High-Throughput Proteomics Processing of Proteins in Polyacrylamide

Mar 17, 2007 - Robert H. Waldo, Edwin D. Cummings, Siva T. Sarva, Janet M. Brown ... Lorne A. Rose, Robert J. Belland, Karl E. Klose, and George M. Hi...
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High-Throughput Proteomics Processing of Proteins in Polyacrylamide in a Multiwell Format Edwin D. Cummings, Janet M. Brown, Siva T. Sarva, Robert H. Waldo, and George M. Hilliard* Center of Excellence in Genomics and Bioinformatics, University of Tennessee Health Science Center, Memphis, Tennessee 38163 Received September 11, 2006

Abstract: Processing multiple protein samples from polyacrylamide at significant sensitivity represents a major chokepoint for raising the success rate in high-volume protein identification projects. A multiwell filterplate method for processing proteins in polyacrylamide was optimized for sensitivity using a protein standard. The results demonstrate this process to be a reliable and reproducible method over a range of gel loadings and suitable for the identification of proteins near the threshold of silver stain. This high-throughput manual method requires a minimum of specialized equipment, and can be performed disconnected from a proteomics infrastructure for the preparation of mass spectrometry-ready samples. Keywords: Polyacrylamide • Proteomics • Multiwell • Trypsin • Extraction

Introduction Recurrent questions asked of institutional proteomics analysis are the amount of protein required in a polyacrylamide band for a successful identity match and how many samples of unknown proteins can be processed for identification. The dynamic ranges of protein expression among biological species of increasing complexity are beginning to be understood, and it is apparent that the relative depth into these ranges that proteomics analysis can reach is limited by multiple factors.1 These limitations increase proportionally with genome size and are relieved partially by proteome fractionation and protein arrays intending to maximize the comparative display of proteins prior to the downstream phases of processing and analysis by mass spectrometry (MS). Despite its limitation to array and detect only the most abundant proteins in a sampled range,2 polyacrylamide consistently remains a popular method for comparative protein display. The early standard for applying proteomics to biological samples in polyacrylamide was set by the development of proteomics processing methods that proved protein silver staining was indeed compatible with subsequent MS analysis of peptides. The primary advancement was to demonstrate that proteins present in the low nanogram range in polyacrylamide could be proteolyzed with trypsin and one or more fragments * Corresponding author. Telephone: (901) 448-6779 (office). Fax: (901) 448-2706. E-mail: [email protected]. 10.1021/pr060472y CCC: $37.00

 2007 American Chemical Society

recovered for analysis by nanospray tandem MS, a sensitive but low sample throughput method.3,4 A batch method with high sensitivity for higher sample throughput in MS analysis is peptide mass fingerprinting (PMF) with matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDITOF MS). In PMF only peptide mass data from an unknown protein is typically made as opposed to peptide sequence data in tandem MS. Furthermore, PMF is an accurate, salt-tolerant MS analysis for protein identification, especially when in combination with modern MALDI-TOF MS instrumentation and PMF database search algorithms.5-7 In practice, the success rate of identifying unknown proteins in polyacrylamide has a dynamic sensitivity threshold essentially inversely related to sample throughput. MS methods that develop more detailed peptide information with decreased amounts of an unknown protein typically take a longer time in analysis (e.g., nanospray tandem MS) with a comparative negative impact on the total sample throughput. In support of proteomics in a research setting, striking a balance between sample throughput and sample success rate is the route of choice to productive proteomics for the most number of investigators. The current sensitivities of high-throughput, batch methods of MS analysis, such as PMF, and database searching are more than sufficient in providing high-confidence protein identifications for progression of competitive proteomics research projects. PMF can provide the bulk of protein identifications quickly, thereby reserving more time-consuming MS procedures for samples of special interest. Currently, processing multiple protein samples from polyacrylamide at significant sensitivity represents a major chokepoint for raising the success rate in high-volume protein identification. In the technical stage of recovering peptides from unknown proteins in polyacrylamide it is very important to manage all factors that influence the success rate of protein identification. Peptide extraction from polyacrylamide is an analytical method, and optimization of peptide recovery during extraction steps ultimately contributes to the production of the highest quality mass spectra for use in database searches. The final quality of the peptide mass measurements dictates the subsequent ability to match the MS data with a protein in a database with high confidence. As an improvement over limitations of sample throughput from existing polyacrylamide peptide extraction methods, a sensitive, high-throughput, manual proteomics processing protocol requiring only a vacuum multiwell plate apparatus is reported here. Polyacrylamide extraction protocols were optiJournal of Proteome Research 2007, 6, 1603-1608

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mized for this multiwell format, and processed samples are suitable for direct PMF analysis with MALDI-TOF MS. Importantly, this processing method is within the capabilities of most biological research laboratories and can be performed disconnected from any proteomics infrastructure within an institution for the purpose of creating mass spectrometry-ready samples. These MS-ready samples could subsequently be submitted for mass spectrometry analysis anywhere it was available.

Materials and Methods Microplate Equipment. Millipore 96-well filter plates (No. MSRLN0410) were evacuated with a vacuum manifold (No. MSVMHTS00); samples were lyophilized in a Thermo Savant (Holbrook, IL) SpeedVac concentrator (No. SC110A-115) with a 96-well plate rotor (No. RH2MP) and a vacuum system equipped with an UltraLow refrigerated trap (No. RVT4104) and digital vacuum gauge (No. DVG50-115). Dual Alcatel Pascal mechanical pumps (No. AV-UM2010SL) and tubing and fittings were from Kurt Lesker Co. (Clairton, PA). Vortex Genie (No. 12-812) and 96-well adaptor (No. 12-812A) were from Fisher Scientific. Multiwell pipettemen were from Liquisystems (No. 01913559). Keratin Control in Gel Casting. Polyacrylamide gels are cast according to Sambrook et al.8 Dust control correlates with keratin control, and all manipulations of acrylamide samples and vacuum manifold are performed in a dust-free hood environment. Dust is cleaned from all work surfaces, and sample handling is reduced by the multiwell vacuum plate, thereby reducing the susceptibility of this process to contamination by keratin. Polypropylene surfaces in contact with the samples are prerinsed with 60% acetonitrile and 0.25% trifluoroacetic acid (TFA) to remove polymers, dust, or releasing agents. Gloves are worn when handling glassware that comes into contact with samples or solutions. Hair is pulled back and secured, and nonlinting clothes are worn under a clean laboratory coat. Gel plates are scrubbed with a stiff brush and soap and cleaned for final assembly using ethanol and lintfree disposable wipes. After assembly, settled dust is blown from between plates using clean pressurized, canned air (Fisher). In casting the gel, acrylamide, Tris, and water are combined and filtered through clean No. 1 Whatman paper in a Buchner funnel prior to addition of sodium dodecyl sulfate (SDS) and polymerization agents. Silver Stain for Gels. Gels are silver-stained by the method of Shevchenko et al.3 Five gel volumes of each solution are freshly made. After electrophoresis, gels are fixed in a solution containing 20% methanol and 10% acetic acid in water for 90 min. The fix solution is removed, and the gel is washed twice in water for 20 min each. The gel is put into 0.02% sodium thiosulfate for 3 min and then washed twice in water for 30 s each. The gel is put into 0.1% silver nitrate for 30 min and then washed twice in water for 30 s each. The silver nitrate is left in up to 2 h for more intense brown color. The gel is developed in 2.5% sodium carbonate/0.02% formaldehyde. Using this silver recipe, bands that begin to appear within 30-60 s are usually identified successfully in this multiwell process. Bands appearing only upon overexposure may not be in the sensitivity range of the total procedure. The developer is removed, and the reaction is quenched with 1% acetic acid for 10 min. The gel is rinsed twice with water for 20 min and stored in water at 4 °C. Band Excision and Destaining. Protein bands for identification are excised and placed into microfuge tubes prewashed 1604

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technical notes with 60% acetonitrile and 0.25% TFA. Control bands for MS are excised from a non-protein containing region of the gel. Bands cut from dried gels are rehydrated with 400 uL of water on ice for 15 min. The paper or cellulose backing swells faster than acrylamide and can be removed by sharp forceps. Destaining of all protein bands results in better comparable signals in MS and is routinely performed for every sample run in the plate. Protein bands stained with coomassie blue or sypro ruby are destained twice in 200 uL of 50% acetonitrile in water at 23 °C and vortexed constantly for 30 min. This step is repeated for coomassie bands until the blue color is gone, usually overnight. Silver-stained bands are destained in the 96well Millipore PTFE 0.45 um filter plate (No. MSRLN0410). The wells of the plate are prewashed with three successive additions of 200 uL of Plate Wash Solution (25% methanol, 0.1% TFA). The wells are evacuated by applying vacuum to the manifold. The excised bands are placed into wells and 200 uL of fresh Farmer’s reagent (30 mM potassium ferricyanide and 100 mM sodium thiosulfate mixed 1:1) is added.9 The plate is removed from the manifold and the under-drain of the 96-well plate is wrapped in Parafilm (keratin shield), and the plate is gently vortexed (Fisher adaptor No. 12-812) until all brown color is removed from the bands (about 5 min, avoid extended incubations in this reagent). The solution is removed by manifold vacuum for 30 s, and the bands are washed in four successive 5 min incubations with gentle vortexing in 200 uL of water. Trypsinization. In the 96-well filter plate, protein bands are dehydrated by addition of 200 uL of neat acetonitrile to each well, and incubated for 5 min at 0 °C (use Parafilm on underdrain as keratin shield while the filter plate is on ice). The solution is removed, and gel pieces are dried by applying vacuum to the manifold for 1 min. Addition of 200 uL of neat acetonitrile is repeated and removed by vacuum. A 100 uL aliquot of fresh digestion buffer (12.5 ng/uL trypsin (Promega No. PRV5111), 25 mM ammonium bicarbonate, and 10% acetonitrile) is added. The bands are completely submerged, and necessary volume is added to cover. The under-drain is wrapped with Parafilm, and the plate is gently vortexed and incubated at 0 °C (ice) for 15 min. The vortex is repeated, and the plate is incubated at 0 °C for no more than 90 min (empirical optimization suggested). The digestion buffer is removed with gel loading tips or by manifold vacuum. A 35 uL aliquot of trypsin-free digestion buffer is added (without trypsin). The bands are completely submerged, and the necessary volume is added to cover. The collection plate (Nunc No. 442587) is taped to the underside of the filter plate, and the top of the filter plate is covered with a Simplate mat (Fisher No. 08408219). The samples are incubated overnight at 37 °C on a temperature block with an adaptor for 96-well plates and covered, and the top of the plate is insulated completely with foam rubber to eliminate condensation on the well covers. Peptide Extraction. The incubation plate assembly is spun briefly to collect condensation off the Simplate mat cover. The assembly is vortexed gently for 15 min at room temperature while the collection plate is still taped to the filter plate. The collection plate is placed in the bottom of the vacuum manifold apparatus, and the filter plate, on top of the grid of the manifold. The solution is collected from each well into the collection plate by applying a vacuum of 4-6 in. of mercury for 60 s to the manifold. The majority of recovered peptides are in this first solution.14 A 25 uL aliquot of fresh extraction solution (50% acetonitrile, 5% TFA, in water) is added; the volume is adjusted for larger acrylamide loads if necessary. For

technical notes

Cummings et al.

Figure 1. Comparison of the multiwell acrylamide peptide extraction process with a solution digest of BSA. Solid lines are BSA peptides extracted from polyacrylamide at each respective gel loading; broken lines are the known amounts of peptides spotted from dilutions of a solution digest of BSA. BSA peptides (M + H) are 927.494 (]), 1439.812 (2), 1479.796 (9), 1567.743 (b), and 1639.938 (×).

15 min the solution is vortexed gently with the collection plate taped to the filter plate, vacuum is applied to collect the solution. A 25 uL addition of extraction solution is repeated, followed by vortexing and collection by vacuum of solution. Unused wells in the filter plate can be subsequently used if the plate is preserved in a zipper-locking bag between runs. No variability within the plate from run to run has ever been noted in the BSA standard as these unused wells remain new until used. The under-drain is wrapped in Parafilm, the Simplate mat is placed on the top, and this is kept in a zipper locked bag at room temperature until the next use. The extracted samples in the collection plate are spun in a SpeedVac capable of 0 Torr (dual pump) until wells are dry (inefficient vacuum associated with losses in sensitivity). A 40 uL aliquot of pellet wash (0.25% TFA) is added; the sample is gently vortexed and spun to dryness for the remainder of the day. A 40 uL aliquot of pellet wash is added; the sample is gently vortexed and spun to dryness overnight. Sample Spotting for MALDI Analysis. The anchor chip (Bruker No. 209513) is washed with 50% methanol. A 1 uL aliquot of resuspend solution (60% acetonitrile, 0.25% TFA) is added to the center of the conical well in the collection plate, where the dried sample is located. The entire volume is transferred to a coordinate on the MALDI target plate. A 1.0 uL aliquot of matrix is added to the sample droplet before it dries, and the mixture is allowed to dry on the plate. The plate may be analyzed at this time. Additionally, the crystals are washed by covering with a 5 uL droplet of spot wash (0.25% TFA) and allowed to stand for a few seconds, and the wash is aspirated off with a pipet tip connected to a vacuum source.10 MALDI TOF Analysis. Spectra were recorded on a Bruker Ultraflex. Internal calibration was on autolytic trypsin fragments 842.509 and 2211.104.11 External calibration was on a 2:1 mixture of des-R-bradykinin (Sigma No. B-1901) and ACTH (Sigma No. A-0673). The number of summed laser shots was 150. Parameter file settings were extraction voltage of 20 kV

and reflector voltage of 11.8 kV. Laser settings were manually optimized for each spectra. Software and Statistical Methods. In each spectrum the signal-to-noise (S/N) ratio at each respective m/z value was calculated using the SNAP algorithm with default parameters within the Bruker FlexAnalysis Version 2.0 software. Searches were done with local ProFound Version 2002.03.01 software on the local nrNCBI database compiled Aug 8, 2005 (2 086 747 sequences). The best four of five peptide ion S/N calculations were used for calculating each mean S/N value used in Figure 1 by discarding the value farthest from the median. The mean S/N, standard deviation (SD), and the relative standard deviation (%RSD) were calculated using a spreadsheet.

Results Calibration of the Process with BSA Standard. Serial dilutions of a commercially available BSA standard (Pierce, 2 mg/mL, No. 23209) were used to calibrate this processing method. A panel of five technical replicates in quantities of 4000, 2000, 1000, 500, and 250 fmol were run in 25 lanes of commercial, precast SDS gels ( Invitrogen No. EC6485), silverstained, excised, and processed as 25 individual samples in a single run of the plate. Mass spectra were internally calibrated on autolytic digest fragments of trypsin present in the mass spectra of each sample. The mass measurement error for BSA peptides ranged from 20 to 100 ppm. The mass resolution of all BSA peptide ions was in the range of 2500-8000. The raw spectra are available online in the Supporting Information. A summary of the recovered peptides of BSA is shown in Table 1. The minimum appearance of a BSA peptide necessary for inclusion in Table 1 was at least four of the set of five mass spectra for each amount (all were five in five except 1439.812 in 250 fmol). The protein quantities loaded in the wells of the gel are presented in moles, but it should be noted that the lowest concentration (250 fmol) is equivalent to 16.5 ng of protein at the molecular weight of BSA. Journal of Proteome Research • Vol. 6, No. 4, 2007 1605

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Multiwell Proteomics Processing Method Table 1. BSA Peptides Recovered from Acrylamide

a

MS bracket, 500-2400 Da. b Z score of 1.65 > 95% confidence.6

The recovery efficiency of BSA peptides from acrylamide was dependent on the amount of protein in the band. In Table 1 it can be seen that a majority of peptides from BSA are recovered from the highest gel loading, but as the gel loadings are reduced, the lowest loadings recover only a few BSA peptides for MALDI-TOF MS analysis. The BSA peptide ions (M + H) that persist down to the 250 fmol gel loading are 927.494, 1439.812, 1479.796, 1567.743, and 1639.938 and are the only S/N values plotted in Figure 1. Consistent performance and recovery are demonstrated by the results of this processing protocol. The range of the coefficient of variation of the mean ion S/N for the 4000 to 500 fmol gel loadings remain consistent and between 3.5 and 31.7 %RSD. Specifically, for ions obtained from the five 500 fmol gel loadings the mean S/N variation of BSA peptides within the single plate run are between 6.9 and 31.7 %RSD. As a comparison, five consecutive plate runs that include a 500 fmol BSA band as a control have a similar variation of the mean S/N in the range of 9.1-40.1 %RSD (data not shown). In contrast, peptide recovery in the lowest gel loading of 250 fmol has a variation of the mean S/N in the range of 35.6-76.4 %RSD. The clear trend is the variation of peptide recovery within-the-plate and plate-to-plate remains consistent in the gel loadings down to 500 fmol, and the highest S/N variability occurs at the lowest gel loading of 250 fmol. Nonspecific adsorption and the occurrence of partial tryptic peptides contribute to the variability in peptide recovery at this lowest gel loading. BSA is still the top match in a database search using ions observed in all loadings, with a proportional loss in confidence score as fewer peptides are recovered as the gel loadings are reduced (Table 1). It is not possible to measure the exact amount of peptides recovered from each gel loading in this or any process, but recovery efficiency can be estimated by comparison of the S/N 1606

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of samples extracted from the gel with a dilution of a known standard loaded directly to the MALDI target. For this control five technical replicates of a BSA solution digest were performed (2 mg/mL, 5% trypsin (w/w)), and dilutions of each digest were subjected to MS analysis after spotting the exact amounts in Figure 1. In the MS data from these solution digests, for clarity, the mean S/N values of only the peptides co-incident with the 250 fmol gel loading are plotted.

Discussion The primary benefit of this protocol over proprietary kits12 or manual methods3,4 is economy of use in repeated highvolume sample processing and optimized performance by complete control of all reagents. Literally thousands of samples have been processed here using this reliable, economic, and sensitive processing method. There are several key steps that are intrinsic to attaining the sensitivity and reproducibility of this multiwell processing method. Proteins can be extracted more efficiently from acrylamide with increased concentrations of trypsin, but this also results in production of prohibitive amounts of autolytic fragments of trypsin that can quench trace peptides in spectra from lighter bands. The activity of trypsin was increased to 300% by using 10% acetonitrile in digestion buffer.13 The net benefit is that trypsin can be reduced up to a factor of 3-fold without changing the net peptide extraction activity, reducing substantially the interference in lightly stained bands from the autolytic fragments of trypsin. The quality and production of MS data can be reduced or completely inhibited by excess buffers or salts included in peptide samples for MALDI-TOF MS analysis. With the exception of the silver destain steps, each solution used in this protocol is volatile and can be removed by lyophilization with a SpeedVac concentrator capable of pulling a very strong

technical notes vacuum (plate begins at 1000 and ends at 0 mTorr). Drying the peptide samples down allows them to be analyzed directly without any loss or selectivity associated with a reverse-phase cleanup step, although irreversible peptide losses probably do occur by drying completely.14 The early protocols recommended small 1 mm cubes of acrylamide, presumably to increase surface to volume ratio, thereby aiding the diffusion and access of trypsin to the protein substrate embedded in the acrylamide matrix.3 Whole bands (typically 1 mm thick × 2 mm × 5 mm) yielded more consistent results from trials here than small cubes or shredded acrylamide in this process. This is similar to results seen by others in controlled studies of peptide recovery,14,15 and further aids throughput by not having to spend time cutting polyacrylamide into pieces. An additional key step was incorporated that probably does relate to making the protein accessible to trypsin, and that was a preincubation of the dehydrated acrylamide band with a trypsin solution.3,4 Presumably, trypsin diffusion into a dehydrated gel piece is enhanced by applying a concentrated trypsin solution during the time a gel reswells to its starting size. Afterward, excess trypsin solution is removed and replaced with only a buffer, resulting in trypsin being applied relative to the volume of acrylamide rather than to the quantity of protein in the band. Since average gel band sizes, exact trypsin digestion buffer concentrations, and time and temperature of trypsin incubation are all laboratory-specific variables, it is important to optimize this step for the routine implementation of this process. Extended incubations with concentrated trypsin solutions can result in significant peptide loss when the trypsin solution is removed. The digestion buffer solution can be monitored by mass spectrometry for the appearance of peptides from a standard protein. When trypsin is in vast excess during the enzyme incubation step, peptides are released and lost when the trypsin digestion buffer solution is removed and replaced with buffer solution. This is probably due to a low percent activity of trypsin (due to inefficient cooling to 0 °C), and is first noticeable as identification failure in lighter stained protein bands. The steps for incubation time and temperature and concentration of trypsin can be optimized for the highest sensitivity of this process as a routine method, or especially for lightly stained protein bands. After resuspension of the extracted peptides, the entire peptide sample is manually spotted for mass spectrometry analysis in this procedure, especially for highest sensitivity. At the lowest gel loading of 250 fmol, the comparison of S/N values with the control suggests that the recovery range of peptides for MS is 10-25 fmol, or approximately 5-10% of what was loaded into the well of the gel. The recovery is substantially better at the higher gel loadings, although these data are not part of the plot in Figure 1 but can be observed in Table 1 by increased protein coverage in the higher gel loadings. The peptide losses are probably cumulative due to nonspecific adsorption to the increased plastic surfaces of the plate, but approximate recovery levels in previous reports3,4,14 even though these processes all have important differences. Peptide recovery efficiency is progressively more severe in this process as gel loadings are reduced. The estimation of peptide recovery at 5-10% is additionally consistent with the performance specification of this model of the MALDI-TOF instrument. In Figure 1 it is evident in the solution digest dilution of 12.5 fmol that

Cummings et al.

peptide S/N corresponds to the vendor specification of obtaining at least a 10:1 signal-to-noise ratio with 10 fmol spotted on the target. The lowest gel loading of 250 fmol of BSA (16.5 ng) exceeds a commonly accepted limit of detection of protein by silver staining: 1 ng and protein- and time-dependent.8 The probability and confidence scores corresponding to protein database search results for each respective gel loading of BSA are shown in Table 1. The MS data obtained from this processing method of this protein standard can be used to successfully distinguish BSA with high probability and confidence in a current nrNCBI collection, near the threshold of detection by MS-compatible silver staining. Protein gel loadings below 250 fmol challenge the high probability and confidence threshold for this multiwell process but often make usable identifications or at minimum encourage hints to protein identity. Troubleshooting procedures in this batch processing method have revealed a few but nonobvious reasons for reduced performance. The first is incomplete washing of protein bands from the silver destain solution. It is possible these bands still contain salts or acetic acid that comparatively inhibits the downstream proteolysis or analysis procedures. Also, extended incubations on ice with a concentrated trypsin solution can result in a failure to identify lighter stained unknown proteins. Reliable protein identifications at these lowest protein levels highlight the absolute requirement of MALDI-TOF MS operation to be at peak efficiency, to include laser performance, focus, and optimal incidence angle on the target. Power supplies and components for extraction voltage and time lag focusing have all been correlated here with decreased performance of the processing method in protein identification, as they would in any method. Additionally, it is an advantage to create parameter files in the mass spectrometer that allow for automatic MS data acquisition of both high and low abundance protein bands. However, the multiwell plate processing method is techniquesensitive, so some variation in parameter file settings is necessary to account for minor inefficiencies or mistakes in the described sample processing. The biggest concern in spectra acquisition is the presence of buffers and salts in the sample after the processing has been completed, and can be attributed mostly to vacuum inefficiency either in the plate evacuation steps on the manifold or the lypohilization stages of the method in the SpeedVac concentrator. An indicator of this occurrence is the improper formation of matrix crystals usually seen as a dense opaque spot. Two solutions to alleviate this problem can be applied. First, a spot wash using 0.25% TFA can be used to remove buffers and salts, but the drawback to this approach is that it can also lead to washing away of peptides.10 To circumvent this problem, a second and more preferable method has been used. It is a field clearing method in which very high laser energy is used (60-80%) to clear an area on the spot for optimal ionization of peptides. The laser burst is very short and usually clears an area of salts and buffers from the spot clean enough for acquisition of peptide spectra using normal operation of the laser (20-30%). Samples that were not perfectly processed are still salvageable by manually acquiring MS data in this way.

Conclusion Protein separations in polyacrylamide are within the capability of virtually all research laboratories. Processing of protein bands for MS analysis is not inherently an analytical technique Journal of Proteome Research • Vol. 6, No. 4, 2007 1607

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that should be exclusive to mass spectrometry laboratories specializing in proteomics. As described, unknown protein samples near the detectable threshold of silver stain can be reliably processed in high volume in a biology research laboratory with very little need for specialized equipment and subsequently delivered anywhere for identification by MS analysis and database search. The processing of archived dried gels or fresh wet gels is equivalent in this method. The method works on bands or spots containing protein mixtures, and the reliable plate-to-plate variability makes it straightforward to calibrate gel loadings of proteomics projects into the sensitivity ranges of this method for the purpose of maximizing the success rates of protein identifications.

Supporting Information Available: Raw spectra of all BSA peptide ions. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Eriksson, J.; Fenyo, D. Predicting the Success Rate of Proteome Analysis by Modeling Protein Abundance Distributions and Mass Spectrometric Dynamic Range Limitations. Proceedings of the 53rd ASMS Conference on Mass Spectrometry and Allied Topics, San Antonio, TX, June 5-9, 2005. (2) Gygi, S. P.; Corthals G. L.; Zhang Y.; Rochon Y.; Aebersold R. Evaluation of Two-Dimensional Gel Electrophoresis-Based Proteome Analysis Technology, Proc. Natl. Acad. Sci. U.S.A. 2000, 97 (17), 9390-5. (3) Shevchenko, A.; Wilm, M.; Vorm, O.; Mann, M. Mass Spectrometric Sequencing of Proteins from Silver-Stained Polyacrylamide Gels. Anal. Chem. 1996, 68, 850-8. (4) Wilm, M.; Shevchenko, A.; Houthaeve, T.; Fotsis, T.; Breit, S.; Schweigerer, L.; Mann, M. Femtomole Sequencing of Proteins from Polyacrylamide Gels by Nano Electrospray Mass Spectrometry. Nature 1996, 379 (6564), 466-9.

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(5) Jensen, O. N.; Podtelejnikov, A. V.; Mann, M. Identification of the Components of Simple Protein Mixtures by High-Accuracy Peptide Mass Mapping and Database Searching. Anal. Chem. 1997, 69, 4741-50. (6) Zhang, W.; Chait, B. T. ProFound: An Expert System for Protein Identification Using Mass Spectrometric Peptide Mapping Information. Anal. Chem. 2000, 72, 2482-89. (7) Eriksson, J.; Fenyo, D. Protein Identification in Complex Mixtures J. Proteome Res. 2005, 4, 387-93. (8) Sambrook, J.; Fritsch, E. F.; Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd ed.; Cold Spring Harbor Laboratory Press: New York, 1989. (9) Gharahdaghi, F.; Weinberg, C. R.; Meagher, D. A.; Imai, B. S.; Mische, S. M. Mass Spectrometric Identification of Proteins from Silver-Stained Polyacrylamide Gel: A Method for the Removal of Silver Ions To Enhance Sensitivity. Electrophoresis 1999, 20, 601-5. (10) Vorm, O.; Roepstorff, P.; Mann, M. Improved Resolution and Very High Sensitivity in MALDI-TOF of Matrix Surfaces Made by Fast Evaporation. Anal. Chem. 1994, 66, 3281-7. (11) Vestling, M. M.; Murphy C. M.; Fenselau, C. Recognition of Trypsin Autolysis Products by High-Performance Liquid Chromatography and Mass Spectrometry. Anal. Chem. 1990, 62, 23914. (12) Pluskal, M. G.; Bogdanova, A.; Lopez, M.; Gutierrez, S.; Pitt, A. M. Multiwell in-Gel Protein Digestion and Microscale Sample Preparation for Protein Identification by Mass Spectrometry. Proteomics 2002, 2, 145-50. (13) Boehringer Mannheim Biochemica. Technical Note on Trypsin, 0892.T31.4. 1419242 B RD 2497. (14) Speicher, K. D.; Kolbas, O.; Harper, S.; and Speicher, D. W. Systematic Analysis of Peptide Recoveries from in-Gel Digestions for Protein Identifications in Proteome Studies. J. Biomol. Tech. 2000, 11, 74-86. (15) Havlis, J.; Thomas, H.; Sebela, M.; Shevchenko, A. Fast-Response Proteomics by Accelerated in-Gel Digestion of Proteins. Anal. Chem. 2003, 75, 1300-6.

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