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Originally published In Press as doi:10.1074/mcp.T600027-MCP200 on July 15, 2006.
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Molecular & Cellular Proteomics 5:1697-1702, 2006.
© 2006 by The American Society for Biochemistry and Molecular Biology, Inc.

Technology

The Identification of Nucleic Acid-interacting Proteins Using a Simple Proteomics-based Approach That Directly Incorporates the Electrophoretic Mobility Shift Assay *

Jonathan A. Stead, Jeff N. Keen and Kenneth J. McDowall{ddagger}

From the Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences, University of Leeds, Leeds LS2 9JT, United Kingdom


   ABSTRACT
TOP
 ABSTRACT
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Proteins that interact with nucleic acids are central to numerous cellular processes, and their continuing characterization represents one of the foremost challenges in the postgenomic era. Here we describe a simple proteomics-based approach for the identification by mass spectrometry of proteins in crude extracts that interact with nucleic acids. It incorporates the electrophoretic mobility shift assay and is based on the finding that when a protein forms a complex with nucleic acid its electrophoretic mobility is affected as well as that of the nucleic acid. Our method should greatly reduce and in some cases may even eliminate the need for extensive protein purification and as such should contribute significantly to the functional annotation of the proteome. Furthermore it requires no prior knowledge of the molecular mass, quaternary structure, or pI of the interacting protein. Proof of principle is demonstrated using a recently discovered transcription factor; however, the approach should also have application in the identification of proteins that interact with RNA.

The study of proteins that interact with nucleic acids is of the utmost importance to understand fundamental organismal level processes such as development and cellular processes including chromatin organization, transcription, RNA processing and decay, replication, recombination and repair, and the packaging of viral genomes. The most widely used method for identifying nucleic acid-binding proteins involves their extensive purification using a combination of several different chromatography steps and often incorporates as the final step some form of selection based on their affinity for a particular nucleic acid target (1, 2). Recent advances in mass spectrometry have greatly reduced the amount of polypeptide required for identification (3–5); therefore, the bottleneck to the study of nucleic acid-binding proteins is often establishing the polypeptide(s) that has the activity of interest.

A technique that has been used extensively for analyzing nucleic acid-protein interactions is the electrophoretic mobility shift assay (EMSA)1 (6–8). This method is based on the finding that the mobility of nucleic acids in non-denaturing gels during electrophoresis is retarded when complexed with protein such that the final position in a gel of a nucleic acid that is bound by protein appears shifted. Given the basis of the EMSA, it is also referred to as the gel shift or retardation assay. EMSAs can be used to resolve even labile complexes due to the "caging" effect of the gel matrix, which prevents protein that dissociates from escaping the DNA; consequently rapid reassociation is favored (6–8). Moreover the EMSA can be used to assay nucleic acid-binding activity in crude extracts and as such has been used extensively for over 2 decades to follow the activity of nucleic acid-binding proteins as they are purified (9).

The separation of a protein(s) of interest from other cellular components is important for not only biochemical and structural studies but to facilitate their identification and reverse genetic approaches that allow studies of function in vivo. Where genome sequence is available for an organism, identification of a protein is readily achieved by, for example, peptide mass fingerprinting (10). Identification not only allows studies of function in vivo, which have importance in their own right, but can assist biochemical and structural studies by facilitating the overexpression and manipulation of the corresponding gene in the natural or a heterologous host and the comparison with entries of known properties or structure in databases. Therefore, methods that allow the rapid identification of proteins that interact with nucleic acids have potential to enhance greatly the study of the many cellular processes to which they are central. One such method uses mass spectrometry to analyze directly proteins that are able to bind a solid support containing a specific nucleic acid sequence (11). The proteins that bind to the solid support are rapidly identified; however, as extensive binding of nonspecific proteins to the support occurs, this approach requires either postidentification the specificity of the interaction to be determined or preidentification at least one chromatography step combined with a series of selections using wild-type and mutant binding sites (12). Another approach that has been described recently starts with the identification of a specific interaction using an EMSA and uses steps in which the pI and mass of the interacting protein are estimated to facilitate the identification of its approximate position on a standard 2D polyacrylamide gel: candidates in this region are then eluted, renatured, and assayed for the original activity in the EMSA (13). Here we describe an equally powerful but much simpler method that also incorporates the EMSA. It does not require any knowledge of the chemical or physical properties of the interacting protein, nor does it require its renaturation after excision from gels, and it is based on the finding that when a protein forms a complex with a nucleic acid target its electrophoretic mobility (14), as well as that of the nucleic acid, is affected (6–8). Proof of principle is demonstrated using AtrA, a recently discovered transcription factor that controls the production of an antibiotic in Streptomyces coelicolor (15).


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Sources of Recombinant AtrA—
Purified recombinant AtrA was obtained using the Escherichia coli pET system (16) as described previously (15). The final protein concentration was determined by measuring the absorbance of samples at 280 nm and using an extinction coefficient of 13,940 m–1 cm–1 (15). The purity of the preparation was greater than 95% as judged by SDS-PAGE analysis. Crude extracts containing AtrA were produced by growing E. coli BL21 (DE3) that contained the pJAS407 expression construct (15) in 500 ml of LB (Luria-Bertani) medium at 37 °C with agitation at 225 rpm for 16 h. The cells were harvested by centrifugation at 3,000 x g for 20 min and resuspended in 5 ml of ice-cold TS (Tris storage) buffer (10 mm Tris-Cl (pH 7.9) and 10% (v/v) glycerol containing 0.2 mg ml–1 of EDTA-free protease inhibitor mixture (Sigma)). The cell suspension was lysed in a precooled French® pressure cell at an internal pressure of 22,500 p.s.i. Insoluble cell debris were removed from the lysate by centrifugation at 10,000 x g at 4 °C for 30 min in a refrigerated bench top centrifuge (MSE, Falcon 6/300). The supernatant was removed and stored as 1-ml aliquots at –80 °C.

Standard Electrophoretic Mobility Shift Assay—
Assays using slab gels were performed as described previously (15). The reaction buffer contained 10 mm Tris-Cl (pH 7.9), 10% (v/v) glycerol, and 50 mm KCl. Each experimental reaction was 25 µl and contained a DNA probe (4 nm), 2 µg of sheared herring sperm DNA (Promega), and either purified AtrA or a sample of crude extract. Control reactions were always included that contained all the components except protein. The binding reactions were allowed to proceed for 20 min at room temperature. Prior to DNA-protein complexes being separated from free DNA and protein using polyacrylamide gel electrophoresis, a 0.5-µl aliquot of a 6x DNA loading dye (0.25% (w/v) bromphenol blue, 0.25% (w/v) xylene cyanol FF, and 40% (w/v) sucrose) was added to simplify loading and facilitate visualization of electrophoretic separation. The standard gel was made of 4% (w/v) 80:1 acrylamide:bisacrylamide in 1x TGE buffer (50 mm Tris-Cl (pH 8.5), 1.9 m glycine, and 10 mm EDTA). The gels were poured between Protean® II xi glass plates (Bio-Rad), and electrophoresis was performed at 120 V for 20 min in 1x TGE buffer with use of a water-cooled central core. The positions of the DNA (see below) were visualized using a Bio-Rad FX Pro molecular imager with an excitation wavelength of 488 nm in combination with a 523-nm filter.

Each of the DNA probes, which were generated by PCR, was labeled at one of its 5'-ends with fluorescein incorporated during primer synthesis. The primer pairs used to generate fragments of increasing size that incorporated Region 2 of the actII-ORF4 promoter region (15) were: I (100 bp), 5'-TTG GGA CGT GTC CAT GTA ATC ACC and 5'-TCG TGC CGC CTG AGG AGC AGC AGC; II (250 bp), 5'-GTT GTA AAA CGA CGG CCA GTG and 5'-GCT CTC CCA TAT GGT CGA CCT; III (500 bp), 5'-GTT GTA AAA CGA CGG CCA GTG and 5'-TTA ATG CAG CTG GCA CGA CAG; and IV (750 bp), 5'-GTT GTA AAA CGA CGG CCA GTG and 5'-CTC TGA CTT GAG CGT CGA TTT. The template for these reactions was a pGEM-T easy-based vector that had inserted at the EcoRV site the 100-bp fragment (designated I) that contains Region 2. The DNA probes were purified following agarose gel electrophoresis using the QIAquick gel extraction kit according to the manufacturer’s instructions (Qiagen). The DNA was eluted using 50 µl of nuclease-free water and stored at –20 °C. The latter was done in foil-covered tubes to prevent photobleaching of the fluorescein dye.

Two-dimensional Electrophoretic Mobility Shift Assays—
The reactions were performed as described above except the volume was 50 µl, the concentration of the fluorescently labeled Region 2 probe was 30 nm, and ~1.5 A600 units of E. coli cell lysate were used. The tube gel was prepared by pouring the polyacrylamide gel mixture described above into 15-cm-long plastic tubes with an internal diameter of 5 mm. The tubes were prepared from 1-ml disposable pipettes (Bibby Sterilin). The tube gels were electrophoresed in a Protean II xi tube gel apparatus (Bio-Rad) using the same conditions described above. The gels were then extracted from the tubes using a 22-gauge needle and transferred into 0.1% (w/v) bromphenol blue, 20% (w/v) SDS, and 125 mm Tris-Cl (pH 6.7) and incubated for 15 min at room temperature. A 10% (w/v) (29:1) polyacrylamide-SDS resolving gel was then poured between Protean II xi plates (Bio-Rad), and the tube gel was incorporated into the 5% (w/v) stacking gel, which was subsequently overlaid with 1% (w/v) molten agarose. Electrophoresis was performed at 200 V for 6 h, and the gel was stained using Coomassie Blue R-250.

Peptide Mass Fingerprinting—
Polypeptide spots of interest excised from Coomassie Blue-stained 2D PAGE gels were destained with 50 mm ammonium bicarbonate and 50% (v/v) acetonitrile for 10 min with sonication in a water bath with the supernatant then being removed and discarded. This step was repeated twice before the gel pieces were dehydrated in acetonitrile for 5 min. The acetonitrile was removed, and the gel pieces were allowed to dry completely in air (30 min). They were then rehydrated in 25 mm ammonium bicarbonate containing 0.02 mg ml–1 trypsin (Promega). Proteolysis was allowed to proceed at 37 °C for 24 h. An aliquot (1 µl) of the digest was mixed with 1 µl of MALDI matrix solution (10 mg ml–1 {alpha}-cyano-4-hydroxycinnamic acid in 50% (v/v) ethanol and 50% (v/v) acetonitrile) and dried onto a MALDI target plate.

The dried target was transferred to a mass spectrometer (M@LDI L/R system, Waters), and each digest was analyzed by MALDI-TOF MS in reflectron mode using standard operating parameters. Briefly the instrument used a N2 laser at 337 nm, source voltage was set at 15,000 V, microchannel plate detector voltage was set at 1950 V, pulse voltage was set at 2450 V, reflectron voltage was set at 2000 V, and coarse laser energy was set to low with fine adjustment used for each sample to optimize the signal. At least 100 laser shots were accumulated and combined to produce a raw spectrum over the m/z range 800–3500. Spectra were processed (background subtraction, smoothing, and peak centroiding) and calibrated externally using a tryptic digest of alcohol dehydrogenase and then internally using a trypsin autolysis product (reducing mass error typically to <50 ppm). The set of monoisotopic peptide masses for each sample was used to search the Swiss-Prot and/or National Center for Biotechnology Information non-redundant (NCBInr) databases using the Mascot search engine (17) (Matrix Science) to identify the parent protein. Searches were performed using an unrestricted protein molecular mass range, searching tryptic peptides from all species, and allowing one missed cleavage site and 50 ppm error tolerance in the peptide mass.


   RESULTS
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
To illustrate that the formation of a complex can cause a significant shift in the electrophoretic mobility of the protein as well as the DNA (14), we used the AtrA transcription factor, which binds to the promoter of actII-ORF4, the cluster-situated activator of the actinorhodin biosynthetic genes in S. coelicolor (15). Purified AtrA binds to a labeled PCR fragment containing a footprinted site (called Region 2) to produce a single complex that is readily identifiable using a standard EMSA (Fig. 1, panel A). Moreover staining of the slab gel with Coomassie Blue shows that the mobility of AtrA increased when bound to the DNA fragment. Indeed in the absence of DNA binding, AtrA remains largely associated with the loading well. When the same promoter fragment was incubated with a crude extract of E. coli cells expressing AtrA, a single complex of the same mobility as that associated with purified AtrA was generated (panel B). However, Coomassie Blue staining of this gel revealed that the position of the complex overlapped the tail of the smear of proteins in the crude extract. The mobility of proteins in native gels is dependent on their size and their pI relative to the pH of the buffer; however, attempts to resolve completely the AtrA-actII-orf4p promoter complex from other proteins of the crude extract were unsuccessful. Although the raising of the pH reduced the amount of bulk protein entering the gel, the mobility of the complex was also reduced (data not shown). The above experiments showed that another form of separation would be required to facilitate the routine identification in a crude extract of a protein band that corresponds to a specific nucleoprotein complex.


Figure 1
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  		FIG. 1. The electrophoretic mobility of a protein is affected when it binds a nucleic acid. The EMSAs were performed as described previously (15) using purified AtrA (panel A) and a crude extract of BL21 (DE3) cells expressing recombinant AtrA (panel B). The binding reactions had a volume of 25 µl, and each contained 100 fmol of DNA corresponding to Region 2 of the actII-ORF4 promoter (15). The DNA was labeled at one of its 5'-ends with fluorescein. The amounts of purified AtrA and crude extract used in panels A and B were 4 µm and 0.8 A600 units, respectively. Lanes 1, 2, and 3 contained DNA, protein, and a mixture of DNA and protein, respectively. The positions of the DNA in the gels were detected using a flatbed scanner that detects fluorescence (see "Experimental Procedures"), whereas the protein was detected in a subsequent step by staining with Coomassie Blue R-250 (images on left and right of each panel, respectively). The position of the complex is indicated by the Roman numeral I.

 
Having used slab gels to establish EMSA conditions that produced a tight band corresponding to the AtrA-actII-orf4p promoter complex, we switched to using tube gels that could be fused in a subsequent step to the top of a standard SDS-polyacrylamide slab gel. The tube gels were prepared using transparent holders that allowed the position of colored loading dyes to be viewed and the same acrylamide mixtures and buffers that were used for the slab gel. The crude extract containing AtrA was incubated separately in the presence and absence of the actII-orf4p promoter fragment, and then the samples were run separately in tube gels that were prepared using the same batch of gel mixture and were assembled in the same electrophoresis apparatus. After running for sufficient time for the AtrA-actII-orf4p promoter complex to migrate to at least the center of the gel, the tube gels were removed and fused separately to identical SDS-polyacrylamide gels that again had been prepared from the same batch of gel mixture (for details of the above, see "Experimental Procedures"). After running in the same electrophoresis apparatus, these gels were stained with Coomassie Blue and compared to identify the spot corresponding to protein that was in the nucleoprotein complex. The latter is a relatively straightforward step as the position of the complex in the first dimension can be estimated from its position relative to a colored dye added at the time of loading or determined directly if the label on the DNA can be detected without damage to the tube gel. For the example described here, this was done by scanning the gel while it was still in its holder for the fluorescein label. Inspection of the 2D gels revealed a spot that was unique to the sample incubated with Region 2 of the actII-orf4p promoter (Fig. 2). Comparison of the gels also identified the position of "free" AtrA in the sample that had not been incubated with the Region 2 fragment. The overall effect was that the position of a single protein band appeared to shift. Peptide mass fingerprinting confirmed the presence of AtrA in the unique spot corresponding to sample incubated with DNA (Fig. 3).


Figure 2
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  		FIG. 2. The direct incorporation of the EMSA into a 2D proteomics-based approach allows the rapid identification of nucleic acid-binding proteins. Samples of crude extract containing AtrA (~1.5 A600 units) were incubated with and without actII-ORF4-promoter DNA (1.5 pmol) in a volume of 50 µl (for conditions, see Fig. 1) and then run in native polyacrylamide tube gels. The tube gels were fused to the top of the stacking portion of SDS-polyacrylamide gels, which were then run under normal denaturing conditions. Panel A corresponds to the sample that had been included without actII-ORF4 promoter DNA, whereas panel B corresponds to the sample that had been incubated with this specific fragment. Both gels were stained with Coomassie Blue R-250. Arrows in panels A and B indicate the position of spots of AtrA (see Fig. 3) that in the first dimension were unbound and bound to actII-ORF4 promoter DNA, respectively.

 

Figure 3
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  		FIG. 3. Peptide mass fingerprinting identifies AtrA in a unique spot from a 2D gel that incorporated an EMSA. A shows the peptide mass fingerprint generated by MALDI-TOF mass spectrometry of the trypsinization products of the spot unique to the sample incubated with the actII-ORF4 promoter DNA (see Fig. 2). The y axis shows species abundance in terms of percent signal, whereas the x axis shows the m/z ratio. Major peaks that correspond to predicted products of AtrA trypsinization are labeled alphabetically according to their molecular mass, which is provided in the inset. Peptide A' is the arginine-extended equivalent of A. Predicted trypsinization products that have a molecular mass identified by peptide mass fingerprint are highlighted in panel B. The labeling above the highlighted peptide segments in AtrA indicates the peak to which they correspond in the peptide mass fingerprint.

 
Although AtrA was produced using the pET system (16), its synthesis was not induced. Indeed standard SDS-PAGE comparison of uninduced cells that contained or lacked the plasmid encoding AtrA did not reveal any detectable differences in the profile of proteins (data not shown). We therefore estimated that AtrA was expressed to a level of <1.0% of total protein. Consistent with this, the intensities of the Coomassie Blue-stained spots of several E. coli proteins in the subset that had sufficient electrophoretic mobility to enter the gel during the EMSA step were similar to that of AtrA (Fig. 2). This illustrates that our technique is sufficiently sensitive to detect in crude extract a specific protein that has been produced to a level that is within the norms for native proteins of at least a bacterial source. In cases where a complex has multiple protein components (for examples, see Refs. 18 and 19), more than one protein band would be expected to shift position in our 2D EMSA.

It is possible that the position of a shifted protein could by chance overlap that of a polypeptide of equal or greater abundance. In this scenario, no shifted band would be detected without altering the mobility of the complex in the first dimension. We have found that a straightforward way to do this is to change the size of the DNA fragment to which the protein binds. This is illustrated using AtrA and fragments ranging in size from 100 to 750 bp: the mobility of the AtrA-containing complex was reduced as the size of its DNA target was increased (Fig. 4).


Figure 4
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  		FIG. 4. The electrophoretic mobility of a nucleoprotein complex is reduced as the size of the DNA target is increased. Panel A shows purified PCR products of different sizes that each contain a single binding site for AtrA. Lanes 1–4 contained fragments of 100, 250, 500, and 750 bp, respectively. Lane M, molecular weight markers. Details of the primers and conditions used for PCR are given under "Experimental Procedures." One primer of each pair was labeled with fluorescein at its 5'-end. The gel, which was composed of 1.8% (w/v) agarose (Tris borate-EDTA buffer), was stained with ethidium bromide following electrophoresis. Panel B shows the results of the corresponding EMSA. Each reaction contained a particular PCR product (20 nm). The size of each fragment is indicated at the top of the panel. Below these sizes, minus and plus signs indicate whether the fragment was incubated without or with 4 µm AtrA. For each fragment, an arrow indicates the position of the nucleoprotein complex. The EMSA assay was performed as described in Fig. 1.

 

   DISCUSSION
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The identification of proteins that interact with specific segments of a nucleic acid is crucial to the investigation of numerous cellular processes by biochemical, genetic, and/or structural studies. Until recently, this required the fractionation of cell extracts using chromatography methods to establish an association between an activity and a particular polypeptide band on an SDS-polyacrylamide gel. Even with advances in mass spectrometry that have reduced the amounts of protein required for successful identification, this approach remains time-consuming and labor-intensive. In contrast, it is often relatively straightforward to determine using the electrophoretic shift assay that a protein in a crude or partially purified extract binds to a nucleic acid target of interest and to investigate without extensive purification the specificity of the interaction via competition experiments, mutagenesis of the target, and footprinting studies (20). Given that the EMSA remains a method of choice for establishing that a particular activity exists, we sought to incorporate this assay into a 2D proteomics-based approach that would allow the source of a particular activity to be identified rapidly. This was achieved by simply replacing isoelectric focusing with the EMSA in the first dimension of 2D PAGE. In this way, the position of bands corresponding to specific nucleic acid-binding proteins of interest can be determined without the need for specialized equipment or expensive reagents. Furthermore identification can be achieved readily by excising bands of interest from SDS-polyacrylamide gels and submitting to specialized facilities for peptide mass fingerprinting. In this way, laboratories with an interest in a particular segment of nucleic acid, such as regulated promoter, can readily take advantage of technical advances in proteomics without committing to a global analysis.

Others have also realized that the EMSA provides an excellent starting point from which to develop proteomic approaches for the identification of nucleic acid-binding activities. However, to our knowledge, the only such approach that had been published prior to our work requires that the molecular mass or pI be estimated so that the position of candidate proteins on a standard 2D gel can be determined (13). The candidate(s) is then excised from the gel, eluted, renatured, and tested for the activity that was originally detected before being identified by mass spectrometry (13). The approach described here is much simpler and does not require knowledge of the molecular mass or pI of the nucleic acid-binding protein.

Once the identity of a protein has been determined using mass spectrometry, our method of choice for confirming it as the source of activity is to express the corresponding gene in a heterologous host such as E. coli and to test for the original activity using EMSA (15). This approach has the added advantage that it can generate sufficient amounts of protein for further biochemical characterization and structural studies. In cases where it is not possible to produce active protein in a heterologous host, the option described above of assaying polypeptide that has been excised from the gel and renatured will be of value (13). Another option is to use antibodies against the candidate to determine whether they cause a supershift of the complex(es) in an EMSA (20). The latter approach might be of particular value in cases where it appears that more than one protein is required to form a complex with a nucleic acid target (18, 19).

The nucleic acid used in this study was DNA; however, we know of no reason why this approach should not work for EMSAs that incorporate RNA. The primary requirement of our approach is that complexes form a tight band during gel electrophoresis. Here we demonstrated the utility of our approach using amounts of complex that were approximately 100 fmol. Thus, the protein component could be detected readily by staining gels with Coomassie Blue (Figs. 1 and 2). It should be noted, however, that the current generation of mass spectrometers can make identifications from subfemtomole amounts of protein (3–5). Therefore, it should also be possible to identify by mass spectrometry shifted proteins that can only be detected following staining with more sensitive reagents such as silver or one of several fluorescent stains (21, 22). To follow complexes in the subfemtomole range via a label on the DNA, we recommend incorporating modified nucleotides that can then be detected enzymatically using an appropriately sensitive fluorescent or chemiluminescent substrate (23). While establishing the technique, we were not averse, however, to using DNA labeled with the radioactive isotope 32P as during the running of the second dimension the DNA exits the gel; consequently radioactivity is not detectable in any gel slices submitted for MS analysis.

EMSAs that are used to determine whether a particular activity exists in a sample tend to maximize the sensitivity by using probes that have been labeled to a high level of specific activity (20). They also tend to use the lowest possible concentrations of probe that allow detection to maximize the possibility that it is substantially lower than the concentration of the protein(s) that binds. Under these conditions the proportion of the probe that is shifted can be used to estimate the amount of activity (20). For the purpose of identifying a protein by the method described here, once it has been established that an activity of interest exists, we recommend using what would be considered for standard EMSA reactions high concentrations of probe (4–40 nm) to drive the formation of complexes and thus increase the probability of detecting unique protein spots in the 2D gels.

We have illustrated the potential of our approach using as the source of crude extract cells of E. coli that were expressing a heterologous protein. This material was chosen as it can be produced readily to provide a standard for other laboratories to establish the technique described here. As for many other transcription factors, the expression of atrA in its native host, S. coelicolor, is dependent on growth conditions and requires specialist experience and knowledge to reproduce. However, it is possible to detect by EMSA the binding of AtrA to DNA in extracts of S. coelicolor that have not been purified using chromatography (15). In some cases, for example the analysis of a transcription factor that is expressed only within a subset of the cells of a multicellular organism, a chromatography step may be required to ensure there is sufficient protein in the small volumes that can be loaded on gels (generally less than 50 µl) to permit its detection after running the second dimension and staining. Nevertheless as shown here our simple proteomics-based approach can be used to analyze sample of complex protein composition and should still greatly reduce the number of chromatography steps required to associate an activity with a specific polypeptide. As such it should contribute significantly to the annotation of the function of the proteome (24, 25).


   ACKNOWLEDGMENTS
 
We thank our colleagues Simon Baumberg, Alison Ashcroft, and Ian Hope for helpful discussions and comments on our manuscript.


  FOOTNOTES
 
Received, May 31, 2006, and in revised form, July 15, 2006.

Published, MCP Papers in Press, July 15, 2006, DOI 10.1074/mcp.T600027-MCP200

1 The abbreviations used are: EMSA, electrophoretic mobility shift assay; 2D, two-dimensional. Back

* This work was supported by Ph.D. Studentship S02/G031 from the UK Biotechnology and Biological Sciences Research Council (BBSRC). Facilities provided by the BBSRC under the JIF (Joint Infrastructure Fund) Initiative were used during this work. Back

{ddagger} To whom correspondence should be addressed. Tel.: 44-113-343-3109; Fax: 44-113-343-2835; E-mail: k.j.mcdowall{at}leeds.ac.uk


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 DISCUSSION
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