MS for Membrane Proteome

In the case of the HT29 membrane protein digest, Figure 2A shows the UV .... Most peptides are eluted with a salt strength of 0.1−0.2 M NaCl. ..... ...
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Liquid Chromatography MALDI MS/MS for Membrane Proteome Analysis Nan Zhang, Nan Li, and Liang Li* Department of Chemistry, University of Alberta, Edmonton, Alberta, Canada T6G 2G2 Received November 27, 2003

Membrane proteins play critical roles in many biological functions and are often the molecular targets for drug discovery. However, their analysis presents a special challenge largely due to their highly hydrophobic nature. We present a surfactant-aided shotgun proteomics approach for membrane proteome analysis. In this approach, membrane proteins were solubilized and digested in the presence of SDS followed by newly developed auto-offline liquid chromatography/matrix-assisted laser desorption ionization (LC/MALDI) tandem MS analysis. Because of high tolerance of MALDI to SDS, one-dimensional (1D) LC separation can be combined with MALDI for direct analysis of protein digests containing SDS, without the need for extensive sample cleanup. In addition, the heated droplet interface used in LC/ MALDI can work with high flow LC separations, allowing a relatively large amount of protein digest to be used for 1D LC/MALDI which facilitates the detection of low abundance proteins. The proteome identification results obtained by LC/MALDI are compared to the gel electrophoresis/MS method as well as the shotgun proteomics method using 2D LC/electrospray ionization MS. It is demonstrated that, while LC/MALDI provides more extensive proteome coverage compared to the other two methods, these three methods are complementary to each other and a combination of these methods should provide a more comprehensive membrane proteome analysis. Keywords: membrane protein • LC/MALDI • SDS • shotgun proteomics

Introduction Membrane proteins play critical roles in many biological functions and are often the molecular targets for drug discovery. However, their analysis presents a special challenge because they are not readily soluble in polar solvents and often undergo aggregation.1-3 Loss of sample during the analysis by protein adhesion to the sample handling surfaces can be particularly problematic. Use of strong surfactants such as SDS in sample workup can assist in protein solubilization and avert the sample adsorption problem. But strong surfactants can interfere with protein characterization in many analytical techniques. Despite several technical shortcomings, gel-based protein separation in combination with mass spectrometry (MS) for protein identification is still a widely used technique for analyzing membrane proteome.4-8 The in-gel approach has some attractive features for analyzing membrane proteins. Strong surfactants can be used in sample preparation and gel separation. In addition, the proteins embedded within the gel matrix after separation establish a localized region for enzymatic or chemical cleavage (i.e., concentration effect) and thus potentially improve the chance for protein digestion. Protein * To whom correspondence should be addressed. E-mail: Liang.Li@ ualberta.ca. 10.1021/pr034116g CCC: $27.50

 2004 American Chemical Society

unfolding, reduced interference after gel separation and subsequent washing are also contributing factors which can lead to better results. Finally, although absolute quantitation of protein expression levels among different proteins displayed in gels is difficult, relative quantitation of proteins can be estimated by examining the stained gel bands or by Western blotting. Recently, the shotgun proteomics approach has emerged as a powerful technique for membrane proteome identification and quantitation.9-12 It is based on the digestion of proteins from extracts of whole cells, organelles, or specific fractions thereof, followed by liquid chromatography (LC) tandem mass spectrometry (MS/MS) analysis of the resulting peptides. For membrane proteome analysis, membrane proteins solubilized in either an organic acid,9 organic solvent,10 or an aqueous solution containing detergents such as SDS11 are subjected to proteolytic or chemical digestion. Alternatively, nonsolubilized membrane proteins are subjected to proteinase K digestion.12 The resulting peptides are separated by one or more dimensions of HPLC and detected by electrospray ionization (ESI) MS. To obtain quantitative information, the isotope-coded affinity tag (ICAT) labeling technique can be used to label the membrane proteins of two different sources.11 These earlier studies addressed the important issues related to protein solubilization/digestion, multidimensional LC separation, as well as quantitation. So far, ESI, due to its readiness to combine Journal of Proteome Research 2004, 3, 719-727

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research articles with MS/MS and LC separation, has been exclusively used for shotgun membrane proteomics applications. Here, we present a method to analyze the membrane proteome using a newly developed technique that combines LC with matrix-assisted laser desorption ionization (MALDI) MS/MS.13 We demonstrate that this LC/MALDI method offers unique advantages over LC/ESI and can analyze a large fraction of the membrane proteome that is not readily detected by existing techniques.

Experimental Section Chemicals and Reagents. R-Cyano-4-hydroxycinnamic acid (HCCA), bovine serum albumin (BSA), bovine trypsin, dithiothreitol (DTT), iodoacetamide, Triton X-100, trifluoroacetic acid (TFA), and sodium dodecyl sulfate (SDS) were purchased from Sigma-Aldrich Canada (Markham, ON, Canada). HPLC-grade acetonitrile and acetone were from Fisher Scientific Canada (Edmonton, Canada). Water was obtained from a Milli-Q Plus purification system (Millipore, Bedford, MA). HCCA was recrystallized from ethanol (95%) before use. Subcellular Fractionation of Adherent Cells. Adherent human HT 29 cells were used as a model system in this work. The HT29 cells were fractionated into cytosolic and membrane fractions by subsequent extractions with saponin and Triton X-100.14 Briefly, the cytosolic proteins were first released by 0.2% saponin lysis buffer containing 50 mM Tris-Cl pH 7.5 and 1 mM phenylmethyl sulfonyl fluoride (PMSF) at 4 °C. Membrane proteins were then extracted by buffer containing 1% TX100, 50 mM Tris-Cl pH 7.5, and 1 mM PMSF at 4 °C. Acetone Precipitation and Protein In-Solution Digestion. Triton X-100 can be removed from the membrane subcellular fraction with acetone precipitation.15 Acetone was pre-cooled at -80 °C and added to the protein extracts to a final concentration of 80% (v/v). After vortexing for 2 min, the mixture was kept cool at -20 °C for 4 h and then centrifuged for 10 min at 4 °C at 16 000 g to obtain the precipitate. The process was repeated with a small volume of acetone before the final pellet was collected and lyophilized for further experiments. For the in-solution digestion, 1 mg of lyophilized protein powder was dissolved in 17 µL 1% SDS (w/v) in 20 mM Tris. After the standard reduction and alkylation procedures,16 the mixture was diluted to a final SDS concentration of 0.1% and pH was adjusted with 100 mM NH4HCO3 to ∼8.5. Finally, 50 µL of 1 µg/µL bovine trypsin and 7 µL of 20 mM CaCl2 were added to the mixture, and the digestion was performed overnight at 30 °C. Additional SDS was added if precipitation was observed during any dilution steps. LC/MALDI MS. Peptide separation was performed on an Agilent (Palo Alto, CA) 1100 series capillary HPLC equipped with an auto sampler. Chromatographic analysis was performed with a reversed-phase 1.0 × 150 mm Vydac C8 column equipped with a 1.0 × 10 mm Nest Group SDS guard cartridge (Southborough, MA). A flow-rate of 40 µL/min was used for separation. Gradient elution was performed with solvent A (0.1%, v/v, aqueous TFA) and B (0.1%, v/v, TFA in acetonitrile). About 30 µg (equivalent to 8 × 105 cells) of peptide mixture was injected and UV wavelength was set at 210 nm. HPLC fractions were directly collected onto a 100-well MALDI target by the heated droplet interface.13 After the fractionation was completed, 1 µL of 50% acetonitrile/water (v/v) saturated by HCCA was added on top of each spot on the sample plate, and allowed to air-dry. The sample spots were then ready for both 720

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MALDI MS and MALDI MS/MS analysis. MS/MS experiments were performed on each peptide peak, starting from the most to least intense peaks and from high to low mass, until the sample was consumed. Usually about 20 MS/MS spectra may be collected from one sample spot. LC/ESI MS. 1D LC/ESI MS/MS was carried out on a ThermoFinnigan LCQ Deca ion trap instrument equipped with a Surveyer LC/MS system (San Jose, CA) and sample was loaded onto a 300 µm × 150 mm C18 capillary column with the same 1.0 × 10 mm SDS guard cartridge used in HPLC/MALDI experiment. For offline 2D LC/ESI MS/MS analysis, the sample was first separated by a Vydac 1.0 × 250 mm cation exchange column (8 µm particle size, 900 Å pore size) (solvent A: 20% Acetonitrile 0.1% TFA (v/v), solvent B: the same as A with 0.5 M NaCl) and then each fraction (1 fraction/minute) was injected on a 75 µm × 150 mm C18 column followed by analysis by a LCQ Deca XP instrument (San Jose, CA). SDS-PAGE and MS. SDS-PAGE was carried out in a Bio-Rad mini-Protein III system using 4%/12% stacking/separating polyacrylamide mini-gels. The SDS sample buffer contained 2% mercaptoethanol (v/v), 1% SDS, 12% glycerol, 50 mM TrisHCl and a trace amount of bromophenol blue. Prior to electrophoresis, the protein sample was mixed in the sample buffer and heated at 95 °C for 4 min. Visualization of protein bands was done using Bio-Rad’s Biosafe Coomassie blue stain reagent. A total of 36 protein bands were excised from the gel and digested with trypsin.16 Briefly, the gel pieces were cut into small segments and washed with fresh water for 20 min. After reduction and alkylation, gel pieces were dehydrated and covered with 10 ng/µL trypsin in 0.1 M NH4HCO3 and 2 mM CaCl2 for overnight digestion at 30 °C. Peptides were extracted 3 times in 60% acetonitrile in 0.25% TFA (volume: enough to cover the gel pieces) and once in 100% acetonitrile with 20 min of shaking each time. The pooled extracts were evaporated to approximately 1 µL. For the analysis of tryptic peptides from in-gel digestion, the two-layer sample preparation method was used for MALDI MS and MS/MS analysis of gel samples.17,18 Peptide mass mapping was performed using the Mascot search engine. The proteins identified from peptide mass mapping were confirmed by MALDI MS/MS. MS/MS experiments were also performed on the unmatched peaks after peptide mass mapping, similar to the HPLC/MALDI MS/MS experiment. Protein Identification from CID Data. Fragmentation information from CID spectra was first submitted to the sequence query program in Matrix Science (http://www.matrixscience.com) for possible identities with a precision tolerance of 0.3 Da for both the parent peptide and MS/MS fragments. These potential protein matches (usually individuals with scores at least close to homology region or better) were then put into the MS-product (http://prospector.ucsf.edu) program and the theoretical fragments were compared with experimental results, which may lead to definite identification of proteins. All the peptide identities were manually analyzed after database search. MALDI MS Instruments. MALDI MS experiments were carried out on a Bruker Reflex III time-of-flight mass spectrometer (Bremen/Leipzig, German) using the reflectron mode of operation. Ionization was performed with a 337-nm pulsed nitrogen laser. MALDI MS/MS experiments were carried out on a MDS Sciex QSTAR Pulsar QqTOF mass spectrometer equipped with an orthogonal UV-MALDI source (Concord, ON,

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Figure 1. Outline of a LC/MALDI MS/MS shotgun proteomics method and its workflow in relation to other existing methods for membrane proteome analysis.

Canada). All data were processed using the Igor Pro Software package (WaveMetrics, Lake Oswego, OR).

Results Figure 1 illustrates the general scheme of analyzing membrane proteome including the new technique of LC/MALDI MS/MS. Membrane proteins enriched from cells are best solubilized in a strong surfactant, SDS, which ensures that proteins are highly charged and, hence, not readily adsorbed or lost to the container wall and other apparatus during the sample workup process. In our work, we solubilized the proteins in 1% SDS. The SDS-solubilized protein mixture can be subjected to the gel-based method for analysis, as depicted in Figure 1. Alternatively or in parallel, a portion of the sample can be used for analysis using the solution-based shotgun proteomics approach. In this case, protein reduction and alkylation are first carried out, followed by 10-fold dilution of the protein solution to reduce the SDS concentration to 0.1%. Trypsin is then added to the solution for overnight digestion. To check the integrity of the membrane protein sample in each step leading to the MS analysis, we used MALDI TOF MS with a two-layer sample/matrix preparation method which can analyze protein and peptide samples containing up to 2% SDS. This method of sample evaluation has been found to be particularly beneficial in ensuring that each step in the sample workup works as expected. Because of the interference of SDS on MS analysis, great effort has been devoted to the removal of SDS. However, removal of SDS often results in the loss of hydrophobic or low abundance proteins.19,20 It would therefore be preferred and more practical to develop a MS method that can tolerate the presence of SDS. In this work, the SDS-dissolved HT29 mem-

brane protein mixture was trypsin-digested in the presence of 0.1% SDS. We used the two-layer sample preparation method to obtain a MALDI MS spectrum of the digested protein mixture (see the Supporting Information, Figure S1). From the MS spectrum, we concluded that the tryptic digestion was effective. The spectrum consists of many peptide peaks, but peptide signal suppression was very severe in the direct analysis of the mixture. A separation of the complex peptides digest mixture is therefore required and essential prior to MS detection to ensure that good ion yields are produced for further MS and MS/MS experiments. In this work, HPLC separation of the SDS-containing tryptic digest was done with the use of a SDS guard column added to a conventional HPLC setup. The performance of this guard column was preexamined with a tryptic digest of BSA standard containing 0.1% SDS (data not shown); the use of a SDS guard column produced a significant improvement in the HPLC separation efficiency. In the case of the HT29 membrane protein digest, Figure 2A shows the UV chromatogram obtained from this sample. Figure 2B-D shows MALDI MS spectra of three HPLC fractions at elution times 23, 42, and 54 min, respectively, corresponding to the fractions labeled with asterisks in Figure 2A. The peptide peaks are mainly from the protonated molecular ions with little or no peptide-Na+ adduct ions, indicating that the LC separation is effective to reduce the salt content in the fractions. In contrast to the whole-digest spectrum, the spectra of Figure 2B-D are less complex. However, there are still many peptides detected in each fraction. This is not surprising in light of the fact that the membrane proteome from HT29 is expected to be highly complex. The heated droplet LC/MALDI system allows the use of microbore column for separation.13 Thus, the total amount Journal of Proteome Research • Vol. 3, No. 4, 2004 721

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Figure 2. (A) Reversed-phase HPLC chromatogram of the tryptic digest of SDS-solubilized membrane protein mixture. MALDI MS spectra obtained from HPLC fractions of (B) 23 min, (C) 42 min, and (D) 54 min.

of peptides injected to HPLC is much larger compared to any capillary LC work, which ensures that a sufficient number of peptides in each fraction have high abundance for both MS and MS/MS analysis. It should also be noted that, in the heated droplet interface, the HPLC fractions are directly concentrated and spotted onto the MALDI target and no vials or further drying steps are needed. Thus sample loss is minimized. This is very helpful in the analysis of hydrophobic peptides where adsorption on the vials during the drying step can be very severe if the fractions are collected in vials followed by solvent evaporation (data not shown). For MALDI analysis, we used a combination of MALDI TOF and MALDI QqTOF instruments to obtain MS and MS/MS spectra. We found that MALDI MS experiments with a conventional MALDI TOF instrument (Bruker Reflex III) should be carried out to obtain the correct tryptic peptide mass information before QqTOF analysis. This is because in-source fragmentation is often observed in the MALDI QqTOF instrument 722

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in MS mode when HCCA is used as the matrix.21-23 These fragmented peptides are not tryptic peptides and will fail a database search if treated as such. Other matrixes may be used, but the MS/MS sensitivity is not as good as HCCA in this instrument. In the LC/MALDI MS/MS experiment, peptide sequencing was performed sequentially from the most to least intense peaks and from high to low mass until the sample spot was consumed or no further peptides could be selected. Generally, up to 20 MS/MS spectra can be obtained from one LC fraction and the actual number of spectra collected per fraction is dependent on the sample amount and complexity on the spot. Figure 3 shows two representative MALDI MS/MS spectra obtained from the HPLC fraction at 50 min. Two peptide sequences AQFEGIVTDLIR and YFPTQALNFAFK were identified based on these MS/MS spectra respectively and led to the identification of stress 70 protein (P38646) and ADP, ATP carrier protein, either from P05141 or P12235, which was further identified as P05141 by additional peptides detected and sequenced from other fractions. A total of 157 peptides from 73 proteins were identified, as shown in Supplementary Table 1. This demonstrates that the developed technique of solubilizing membrane proteins with SDS, digesting tryptically in SDS followed by auto-offline LC/MALDI MS and MS/MS is efficient and successful. Fifty-three of the seventy-three identified proteins were membrane or membrane associated proteins, indicating this approach is very useful in membrane protein identifications. For comparison, LC/ESI MS/MS was also used to examine the HT29 membrane protein digest. However, in ESI MS, the nonvolatile surfactant SDS severely destroys the analyte signals.24-28 Hydrophobic proteins or peptides are usually solubilized in organic solvents for ESI analysis.29-32 Several groups have reported their studies on the effect of SDS on ESI signals. For examples, Ogorzalek Loo and co-workers examined the effect of SDS on ESI by flow injection and showed that with 0.01% and 0.1% SDS in 6 pmol/µL myoglobin, 10% of the original myoglobin signal was retrieved.28,33 Kirby and coworkers obtained ESI signals from 150 mM peptides with 1.25 mM (0.04%) SDS using 1.25 mM Genapol C-100 to shield the surfactant’s effect.34 Vissers, Salzmann, and co-workers reported that they successfully obtained LC/MS/MS data from 8 pmol cytochrome c tryptic digest in the presence of 0.1% SDS using a SDS removal cartridge coupled with a LC column using a triple quadruple ESI instrument.35 More recently, Hixson and co-worker analyzed bacteriorhodopsin peptides by LC/ESI MS and MS/MS after digestion in 0.015% SDS and overnight dialysis.36 Han and co-workers successfully analyzed a tryptically digested protein mixture containing less than 0.05% SDS by a 2D LC/ESI MS/MS method.11 In our work, when the SDS-containing digest from HT29 cells was analyzed by using the LCQ Deca ion trap mass spectrometer equipped with a Surveyer LC system, a 300-µm i.d. reversed-phase column, and a SDS removal guard column, only 10 peptides were detected. This poor performance is mainly due to the fact that the SDS-removal guard column is not 100% efficient and the remaining SDS coeluting with the analyte can still severely interfere with the analyte signal.24,34 We then tried to use a Millipore HPL hydrophobic or C18 ZipTip to remove the SDS; we used poly(ethylene glycol) lauryl ether to reduce the SDS suppression in ESI;34 and we tried to load various sample amounts into the capillary column. All attempts failed to produce any improvement.

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Figure 3. MALDI MS spectrum of HPLC 50 min fraction (A), and MALDI MS/MS spectra of peptides 1446.6 Da (B) and 1361.7 Da (C).

To determine the reason for the low number of proteins identified by ESI MS, another control experiment was performed. The chromatograms from the ESI analysis on 8 and 0.8 pmol cytochrome c tryptic peptides with or without SDS are shown in the Supporting Information, Figure S2. In the analysis of the 8-pmol cytochrome c tryptic peptide containing no SDS, twelve peptides were identified. With 0.1% SDS added into the digested sample and SDS removal guard cartridge coupled onto the LC column, the intensity of base peak chromatogram decreased, especially for the hydrophobic peptides which were eluted at high acetonitrile composition and only five peptides were identified. For the 0.8-pmol digest, 3 peptides were observed with high intensity in the absence of SDS in the sample. However, when 0.1% SDS was present in the sample, no peptide peaks were identifiable. Thus, as the protein (and peptide) concentration decreases, the suppression effect from SDS becomes dominant and deteriorates ESI MS and MS/MS signals. For a HT 29-cell extract which may contain hundreds of proteins, even though the total protein concentration is high, the individual protein concentration in the mixture can be very low. As a result, the presence of SDS may have a

strong negative effect on ESI analysis and only a few peptides from the high abundant proteins were identified by using the 1D ESI MS and MS/MS approach. To further study this sample by ESI, a 2D LC separation method coupled with ESI MS and MS/MS was applied. The same amount of sample as in the LC/MALDI experiment was used for the 2D LC/ESI experiment. The 2D separation was achieved by a cation exchange fractionation followed by reversed-phase separation. In cation exchange, cations are adsorbed onto the resin until eluted by a salt gradient. Most peptides are eluted with a salt strength of 0.1-0.2 M NaCl. Therefore, for a peptide mixture containing 0.1% SDS (3 mM), the sodium amount in this sample itself (3 mM) can be ignored compared to the sodium concentration in the mobile phase. Figure 4A shows the UV chromatogram of the cation exchange (IE) separation of the tryptic digest from 30 µg of membrane protein mixture. Eighteen fractions (2 to 20 min) were collected in 1 min intervals, and each fraction was individually injected onto a reversed-phase column followed by ESI MS/MS analysis. The reversed-phase LC separation base peaks of IE fractions at 6 and 11 min were shown in Figure 4B. The corresponding Journal of Proteome Research • Vol. 3, No. 4, 2004 723

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Figure 4. (A) UV chromatogram of the cation exchange separation of the tryptic digest from the membrane protein mixture. (B) LC/ESI base peaks and (C) selected peptide MS/MS spectra from fractions 6 and 11 min, respectively.

MS/MS spectra were recorded and used to search for protein identities using the Mascot engine (Figure 4C). In this approach, a total of 51 peptides were detected, leading to the identification of 37 unique proteins. Among the proteins identified, 20 had already been identified by the LC/MALDI MS and MS/MS analysis and 17 were newly identified with this 2D LC/ESI approach. The proteins identified by 2D LC/ESI are also listed in Supplementary Table 1. Finally, a comparison experiment using the same membraneenriched protein sample was performed using the in-gel approach. In this experiment, one-dimensional SDS-PAGE was used to separate the membrane protein sample to avoid the possible solubilization problems often encountered in the rehydration step in 2D gel separation.5,37,38 The same amount of sample injected for the LC/MALDI and LC/ESI experiments, 30 µg of protein extract, was mixed with SDS sample buffer and loaded onto a 1D gel. The coomassie blue stained gel is shown in Figure 5A. All visible gel bands were excised, in-gel tryptically digested, and identified with MALDI MS and MALDI MS/MS. It should be noted that the total amount of proteins loaded to the gel is relatively small and many observed bands may only contain a very small amount of protein or proteins. It is our experience that, with this type of sample at this level of sample loading, LC/ESI MS/MS of in-gel digest from a band generally does not give as good results as that obtained by direct MALDI MS and MS/MS, which is more sensitive. In our lab, we have been using a strategy of result-dependent data collection and analysis for protein identification from in-gel digests. As an example, Figure 5B shows the MALDI MS spectrum of the tryptic peptides digested from the gel band 724

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indicated with the arrow in Figure 5A. All strong peptide peaks were loaded into the database for protein identification by peptide mass mapping. The unmatched peptide peaks from peptide mass mapping were sequenced by MALDI MS/MS for possible identification of other proteins in the band. In this particular gel band, protein ATP synthase B chain, P24539, was first tentatively identified by peptide mass mapping and all peptides that may belong to this protein were marked as “1” in Figure 5B. This identification was confirmed by the MALDI MS/MS spectrum of a peptide HYLFDVQR (m/z 1077.5) specific to protein ATP synthase B chain as depicted in Figure 5C. For the unmatched peaks, MS/MS sequence query was used for protein identification. For example, Figure 5D is a MALDI MS/ MS spectrum from an unmatched peptide from peptide mass mapping results, which led to the identification of the protein “membrane associated progesterone receptor component 1 (mPR)”, O00264. A theoretical trypsin digest was performed on this newly identified protein and one of the unmatched peptides that matches the newly identified protein (from peptide 1644.8 Da) was subjected to MALDI MS/MS sequencing and identification, followed by labeling all peptides that belong to the second protein as “2”. By using this strategy of resultdependent data collection/analysis, fewer samples are consumed to identify more proteins present in this band. This process was repeated several times until sample was consumed or no more new protein could be identified. By this gel approach, 45 proteins are identified and among them 33 proteins are membrane or membrane-associated proteins and 12 are cytoplasmic proteins (see Supplementary Table 1).

Discussion Supplementary Table 1 lists the proteins identified from this membrane protein fraction of HT29 cells using the in-gel, LC/ MALDI, and 2D LC/ESI methods. Figure 6 summarizes the results obtained from the three methods. As Figure 6A shows, a total of 112 proteins were identified. The low number of proteins identified from this sample is mainly due to the small amount of starting materials used for the experiments as well as the nature of the sample, i.e., a highly enriched membrane protein fraction containing many hydrophobic proteins that may not be digested by trypsin effectively compared to watersoluble proteins in other samples. Among the 112 proteins identified, LC/MALDI identified 73 proteins, compared to 37 from 2D LC/ESI and 45 from the gel-based method. There were proteins identified by more than one method, as well as a number of proteins identified by a single method. This finding is not surprising considering the significant differences in experimental conditions used among the three methods. It is clear that for comprehensive proteome profiling, these methods complement each other. One significant observation is that, with 1D LC/MALDI, a greater number of proteins were identified compared to the other methods. This difference is even more striking when only the membrane proteins are considered. Among the 112 proteins identified, 76 proteins are membrane and membrane-associated proteins including 28 integral membrane proteins. Figure 6B illustrates that 50 of the 76 membrane proteins were identified by LC/MALDI vs 24 from 2D LC/ESI and 33 from the in-gel method. For the 28 integral membrane proteins (Figure 6C), 22 were identified by LC/MALDI, 7 were found by 2D LC/ESI, and 7 were detected by the in-gel method. The challenge of detecting membrane proteins, particularly integral membrane proteins, is well documented1,5 and thus the

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Figure 5. (A) SDS-PAGE gel image of separated membrane proteins. (B) MALDI MS spectrum of tryptic peptides from gel band marked with arrow in A. MALDI MS/MS spectra of peptides 1077.5 Da (C) and 1644.8 Da (D) along with their sequences and the protein Ids.

identification results from the in-gel method are not unexpected. But it is surprising to discover that there is a great difference in membrane protein detection ability between the ESI and MALDI proteomic methods. This difference may be attributed to several factors including the possibility of higher efficiency of MALDI in ionizing more hydrophobic peptides and the concentration effect during LC fractionation in LC/MALDI that enriches the peptides cleaved from low abundance membrane proteins and thus increases the probability of their detection. As Figure 6 shows, ninety proteins were identified with insolution approach while only 45 proteins were identified with in-gel approach. Proteins only identified by the in-gel approach might be a result of digestion bias by in-solution digest. Proteins in solution can have very different concentrations and some may be much easier to digest than others. The relatively low abundance proteins or difficult-to-digest proteins may not be able to compete for trypsin effectively with those proteins with high concentrations or easier-to-digest ones. As a result, digestion is not complete for some proteins, preventing them from identification by peptide analysis. However, in the in-gel approach, proteins are first separated and localized in different positions of the gel. Each protein, to a certain degree, is “enriched” in its own molecular weight region for digestion. Higher protein concentration generally results in better diges-

tion. The proteins in the gel band are also relatively free from surfactants or other impurities that may interfere the digestion, compared to the solution digestion containing SDS. There are several possible reasons for detecting more proteins in the in-solution methods. First, hydrophobic proteins might not be fully denatured by the gel sample buffer, or the conformation might be restored during the sample loading process. As a result, these proteins may not migrate into the gel or only a small portion of the total amount of these proteins is actually being separated. This might be the reason a dark band was observed at the top of the separating gel in the sample lane, while there is nothing in the same region in the protein standard marker lane (see Figure 5A). Second, in the in-solution identification approach, proteins are not only solubilized by SDS but also digested in the presence of SDS. SDS will maintain proteins in the denatured condition during the entire digestion process which in turn aids the enzyme to work on the proteins, especially hydrophobic proteins. This is opposite to the in-gel digestion process where extensive washes are applied to remove SDS and other contaminants. Third, extraction of peptides from gel pieces after in-gel digestion, especially those hydrophobic peptides, may not be very efficient and adsorption on vial walls of these peptides can also be a severe problem, while these factors are not present in this developed in-solution technique. Finally, comigrated hydroJournal of Proteome Research • Vol. 3, No. 4, 2004 725

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Figure 6. Schematic graph showing the overlap of proteins identified by solution-based LC/MALDI MS/MS method, solutionbased 2D LC/ESI MS/MS, and in-gel MALDI MS/MS. (A) A total of 112 proteins were identified. Among them, (B) 76 are membrane and membrane-associated proteins, which include (C) 28 integral membrane proteins.

philic proteins in a gel band can suppress the digestion, extraction and detection of hydrophobic peptides,39,40 leading to the failure to identify these hydrophobic proteins. Although LC-MS/MS after in-gel digestion can minimize the detection bias problem, the sample amount is often limited from the ingel digest and LC-MS/MS can still have difficulty analyzing the low abundance peptides after separation. One protein identified in this in-solution method, ADP/ATP carrier protein (SwissProt P05141) can be used as an example for the above discussion. This protein is an integral membrane protein, has a molecular weight of 32.9 kDa, and contains 6 trans-membrane regions. Six peptides including three in the transmembrane regions were detected by the in-solution method which conclusively identified this protein. Although this protein was not identified in the gel experiment, we found two small peptide peaks that match the masses of ADP/ATP carrier protein’s theoretical tryptic digest after careful examination of the MALDI MS spectra from the band with molecular weight around 33 kDa. These two peptides may belong to this membrane protein, but they were too weak to obtain good MS/ MS spectra for a positive identification. Again, as a hydrophobic membrane protein, it might partially migrate into the gel with not enough quantity for a successful digestion. Trypsin may not successfully target this membrane protein and the extraction and detection of hydrophobic peptides are problematic as well. More interestingly, the band at the 33 kDa region which 726

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gave the two small peaks that may belong to this membrane protein was identified to be Prohibitin (Swiss-Prot P35232), a very hydrophilic protein. The presence of this protein in the band may hinder the digestion, extraction, and detection of the hydrophobic protein. At present, proteome data generated by three protein identification methods are still very small in scope. Future work of membrane proteome analysis involving in the use of these methods will surely increase the number of proteins identified, which should facilitate data comparison and understanding of the reasons underlying the difference of protein identification results generated by the three methods. For now, it appears that the three methods can provide complementary data for proteome analysis. For LC/MALDI, this present work involves the use of 1D LC for peptide separation. More efficient peptide separation with, for example, 2D LC, should decrease the complexity of the peptides in each fraction and thus increase the total number of detectable peptides. In conclusion, we have demonstrated a new proteome analysis platform for membrane proteomics research. It involves the use of SDS in sample workup and digestion to facilitate membrane sample handling and minimize sample loss, and the use of shotgun proteomics methods based on LC/ MALDI MS/MS as well as 2D LC/ESI MS/MS, and the in-gel approach to generate a more comprehensive proteome map. It is shown that the LC/MALDI shotgun proteomics method offers several unique advantages over LC/ESI MS/MS such as compatibility with SDS-containing digest and more extensive detection of integral membrane proteins. This work illustrates that, for comprehensive analysis of membrane proteomes of interest, a preferred proteomics approach should incorporate all the three technologies.

Acknowledgment. This work was funded by the Natural Science and Engineering Research Council of Canada and the Alberta Cancer Board. Supporting Information Available: A list of proteins identified from this membrane protein fraction of HT29 cells using the in-gel, LC/MALDI, and 2D LC/ESI methods (Supplementary Table 1). MALDI MS spectrum of tryptic digest of HT29 membrane protein mixture with 0.1% SDS present (Supplementary Figure S1). Ion chromatograms of cytochrome c tryptic digest with and without the presence of 0.1% SDS (Supplementary Figure S2). These materials are available free of charge via the Internet at http://pubs.acs.org. References (1) Wu, C. C.; Yates J. R., III Nat. Biotechnol. 2003, 21, 262-267. (2) Rabilloud, T. Nat. Biotechnol. 2003, 21, 508-510. (3) Shaw, A. R.; Li, L. Curr. Opin. Mol. Therapeutics 2003, 5, 294301. (4) Henningsen, R.; Gale, B. L.; Straub, K. M.; DeNagel D. C. Proteomics 2002, 2, 1479-1488. (5) Santoni, V.; Molloy M.; Rabilloud T. Electrophoresis 2000, 21, 1054-1070. (6) Simpson, R. J.; Connolly, L. M.; Eddes, J. S.; Pereira, J. J.; Moritz, R. L.; Reid, G. E. Electrophoresis 2000, 21, 1707-1732. (7) van Montfort, B. A.; Doeven, M. K.; Canas, B.; Veenhoff, L. M.; Poolman, B.; Robillard, G. T. Biochim. Biophys. Acta 2002, 1555, 111-115. (8) Quach, T. T. T.; Li N.; Richards, D. P.; Zheng, J.; Keller, B. O.; Li, L. J. Proteome Res. 2003, 2, 543-552. (9) Washburn, M. P.; Wolters, D.; Yates, J. R., III Nat. Biotechnol. 2001, 19, 242-247. (10) Blonder, J.; Goshe, M. B.; Moore, R. J.; Pasa-Tolic, L.; Masselon, C. D.; Lipton, M. S.; Smith, R. D. J. Proteome Res. 2002, 1, 351360.

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