Special Issue Papers |
Comparison of a Salmonella typhimurium proteome defined by shotgun proteomics directly on an LTQ-FT and by proteome pre-fractionation on an LCQ-DUO
Corresponding author. David R. Goodlett, Medicinal Chemistry Department, University of Washington, Box 335351, Seattle, WA 98195, USA. E-mail: goodlett{at}u.washington.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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Shotgun proteomics is rapidly becoming one of the most efficient and popular tools to examine protein expression in cells. Numerous laboratories now have a wide array of low- and high-performance mass spectrometry instrumentation necessary to complete proteome-wide projects. Often these laboratories have time and financial constraints that prohibit all projects from being conducted on high-performance state-of-the-art mass spectrometers. Here, we compare shotgun proteomic results using a direct ‘lyse, digest and analyse’ approach on a high-performance mass spectrometer (i.e. the LTQ-FT) with the results from a much lower-performance instrument (i.e. the LCQ-DUO) where, for the latter, various traditional protein pre-fractionation steps and gas-phase fractionation were used to increase the proteome coverage. Our results demonstrate that shotgun proteomic analyses conducted on the lower-performance LCQ-DUO mass spectrometer could adequately characterize a PhoP constitutive strain of Salmonella typhimurium if proteome pre-fractionation steps and gas-phase fractionation were included.
Keywords: shotgun proteomics, ion trap mass spectrometer, Salmonella, linear ion trap and Fourier transform ion cyclotron resonance mass spectrometer, membrane proteome
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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Shotgun proteomics, also generally known as non-polyacrylamide-based proteomics, involves the direct analysis and identification by mass spectrometry of peptides resulting from enzymatic digestion of a complex mixture of proteins in order to characterize the proteins of a cell. The emergence of this technique to identify and verify the expression of numerous proteins from a complex mixture of peptides represents a biochemical breakthrough, that rapidly improved the understanding of biological states [1–4]. As an analytical technique, shotgun proteomics has a great amount of potential; already, since its emergence in the 1990s, it has been used for the identification of previously uncharacterized proteins [5], understanding biological functions (e.g. cell death, virulence), biological characterization [6], genomic verification [7], drug discovery [8], biomarker discovery [9] and post-translational modifications [10]. The technique is rapidly being adopted by a wide variety of laboratories because of the great amount of biological information that can be gained by this single approach.
There are numerous types of mass spectrometers available to researchers who wish to conduct shotgun proteomics. For the identification and sequencing of large proteomes, the mass spectrometer is the limiting step as it has to analyse, isolate, fragment and record data from a dynamic sequence of consecutive co-eluting peptides from the chromatography column. Although constant improvements in mass spectrometry technology are rapidly expanding the capabilities for large shotgun proteomic experiments [3], it is unreasonable to expect individual laboratories to keep up with the financial burden of constantly upgrading instrumentation. Even well-established proteomic laboratories with top-end, high-performance mass spectrometers must economically manage the time of these instruments because they are always in higher demand than lower-performance instruments in the same laboratory. Here, we examined how many layers of protein fractionation are required to get equivalent proteome results from an economical, more readily available, lower-performance ion trap mass spectrometer with an ion scan rate one-fifth the speed (i.e. duty cycle) of the highest performance mass spectrometer in our laboratory.
This manuscript evaluates the use of two very different mass spectrometers to define the Salmonella typhimurium PhoP constitutive proteome: an advanced hybrid linear ion trap-Fourier transform-ion cyclotron resonance (FT-ICR) mass spectrometer (LTQ-FT, Thermo Electron; San Jose, CA) and an older quadrupole ion trap (LCQ-DUO, Thermo Electron; San Jose, CA). The linear ion trap of the LTQ-FT allows greater ion capacity and dynamic range than the prior generation of ion traps embodied by the LCQ-DUO, has two electron multipliers to enhance sensitivity and scans approximately five times faster than the LCQ-DUO [11]. Although the FT-ICR cell yields better mass accuracy, making this an excellent instrument selection for large shotgun proteomic experiments, it is a costly alternative to more commonly available mass spectrometers and can be more difficult to maintain. From recent price estimates, an LTQ-FT can cost up to 20 times more than a readily available second-hand LCQ-DUO available from used instrument vendors. To new investigators, this financial aspect of proteomics, along with the fact that many established laboratories already have LCQ-DUO or equivalent instruments, encourages the question, ‘Can we even compete?’ using older technology.
To increase proteome coverage with the older ion trap mass spectrometer, we employed the use of some traditional pre-fractionation techniques. We then compared proteomic results from a mutant variety of a common Gram-negative bacterium using the combined efforts of protein pre-fractionation and peptide-ion gas-phase fractionation (GPF) on the LCQ-DUO to whole-cell analyses on a state-of-the-art LTQ-FT. In addition to increasing protein identifications from the whole proteome and improving protein sequence coverage using the shotgun proteomic technique, we also explored improving membrane proteome coverage.
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CELLULAR PREPARATION, PROTEIN ISOLATION, DIGESTION AND ANALYSES |
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TOPABSTRACTINTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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Cellular preparation
Salmonella typhimurium PhoP constitutive (CS022) mutants were used in this study [12]. Cells were grown in Luria-Bertani broth to late-log phase (OD600
0.9) before being harvested. Salmonella cell pellets were resuspended in 10 mM Tris (pH 7.5), 5 mM MgCl2, 1 mM dithiothreitol (DTT), DNAse and 2 mg lysozyme. Cells were lysed using a 100 W MSE 20 kHz sonicater (MSE Ltd, Crawley, Surrey, UK) with a titanium micro-tip (3 x 10 s, 4°C). Unlysed cells were removed by centrifugation (4700 x g 10 min), resulting in a whole-cell lysate. To isolate the membrane fraction from lysed cells, the whole lysate was centrifuged (17 000 x g, 30 min) to pellet membranes and the supernatant was removed. Several protein fractionation steps were completed to improve proteome coverage prior to analysis with the LCQ-DUO (Figure 1) and are discussed in detail subsequently.
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Trypsin digestion and mass spectrometry
Protein pellets were first solubilized in 300 µl of 6 M urea, followed by the addition of 20 µl of 1.5 mM Tris (pH 8.8) to maintain an alkaline pH while the sample was brought to a final concentration of 5 mM tris-(2-carboxylethyl)phosphine (TCEP) (37°C, 1 h). Disulphide bonds were reduced with dithiothreitol (DTT), alkylated with 60 µl of 200 mM iodoacetamide (IAM) and vortexed and stored in the dark for 1 h (25°C); excess iodoacetamide was neutralized with 60 µl of 200 mM DTT for 1 h (25°C). Each sample (150 µl) was aliquoted into three tubes and 800 µl of NH4HCO3 was added to dilute the urea prior to the addition of 200 µl of MeOH and sequence grade trypsin (Promega, Madison, WI) at 50:1 substrate:enzyme(w/w). Trypsin digestions were vortexed and incubated overnight at 37°C. Samples were then taken to near dryness in a speed-vac and pooled together. To reduce the NH4HCO3, 200 µl of MilliQ-H2O was added to each tube three times and evaporated. Samples were stored at –80 C until mass spectrometry (MS) analysis.
Just prior to all MS analysis, trypsin digestions were desalted using a micro-spin C18 column (NestGroup) following the manufacturers guidelines. The final eluted sample was then evaporated to a small volume in a speed-vac and resuspended in 5% ACN containing 0.1% formic acid (v/v). Samples were analyzed on either the LCQ-DUO or LTQ-FT with a 11 cm long, 75 µm i.d. silica capillary column packed with C18 particles (Magic C18AQ, 100 A, 5 µ. Michrom, Bioresources, Inc., CA) fitted with a 2 cm long, 100 µm i.d. pre-column (Magic C18AQ, 200A, 5u. Michrom) following the procedures outlined in Zhang et al. [13]. All samples were analysed a minimum of two times (A: H2O, 0.1% formic acid, B: ACN, 0.1% formic acid; gradient: LCQ-DUO 10–60% in 40 min and LTQ-FT 5–35% in 60 min). All mass spectral results for this manuscript were interpreted and searched with SEQUEST [14] with the S. typhimurium [LT2] genome annotation (National Center for Biotechnology Information, NCBI). A false-positive rate of <1% was estimated using the PeptideProphet tool [15].
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SIMPLIFYING SHOTGUN PROTEOMICS WITH AN EXPENSIVE TOOL, THE LTQFT |
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TOPABSTRACTINTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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Triplicate LC-MS/MS analyses were completed on the whole-cell lysates of S. typhimuirium on an LTQ-FT. Survey scans were completed in the FT-ICR cell while MS/MS data-dependent scans were acquired in the linear ion trap. From this analysis, 4693 peptide tandem mass spectra were matched to sequences in a database, yielding 655 unique protein identifications with >0.99 probability. The average protein sequence coverage was 29%. Based on the cellular location prediction model PSORT (www.psort.org), the identified proteins were 64% cytosolic, 9% periplasmic and 27% membrane-associated (Figure 2A, Table 1). Over 14% of the known S. typhimurium proteins were identified from triplicate LC-MS/MS runs of a whole-cell tryptic digest on the LTQ-FT.
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USING PROTEOME FRACTIONATION TO IMPROVE LCQ-DUO RESULTS |
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TOPABSTRACTINTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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What if your laboratory does not have access to an LTQ-FT? We asked this question because even in our laboratory where an LTQ-FT operates nearly continuously, not all of our samples can be analysed by this instrument. Here, we show results that answer the question how much more effort is involved in characterizing a simple bacterial proteome on the LCQ-DUO compared with the LTQ-FT. As with the LTQ-FT, the LCQ-DUO has data dependent (DD) ion selection, selecting the most intense peaks for MS/MS from the full m/z range (e.g. 400–1800), and dynamic exclusion which minimizes redundancy of the MS/MS data by excluding previously selected precursors, ions, from being repeatedly interrogated by MS/MS. The effect of the latter allows the DD process to dig deeper into the dynamic range of the mass spectrum, and hence, expands the dynamic range of the ions selected for MS/MS. This is relevant because in a typical experiment, thousands of peptides resulting from a whole-cell lysate are chromatographically ‘separated’ prior to MS analysis but are rarely baseline resolved. Thus, the scan rate of the instrument defines the sampling efficiency of the mixture, and thus effects the observed proteome coverage. In the case of the LCQ-DUO, the duty cycle or scan rate for MS/MS event is approximately one-fifth that of the LTQ in the LTQ-FT. As a result of this difference in duty cycle, in any single LC analysis, the LCQ-DUO will identify fewer proteins. We will ignore in our subsequent discussions issues of ‘sensitivity’ because this is a much abused and often ill-defined term. When a complex mixture is being ‘under-sampled’ by the DD process, ion selection within the mass spectrometer is a somewhat random process and two runs of an identical sample may result in different results in terms of the unique peptides and proteins identified. As the scan rate for MS/MS is increased, and coupled with dynamic exclusion, the ability to more completely sample the available ions at a given point in time is approached. Thus, this desirable attribute of tandem mass spectrometers has huge implications for a laboratory's ability to define an organism's proteome.
Prior to starting this work, we expected that some of the disadvantages of the LCQ versus the LTQ-FT could be overcome, in part, by protein fractionation prior to MS analysis. To simplify the analysis of a complete cell lysate, there are a variety of pre-fractionation steps that can be performed to either reduce the total number of proteins analysed via shotgun proteomics or to isolate a subset of proteins for shotgun analyses.
Dropping out of solution—revisiting (NH4)2SO4 precipitations
One of the most readily available and straightforward fractionation steps that can be completed on whole-cell lysates is by ‘salting-out’ of proteins using ammonium sulphate [i.e. (NH4)2SO4]. The method works by increasing the ionic strength of the solution slowly to a given point [i.e. % (NH4)2SO4] where proteins begin to aggregate and precipitate. In theory, this works because proteins with a greater number of (or larger) hydrophobic moieties will precipitate out at lower (NH4)2SO4 concentrations, resulting in a gradual fractionation as salt saturation increases.
Ammonium sulphate was used in this study to fractionate because it is readily available, inexpensive and allows for the Bradford assay to be performed with no additional ‘clean-up’ steps. Saturated (NH4)2SO4 (0°C) solution was added to 1 ml of whole-cell lysate to yield 30% (NH4)2SO4. The sample was then mixed on a vortex device and maintained at 4°C for 1 h prior to centrifugation (17000 x g, 30 min). The supernatant was set aside and the resulting pellet from the 30% (NH4)2SO4 precipitation was lightly dried in the speed-vac and stored at –80°C prior to trypsin digestion and MS analysis. Subsequent additions of saturated (NH4)2SO4 were added to the supernatant to yield 50, 70 and 90% (NH4)2SO4.
The combined results of the four (NH4)2SO4 fractions analysed in a total of eight LC-MS/MS runs on the LCQ-DUO led to 817 peptide tandem mass spectra with confident matches to the S. typhimurium proteome (probability >0.99). A total of 255 proteins were identified from these experiments with an average protein sequence coverage of 20.7% (Table 1). The distribution of proteins identified from different cellular localizations was similar to those identified with the LTQ-FT (Table 1, Figure 2B; 61% cytosolic, 28% membrane and 11% periplasmic). (NH4)2SO4 fractionation yielded a total of 6% of the S. typhimurium proteome, corresponding to 26% of the total proteins identified with the LTQ-FT (Figure 3A). Analysis of these fractions identified 20 unique proteins that were not identified in the whole-cell lysate analysed on the LTQ-FT (Figure 3B). However, given the unpredictable character of ion selection for MS/MS by DD methods that results in under sampling, we cannot state that these protein identifications are a result of the (NH4)2SO4 fractionation itself.
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A closer examination of the proteins identified in the four (NH4)2SO4 precipitations reveals redundancy in proteins detected between fractions and that the (NH4)2SO4 precipitation process is not an effective means of protein fractionation. Of the 255 proteins identified, only 133 proteins were unique to one of the four (NH4)2SO4 concentrations; 122 proteins were observed in two or more of the (NH4)2SO4 fractions and 15 proteins were identified in all four (NH4)2SO4 fractions (Figure 3C). Interestingly, there was no statistical difference in the cellular distribution or the isoelectric points (pIs) of the four (NH4)2SO4 fractions (Figure 4A, C, Table 1). This was clearly a disappointing result; we had hoped to be able to use this method to effectively separate protein mixtures. The results of this type of fractionation, however, may be different for more complicated mammalian cells.
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Gaining an advantage with gas-phase fractionation
As mentioned earlier, one of the primary limitations of shotgun proteomics is the DD scan rate of the mass spectrometer. Given the pace of change in mass spectrometry technology, it is common for even advanced proteomic laboratories to have older models, like the LCQ-DUO, that have much lower performance specifications than the newer models. The primary shortcoming of the LCQ-DUO compared with the LTQ is the slower MS/MS scan rates for DD ion selection during high pressure liquid chromatography (HPLC) introduction of a sample. Because of the slower scan rate that can result in thousands of missed opportunities for MS/MS during LC sample introduction, scientists in our laboratory with open access to both instruments choose the LCQ only as a last resort. Recently, it has been shown that the exploitation of on-line GPF of peptide ions introduced by LC to a tandem mass spectrometer operating in DD ion selection mode yielded greater proteome coverage of complex mixtures [16,17]. GPF directly expands the dynamic range of the selected peptides for MS/MS by limiting the DD sampling to smaller, discrete m/z windows of the mass spectrum. This is accomplished by performing repeat analyses of the sample across several, often overlapping, narrow m/z ranges (e.g. 500–600) rather than one wide m/z range (e.g. 400–2000). As a method of sample fractionation, GPF avoids the loss of proteins or peptides from ‘over-handling’ of a sample, is easily implemented on all commercially available tandem mass spectrometers equipped with some form of DD operation, and is easily automated. Unlike the popular MudPIT method [18], no special columns are required; the only requirement for GPF is that the sample not be limited. This could be argued to be a severe limitation for some applications, but for many of our in vitro proteome profiling projects, it was not an issue. Therefore, we investigated the use of GPF on the LCQ to catalogue proteins from (NH4)2SO4 precipitation of a whole-cell S. typhimurium lysate to determine if we could increase the number of proteins identified or the amount of individual protein sequence coverage.
The same fractions from (NH4)2SO4 precipitation of the S. typhimurium described earlier were analysed using GPF on the LCQ-DUO in duplicate. GPFs were completed with four narrow m/z ranges: 600–1000, 900–1250, 1200–1650 and 1500–2000, resulting in a total of 40 LC-MS/MS analyses for the four (NH4)2SO4 fractions.
Rather than a single wide m/z range scan, the use of narrow m/z scan ranges with GPF positively identified 1.8x more proteins (>0.99 probability), a total of 472 unique proteins, with little additional effort because the runs were entirely automated. 1918 peptide tandem mass spectra were matched to sequences in the S. typhimurium database, providing an average protein sequence coverage of 26.4%. The majority of proteins identified without GPF were also seen in the analyses of (NH4)2SO4 precipitations with GPF (237 out of 255 proteins), the distribution of (NH4)2SO4 fractionated proteins identified from different cellular locations is indistinguishable between the analyses (Table 1, 61% cytosolic, 28% membrane and 11% periplasmic). GPF revealed an additional 4% S. typhimurium proteome when compared with the (NH4)2SO4 precipitations with no GPF, covering a total of 10.4% of the whole proteome. Compared with the whole-cell shotgun analysis on the LTQ-FT, the LCQ-DUO analyses of (NH4)2SO4 precipitations using GPF identified 396 of the same proteins and 76 unique proteins (Figure 3B). These results suggest that the data from (NH4)2SO4 precipitation without GPF were biased towards high-abundance proteins and that the use of GPF on these same fractions allowed ‘access’ to unique proteins present but undetected in either the LTQ-FT analysis of the whole-cell lysate or the LCQ-DUO analysis of the four (NH4)2SO4 precipitation fractions.
Towards better membrane protein coverage with shotgun proteomics
By prediction [19, 20], membrane proteins typically make up two-thirds of typical proteomes, but are consistently under-represented in shotgun proteomic results [21]. This is thought to be due to the insolubility resulting from the high number of hydrophobic amino acids present in membrane proteins. However, because of their increasing recognition as the primary players in cellular signalling, cell-to-cell interactions, ion transport, drug resistance and many other key cellular functions, membrane proteomes are greatly sought after. Traditionally, protein studies were initiated with the isolation of proteins using two-dimensional (gel electrophoresis (2DE) followed by the excision of individual protein-spots for MS/MS analyses. This method has great limitations for membrane protein studies because proteins with multiple transmembrane domains precipitate when they reach high concentrations at their respective pIs. In the interest of completing large membrane proteome studies in a more efficient manner with shotgun proteomics, two issues need to be addressed: (i) the solubility of membrane proteins must be increased prior to proteolysis and (ii) elimination of the use of 2D-gel electrophoresis to fractionate proteins prior to MS analysis. Recent studies have demonstrated that organic co-solvents can help increase the solubility of hydrophobic membrane proteins [22–24], and Deshusses et al. [25] demonstrated that the use of trifluoroethanol (TFE) can be used to increase the extraction efficiency of intrinsic membrane proteins. Combining the efforts of these studies increases membrane proteome coverage by allowing membrane proteins to maintain adequate solubility during isoelectric focusing (IEF). To avoid the use of polyacrylamide gel-based medium for separation, we investigated the effectiveness of a liquid IEF method to fractionate membrane proteins [26]. The use of a liquid IEF has several advantages over isoelectric strips: proteins are in a solution amenable to MS analysis, the solution can easily be digested by proteases, and a relatively inexpensive device is commercially available allowing up to 1.5 mg of proteins to be isolated per pI range. It has been noted that there is a significant loss in proteins with multiple transmembrane domains because of precipitation at high concentrations during IEF [27]. We therefore combined the two methods using TFE extraction to try to increase the solubility of membrane proteins prior to liquid IEF [26], with the ultimate goal of achieving greater membrane proteome coverage via shotgun proteomics on the LCQ-DUO mass spectrometer.
To pellet the membrane fraction from lysed cells, the whole-cell lysate was centrifuged (17 000 x g, 30 min) and the supernatant removed. Membranes were further purified using a TFE/CHCl3 phase separation [25] where extract intrinsic membrane proteins are isolated in the upper aqueous TFE phase and the insoluble middle-phase (Figure 1). Briefly, the 17 000 x g pellet (300–1000 µg) was re-suspended in 50 mM ammonium bicarbonate (100 µl) and added 1 ml of TFE:CHCl3 (2:1). The mixture was stored at 0°C for 1 h and occasionally mixed. This was followed by centrifugation (9000 x g, 4 min) to yield three visible phases (Figure 1). Deshusses et al. [25] reported finding very little protein in the lower lipid-rich chloroform phase, therefore, only the two upper phases were collected for protein analysis. The membrane fractions (TFE-soluble and TFE-insoluble) were dried in a speed-vac and re-solubilized in a mixture of 5 M urea, 2 M thiourea, 4% CHAPS (3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate) (UTC), 40% TFE and 10 mM Tris HCl (pH 8.8–9.0) [23, 25, 28]. Proteins were reduced for 1 h at room temperature with a final concentration of 50 mM DTT. Alkylation of membrane proteins was conducted with acrylamide at a concentration of 100 mM [29].
Trifluoroethanol extraction of membrane proteins prior to shotgun proteomics
Duplicate LC-MS/MS analyses of each of the TFE phases (insoluble and soluble; Figure 1) on the LCQ-DUO yielded a total of 607 good quality peptide tandem mass spectra. Reduction of this peptide data lead to the identification of 203 proteins (probability >0.99) with an average protein sequence coverage of 21%. An examination of the number of protein identifications made from different cellular locations indicates that 41% of the TFE-extracted proteins were membrane-associated (Figure 4A; 48 inner, 34 outer membrane proteins). A comparison of the number of unique membrane proteins identified from either of the TFE-soluble or TFE-insoluble phases, suggest that the phases should be considered as a whole fraction since neither phase was from a specific cellular location and there was significant crossover in the identifications of each fraction (Figure 4B, Table 1). Examination of the membrane proteins identified with the TFE extraction, demonstrates that the extraction does improve membrane proteome coverage. Compared with the (NH4)2SO4 fractionation, the TFE extraction identified 53 additional membrane proteins. From the twelve tandem LC-MS/MS analyses of (NH4)2SO4 (8) and TFE (4) extractions, a total of 121 membrane proteins were identified (Figure 4C).
GPF of the two TFE phases increased the average protein sequence coverage and the number of proteins identified to 24 and 298, respectively. By narrowing the m/z scan window in a total of 20 LC-MS/MS analyses, we were able to identify an additional 70 proteins not observed with the wide m/z scan. GPF increased the membrane proteome coverage by identifying 111 membrane proteins, 41 unique to GPF analyses (Figure 4D). As expected from the previous results, GPF did not specifically enhance membrane protein identifications, suggesting that membrane protein solubility and abundance are the primary factors dictating identification, and not ionization potential or chromatographic resolution of trans-membrane domain peptides. By merging the results from GPF analyses of (NH4)2SO4 precipitations (40 LC-MS/MS analyses) and the TFE extractions (20 LC-MS/MS analyses), we identified 11% (207 membrane proteins) coverage of the predicted S. typhimurium membrane proteome (1577 proteins) (Figure 5).
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Isoelectric focusing of TFE extracts to improve membrane proteome coverage
To determine if we could increase membrane proteome coverage by pre-fractionating the proteome, we used a commercially available liquid IEF unit (Invitrogen). The liquid IEF unit was designed from the research completed by Zuo and Speicher [26] as a means to reduce proteome complexity of eukaryotic cells by focusing abundant albumin proteins into one pI chamber for eventual MS analyses. Unfortunately, the S. typhimurium proteome does not have a distinct bimodal isoelectric pI distribution of proteins relating to cellular localization as suggested by Schwartz et al. [30] (Figure 6A); therefore, prior to IEF, membrane proteins were isolated using the TFE phase separation technique [16, 25]. We anticipated that the liquid IEF unit would fractionate membrane proteins into one of five pI ranges to increase both the number of membrane proteins identified on the LCQ-DUO and the membrane protein sequence coverage.
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) inner membrane, (
) outer membrane, (•) periplasmic, (x) cytosolic. (B) Distribution of proteins identified with the LTQ-FT and from the LCQ-DUO analyses of the (NH4)2SO4 precipitations with and without GPF (C), of the TFE extracts with and without GPF (D) and of the TFE extracts fractionated by IEF (E). Protein molecular weights predicted from the genome, pI calculated as reported previously [Each membrane protein fraction (TFE-soluble and TFE-insoluble) was individually subjected to IEF using the liquid IEF unit (Figure 1) (Invitrogen Zoom) (pH ranges 3–4, 4–5, 5–6, 6–7, 7–10) [31, 32]. Briefly,
0.5 mg of protein solubilized in UTC and TFE was added to two focusing buffers (Invitrogen), DTT and bromophenol blue prior to running the unit. IEF was carried out using 3 voltage steps (100 V, 20 min; 200 V, 80 min; 800 V, 80 min). Samples were then removed and the chambers were rinsed with 300 µl of UTC and TFE to remove any proteins adsorbed to the chamber walls. The IEF-fractionated membrane proteins were precipitated with cold acetone and then air-dried prior to trypsin digestion and mass spectrometry on the LCQ-DUO.
Duplicate LC-MS/MS analysis of 10 IEF fractions (5 pI ranges for each TFE-soluble and TFE-insoluble) were completed (20 LC-MS/MS analyses), yielding positive matches to 325 S. typhimurium proteins (probability >0.99). By combining all the IEF analyses, average percent protein sequence coverage increased to 23%. A total of 157 membrane proteins were identified (Table 1; 115 inner membrane and 42 outer membrane proteins), making it the most extensive list of membrane proteins identified from a fractionation technique with the LCQ-DUO. By combining the results of the LCQ-DUO shotgun analysis of membrane proteins identified with IEF and the analysis of the (NH4)2SO4 GPF, and TFE GPF 14% of S. typhimurium membrane proteome is confirmed (Figure 5).
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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Proteome pre-fractionation provides substantial advantage for low performance mass spectrometers. The increasing demand for proteomic information to answer complex biological problems requires the expertise and efforts of multiple laboratories. To increase laboratory throughput of samples for shotgun proteomic data, we have presented the use of easily accessible protein pre-fractionation techniques that can be used in series with older, inexpensive, ion-trap mass spectrometers found in a variety of protein laboratories. The use of these techniques can help investigators to improve the proteome coverage and produce data comparable in quantity to the data produced from state-of-the-art LTQ-FT instrumentation. Three tandem MS/MS analyses of whole-cell lysates using the LTQ-FT yielded the identification of 655 S. typhimurium proteins. If we combine the data from the 92 MS/MS analyses collected with the LCQ-DUO, we achieve positive identification of 648 proteins from
30x more instrument time (Figure 7). Given the many time constraints for access to high-performance instruments, this study demonstrates that quite a lot of initial work may be done on lower-performance instruments. Although these results hardly seem worth comparing when examining protein ID's per MS/MS analysis, we must take into account that many of the MS/MS analyses completed on the LCQ-DUO came from automated GPF which greatly increased the average time spent on the DUO, but also provided a great advantage.
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We recommend to proteomic practitioners trying to optimize proteomic coverage using lower-performance instruments like the LCQ-DUO to employ the use of GPF. This simple method provided more proteome per unit of MS time and wet laboratory effort than any other method we examined here. Specifically, we found that repeat analysis of the same fractions [TFE or (NH4)2SO4] under the same chromatographic and MS conditions, with the exception of using five narrow m/z scan ranges, yielded over 1.5x more protein identifications (e.g. Figure 3A, B). As suggested by Yi et al. [16], the automation of GPF with DD LC-MS/MS methods provides dramatic improvement in proteome coverage making GPF the most appealing addition to a shotgun proteomic experiment.
Regrettably, the (NH4)2SO4 precipitations did not effectively isolate one group of proteins based on the metrics of hydrophobicity, cellular location or pI. LC-MS/MS analysis of all four (NH4)2SO4 fractionations revealed some level of overlap and redundancy in the proteins identified (Table 1, Figure 3C). In spite of this, analysis of these same samples on the LCQ-DUO with GPF identified nearly three-quarters the number of S. typhimurium proteins (472) identified with the LTQ-FT (655), of which 76 proteins were exclusively identified by GPF from the (NH4)2SO4 fractions. This simple technique of gradually isolating different suites of proteins by salting-out is clearly a fast, inexpensive and easy fractionation step that can be adopted by any laboratory. Unlike the use of the liquid IEF unit, this procedure does not have limitations on the amount of protein used nor does it produce large amounts of chemical waste as with the IEF method [32]. Therefore, this method allows the investigator to make as many, or as few, (NH4)2SO4 fractionation steps, which may greatly improve the proteome coverage depending on the question and organism of interest. Since this method does not enhance protein coverage of a particular cellular fraction in S. typhimurium relative to whole-cell analyses of Gram-negative bacteria (Figure 2A, B), it can also be used to increase the general proteome coverage without the exclusion of any of the components.
To improve membrane protein identifications, the use of TFE was investigated to determine if it was amenable to direct shotgun proteomic analysis. The solubilization of membranes by TFE only slightly increased the percentage of membrane proteins identified using the shotgun technique relative to the other methods examined (Figures 4A, C and 5). Where the LTQ-FT identified only 22 outer membrane proteins and the (NH4)2SO4 fractionations identified only four outer membrane proteins, solubilization with TFE positively identified 41 and 34 with and without GPF, respectively. For the purpose of achieving the greatest membrane protein coverage, we would advise solubilization of the membrane fraction with TFE into the two phases, followed by the use of the liquid IEF device. To improve membrane protein coverage results, we would also suggest the use of GPF to increase the number of ions selected for tandem MS/MS analysis. Because the IEF unit partitions proteins into different pI ranges, it would make sense to complete more MS/MS analyses on the upper pI ranges in hopes of achieving better coverage of basic proteins (Figure 6B, C and D).
Although this was a narrow study of one Gram-negative bacterial strain, the results demonstrate that the use of the LCQ-DUO provided comparable proteome coverage to the much higher-performance LTQ-FT from 10x more LC-MS/MS analyses. The single most important key to this success appears to have been the inclusion of automated GPF. Thus, where sample is not limiting, we now recommend to users in our own laboratory and collaborators that they consider the use of GPF and well-designed pre-fractionation techniques followed by analysis on the LCQ-DUO. Of course if sample is limiting, then the LTQ-FT is the instrument of choice to maximize proteomic (or individual protein sequence coverage) content from minimal LC-MS/MS analyses.
Key Points
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Acknowledgements |
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TOPABSTRACTINTRODUCTIONCELLULAR PREPARATION, PROTEIN...SIMPLIFYING SHOTGUN PROTEOMICS...USING PROTEOME FRACTIONATION TO...CONCLUSIONSAcknowledgementsReferences |
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For partial support, we thank the University of Washington Center for Ecogenetics and Environmental Health NIEHS grant P30ES07033 and the University of Washington WWAMI Regional Center of Excellence for Biodefense and Emerging Infectious Disease NIAID grant 1U54AI57141-01. Also for support, BLN thanks the NSF Office of Polar Programs Postdoctoral Fellowship grant 0444148 and SIM NIH grant AI-30479. All protein identifications available upon request.
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Brook L. Nunn, NSF postdoctoral fellow in oceanography examining Southern Ocean dissolved proteins and phytoplankton proteomics. Trained by Dr RG Keil in oceanography and Dr DR Goodlett in proteomics/mass spectrometry.
Scott A. Shaffer, trained by Dr F Turecek (1991–1995) and Dr RD Smith (1996–1998) in mass spectrometry, is a Research Scientist in the Goodlett laboratory.
Alexander Scherl, trained by Dr Hochstrasser and Dr Hopfgartner at the University of Geneva, is a postdoctoral researcher in the Goodlett laboratory.
Byron Gallis is a protein chemist/kinase expert trained by Dr Edwin G. Krebs.
Manhong Wu is studying Pseudomonas aeruginosa biology with Dr Miller and Dr Goodlett.
Samuel I. Miller, Professor of Medicine, is an internationally recognized expert in Gram negative bacteria.
David R. Goodlett, trained by Dr RB van Breemen (1987–1991) and Dr RD Smith (1991–1993) in mass spectrometry and Dr JL Aull, Dr FF Bartol and Dr HH Darron (1982–1986) in protein chemistry, is Associate Professor in Medicinal Chemistry.
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