the remaining methanol to evaporate by leaving the sample tubes open on the bench for 20 min at room temperature. The dried pellet will be a gray/white color.

Resolubilize proteins for SDS-PAGE 12. Add 50 μl of 4% SDS buffer with EDTA-free protease inhibitors to dissolve the protein pellets. A bath sonicator may be used at this step to speed the solubilization process. Pellets are generally dissolved within 5 sec using a sonicator.

13. Add 17.5 μl of 4× loading buffer and 3.5 μl of 2-ME to each sample (5% 2-ME final concentration), and vortex the samples. DTT has been used successfully as an alternative to 2-mercaptoethanol.

14. Denature the proteins for 5 min at 95◦ C, using a heating block. 15. Vortex, and centrifuge 1 min at 5000 × g. room temperature. 16. Load 20 μl of samples and diluted fluorescent protein ladder onto an 18-well 4% to 20% Tris·Cl gel, and electrophorese 1 hr at 200 V. A fluorescent protein ladder should be used as a molecular weight standard. Commercially available protein ladders may require as much as a 1:10,000 dilution with 4% SDS buffer and appropriate addition of 4× loading buffer to avoid saturation and bleed-through of the standard into other lanes of the gel.

Destain gel and scan 17. Destain the gel by rocking for 1 hr at room temperature in fluoresecence gel destaining solution to remove remaining traces of loading buffer and unreacted rhodamine. Visualization of Fatty Acylated Proteins

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18. Scan the gel using a fluorescence gel scanner. For the azido-rhodamine derivative shown in Figure 1, an Amersham Biosciences Typhoon 9400 scanner has been successfully used with 532-nm excitation and 580-nm detection filters with 30-nm band-pass. For more information about azido-rhodamine visualization, see Charron et al. (2009b).

19. Stain the gel with Coomassie blue (e.g., see Sasse and Gallagher, 2009) to demonstrate equal protein loading (see Fig. 2).

FLUORESCENT VISUALIZATION OF FATTY ACYLATION ON CANDIDATE PROTEINS

BASIC PROTOCOL 3

Proteins labeled with chemical reporters of fatty acylation are immunoprecipitated prior to click chemistry ligation with azido-rhodamine and SDS-PAGE.

Materials Chemical reporter–labeled cells (Basic Protocol 1), e.g., ∼2 × 106 HeLa cells/pellet 1% (w/v) Brij97 buffer with EDTA-free protease inhibitors (see recipe) BCA assay reagents (Pierce Protein Research) 4% (w/v) SDS buffer with EDTA-free protease inhibitors (see recipe) 5 mM azido-rhodamine (Charron et al., 2009b) in DMSO or tetramethylrhodamine-5-carbonyl azide (Invitrogen; see Martin and Cravatt, 2009) in DMSO 50 mM tris(2-carboxyethyl)phosphine (TCEP): prepare fresh 2 mM tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl] amine (TBTA) in 1:4 (v/v) dimethyl sulfoxide (DMSO)/butanol 50 mM CuSO4 : prepare fresh 4× loading buffer (see recipe) 2-mercaptoethanol (2-ME) 25% (w/v) hydroxylamine (NH2 OH), pH 7.0 (optional) Fluorescent protein molecular weight ladder (10 to 250 kDa, dual color; Bio-Rad), diluted as necessary 18-well 4% to 20% Tris·Cl protein gels (Bio-Rad) Fluorescence gel destaining solution: 50% (v/v) methanol/10% (v/v) acetic acid/40% (v/v) H2 O 1.5-ml microcentrifuge tubes 95◦ C heating block Fluorescence gel scanner with a 532-nm excitation and 580 nm detection filter and 30 nm band-pass (e.g., Typhoon Amersham Biosciences 9400, scanner) Additional reagents and equipment for quantifying protein concentration using BCA (Olson and Markwell, 2007), performing immunoprecipitation techniques (Bonifacino et al., 2006), carrying out SDS-PAGE (Gallagher, 2006), staining proteins on gels (Sasse and Gallagher, 2009), and performing immunoblotting (Gallagher et al., 2008) 1. Lyse chemical reporter–labeled cells (e.g., ∼2 × 106 HeLa cells) by adding 100 μl of 1% Brij97 buffer with EDTA-free protease inhibitors to the cell pellet. Centrifuge 5 min at 1000 × g, 4◦ C, to remove cellular debris. Other lysis buffers compatible with immunoprecipitation may be substituted for Brij97 buffer. Protease inhibitors containing EDTA are not compatible with click chemistry.



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2. Quantify protein concentrations using a standard BCA assay (e.g., see Olson and Markwell, 2007). 3. Dispense 500 μg of protein for each sample into 1.5-ml microcentrifuge tubes. Bring the volumes to 100 μl with 1% Brij97 buffer with EDTA-free protease inhibitors. Increased quantities of proteins may be needed, particularly when examining proteins of low cellular abundance. Generally, 500 μg is adequate for the study of overexpressed proteins and most native proteins.

4. Carry out standard immunoprecipitation techniques for the protein of interest (e.g., see Bonifacino et al., 2006), and wash the agarose beads with the antibody-linked protein at least three times with lysis buffer. 5. Add 22.25 μl of 4% SDS buffer with EDTA-free protease inhibitors to the pelleted agarose beads used for the immunoprecipitations. The addition of 4% SDS buffer will dissociate antibody-protein complexes. Thus, proteins will no longer be associated with beads and additional washes subsequent to this step should not be performed.

6. Prepare a click chemistry master mix, by combining the following volumes of components per sample:

0.5 μl of 5 mM azido-rhodamine in DMSO 0.5 μl of 50 mM TCEP 1.25 μl of 2 mM TBTA in 1:4 (v/v) DMSO/butanol 0.5 μl of 50 mM CuSO4 . Tetramethyl rhodamine-5-carbonyl azide has also been used instead of the azidorhodamine in click chemistry reactions (Martin and Cravatt, 2009) and is commercially available.

7. Add 2.75 μl of the click chemistry reagent master mix to each sample (containing the bead pellets and the dissociated protein) for a final volume of 25 μl, and vortex to mix. 8. Incubate 1 hr at room temperature. 9. Add 9 μl of 4× loading buffer and 1.8 μl of 2-ME to each sample (5% 2-ME final concentration), and vortex. 10. Optional: Split the sample into two microcentrifuge tubes, and add 2 μl of 25% neutral NH2 OH to one tube (2.5% final concentration) and 2 μl of water to the second tube, and mix by vortexing. Continue to step 11. NH2 OH is known to cleave thioester bonds characteristic of protein palmitoylation, and thus fluorescence signal should be lost or greatly reduced for proteins that are labeled with alkyne-palmitate reporter on cysteine residues.

11. Denature the proteins 5 min at 95◦ C, using a heating block. 12. Vortex, and centrifuge 1 min at 9000 × g, room temperature, to pellet agarose beads used earlier for immunoprecipitation. 13. Load 15 μl of the samples (avoiding the pelleted beads) and a diluted fluorescent protein ladder onto duplicate 18-well 4% to 20% Tris·Cl gels, and electrophorese 1 hr at 200 V (e.g., see Gallagher, 2006). Gel 1 will be used for fluorescent gel scanning, and gel 2 will be used in immunoblotting for the protein of interest as a loading control. Visualization of Fatty Acylated Proteins

If samples were split for NH2 OH treatment, additional lysis buffer and 4× loading buffer can be added before loading onto gels. Alternatively, lower volumes can be loaded.

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A fluorescent protein ladder should be used as a molecular weight standard. Commercially available protein ladders may require as much as a 1:10,000 dilution with 4% SDS-buffer and appropriate addition of 4× loading buffer for the fluorescent gel to avoid saturation and bleed-through of the standard into other lanes of the gel.

14. To visualize fluorescence: Destain gel 1 with fluorescence gel destaining solution, and perform fluorescence gel scanning as described in Basic Protocol 2, steps 17 and 18. 15. To perform an immunoblot: Transfer proteins from gel 2 onto a nitrocellulose membrane, and probe for the protein of interest using standard immunoblotting techniques (e.g., see Gallagher et al., 2008).

PROTEOMIC IDENTIFICATION OF PROTEINS LABELED WITH CHEMICAL REPORTERS OF FATTY ACYLATION

BASIC PROTOCOL 4

Proteins labeled with alkyne-bearing chemical reporters of fatty acylation are reacted with the azide-bearing detection tag azido-azo-biotin via click chemistry, allowing selective retrieval of the labeled proteins using streptavidin beads. Na2 S2 O4 cleavage of the azo group allows specific elution of labeled proteins from beads, without boiling.

Materials Chemical reporter–labeled cells (Basic Protocol 1), e.g., ∼20 × 106 HeLa cells/pellet 4% (w/v) SDS buffer with EDTA-free protease inhibitors (see recipe) 250 U/μl Benzonase BCA assay reagents (Pierce Protein Research) 5 mM azido-azo-biotin (Yang et al., 2010b) or azide-biotin (Invitrogen; also see Martin and Cravatt, 2009) 50 mM and 200 mM tris(2-carboxyethyl)phosphine (TCEP): prepare fresh 2 mM tris[(1-benzyl-1H -1,2,3-triazol-4-yl)methyl] amine (TBTA) in 1:4 (v/v) dimethyl sulfoxide (DMSO)/butanol 50 mM CuSO4 : prepare fresh Methanol, ice-cold Chloroform, ice-cold Water, ice-cold 0.5 M EDTA 1% Brij97 buffer with EDTA-free protease inhibitors (see recipe) Streptavidin agarose beads (Thermo Scientific) PBS, ice cold, pH 7.4 (see recipe) PBS/0.2% (w/v) SDS, pH 7.4 (ice-cold) ABC buffer (250 mM ammonium bicarbonate), ice-cold 8 M urea 400 mM iodoacetamide: prepare fresh 1% (w/v) SDS Na2 S2 O4 elution buffer (see recipe) NuPAGE 4× LDS buffer (Invitrogen): dilute to 1× before use) 2-mercaptoethanol (2-ME) Fluorescent protein molecular weight ladder (15 to 250 kDa; Bio-Rad) 18-well 4% to 20% Tris·Cl protein gels (Bio-Rad) Coomassie blue staining reagents (Pierce Protein Research) 15-ml conical, polypropylene tubes Nutating mixer 2-ml dolphin microcentrifuge tubes Green Centricon devices (YM-10 membranes with 10 kDa molecular weight cutoff; Millipore)



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Rotary evaporater (e.g., SpeedVac) 95◦ C heating block Additional reagents and equipment for quantifying protein concentration using BCA (Olson and Markwell, 2007), carrying out SDS-PAGE (Gallagher, 2006), and staining gels for protein (Sasse and Gallagher, 2009) 1. Lyse chemical reporter–labeled cells (e.g., ∼20 × 106 HeLa cells) by adding 1 ml of 4% SDS buffer with EDTA-free protease inhibitors and 1 μl (250 units) of Benzonase to the cell pellet. Protease inhibitors containing EDTA are not compatible with Benzonase or click chemistry reactions.

2. Quantify protein concentrations using a standard BCA assay. 3. Dispense equal amounts (>10 mg) of protein for each sample into 15-ml conical, polypropylene tubes and bring the volume to 4.45 ml with 4% SDS buffer with EDTA-free protease inhibitors. A minimum of 10 mg total protein is recommended.

Ligate chemical reporter–labeled proteins with azido-azo-biotin 4. Prepare a click chemistry master mix by combining the following volumes of components per sample: 100 μl of 5 mM azido-azo-biotin 100 μl of 50 mM TCEP 250 μl of 2 mM TBTA in1:4 DMSO/butanol 100 μl of 50 mM CuSO4 . Solutions of TCEP and CuSO4 should be prepared fresh for each experiment. Azide-biotin is available from Invitrogen and has also been successfully used in click chemistry reactions and proteomic studies (Martin and Cravatt, 2009).

5. Add 550 μl of the click chemistry master mix to each protein sample (final reaction volume of 5 ml), and vortex to mix. 6. Incubate the click chemistry reaction 1.5 hr at room temperature. 7. Perform a methanol/chloroform precipitation of the protein as in Basic Protocol 2, steps 7 to 11, scaling up the volumes proportionately (100×), and washing the protein pellet twice with 50 ml ice cold methanol, dissociating the protein pellet by pipeting. 8. Resuspend the protein pellet in 1 ml of 4% SDS buffer with or without EDTA-free protease inhibitors, supplemented with 20 μl of 0.5 M EDTA solution. EDTA will chelate remaining copper and prevent any further click chemistry reaction.

9. Quantify the resuspended protein by BCA assay (e.g., see Olson and Markwell, 2007). Generally, 50% to 80% of proteins are recovered after precipitation.

Bind reporter-labeled and biotin-ligated proteins to streptavidin beads 10. Add 5 mg protein in 1 ml of 4% SDS buffer with EDTA-free protease inhibitors to individual 15-ml Falcon tubes. Visualization of Fatty Acylated Proteins

11. Add 2 ml of 1% Brij97 buffer with EDTA-free protease inhibitors to dilute the SDS concentration to a level compatible with biotin/streptavidin binding (i.e., less than 1.5% SDS).

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12. Wash 100 μl of streptavidin agarose beads per 5 mg protein with 50 ml PBS three times, centrifuging the beads for 2 min at 4000 × g, room temperature, to remove the supernatant after each wash. 13. Resuspend the washed streptavidin agarose beads in 100 μl of 1% Brij97 buffer with EDTA-free protease inhibitors per sample, and add 100 μl of the bead suspension to each protein sample. 14. Rock the protein/bead mixture on a nutating mixture 1 hr at room temperature.

Wash streptavidin beads 15. Wash beads by inverting several times in 10 ml PBS/0.2% SDS, and centrifuge 2 min at 4000 × g, 4◦ C. 16. Discard the supernatant, and wash the beads three times with 10 ml PBS, centrifuging 2 min at 4000 × g, 4◦ C, between washes. 17. Discard the supernatant, and wash the beads twice with 10 ml of 250 mM ABC buffer, centrifuging 2 min at 4000 × g, 4◦ C, between washes. Washing minimizes nonspecific pulldown of proteins.

Cap cysteines for mass spectrometry analysis 18. Remove the supernatant, and add the following: 500 μl 8 M urea 25 μl 200 mM TCEP 25 μl 400 mM iodoacetamide. Mix by pipetting, and incubate 30 min at room temperature. This reaction alkylates cysteines, thereby preventing formation of high molecular weight aggregates by random disulfide bonding.

19. Add 10 ml ABC buffer and centrifuge 2 min at 4000 × g, 4◦ C. 20. Remove the supernatant, resuspend the beads in 1 ml ABC buffer, and transfer the suspension to 2-ml dolphin tubes. 21. Centrifuge 2 min at 4000 × g, 4◦ C.

Elute labeled proteins from streptavidin beads 22. Remove the supernatant. add 250 μl Na2 S2 O4 elution buffer to the beads, resuspend the beads by pipetting, and incubate 30 min at room temperature. 23. Centrifuge 2 min at 4000 × g, 4◦ C, and save the supernatant containing the eluted proteins. 24. Add an additional 250 μl Na2 S2 O4 elution buffer to the beads and incubate 30 min at room temperature. 25. Centrifuge 2 min at 4000 × g, 4◦ C, collect the eluent, and combine with the previously eluted proteins.

Concentrate eluted proteins 26. Wash a green Centricon device with 500 μl of 1% SDS and centrifuge 30 min at 9000 × g, room temperature. 27. Wash the Centricon with 500 μl water and centrifuge 30 min at 9000 × g, room temperature.



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28. Add 500 μl of eluted proteins to the Centricon, and centrifuge 30 min at 9000 × g, room temperature. 29. Turn the Centricon upside down and centrifuge into a collection tube 3 min at 1000 × g, room temperature. 30. Add 50 μl of 1% SDS supplemented with 75 mM of 2-ME (5.3 μl 2-ME in 1 ml SDS solution)to the Centricon reservoir, pipet up and down several times, and centrifuge the upside-down Centricon into the same collection tube 3 min at 1000 × g, room temperature. 31. Repeat step 30. The final volume will be approximately 125 μl.

32. Lyophilize the samples to a solid white powder using a rotary evaporator.

Perform SDS-PAGE of eluted proteins 33. Resuspend the white powder in 25 μl of NuPAGE 1× LDS containing 5% (v/v) of 2-ME, and vortex vigorously. 34. Incubate the samples for 5 min on a 95◦ C heating block. 35. Centrifuge 1 min at 1000 × g, room temperature. 36. Load 20 μl protein mixture for each sample onto a 4% to 20% Tris·Cl gel. For gel slices to be cut for proteomic analysis, blank lanes should be placed between samples. This will aid in preventing contamination during gel slicing for gel-based proteomics. Blanks should contain LDS buffer with ∼20% SDS and 5% 2-mercaptoethanol. If SDS is not added to the blank lanes, the sample proteins may expand horizontally in the gel during electrophoresis. A fraction of the retrieved proteins can be saved for immunoblotting to confirm that the proteins identified by mass spectrometry (step 38) are indeed selectively retrieved.

37. Electrophorese the gel 1 hr at 200 V (e.g., see Gallagher, 2006), and stain with Coomassie blue (e.g., see Sasse and Gallagher, 2009). 38. Cut gel slices for proteomic analysis according to standard procedures (see Jiménez et al., 1998).

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Brij97 buffer (1%) with EDTA-free protease inhibitors, pH 7.4 1% (w/v) Brij97 150 mM NaCl 50 mM triethanolamine (dilute 1 M triethanolamine stock, pH 7.4) Sterilize by passing through a 0.22-μm filter, and store up to 1 year at room temperature. Just before use, add EDTA-free protease inhibitor cocktail tablets (Roche) according to the manufacturer’s directions. Loading buffer, 4×


40% (v/v) glycerol 240 mM Tris·Cl, pH 6.8 8% (w/v) sodium dodecyl sulfate (SDS) 0.04% (w/v) bromphenol blue Store up to 1 year at room temperature

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Na2 S2 O4 elution buffer 250 mM ammonium bicarbonate 0.1% (w/v) sodium dodecyl sulfate (SDS) 25 mM Na2 S2 O4 Prepare fresh for each experiment Phosphate-buffered saline (PBS), pH 7.4 144 mg/liter KH2 PO4 9 g/liter NaCl 95 mg/liter Na2 HPO4 ·7H2 O Sterilize by passing through a 0.22-μm filter, and store up to 1 year at 4◦ C SDS buffer (4%) with EDTA-free protease inhibitors, pH 7.4 4% (w/v) sodium dodecyl sulfate (SDS) 150 mM NaCl 50 mM triethanolamine (dilute 1 M triethanolamine stock, pH 7.4) Sterilize by passing through a 0.22-μm filter, and store up to 1 yr at room temperature. Just before use, add EDTA-free protease inhibitor cocktail tablets (Roche) according to the manufacturer’s directions. COMMENTARY Background Information The use of chemical reporters and bioorthogonal ligation chemistries has a rich history in the field of protein glycosylation (Prescher and Bertozzi, 2006) and has recently been extended to the study of protein myristoylation, S-palmitoylation (Charron et al., 2009b; Yount et al., 2010; Zhang et al., 2010), prenylation (Charron et al., 2010; Fig. 1A,B), and acetylation (Yang et al., 2010a). Protein acylation has classically been studied using radiolabeled lipids. However, this method is hazardous and provides low sensitivity, often requiring film exposure times of weeks to months. For example, the bacterial effector protein SifA was demonstrated to be prenylated using radiolabeled lipids in an in vitro translation system, but radiolabeled lipids did not provide the sensitivity needed to detect SifA prenylation in living cells (Reinicke et al., 2005). In contrast, study of protein acylation with alkyne-bearing chemical reporters and bioorthogonal ligation with azido-rhodamine provides a robust and sensitive way to visualize these modifications (Charron et al., 2009a). For example, SifA prenylation in living cells was recently confirmed using a prenylation reporter, allowing identification of classes of cellular transferases responsible for SifA prenylation (Charron et al., 2010). The increased sensitivity of chemical reporters is also accompanied by an increase in efficiency of experimental workflow, as a typical cellular labeling and visualization procedure can be

completed in a single workday without long exposure times. Chemical reporters coupled with biotin and selective elution of labeled proteins allows the identification of novel acylated proteins (Martin and Cravatt, 2009; Yount et al., 2010). This method has proved fruitful in the identification of new palmitoylated proteins, which has traditionally been difficult, as this modification does not occur on a predictable motif. The interferon-induced transmembrane protein 3 (IFITM3) was discovered in this way to be palmitoylated and, using subsequent labeling and visualization of IFITM3 mutants, the sites of palmitoylation were mapped (Yount et al., 2010). Palmitoylation was found to be essential for full antiviral activity of IFITM3 against influenza virus, demonstrating the unique and interesting biological findings that can be elucidated with a chemical reporter approach (Yount et al., 2010). Other methods for identifying novel palmitoylated proteins have been described e.g., acyl-biotin exchange (ABE) chemistry, which has been important in expanding our knowledge of the multitude of specific proteins on which palmitoylation can occur (Roth et al., 2006). ABE chemistry and chemical reporters afford complementary tools for analyzing S-fatty acylation. Chemical reporters of acylation are likely to be especially useful for identifying novel acylated proteins during different cellular states, e.g., differentiation, apoptosis, or cellular



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infection. For example, chemical reporters of myristoylation, which occurs predictably on N-terminal glycines, could allow identification of new myristoylation sites that are revealed upon caspase cleavage of proteins during apoptosis. Likewise, C-terminal CaaX motifs may also be identified after protein cleavage events or on uncharacterized protein isoforms with the use of chemical reporters and proteomic methods.

Critical Parameters and Troubleshooting


Cellular labeling of proteins with reporters of fatty acylation requires complete solubilization of the reporter, as precipitates are toxic to most cell types. To this end, chemical reporter should always be added to media at 37◦ C and mixed thoroughly. Washes of labeled cells should be done quickly and with ice-cold PBS to avoid turnover of some types of acylation, thus maximizing the detected lipidation signal. The choice of detergents for lysing cells is critically important to the success of the experiment, particularly when performing immunoprecipitations. While SDS-containing buffer maximally solubilizes proteins and is especially useful for acylated proteome profiling, Brij97-containing buffer also solubilizes many lipidated membrane proteins and is compatible with immunoprecipitations. When performing the click chemistry reaction, it is imperative that fresh solutions of TCEP and CuSO4 be used. TCEP may become oxidized over time, and Cu(I) is required for correct coordination of the reaction. Upon completion of SDS-PAGE, destaining the gel destined for fluorescence gel scanning greatly improves signal to noise, as residual azidorhodamine in the lanes may cause background signal. High quality reagents should be used to prepare the destain solution to avoid any contaminants that may produce a fluorescent signal. Likewise, the gel should be handled only on the edges, as physical manipulations of the gel can often appear as smudges or fingerprints in fluorescence gel scans even when gloves are worn. The fluorescence gel molecular weight standard should also be titrated so that bleedthrough into other lanes does not occur. Wiping the pipet tip before loading and starting the current immediately upon addition may also help if bleedthrough of the standard is observed. For selective retrieval of acylated proteins, protein input of 5 to 10 mg is a good starting point. Smaller amounts of protein may limit

detection to only the most abundant proteins. In the precipitation step following click chemistry with azido-azo-biotin, it is important that the protein pellet is washed twice with icecold methanol so that unreacted biotin does not block streptavidin binding sites on the beads used for pulldown of labeled proteins. Following the methanol washes, using 4% SDS buffer and sonication is an effective way to completely resolubilize the protein pellet. Because 4% SDS is not compatible with the biotinstreptavidin binding, the SDS concentration must be diluted for this step. It has also been reported that two elutions using Na2 S2 O4 are more effective than one longer elution (Verhelst et al., 2007), and improved protein recovery from the streptavidin beads is observed when performing two elutions. Concentration of the eluted protein with a Centricon is necessary for buffer exchange prior to SDS-PAGE.

Anticipated Results Using alkyne-bearing chemical reporters and azido-modified detection reagents, negligible signal should be seen in the DMSO-treated control lanes. An experiment is deemed successful if signal is detected above these control lanes for both rhodamine visualization and Coomassie staining. The magnitude of the signal can be increased or decreased as necessary by changing the concentration of chemical reporter used, changing the labeling time, or by modifying the amount of protein added to each lane of the gel. If signal is undetectable, the investigator should label Jurkat T cells with the alk-16 reporter as a positive control, as these cells are efficiently labeled with this reporter at 20 μM for 1 hr. Likewise, an immunoprecipitation for endogenous palmitoylated Lck from alk-16 labeled Jurkat cell lysate can also serve as a positive control using a mouse anti-Lck monoclonal antibody (Clone 3A5 from Invitrogen; Zhang et al., 2010). For proteomic experiments, the authors have routinely identified more than 100 selectively retrieved lipidated proteins when starting with 5 mg or more of total protein in cell lysate. When using lipidation reporters of different chain lengths, alk-12 preferentially labels myristoylated proteins while alk-16 preferentially labels palmitoylated proteins. Minimal, but observable, cross-talk has been observed for both reporters (Charron et al., 2009b; Martin and Cravatt, 2009; Wilson et al., 2010; Yount et al., 2010), likely due to substrate promiscuity of the respective acyltransferases.

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Time Considerations Basic Protocol 1 can typically be completed in 2 hr, and cell pellets can be frozen, or cells can be utilized in Basic Protocols 2 or 3 in the same day. Both protocols typically require ∼6 hr before fluorescence gel scans are obtained. Loading controls for these experiments, i.e., Coomassie gel staining and immunoblotting, respectively, can also be completed the same day. Alternatively, membranes for blotting can be blocked overnight and probed the following day. Likewise, the Coomassie stain can be applied overnight, or the destain procedure can be performed overnight at 4◦ C. Throughout Basic Protocols 2 and 3 there are at least three 1-hr incubation periods during which the experimenter can focus on other tasks. Basic Protocol 4 is a more lengthy procedure, and requires approximately 15 hr to prepare samples for proteomic analysis, although this can easily be spread over 2 days, as there are several stopping points within the protocol. There are also several 1-hr or 30-min incubation periods. If gelbased proteomics is desired, then another 2 days of sample preparation time should be planned.

Literature Cited Bonifacino, J.S., Dell’Angelica, E.C., and Springer, T.A., 2006. Immuunoprecipitation. Curr. Protoc. Neurosci. 5.24.1-5.24.28. Charron, G., Wilson, J., and Hang, H.C. 2009a. Chemical tools for understanding protein lipidation in eukaryotes. Curr. Opin. Chem. Biol. 13:382-391. Charron, G., Zhang, M.M., Yount, J.S., Wilson, J., Raghavan, A.S., Shamir, E., and Hang, H.C. 2009b. Robust fluorescent detection of protein fatty-acylation with chemical reporters. J. Am. Chem. Soc. 131:4967-4975. Charron, G., Tsou, L.K., Maguire, W., Yount, J.S., and Hang, H.C. 2010. Alkynyl-farnesol reporters for detection of protein S-prenylation in cells. Mol. Biosyst. 7:67-73. Gallagher, S.R. 2006. One-dimensional SDS gel electrophoresis of proteins. Curr. Protoc. Mol. Biol. 75:10.2A.1-10.2A.37. Gallagher, S.R, Winston, S.E., Fuller, S.A., and Hurrell, J.G.R. 2008. Immunoblotting and immunodetection. Curr. Protoc. Mol. Biol. 10.8.110.8.28. Jiménez, C.R., Huang, L., Qiu, Y., and Burlingame, A.L. 1998. In-gel digestion of proteins for

MALDI-MS fingerprint mapping. Curr. Protoc. Protein Sci. 14:16.4.1-16.4.5. Linder, M.E. and Deschenes, R.J. 2007. Palmitoylation: Policing protein stability and traffic. Nat. Rev. Mol. Cell Biol. 8:74-84. Martin, B.R. and Cravatt, B.F. 2009. Large-scale profiling of protein palmitoylation in mammalian cells. Nat. Methods 6:135-138. Olson, J.S.C. and Markwell, J. 2007. Assays for determination of protein concentration. Curr. Protoc. Protein Sci. 3.4.1-3.4.29. Prescher, J.A. and Bertozzi, C.R. 2006. Chemical technologies for probing glycans. Cell 126:851854. Reinicke, A.T., Hutchinson, J.L., Magee, A.I., Mastroeni, P., Trowsdale, J., and Kelly, A.P. 2005. A Salmonella typhimurium effector protein SifA is modified by host cell prenylation and S-acylation machinery. J. Biol. Chem. 280:14620-14627. Roth, A.F., Wan, J., Bailey, A.O., Sun, B., Kuchar, J.A., Green, W.N., Phinney, B.S., Yates, J.R., 3rd, and Davis, N.G. 2006. Global analysis of protein palmitoylation in yeast. Cell 125:10031013. Sasse, J. and Gallagher, S.R. 2009. Staining proteins in gels. Curr. Protoc. Molec. Biol. 10.6.110.6.27. Verhelst, S.H., Fonovic, M., and Bogyo, M. 2007. A mild chemically cleavable linker system for functional proteomic applications. Angew. Chem. Int. Ed. Engl. 46:1284-1286. Wilson, J.P., Raghavan , A.S., Yang, Y.Y., Charron, G., and Hang, H.C. 2010. Proteomic analysis of fatty-acylated proteins in mammalian cells with chemical reporters reveals S-acylation of histone H3 variants. Mol. Cell Proteomics 10:M110.001198. Yang, Y.Y., Ascano, J.M., and Hang, H.C. 2010a. Bioorthogonal chemical reporters for monitoring protein acetylation. J. Am. Chem. Soc. 132:3640-3641. Yang, Y.Y., Grammel, M., Raghavan, A.S., Charron, G., and Hang, H.C. 2010b. Comparative analysis of cleavable azobenzene-based affinity tags for bioorthogonal chemical proteomics. Chem. Biol. 17:1212-1222. Yount, J.S., Moltedo, B., Yang, Y.Y., Charron, G., Moran, T.M., Lopez, C.B., and Hang, H.C. 2010. Palmitoylome profiling reveals Spalmitoylation-dependent antiviral activity of IFITM3. Nat. Chem. Biol. 6:610-614. Zhang, M.M., Tsou, L.K., Charron, G., Raghavan, A.S., and Hang, H.C. 2010. Tandem fluorescence imaging of dynamic S-acylation and protein turnover. Proc. Natl. Acad. Sci. U.S.A. 107:8627-8632.



Volume 3

Miniaturized High-Throughput Fluorescent Assay for Conversion of NAD(P)H to NAD(P) Andrew D. Napper1,2 and Sharmila Sivendran1,3 1

Penn Center for Molecular Discovery, Institute for Medicine and Engineering and Department of Chemical and Biomolecular Engineering, University of Pennsylvania, Philadelphia, Pennsylvania 2 Nemours Center for Childhood Cancer Research, Wilmington, Delaware 3 GlaxoSmithKline, Collegeville, Pennsylvania

ABSTRACT This unit describes a miniaturized fluorescence assay that monitors the conversion of NADPH to NADP+ . The same assay format may also be used to measure NADH to NAD+ conversion. Examples of assay development and validation results are presented to illustrate the use of this protocol to screen an enzyme that consumes NADPH as a cofactor during conversion of substrate to a reduced product. Enzymatic assays are carried out in low-volume 384-well plates, in which the turnover of NADPH is monitored by the decrease in fluorescence emission at 460 nm between an initial measurement and a second reading after 90 min. A follow-up assay is used to rule out false-positive artifacts C arising from compounds that fluoresce at 460 nm. Curr. Protoc. Chem. Biol. 3:81-97 2011 by John Wiley & Sons, Inc. Keywords: NADPH detection r NADH detection r fluorescence r high-throughput screening r oxidoreductase r enzyme assay r fluorescent artifact

BASIC PROTOCOL We have optimized a miniaturized high-throughput screening assay that monitors the decrease in fluorescence at 460 nm upon conversion of NADPH to NADP+ . To correct for the intrinsic fluorescence at 460 nm of some test compounds, two readings of each plate are obtained: one immediately after compound and reagent addition, and the second after a 90-min incubation with the target enzyme. Nevertheless, a slight increase in compound fluorescence readings over 90 min is sufficient to give rise to a number of false positives (compounds that appear to be inhibitors of the target enzyme but in fact are not). A counterscreen is used to identify these false positives and exclude them from further study. Two assays are described here: Primary screening assay: NADPH fluorescence assay: Enzyme activity is monitored by the decrease in NADPH fluorescence over 90 min. Counterscreen assay: Fluorescence change in the absence of enzyme and substrate: Fluorescence change due to test compounds is monitored in assay buffer alone. The optimization of these assays and their use to confirm screening hits is exemplified by assay validation and HTS to discover inhibitors of two enzymes, RmlC and RmlD, essential for cell wall biosynthesis in Mycobacterium tuberculosis (Sivendran et al., 2010). The two enzymes are assayed together as a mixture: RmlC epimerizes the keto sugar nucleotide, dTDP-4-keto-6-deoxy-D-xylo-hexulose, and RmlD then uses the cofactor NADPH to reduce the epimerized keto sugar nucleotide to dTDP-rhamnose (Fig. 1).

Current Protocols in Chemical Biology 3: 81-97, June 2011 Published online June 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470559277.ch100155 C 2011 John Wiley & Sons, Inc. Copyright

HTS Fluorescent Assay for NAD(P)H to NAD(P)

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O O HO

CH3

O

HN

O OH O O P O P O O

O

TDP-KDX

OH

O RmlC

H

CH3

HN

N

O

O

O CH3

RmlD

H3C HO HO

O

O

NADPH

P O P O O

OH

O

O

N

O

O OH

H

NADP

Figure 1 RmlC/D enzyme assay. RmlC epimerizes the keto sugar nucleotide dTDP-4-keto-6-deoxy-D-xylo-hexulose (TDP-KDX), and RmlD then uses the cofactor NADPH to reduce the epimerized keto sugar nucleotide to dTDP-rhamnose. Enzyme activity is determined by monitoring the decrease in NADPH fluorescence.

To simplify and to provide a clear example, the protocol describes the method used for the Rml enzymes assay. The Critical Parameters and Troubleshooting section describes how the Rml protocol may be modified and broadened to be generally applicable to a wider range of NADPH- or NADH-utilizing enzymes.

Materials NADPH (see recipe) Assay buffer (see recipe) Target enzyme(s): For the study used as an example here, the M. tb rhamnosyl biosynthetic enzymes dTDP-6-deoxy-D-xylo-4-hexulose 3,5-epimerase (RmlC) and dTDP-6-deoxy-L-lyxo-4-hexulose reductase (RmlD) were cloned and expressed in E. coli and purified by Michael McNeil and co-workers (Ma et al., 2001); these enzymes are not commercially available Substrates: TDP-6-deoxy-D-xylo-hexopyranosid-4-ulose (TDP-KDX) was synthesized enzymatically and stored frozen at a concentration of 1 mg/ml in 50 mM MOPS buffer (Sigma), pH 7.4, at −80◦ C as previously described (Sivendran et al., 2010); this substrate is not commercially available Positive control inhibitor: thymidine diphosphate (TDP; Sigma) Analytical-grade dimethyl sulfoxide (DMSO; Fisher or VWR), anhydrous Test compounds: Store dissolved in DMSO (e.g., at 10 mM stock concentration) in 384-well polypropylene plates (room temperature storage is recommended if the compounds are to be reused within a period of less than one week; for longer term storage, compounds stocks should be frozen at 100 calculated using Eq. 2) are not meaningful in the context of bona fide enzyme inhibition. Therefore, the appearance of a dose-response curve such as the one shown in panel B immediately suggests that the compound “activity” is an artifact of its intrinsic fluorescence.

(see Eq. 2). Examples of highly fluorescent compounds (e.g., coumarins) giving percent inhibition values up to 1500% can be found in PubChem assay IDs 1532 and 1533 (see Internet Resources). Factors to consider in applying this protocol to other enzymes A significant number of compounds in screening libraries are fluorescent under the detection conditions used here (excitation 340 nm, emission 460). Therefore, it is important to follow HTS based on NAD(P)H fluorescence with the counterscreen assay.

Anticipated Results HTS Fluorescent Assay for NAD(P)H to NAD(P)

Enzyme titration and timecourse Figure 7 shows an enzyme timecourse in which the selected dilution of Rml enzymes mixture gives a linear decrease in NADPH

fluorescence over 180 min. The amount of NADPH converted to NADP+ may be calculated from the fluorescence change using the NADPH standard curve (steps 1 to 5). Km determinations Figure 3 shows Km determinations for TDPKDX and NADPH using the Rml enzymes. Note that the sigmoidal curves, as seen in this figure, are fairly unusual (see also Km determinations under Critical Parameters and Troubleshooting). Positive control inhibitor dose-response and IC50 determination Figure 8 shows a dose-response curve for TDP inhibition of RmlC/D. Nonlinear regression analysis (steps 33 to 36) gave the curve fit shown on the figure and an IC50 of 388 μM.

94 Volume 3


NADPH fluorescence

900,000 800,000 700,000 600,000 500,000 Blank Control (Rxn) assay

400,000 300,000 0

30

60

90 120 Time (min)

150

180

Figure 7 Enzyme activity timecourse. Example of dilution of Rml enzymes mixture that gives linear timecourse over 180 min. The protocol was based on that described under steps 6 to 10, Enzyme titration and timecourse. Data points represent the mean of triplicate determinations; error bars depict standard deviation. Control assay: The fluorescence of a mixture of 2.63 × 10−4 μg/μl Rml C, 6.53 × 10−4 μg/μl Rml D, 25 μM NADPH, and 200 μM TDP-KDX in 1× assay buffer was monitored over 180 min at 25◦ C. Blank: The Control mixture without TDP-KDX was used to determine the change in NADPH fluorescence in the absence of enzymatic activity. (The Rml enzymes do not convert NADPH to NADP in the absence of TDP-KDX.)

Change in NADPH fluorescence

240,000 200,000 160,000 120,000 80,000 40,000 0 10

100

1000

10,000

TDP ( M)

Figure 8 Dose-response curve for TDP inhibition of enzymes RmlC/D. Data points represent a single well at each concentration. The protocol is described under steps 25 to 36, Positive control inhibitor dose-response and IC50 determination. TDP was tested at concentrations ranging from 1 μM to 30 mM. From these data, the IC50 for TDP inhibition was determined to be 388 μM.

Quality control (QC) validation plate Figure 4 shows representative QC plate data. TDP at a concentration of 500 μM gave 66% inhibition, essentially identical to the potency predicted based on the IC50 value of 388 μM determined from the dose-response curve shown in Figure 8. Summary of RmlC/D screening results This section provides a brief summary of the outcome of the RmlC/D screening, doseresponse testing, and counterscreening. A to-

tal of 201,368 unique compounds (265,000 including duplicates) from the Molecular Libraries Small Molecule Repository (MLSMR) were screened at a concentration of 5.5 μM as mixtures of four per well in 384-well plates. A total of 2328 mixtures that showed >30% inhibition were selected as hits. (The standard deviation of the mean of the negative controls in each plate was 30% inhibition. The very low retest rate is to be expected given the likelihood that the activity of each mixture stemmed from only one compound in that mixture. Only 388 out of the 470 compounds active at 5.5 μM were available for dose-response testing; of these, 372 gave an IC50

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