Within its approximately 20-year history, pro-teome analysis has been aided primarily through instrumental advances. Relative to the instru-mentation used in the first peptide mass finger-printing and peptide sequencing experiments, today’s mass spectrometers scan faster, and are more sensitive, selective and versatile than ever before. Sparked by the development of soft ionization (i.e., MALDI, ESI), biological mass spectrometry (MS) has evolved through novel fragmentation strategies (electron-capture dis-sociation, electron-transfer dissociation), and through commercialization of new or improved mass analyzers such as the Orbitrap, linear traps, quadrupole TOFs or Fourier transform ion cyclotron resonance. These advances are in direct response to the incredible demands imposed by proteome characterization.
Of course, MS rarely acts alone for proteome profiling; MS typically follows extensive sepa-ration (single or multidimensional) to reduce sample complexity. Exceptions include the rapid profiling of whole organisms by Fenselau and Demirev through MALDI MS [1] or, most recently, through top-down MS [2]. Nonetheless, extending the characterization beyond the most abundant sample components requires separation.
The limitations of MS for proteome ana lysis are illustrated through a recent study by Mann et al., which delved into the complexity of a pro-teome sample [3]. Focus was on the unidentified portion of the proteome, referring to peptides for which no tandem MS (MS/MS) sequence deter-mination was made. With over 100,000 ‘peptide features’ observable in a single MS experiment, the vast majority of ions selected for MS/MS sequencing represent mixed spectra from co-eluting peptides (reported as a median 14% of the isolated signal for MS/MS coming from a single component). While increased MS scan rates and improved sensitivity would improve detection, this would not eliminate the peptide co-elution problem. One possible solution is to further reduce sample complexity by introducing high-resolution proteome fractionation.
Peptide- versus protein-level separationA distinction between peptide- versus protein-level separation could just as easily take on a discussion of solution versus gel approaches, or alternatively chromatographic versus electrophoretic plat-forms for proteome fractionation. Characteristic differences exist between peptides and proteins, which warrant optimization of distinct separation
Alan A Doucette*1, John C Tran2, Mark J Wall1 and Shayla Fitzsimmons1
1Department of Chemistry, Dalhousie University, 6274 Coburg Road, Halifax, NS, B3H 4R2, Canada 2Department of Molecular Biosciences, Proteomics Center of Excellence, Northwestern University, 2170 Campus Drive, Evanston, IL 60208, USA *Author for correspondence: Tel.: +1 902 494 3714 Fax: +1 902 494 1310 [email protected]
Proteome fractionation refers to separation at the level of intact proteins. Proteome fractionation may precede sample digestion and subsequent peptide-level separation and detection (i.e., bottom-up mass spectrometry [MS]). For top-down MS, proteome fractionation acts as a stand-alone separation platform, since intact proteins are directly analyzed by the mass spectrometer. Regardless of the MS identification strategy, separation of intact proteins has clear benefits as a result of decreasing sample complexity. However, this stage of the workflow also creates considerable challenges, which are generally absent from the counterpart peptide separation experiment. For example, maintaining protein solubility is a key concern before, during and after separation. To this end, surfactants such as sodium dodecyl sulfate may be employed during fractionation, so long as they are eliminated prior to MS. In this article, current strategies for proteome fractionation in a MS-compatible format are reviewed, illustrating the challenges and outlooks on this important aspect of proteomics.
Keywords: bottom-up MS • gel electrophoresis • gel-free separation • protein solubilization • proteome fractionation • SDS removal • surfactants • top-down MS
Intact proteome fractionation strategies compatible with mass spectrometryExpert Rev. Proteomics 8(6), 787–800 (2011)
platforms. In the classical MS protein identification experiment (i.e., peptide mass fingerprinting), proteins are required in relatively high purity, and so proteome fractionation is critical. Depending on sample complexity, this was traditionally accomplished through SDS PAGE or 2DE. The coupling of electrophoretic separation, be it through SDS PAGE or 2DE, with MS is ideal for targeted analysis of proteins, and remains a popular approach for protein identification.
Compared with the slow and laborious nature of the classic gel-based experiment, peptide-level separation had an initial focus on automation and increased throughput. With MS/MS peptide sequencing, pure protein samples are no longer required for identi-fication. Given the availability of electrospray, evolving into nano-spray, the liquid chromatography (LC)-MS experiment became an obvious choice for increased proteome coverage at high throughput [4]. Multidimensional chromatography can further enhance the depth of proteome coverage [5]. Peptide-level separation strategies have been reviewed extensively [6], and so a detailed description of the available techniques will not be presented in this article.
Challenges to protein separationProteome separation involves partitioning of a significantly smaller number of components than its peptide-level counterpart (each protein gives rise to multiple peptide fragments). Therefore, from a purely numerical standpoint, the intact proteome could be considered a simpler mixture, and thus require a lower resolu-tion separation. Unfortunately, the number of sample compo-nents does not address the practical limitations of peptide- ver-sus protein-level separation. Highlighted below are some of the challenges of proteome fractionation.
Greater sample diversity at protein levelIn comparison to peptides, proteins have a more diverse range of chemical properties (i.e., charge, hydrophobicity and size). This makes it difficult to create a single method that is universally appli-cable to the entire proteome. For example, in chromatographic separation of proteins, it is difficult to find a suitable solvent system that first permits the majority of the sample components to first bind to the column during loading and subsequently elute from the column. This was noted by our laboratory when poor recovery was observed during ion exchange fractionation of a proteome [7].
Poor protein solubilityMaintaining a high level of protein recovery, not only during the separation, but prior to and following fractionation, is a significant challenge. Solubilizing additives such as SDS are commonly used to improve protein solubility. However, the addition of such additives places constraints on the available platforms employed for protein separation. Furthermore, SDS is incompatible with subsequent MS detection. Fortunately, as described in later sections, there are strategies that permit the use of SDS.
Lack of proteotypic proteinNot only are peptides generally more soluble and thus easier to recover through peptide separation, but the bottom-up experiment
also implies a large degree of ‘acceptable loss’. The loss of a signifi-cant number of peptides during separation is perhaps favored in the bottom-up experiment, with a goal of protein identi fication as opposed to complete protein sequence characterization [8]. Considering MS peptide sequencing, a representative protein frag-ment is sufficient to identify that component. While the physical properties of proteins vary considerably in a proteome mixture, it is hoped (and assumed) that there will be at least one peptide hav-ing desirable properties that are compatible with separation (and detection). The term ‘proteotypic peptide’ has been used to quan-tify peptides which are more favorably detected in a given MS experiment [9,10]. For example, Beavis et al. suggested that only 6% of all tryptic peptides generated from the human proteome are frequently observed by MS, although only 4% were required to generate a library of approximately 15,000 unique proteins [9].
To date, an equivalent predictor of proteotypic proteins (as per top-down MS, for example) has not been presented. Nonetheless, it can be assumed that the propensity to detect proteins will decrease at extremities of molecular weight (MW), hydropho bicity and charge, as such proteins are more difficult to handle during fractionation. For example, membrane proteins generally have poor solubility in aqueous solvents and are therefore difficult to separate via chromato-graphic approaches. Considering the character istic properties of the sample components, a protein mixture is far more diverse than its peptide counterpart. Consequently, it becomes a challenge to pro-vide a single separation platform which can accomodate this sample diversity. If a protein component is lost during fractionation, no MS instrument will recover the lost information (Figure 1).
Decreased chromatographic resolutionIn chromatography, under comparable conditions, peptides gener-ally separate with a much higher degree of resolution than their intact protein counterparts. Given the size of a protein molecule, multiple interactions can occur with the stationary phase, depend-ing on protein folding and orientation. This, among many other causes, has a net effect of broadening chromatographic peaks. Peptides are generally well-behaved molecules during chromato-graphic separation, and thus high-resolution separation is possible. For example, Smith et al. demonstrated a peak capacity of approxi-mately 1100 over a 100-min reversed-phase separation [11]. A peak capacity on the order of 30 is more typical for reversed-phase protein-level separation [12].
Lower throughput & reproducibility concernsConsidering a bottom-up experiment, it is simpler to perform a global digest of the proteome than to digest multiple fractions. A single sample processing step also implies greater consistency, which is particularly important for quantitative investigations. For example, in an isotopic labeling experiment, the proteome samples to be compared are globally digested, chemically labeled and then combined prior to separation. This strategy alleviates any differ-ences instilled during fractionation. Isotopic tagging at the protein level is also possible through chemical means [13], but has yet to be generally adopted over peptide-based labeling strategies. Metabolic incorporation of isotopes into proteins is feasible through stable
Doucette, Tran, Wall & Fitzsimmons
www.expert-reviews.com 789
Review
isotope labeling with amino acids in cell culture [14], and the method has recently been extended to samples from human subjects [15].
These arguments provide strong support in favour of bottom-up strategies involving peptide separations, and have given rise to many proteome detection platforms that rival the use of 2DE for proteome analysis.
Motivation of proteome fractionationGiven a landscape apparently favored to separation at the peptide level, one may question if protein-level separation is worthwhile. In fact there are several reasons why proteome fractionation is desired in a proteome workflow, as outlined below.
Orthogonal (or complementary) to peptide separationMeyer et al. recently overviewed the possibility of obtaining 100% proteome coverage [16]. With several technologies available for
proteome analysis no single strategy can be declared a complete platform for comprehensive characterization. Thus, proteome cov-erage can be improved by combining complementary technologies. With a focus on separation, it is recognized that intact separations are complementary to peptide-level separation. When combined in tandem, protein and peptide separations are fully orthogonal, a critical parameter in designing a multidimensional platform.
High-abundance protein depletion & targeted analysisRecognizing the dynamic range of concentration in a proteome, mining the low-abundance proteome can be enhanced by deplet-ing the most abundant sample components. Protein Equalizer Technology (ProteoMiner) protein-enrichment kits are available as a means of ‘normalizing’ the proteome [17]. This technology functions through selective capture of a defined quantity of each protein using a combinatorial library of short peptides bound to
Non-extracted proteins
SampleExtraction
Loss ofinsoluble proteins Buffe
ring
Digestion
Peptide separationProtein separation
Peptide lossduring clean-up
Sample lossduring separation
Precipitation
MS analysis
Chr
omat
ogra
phy Chromatography
Impact of loss onprotein identification
Moderate
Severe
Figure 1. A simplified representation of two proteomics workflows, highlighting the impact of sample loss during proteome fractionation versus peptide separation. Following extraction, the workflow is shown to break into two streams: proteome fractionation strategy (left) and peptide separation strategy (right). Colored flags highlight the steps within each workflow in which protein or peptide loss can occur. The severity of loss, as indicated by white (moderate) or black (severe), is defined according to the impact on protein identification through MS. MS: Mass spectrometry.
Intact proteome fractionation strategies compatible with mass spectrometry
Expert Rev. Proteomics 8(6), (2011)790
Review
a support. Similarly to high-abundance protein depletion, low-abundance protein components can be targeted through proteome fractionation. Affinity approaches are again available to enrich, for example, glyco- or phospho-proteins [18]. Through a process called analysis-driven experimentation, Aebersold et al. addressed the under-represented classes of (small/basic) Drosophila proteins [19]. Here, a targeted analysis followed proteome fractionation of low-MW and acidic proteins, with a net result of detecting approximately 63% of the Drosophila proteome (>9000 proteins).
Preservation of a protein’s primary structurePerhaps the greatest criticisms of the shotgun approach for pro-teome characterization is the loss of protein integrity induced by global sample digestion. One must infer that the presence of a peptide (or multiple peptides) indicates the presence of the intact protein in the sample. Issues around biological protein degrada-tion cannot be addressed. Histones are an example where protein preservation is important. Not only is the detection of protein modifications important, but tracking the synergistic effects of multiple modifications on a single protein may prove to be key in understanding the regulatory process of these proteins [20]. By preserving the protein’s primary structure during separation, the intrinsic information afforded by the separation (e.g., protein isoelectric point [pI] and MW from 2D gels) can also be used to better understand the biological system.
The emergence of top-down MSThe potential of top-down proteomics is enormous. First, it enables unambiguous protein identification through its ability to provide 100% sequence coverage. This is particularly important in fully characterizing protein isoforms, which has been problem-atic with bottom-up approaches. Arguably, the greatest advantage of top-down analysis is its ability to localize post-translational modifications (PTMs), especially if they occur in combination. Further, top-down has the ability to preserve labile PTM sites [21–23], which are often lost in bottom-up proteomics. Significant advances in recent years include the ability to sequence a 200 kDa protein [24], and providing top-down analysis compatible with chromatographic time scales [25] and at high mass [26]. As outlined below, extensive research is currently being conducted to improve the front-end introduction of proteins to the mass spectrometer. Most importantly, the need for MS-compatible alternatives to SDS-PAGE for separation of intact proteins has renewed interest in the development of novel proteome fractionation strategies.
Proteome fractionation platformsProteome fractionation can take on a definitively low-resolution format (two fractions as seen through immunoaffinity cap-ture of a single protein), or can offer extremely high resolution (~10,000 spots, considering 2DE from multiple narrow-range pH Immobiline™ polyacrylamide gel strips). Techniques for affin-ity enrichment or depletion of the proteome have recently been reviewed [18,27]; immunoaffinity strategies will therefore not be covered in this article. As applied in this article, proteome frac-tionation entails the (potentially) comprehensive fractionation of
the complete proteome. Thus, all sample components are retained across the collected fractions.
Before outlining some recent strategies for proteome frac-tionation, we should also mention protein recovery. In report-ing recovery, one typically refers to the total mass of a complex mixture that remains following a given separation. However, an assessment of possible bias (e.g., towards loss of hydrophobic proteins) should also be made. Discovery-based proteome experi-ments do not assume any knowledge of which components are of interest, and thus it is important to maintain high recovery throughout the workflow.
SDS-PAGE & 2DE2DE combines two orthogonal modes of separation. Prior to MW separation via SDS PAGE, proteins are focused according to their pI, which relies on the establishment of a pH gradient in an applied electric field. Charged proteins migrate along the pH gradient to a point where their net charge is zero (defined as the proteins’ pI). It is noted that this stage of separation must avoid the inclusion of SDS (or other charged surfactants) to any appre-ciable concentration, which otherwise overwhelm and deteriorate the focusing step.
The dominance of 2DE as a proteomic tool is attributed to its unrivalled high resolution power. Up until the last decade, 2DE with MS fingerprinting was the definitive platform for large-scale proteome identification. Using this technology, impres-sively large numbers of proteins were identified [28]. However, like all technologies, there are disadvantages to 2DE that should be highlighted.
A notable difficulty is the limited loading capacity of 2DE where, depending on the sample, as little as 1 mg of protein on a single gel can result in band smearing and degraded resolution [29]. This limited loading capacity hinders the detection of low-abundance proteins. An evaluation of 2DE for analysis of yeast by Gygi and Aebersold suggests that the large dynamic range of protein expres-sion limits the ability to visualize (through staining), and conse-quently detect (through MS) even the medium-abundance proteins of yeast [30]. In addition, the loss of proteins at extreme pI and MW will result in detection of only a portion of the proteome population. Thus, the 2DE + MS experiment results in biased identification of proteins. Finally, analytical throughput, being an important con-sideration of comprehensive proteomics, mainly relates to the ability to automate a technology. Robotic systems are available for spot picking and digestion. Nonetheless, the need for excision, digestion and subsequent analysis of hundreds, if not thousands, of gel spots characterizes the 2DE + MS workflow as a time-consuming task. Taken together, these limitations gave rise to the development of gel-free alternatives for proteome processing, as seen through the emergence of shotgun approaches to proteome analysis.
Perhaps ironically, many of the limitations imposed by the 2DE-MS experiment can be alleviated by reverting to a lower resolution separation platform. As a standalone platform, SDS-PAGE is the single most widely used protein separation tool in proteomics [31]. Its ability to separate with extremely high resolution and in a highly cor-relative fashion makes it one of the most valuable protein separation
Doucette, Tran, Wall & Fitzsimmons
www.expert-reviews.com 791
Review
tools available. In the SDS-PAGE procedure, proteins are first solu-bilized in a sample buffer containing the anionic surfactant SDS. SDS binds and denatures individual protein molecules imparting a similar negative charge density for all proteins. The benefits of using SDS are twofold: it imparts a general (logarithmic) relationship between the mobility and the MW of the protein; and it assists in maintaining protein solubility during separation.
In the ‘GeLC-MS’ experiment [32], an SDS-PAGE separation of the proteome is first performed. By omitting the isoelectric focusing (IEF) stage, the previously imposed bias surrounding protein pI and hydrophobicity are alleviated. Furthermore, with GeLC-MS the entire lane is excised into approximately 10–20 even-sized fractions, thus the bias towards protein abundance is significantly reduced. Each fraction is systematically processed and subject to bottom-up peptide sequencing using LC-MS/MS. The GeLC-MS experiment is the clear standard for bottom-up proteome processing [32].
Irrespective of the laborious nature surrounding the GeLC exper-iment, there are several motivators to other modes of proteome frac-tionation. Solution-based separations are much easier to automate thereby making them less labour intensive. Difficulties surrounding the extraction of intact proteins from a gel mean that the technique is not amenable to top-down MS. There are a wide range of sepa-ration strategies that operate in solution. When used collectively (although not necessarily coupled in sequence), these platforms can potentially provide full proteome coverage. These alternative solution-phase fractionation platforms are discussed below.
Reversed-phase LCAs stated above, the greater diversity of protein properties pre-sented a challenge in maintaining high recovery during separa-tion. Nonetheless, this diversity of properties has enabled mul-tiple forms of chromatography to be adopted into intact proteome prefractionation platforms.
Optimization of reversed-phase LC (RPLC) for fractionation of intact proteins takes on two main research focuses: column chemis-try and solvent selection. RPLC is perhaps the most important mode of chromatography to consider as this system is fully compatible with MS and can thus be coupled through ESI in an online format. Hence, RPLC-MS is the favored platform for top-down proteome profiling. Considering the column construction, it was recognized that larger protein molecules can be lost to the relatively smaller pores of silica supports, which have been optimized for peptide-level separation. The adoption of non-porous material by Lubman and colleagues presented a method to avoid loss of large components [33]. Support material with larger pores (>300 Å, and as high as 4000 Å) are available commercially, being generally marketed for protein separation [12]. The transition from silica-based to polymeric sup-ports has also proved to be favorable for protein separation [12]. The selection of less hydrophobic, shorter chain length bonded phases (C
4 or C
8, over C
18) remains a favored strategy for protein separa-
tion. RPLC (10 cm long PLRP-S columns [Agilent Technologies] with 5 µm particle size) peak widths below 1 min can be typical for intact proteins <25 kDa. However, there is much room for improve-ment in RPLC separations of intact proteins >50 kDa in regards to both recovery and resolution. The difficulty in separating large
MW intact proteins using RPLC remains one of the most notorious bottlenecks in separations for top-down proteomics
The acetonitrile–water gradient system, which is standard for peptide separation, can also be used for protein separation. The switch to stronger eluents, such as isopropanol or even formic acid–isopropanol, has been adopted as a favorable approach to recover hydrophobic analytes such as membrane proteins [34]. Ion-pairing agents such as trifluoroacetic acid are generally avoided in LC-MS experiments. However, the need for improved chromato-graphic performance may outweigh the suppression of these agents during ionization.
Other modes of chromatographyProtein separations based on charge have historically been domi-nated by ion-exchange chromatography (IEC), being due in part to the relatively high resolution and high loading capacity of IEC. Protein elution in IEC can occur by increasing either the ionic (salt concentration) or pH strength of the mobile phase. Although using a salt gradient is the most popular strategy, it does not sepa-rate proteins predictably according to pI [7]. Chromatofocusing, a variant of IEC that utilizes pH gradients, is capable of separat-ing with high pI correlation [35]. However, it is susceptible to low protein recovery owing to isoelectric precipitation.
Size-exclusion chromatography (SEC) is a favored approach to size separation. SEC columns are made up of porous gel beads with selectivity for a range of protein size. SEC is a favorable technique since it separates proteins with reasonable correlation to MW [36,37], a universal parameter, and retains protein activity. However, the technique suffers from detrimental dilution as fractions are recov-ered. Furthermore, the resolution of the technique has remained notoriously poor, and therefore its use has been limited in proteome prefractionation.
Solution IEFOne of the benefits of intact proteome fractionation is the potential for predictable separations (i.e., according to size or charge). The importance of separating under predictive properties cannot be undervalued since it contributes to additional confidence in the identification process, enabling one to add constraints based on information from the separation. As a focusing technique, IEF is not only predictable but can potentially surpass the resolution afforded by any other charge-based platform. The IEF resolution using an immobilized pH gradient (IPG) can approach 0.001 pH units [38]! In the past decade, IEF in solution has been widely exploited for its high resolution and its ability to separate with high pI correlation.
The principles of solution IEF are similar to those performed in gels, where the usual electrolytic buffers and pH gradient are supplied. Capillary IEF employs carrier ampholytes, with the protein sample focused at high voltages, providing extremely fast (<10 min) and high-resolution focusing. The high surface area to volume ratio of the capillary column acts to minimize Joule heating, which would otherwise lead to band broadening. As a demonstration of the high resolution, Jensen and coworkers coupled capillary isoelectric focusing directly to Fourier trans-form ion cyclotron resonance and generated 2D maps of some
Intact proteome fractionation strategies compatible with mass spectrometry
Expert Rev. Proteomics 8(6), (2011)792
Review
1200 components from the proteome of Deinococcus radiodu-rans [39]. Although the dimensions of the capillary are beneficial to the resolution, they also limit loading capacity. Therefore, capil-lary isoelectric focusing has not been as popular as the larger IEF devices, which provide capacities ranging from low milliliters to tens of milliliters [40]. Such ‘semi-preparative’ IEF devices have emerged as a compelling competitor to IEC.
The Digital ProteomeChip (dPC) Fractionator employs IEF to separate proteins [41]. In the commercial system, a set of 41 discrete IPG gel ‘plugs’ embedded within a separation cartridge span a fixed pH range. Proteins initially in solution are focused within these gel plugs according to pI. Gel-bound proteins can then be processed in much the same way as excised segments of an IPG strip or SDS-PAGE bands.
The Agilent OFFGEL Fractionator device [42] contains a 24-com-partment platform that is tightly sealed over an IPG strip. Contrary to the dPC Fractionator, focusing occurs through the IPG gel strip; however, once the proteins reach the pH region matching their pI, they can permeate into the solution-filled compartment above the gel strip. Therefore, while proteins are separated in a gel, they are ulti-mately recovered in solution. Relatively high resolution is possible; Michel and coworkers demonstrated that simple protein standards are focused almost entirely in single compartments of the OFFGEL device [43]. However, they noted poor recoveries of standard proteins; four out of the eight proteins had recoveries below 9%, the lowest being only 3.7% (trypsin inhibitor) [43]. The authors speculated that the low recoveries were owing to isoelectric precipitation in-gel. Thus, OFFGEL separations have limited use for proteome fraction-ation, but are extremely effective at separating peptides, where their use has afforded some of the highest proteome coverages [44].
Popular solution IEF designs make use of multicompartment chambers, however unlike the OFFGEL or dPC chip, nearly all focusing occurs in solution. In multicompartment solution IEF (sIEF), the medium contains the protein sample and solubiliz-ing reagents, such as urea, dithiothreitol, thiourea and CHAPS (3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate), dissolved in water [45]. Different sIEF systems can be classified according to the method of establishing the pH gradient. For sys-tems that strictly use carrier ampholytes, Biorad’s Rotofor® remains popular. The Rotofor is favored for its simplicity; however, since the device only uses a water permeable screen to isolate chambers, bulk flow can result in convective mixing, which leads to poor resolu-tion. The alternative carrier ampholyte sIEF device is Free Flow Electrophoresis, introduced by Hannig [46]. This device continu-ously pumps protein sample downstream, while a perpendicular electric field is used to separate the proteins according to pI. Up to 96 fractions can be aspirated and collected [47]. Although this separa-tion is capable of higher resolution and higher loading capacity than the Rotofor, its high expense has limited its use to selected research groups. In another approach, an eight-chamber carrier ampholyte sIEF device makes use of restriction channels between chambers to limit the bulk flow of liquid between chambers (Figure 2), allowing isolation between chambers [48]. This approach enables the use of washes (e.g., using acid and SDS) in the chambers after focusing and collection, to enhance the recovery of precipitated proteins.
In a multicompartment electrolyzer designed by Righetti and coworkers, IPG membranes are used to define each compart-ment [49,50]. These membranes not only minimize bulk flow but enable higher loading capacity and resolution, and minimize cathodic drifts compared with the Rotofor. The original design has been commercialized by Amersham as the IsoPrime device and a miniature version is commercialized by Proteome Systems as IsoelectrIQ. While isoelectric membranes have their benefits, they also decrease protein recovery, where up to 20% of all pro-teins are lost to the membranes through isoelectric precipitation [45]. Furthermore, isoelectric membranes require longer runs to prevent overheating of the system compared with carrier ampholyte IEF devices.
Zuo and Speicher designed their sIEF device by combining the principles of the Rotofor and the Isoprime by using both carrier ampholytes and isoelectric membranes [29]. This device is commer-cialized by Invitrogen as the ZOOM® IEF, with custom-designed systems reported as well [29]. The ZOOM IEF was the smallest scale sIEF device at the time (three to six compartments of 650 µl each), making it more suitable for proteomics applications. This was a significant achievement, and since then, both the Rotofor and the Isoprime have been modified with miniature versions as mentioned.
Mass-based electrophoretic separationSince SDS-PAGE offers the highest resolution, it is logical to attempt to recover intact proteins (in solution) following SDS-PAGE. This strategy is analogous to an OFFGEL fractionation of proteins by pI. Intact proteins can be extracted from a poly-acrylamide gel using solvents [51]. However, this strategy inevi-tably results in very low recoveries [52]. A more efficient way to extract intact proteins is through electroelution, which forces the migration of proteins out of the gel using electric fields.
Two commercial devices for preparative gel electrophoresis are offered by Biorad: the Prep Cell and Whole Gel Eluter, which separate through SDS-PAGE but recover proteins in solution by electroelution. The Prep Cell takes its beginning from several research groups in the 1960s [53,54]. It is a form of continuous elu-tion tube gel electrophoresis first described by Ornstein [55] and Davis [56], and put into practice as a preparative method by Lewis and Clark [53]. In this strategy, separation occurs in a tubular gel. Rather than terminate the run while (separated) proteins remain in the gel, the applied voltage causes proteins to migrate from the gel into solution. A dialysis membrane prevents further migration of proteins into the anode. Sample collection involves continuous pumping of elution buffer to move proteins into discrete frac-tions. The inclusion of a cold water cooling system through the donut-shaped SDS-PAGE column reduces Joule heating inside the column [54].
The Prep Cell is capable of relatively high mass resolution, being on the order of a few kilodaltons. As such, the Prep Cell is intended for protein purification and has proven successful for such an appli-cation [57]. As a tool for proteome fractionation, the long gel col-umn contributes to extremely long migration times and imposes a practical upper limit on mass. Meng and coworkers reported a separation mass range limit of 45 kDa after an 8 h separation [58].
Doucette, Tran, Wall & Fitzsimmons
www.expert-reviews.com 793
Review
Furthermore, the continuous elution strategy for protein recov-ery causes detrimental dilution of sample (100-fold dilution) [58]. Sample dilution is more severe for larger proteins, and contributes to collection of the larger proteins across multiple fractions.
In 1993, Andersen and Heron introduced the Whole Gel Eluter [59]. In this approach, separations are performed by SDS-PAGE, using the entire width of a gel for increased loading capacity.
Following separation, a transverse electric field is used to electro-elute the proteins into several discrete chambers (14–30 in total). While the Prep Cell is designed for protein purification, the Whole Gel Eluter is ideal for proteome fractionation, since it has the ability to collect a broad mass range with minimal dilution.
The Gelfree® platform [60] is another electroelution device, and is available from Protein Discovery as the Gelfree 8100
Intact proteome fractionation strategies compatible with mass spectrometry
Expert Rev. Proteomics 8(6), (2011)794
Review
fractionation system. The principles of Gelfree separation are not unlike those of the Prep Cell. However, as described by Tran and Doucette, the use of short gel columns, combined with a stop-and-go strategy to collect fractions, provides broad mass range (~10–150 kDa) protein separation in as little as 90 min [60]. Furthermore, the composition of the gel column can be tailored to deliver approximately 20 fractions over a narrower mass range, resulting in high-resolution protein separation. The recovered proteins can be analyzed through bottom-up [60–62] or top-down approaches [12,63]. However, the Gelfree platform will result in the recovery of intact protein fractions in solutions containing SDS (~0.1%). Thus, methods for removal of this interfering surfactant are critical to the success of the Gelfree-MS platform (Figure 3).
Multidimensional separationsGiddings stated that when two orthogonal separations are cou-pled, the peak capacity is the factor of their individual peak capac-ities [64]. Assuming a 3D separation platform, with each mode having a peak capacity of 30, then the total peak capacity of this platform will be 27,000! However, coupling multiple dimensions of separation amplifies the challenges previously mentioned for proteome fractionation. Considering throughput, one can quickly generate more fractions than can be feasibly analyzed. In the above example, if one collected 30 fractions per dimension, and assuming 90 min LC-MS per fraction (RPLC being the third dimension), almost 2 months would be required to process a single sample. Such lengthy experiments are increasingly becom-ing the norm for deep proteome profiling. It is noted that it is extremely difficult to find three modes of separation that are truly orthogonal, although 3D couplings have been demonstrated [65].
The second concern of a multidimensional separation relates to protein recovery. While peak capacities are multiplicative, so too are recovery rates; an 80% recovery (a generous rate for proteins) in each of three dimensions of separation translates into 50% total recovery. Nearly every combination of separation mode has been coupled in multidimensional platforms: Zuo and Speicher used the ZOOM IEF for serum fractionation ahead of 2DE. They claimed a 6–30-fold higher protein load using narrow pH range IPG gels [45]. Opiteck and coworkers first coupled IEC to RPLC for their com-prehensive analysis of Escherichia coli [66]. Assiddiq et al. compared protein and peptide separations using their IEC/RPLC platform and concluded that a similar number of proteins can be identified after-wards using LC-MS/MS from both methods [67]. Kelleher’s group used IEC/RPLC followed by top-down MS characterization in a technique they referred to as multidimensional protein characteriza-tion by automated top down (MudCAT) [25]. Chromatofocusing has been coupled to RPLC [68], resulting in high pI correlation, in contrast to IEC–RPLC coupling. This tandem coupling has become a popular choice through the introduction of Beckman Coulter’s Protein Fractionation 2D system. Wall and coworkers coupled the Rotofor to RPLC, resolving approximately 700 proteins [33]. The platform has also been used in cancer profiling [68,69]. Moritz and coworkers first coupled free flow electrophoresis to RPLC in 2004 and calculated their peak capacity to be 6720 [70]. However, this peak capacity is an overstatement of the true peak capacity since it
assumes that proteins are found in every given fractions. Free flow electrophoresis has also been used to resolve histone proteins [71].
While size–hydrophobic coupling can be conducted, these modes are not truly orthogonal. However, the benefits of a size-based separation followed by a MS-compatible RPLC make it a very favorable approach. The Kelleher group has recently coupled Gelfree to RPLC for top-down MS [12,63].
SDS for proteome fractionationAlthough less concerning for peptide-level separation, maintaining protein solubility is a key issue surrounding proteome fractionation. Protein solubility must be maintained throughout the fractionation stage; effective proteome extraction prior to separation is equally important. SDS is the most widely used surfactant for proteome processing. SDS promotes the disruption of lipid cell membranes in protein extraction, aids in protein solubilization [72] and facilitates protein separation according to MW (e.g., SDS-PAGE and Gelfree). Moreover, SDS can enhance digestion efficiency [73] or be used as a disaggregating agent [74]. Unfortunately, with all the positive attri-butes of SDS, this compound cannot be tolerated to any appreciable level in the ESI-MS experiment (exceptions noted in MALDI MS [75]); peptide as well as protein ionization in ESI experiences severe suppression when conducted in the presence of SDS.
MS & LC suppression problemThe mechanism of SDS suppression in MS is presented in detail elsewhere [76]. It suffices to say that the anionic surfactant competes very effectively with positive charges, particularly at the surface of ESI droplets. It has been reported that complete removal of detergent is necessary in order to obtain reliable mass spectra [77]. Investigations into the effect of background ions on the ESI process have shown that ion concentrations above 10–5 M (~3 × 10-4%) cause severe signal suppression [78]. However, the values above refer to signal suppression observed via direct infusion MS, while proteome analysis is primarily through LC-ESI-MS. Referring to a bottom-up LC-ESI-MS experiment, SDS was well tolerated at levels up to 0.01% [62].
Focusing on separation directly, SDS has a well-known nega-tive effect on RPLC separations. SDS adsorbs strongly onto the reversed-phase column, where the anionic head group works as an ion-exchanger, attracting proteins/peptides, retarding elution and decreasing resolution [79]. SDS has similar consequences on IEC, and will also negatively impact IEF. While SDS has clear benefits during some forms of proteome fractionation, it must be used with caution. The concentration of SDS must be reduced prior to pro-ceeding to the MS stage of the proteome workflow. Alternatively, it may be favorable to substitute SDS for another suitable reagent.
Methods for SDS removalDespite a number of alternative detergents available for use in proteomics experiments, SDS remains the top choice for many researchers. As such, a considerable number of strategies have been proposed for the depletion of SDS from protein-containing samples. The classic approach is through extensive dialysis of the sample [80], although such a method is not effective for the removal
Doucette, Tran, Wall & Fitzsimmons
www.expert-reviews.com 795
Review
of bound SDS. An overview of current SDS removal methods is provided below.
SDS depletion methods which rely on precipitation fall into two categories: pro-tein precipitation or detergent precipita-tion. The latter exploits the insolubility of dodecyl sulfate potassium and divalent salts to precipitate free dodecyl sulfate [81–82]. The commercial product SDS-Out™ (Pierce) induced precipitation of free SDS, although the agent used for precipitation is proprietary. Traditionally, SDS precipita-tion is effective at reducing the bulk of free SDS, but the inherent solubility of potas-sium dodecyl sulfate precludes significant reduction of sample.
Organic solvent protein precipitation is a popular technique for the removal of many contaminants, including surfactants. Thus, protein precipitation through various solvents, including acetone or chloroform/methanol/water, can be used to reduce the level of SDS in the sample [62,83,84]. Reported protein recoveries following acetone precipitation were shown to be reasonably high (>80%) [84]. More recent studies, as conducted by the authors, dem-onstrate even higher recoveries through acetone precipitation (>95%) (Figure 3). Organic solvent precipitation must gener-ally be applied prior to sample digestion, since recovery of peptides is ineffective with these techniques.
Several solid-phase formats to protein extraction have been presented, encompassing the methods of size exclusion [85], ion exchange (anion/cation) [86], ion-retardation resin [87], ceramic hydroxyapatite [88,89], reversed-phase [90], hydrophilic interaction LC [91], and other commercialized strategies including Pierce Detergent Spin Removal Columns (Pierce), Detergent-OUT™ (Millipore) and QuickSpin Detergent Removal Kit (DualSystems Biotech AG). These supports explore the intrinsic differences between the SDS molecule and the proteins themselves. The suc-cess of these approaches varies, but again the main concern lies in breaking the SDS–protein interactions, so as to avoid loss of protein during SDS depletion.
As already seen through SDS-PAGE, a number of approaches have exploited the separation of protein from SDS through differ-ential extraction from polyacrylamide gels. For example, overnight incubation of an SDS-containing polyacrylamide gel in methanol/acetic acid/water removes SDS by diffusion [92]. If one wishes to avoid the gel-based separation step, SDS-containing protein sam-ples can be added to a polyacrylamide gel during polymerization, or alternatively to a dried gel plug, and then the sample can be sub-jected to in-gel washing and digestion [93,94]. As with SDS-PAGE, the method is only suitable for bottom-up MS analysis.
Alternatives to SDSWhile SDS is the most popular detergent for use in proteomics experiments, other additives can be exploited that are more amenable to the separation stage, or alternatively to the detec-tion stage. These include nonsurfactant chaotropic agents (i.e., urea, thiourea and guanidine), which have also been shown to weaken the interaction between protein and SDS. In an innova-tive approach known as filter-aided sample preparation, Mann et al. exchanged SDS with urea using a standard filtration device enabling an unprecedented 7093 proteins to be identified from HeLa cells [95]. In addition, extractable [96,97], volatile [98] and cleavable surfactants [99,100] show considerable promise as MS-compatible solubilizing agents, given their ease of removal following separation. Ammonium perfluorooctanoate, for exam-ple, is a (semi-) volatile surfactant, which can therefore be elimi-nated during an extended process of solvent evaporation [101]. The surfactant has been applied in a ‘one-pot protocol’, along with other volatile reagents, to facilitate membrane proteome pro-cessing [101]. Sodium deoxycholate can also easily be eliminated from the sample through liquid extraction [97]. As a bile salt, this detergent is a natural pairing for trypsin and has been shown to improve membrane proteome coverage through detergent-assisted
Intact proteome fractionation strategies compatible with mass spectrometry
Expert Rev. Proteomics 8(6), (2011)796
Review
digestion [97,102]. Finally, acid-cleavable surfactants are commer-cially available (e.g., Rapigest™ SF Surfactant from Waters and PPS™ Silent Surfactant from Protein Discovery) and can be substituted for SDS. Being ionic, these act as efficient protein solubilizers. The MS suppression phenomenon that would other-wise be experienced from ionic surfactants is neutralized through chemical cleavage of the surfactant. Thus, samples may be applied directly to MS without further work-up.
Expert commentaryDefining the characteristics of an MS compatible separation plat-form will depend on the MS detection strategy (e.g., top-down or bottom-up; LC-ESI-MS or MALDI MS). Thus, the purity of the protein(s), presence of interferences or state of the sample (gel/liquid) varies according to the experiment. Considering that proteome fractionation represents just one part of an often lengthy analytical workflow, additional steps can be introduced to trans-form a seemingly incompatible separation (e.g., one that employs SDS) into a favorable strategy (e.g., SDS-PAGE-MS). With sev-eral strategies available for proteome fractionation, one needs only to make a selection. However, one must also be cautious of the potential risk of protein loss, or insufficient fractionation of sample components of interest.
Chromatographic separation is the natural choice for peptide-level separation, as seen through the online pairing of RPLC with MS. A long history exists for using various modes of chromato-graphy (e.g., ion exchange, size exclusion and affinity) to purify proteins. However, the constraints imposed by proteomics, namely to distribute all components across multiple fractions, all in high yield, implies that the chromatographic fractionation of a pro-teome is still in its infancy. When applied to proteins, chromato-graphy introduces a high risk of sample loss, particularly for poorly soluble protein components. Gel-based strategies, in particular SDS PAGE and 2DE, alleviate the issue of solubility during the separation. However, the recovery of intact proteins following the separation presents yet another issue [52]. New electro phoretic strategies have been introduced to overcome these limitations.
Maintaining solubility during the fractionation stages is a criti-cal aspect of the experiment. To this end, one may benefit from the inclusion of SDS or other solubilizing agents. Whenever possible, the surfactant should be added to the sample from the point of extrac-tion and included in the sample until the latest possible step. For bottom-up MS, this may even occur following digestion, as long as the concentration of surfactant does not suppress digestion efficiency [103]. It is noted that the addition of trypsin to a precipitated protein sample will promote the solubilization of the sample (as peptide fragments) [104]. In the top-down experiment, detergent removal must occur prior to the penultimate stage of RPLC (online with MS). Thus, development of efficient strategies for intact proteome separation remains a critical aspect in this workflow.
Five-year viewAdvances in MS instrumentation continue to push the bound-aries of proteome analysis. It is not only possible to identify a much greater percentage of all protein components in a given
proteome, but it is also possible to monitor changes between pro-teomes through quantitative profiling. The bottom-up strategy for protein identification has clearly matured, although a greater depth of characterization can undoubtedly be benefited by incor-poration of top-down (and middle-down) MS experiments. As recognized from the beginning, the most important components of a sample are rarely the most abundant. Separation of high- from low-abundance components has clear benefits for unmask-ing these samples for MS detection. Despite improvements in MS instrumentation, proteome fractionation will continue to play a critical role in defining the proteome experiment.
While recognizing that higher resolution is always desired dur-ing separation, there is also a need to focus on the reproducibility of the experiment. Clinical profiling brings with it the importance of including biological replicates and so all aspects of the ana-lytical workflow must be precisely controlled so that data from multiple experiments can be compared. If a proper comparison is to be made, then there is a pressing need to standardize the strategies for proteome extraction and preservation (i.e., ahead of separation). Thus, an objective of achieving complete (or near-complete) proteome solubilization can surely benefit from the inclusion of solubilizing additives such as SDS. With a focus on MS-compatible surfactants, one may also consider ‘separation compatible’ solubilizing additives, as it is most important that this stage of the workflow accommodate such additives.
With two clear strategies for proteome fractionation (i.e., chro-matographic vs electrophoretic), one should also recognize that both can contribute to a more complete separation of the sam-ple. Electrophoretic and chromatographic approaches are often orthogonal, and thus a multidimensional approach to proteome fractionation should consider both strategies. The direct cou-pling of chromatography with MS (i.e., RPLC-MS) epitomizes a fully MS-compatible fractionation approach. Similar coupling of electro phoresis to chromatography, as well as electrophoresis to MS, may prove beneficial for proteome fractionation.
Finally, the throughput of a proteomic experiment must become a forefront consideration of strategies which incorporate proteome fractionation. The generation of a greater number of fractions may be possible through inclusion of added dimensions of separa-tion. However, the practicality of such an experiment is brought to question, particularly if multiple samples are to be profiled. Increased throughput in the MS detection strategy can be realized through use of parallel mass analyzers [105,106], or perhaps more simply through a focused approach which would recognize and prioritize fractions of interest [19].
Financial & competing interests disclosureThis work was supported in part by the National Sciences and Engineering Research Council of Canada. The Gelfree technology described in this article is licensed by Dalhousie University to Protein Discovery Inc. The authors have no other relevant affiliations or financial involvement with any organi-zation or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript apart from those disclosed.
No writing assistance was utilized in the production of this manuscript.
Doucette, Tran, Wall & Fitzsimmons
www.expert-reviews.com 797
Review
ReferencesPapers of special note have been highlighted as:• of interest•• of considerable interest
1 Demirev PA, Fenselau C. Mass spectrometry for rapid characterization of microorganisms. Ann. Rev. Anal. Chem. 1, 71–93 (2008).
2 Wynne C, Fenselau C, Demirev PA, Edwards N. Top-down identification of protein biomarkers in bacteria with unsequenced genomes. Anal. Chem. 81(23), 9633–9642 (2009).
3 Michalski A, Cox J, Mann M. More than 100,000 detectable peptide species elute in single shotgun proteomics runs but the majority is inaccessible to data-dependent LC-MS/MS. J. Proteome Res. 10(4), 1785–1793 (2011).
4 Aebersold R, Mann M. Mass spectrometry-based proteomics. Nature 422(6928), 198–207 (2003).
5 Washburn M, Wolters D, Yates J. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19(3), 242–247 (2001).
7 Tran JC, Wall MJ, Doucette AA. Evaluation of a solution isoelectric focusing protocol as an alternative to ion exchange chromatography for charge-based proteome prefractionation. J. Chromatogr. B 877(8–9), 807–813 (2009).
8 Gygi S, Rist B, Gerber S, Turecek F, Gelb M, Aebersold R. Quantitative analysis
of complex protein mixtures using isotope-coded affinity tags. Nat. Biotechnol. 17(10), 994–999 (1999).
9 Craig R, Cortens J, Beavis R. The use of proteotypic peptide libraries for protein identification. Rapid Commun. Mass Spectrom. 19(13), 1844–1850 (2005).
10 Mallick P, Schirle M, Chen SS et al. eComputational prediction of proteotypic peptides for quantitative proteomics RID A-5619–2008. Nat. Biotechnol. 25(1), 125–131 (2007).
11 Shen YF, Zhao R, Belov ME et al. Packed capillary reversed-phase liquid chromatography with high-performance electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry for proteomics. Anal. Chem. 73(8), 1766–1775 (2001).
12 Vellaichamy A, Tran JC, Catherman AD et al. Size-sorting combined with improved nanocapillary liquid chromatography-mass spectrometry for identification of intact proteins up to 80 kDa. Anal. Chem. 82(4), 1234–1244 (2010).
13 Schmidt A, Kellermann J, Lottspeich F. A novel strategy for quantitative proteomics using isotope-coded protein labels. Proteomics 5(1), 4–15 (2005).
14 Ong SE, Blagoev B, Kratchmarova I et al. Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol. Cell. Proteomics 1(5), 376–386 (2002).
15 Geiger T, Cox J, Ostasiewicz P, Wisniewski JR, Mann M. Super-SILAC mix for quantitative proteomics of human tumor tissue. Nat. Methods 7(5), 383–385 (2010).
16 Meyer B, Papasotiriou DG, Karas M. 100% protein sequence coverage: a modern form of surrealism in proteomics. Amino Acids 41(2), 291–310 (2011).
17 Righetti PG, Boschetti E, Lomas L, Citterio A. Protein Equalizer (TM) Technology: The quest for a ‘democratic proteome’. Proteomics 6(14) 3980–3992 (2006)
20 Strahl BD, Allis CD. The language of covalent histone modifications. Nature 403(6765), 41–45 (2000).
21 Kelleher R, Zubarev R, Bush K et al. Localization of labile posttranslational modifications by electron capture dissociation: the case of g-carboxyglutamic acid. Anal. Chem. 71(19), 4250–4253 (1999).
22 Mirgorodskaya E, Roepstorff P, Zubarev R. Localization of O-glycosylation sites in peptides by electron capture dissociation in a fourier transform mass spectrometer. Anal. Chem. 71(20), 4431–4436 (1999).
Key issues
• While multiple separation strategies have been optimized for peptide-level separation, it should not be assumed that these approaches can be directly translated to fractionate proteins with equivalent efficiency. Thus, platforms that present poor recovery of proteins should be avoided in favor of others that have been optimized for protein fractionation.
• Concerning protein recovery, one must consider maintaining the solubility of all protein components not only during the fractionation step, but also before and immediately after separation.
• Gel-free approaches can maximize protein recovery as they permit intact proteins to be isolated in solution phase ahead of top-down or bottom-up mass spectrometry (MS).
• Intact separation platforms are most beneficial when sample fractionation occurs according to a predictable property of the protein. This would include molecular weight, hydrophobicity, isoelectric point or other intrinsic properties of the sample.
• Solubilizing additives have an important role to play ahead of MS. These additives maintain protein solubility during separation, among other benefits.
• While MS-friendly surfactants are available, several strategies exist for efficient removal of SDS. When selecting an appropriate sample clean-up strategy, one should be concerned with the efficiency of purification, as well as the recovery rate of the proteins.
• A greater number of fractions implies more MS analysis time. One should optimize the fractionation strategy according to the required throughput of the experiment and avoid collecting more fractions, particularly if the resolution of the platform does not justify it.
Intact proteome fractionation strategies compatible with mass spectrometry
Expert Rev. Proteomics 8(6), (2011)798
Review
23 Cooper H, Hakansson K, Marshall A. The role of electron capture dissociation in biomolecular analysis. Mass Spectrom. Rev. 24(2), 201–222 (2005).
24 Han X, Jin M, Breuker K, McLafferty FW. Extending top-down mass spectrometry to proteins with masses greater than 200 kilodaltons. Science 314(5796), 109–112 (2006).
25 Parks BA, Jiang L, Thomas PM et al. Top-down proteomics on a chromatographic time scale using linear ion trap Fourier transform hybrid mass spectrometers. Anal. Chem. 79(21), 7984–7991 (2007).
26 Roth MJ, Parks BA, Ferguson JT, Boyne MT II, Kelleher NL. ‘Proteotyping’: population proteomics of human leukocytes using top down mass spectrometry. Anal. Chem. 80(8), 2857–2866 (2008).
27 Fang X, Zhang WW. Affinity separation and enrichment methods in proteomic analysis. J. Proteomics 71, 284–303 (2008).
28 Shevchenko A, Wilm M, Vorm O, Mann M. Mass spectrometric sequencing of proteins from silver stained polyacrylamide gels. Anal. Chem. 68(5), 850–858 (1996).
29 Zuo X, Speicher DW. A method for global analysis of complex proteomes using sample prefractionation by solution isoelectrofocusing prior to two-dimensional electrophoresis. Anal. Biochem. 284(2), 266–278 (2000).
30 Gygi SP, Aebersold R. Mass spectrometry and proteomics. Curr. Opin. Chem. Biol. 4(5), 489–494 (2000).
31 Laemmli UK. Cleavage of structural proteins during assembly of head of bacteriophage-T4. Nature 227(5259), 680–685 (1970).
32 Schirle M, Heurtier MA, Kuster B. Profiling core proteomes of human cell lines by one-dimensional PAGE and liquid chromatography-tandem mass spectrometry. Mol. Cell. Proteomics 2(12), 1297–1305 (2003).
33 Wall DB, Kachman MT, Gong SY et al. Isoelectric focusing nonporous RP HPLC: a two-dimensional liquid-phase separation method for mapping of cellular proteins with identification using MALDI-TOF mass spectrometry. Anal. Chem. 72(6), 1099–1111 (2000).
34 Whitelegge J, Gundersen C, Faull K. Electrospray-ionization mass spectrometry of intact intrinsic membrane proteins. Prot. Sci. 7(6), 1423–1430 (1998).
35 Chong BE, Yan F, Lubman DM, Miller FR. Chromatofocusing nonporous reversed-phase high-performance liquid chromatography/electrospray ionization time-of-flight mass spectrometry of proteins from human breast cancer whole cell lysates: a novel two-dimensional liquid chromatography/mass spectrometry method. Rapid Commun. Mass Spectrom. 15(4), 291–296 (2001).
36 Lecchi P, Abramson FP. Analysis of biopolymers by size-exclusion chromatography mass spectrometry. J. Chromatogr. A 828(1–2), 509–513 (1998).
37 Lecchi P, Gupte AR, Perez RE, Stockert LV, Abramson FP. Size-exclusion chromatography in multidimensional separation schemes for proteome analysis. J. Biochem. Biophys. Methods 56(1–3), 141–152 (2003).
38 Cossu G, Righetti PG. Resolution of g-gamma and a-gamma fetal hemoglobin tetramers in immobilized pH gradients. J. Chromatogr. A 398, 211–216 (1987).
39 Jensen PK, Pasa-Tolic L, Peden KK et al. Mass spectrometic detection for capillary isoelectric focusing separations of complex protein mixtures. Electrophoresis 21(7), 1372–1380 (2000).
40 Righetti PG, Castagna A, Antonioli P, Boschetti E. Prefractionation techniques in proteome analysis: the mining tools of the third millennium. Electrophoresis 26(2), 297–319 (2005).
42 Ros A, Faupel M, Mees H et al. Protein purification by off-gel electrophoresis. Proteomics 2(2), 151–156 (2002).
43 Michel PE, Reymond F, Arnaud IL, Josserand J, Girault HH, Rossier JS. Protein fractionation in a multicompartment device using Off-Gel™ isoelectric focusing. Electrophoresis 24(1–2), 3–11 (2003).
44 de Godoy LMF, Olsen JV, Cox J et al. Comprehensive mass-spectrometry-based proteome quantification of haploid versus diploid yeast. Nature 455(7217), 1251–1260 (2008).
45 Zuo X, Speicher DW. Comprehensive ana lysis of complex proteomes using microscale solution isoelectrofocusing prior to narrow pH range two-dimensional electrophoresis. Proteomics 2(1), 58–68 (2002).
46 Hannig K. Continuous free-flow electrophoresis as an analytical and preparative method in biology. J. Chromatogr. 159(1), 183–191 (1978).
47 Ouvry-Patat SA, Torres MP, Quek H et al. Free-flow electrophoresis for top-down proteomics by Fourier transform ion cyclotron resonance mass spectrometry. Proteomics 8(14), 2798–2808 (2008).
48 Tran JC, Doucette AA. Rapid and effective focusing in a carrier ampholyte solution lsoelectric focusing system: a proteome prefractionation tool. J. Proteome Res. 7(4), 1761–1766 (2008).
49 Righetti PG, Wenisch E, Faupel M. Preparative protein-purification in a multi-compartment electrolyzer with Immobiline membranes. J. Chromatogr. A 475, 293–309 (1989).
50 Herbert B, Righetti PG. A turning point in proteome analysis: sample prefractionation via multicompartment electrolyzers with isoelectric membranes. Electrophoresis 21(17), 3639–3648 (2000).
51 Nesatyy VJ, Ross NW. Recovery of intact proteins from silver stained gels. Analyst 127(9), 1180–1187 (2002).
52 Rabilloud T. Two-dimensional gel electrophoresis in proteomics: old, old fashioned, but it still climbs up the mountains. Proteomics 2(1), 3–10 (2002).
53 Lewis U, Clark M. Preparative methods for disk electrophoresis with special reference to isolation of pituitary hormones. Anal. Biochem. 6(4), 303–315 (1963).
54 Jovin T, Naughton M, Chrambach A. Apparatus for preparative temperature-regulated polyacrylamide gel electrophoresis. Anal. Biochem. 9(3), 351–369 (1964).
55 Ornstein L. Disc electrophoresis – I: Background and theory. Ann. NY Acad. Sci. 121(A2), 321–351 (1964).
56 Davis B. Disc electrophoresis – 2: Method and application to human serum proteins. Ann. NY Acad. Sci. 121(A2), 404–427 (1964).
Doucette, Tran, Wall & Fitzsimmons
www.expert-reviews.com 799
Review
57 Masuoka J, Glee PM, Hazen KC. Preparative isoelectric focusing and preparative electrophoresis of hydrophobic Candida albicans cell wall proteins with in-line transfer to polyvinylidene difluoride membranes for sequencing. Electrophoresis 19(5), 675–678 (1998).
58 Meng F, Cargile B, Patrie S, Johnson J, McLoughlin S, Kelleher N. Processing complex mixtures of intact proteins for direct analysis by mass spectrometry. Anal. Chem. 74(13), 2923–2929 (2002).
59 Andersen P, Heron I. Simultaneous electroelution of whole SDS-polyacrylamide gels for the direct cellular analysis of complex protein mixtures. J. Immunol. Methods 161(1), 29–39 (1993).
60 Tran JC, Doucette AA. Gel-eluted liquid fraction entrapment electrophoresis: an electrophoretic method for broad molecular weight range proteome separation. Anal. Chem. 80(5), 1568–1573 (2008).
61 Tran JC, Doucette AA. Multiplexed size separation of intact proteins in solution phase for mass spectrometry. Anal. Chem. 81(15), 6201–6209 (2009).
62 Botelho D, Wall MJ, Vieira DB, Fitzsimmons S, Liu F, Doucette A. Top-down and bottom-up proteomics of SDS-containing solutions following mass-based separation. J. Proteome Res. 9(6), 2863–2870 (2010).
63 Lee JE, Kellie JF, Tran JC et al. A robust two-dimensional separation for top-down tandem mass spectrometry of the low-mass proteome. J. Am. Soc. Mass Spectrom. 20(12), 2183–2191 (2009).
65 Wang H, Clouthier SG, Galchev V et al. Intact-protein-based high-resolution three-dimensional quantitative analysis system for proteome profiling of biological fluids. Mol. Cell. Proteomics 4(5), 618–625 (2005).
66 Opiteck GJ, Lewis KC, Jorgenson JW, Anderegg RJ. Comprehensive on-line LC/LC/MS of proteins. Anal. Chem. 69(8), 1518–1524 (1997).
67 Assiddiq BF, Snijders AP, Chong PK, Wright PC, Dickman MJ. Identification and characterization of Sulfolobus
solfataricus P2 proteome using multidimensional liquid phase protein separations. J. Proteome Res. 7(6), 2253–2261 (2008).
68 Kreunin P, Urquidi V, Lubman DM, Goodison S. Identification of metastasis-associated proteins in a human tumor metastasis model using the mass-mapping technique. Proteomics 4(9), 2754–2765 (2004).
69 Kreunin P, Zhao J, Rosser C, Urquidi V, Lubman DM, Goodison S. Bladder cancer associated glycoprotein signatures revealed by urinary proteomic profiling. J. Proteome Res. 6(7), 2631–2639 (2007).
70 Moritz RL, Ji H, Schutz F et al. A proteome strategy for fractionating proteins and peptides using continuous free-flow electrophoresis coupled off-line to reversed-phase high-performance liquid chromatography. Anal. Chem. 76(16), 4811–4824 (2004).
71 Sneekes E, Han J, Elliot M et al. Accurate molecular weight analysis of histones using FFE and RP-HPLC on monolithic capillary columns. J. Sep. Sci. 32(15–16), 2691–2698 (2009).
72 Speers AE, Wu CC. Proteomics of integral membrane proteins-theory and application. Chem. Rev. 107(8), 3687–3714 (2007).
73 Zhang N, Chen R, Young N et al. Comparison of SDS- and methanol-assisted protein solubilization and digestion methods for Escherichia coli membrane proteome analysis by 2-D LC-MS/MS. Proteomics 7(4), 484–493 (2007).
74 Breaux GA, Green-Church KB, France A, Limbach PA. Surfactant-aided, matrix assisted laser desorption/ionization mass spectrometry of hydrophobic and hydrophilic peptides. Anal. Chem. 72(6), 1169–1174 (2000).
75 Tummala R, Limbach PA. Effect of sodium dodecyl sulfate micelles on peptide mass fingerprinting by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 18(18), 2031–2035 (2004).
76 Rundlett KL, Armstrong DW. Mechanism of signal suppression by an ionic surfactants in capillary electrophoresis electrospray ionization mass spectrometry. Anal. Chem. 68(19), 3493–3497 (1996).
77 Beavis RC, Chait BT. Rapid, sensitive analysis of protein mixtures by mass-spectrometry. Proc. Natl. Acad. Sci. USA 87(17), 6873–6877 (1990).
78 Ikonomou MG, Blades AT, Kebarle P. Investigations of the electrospray interface
for liquid-chromatography mass-spectrometry. Anal. Chem. 62(9), 957–967 (1990).
79 Kawasaki H, Suzuki K. Separation of peptides dissolved in a sodium dodecyl-sulfate solution by reversed-phase liquid-chromatography – removal of sodium dodecyl-sulfate from peptides using an ion-exchange precolumn. Anal. Biochem. 186(2), 264–268 (1990).
80 Pitt-Rivers R, Impiombato AF. The binding of sodium dodecyl sulphate to various proteins. Biochem. J. 109(5), 825–830 (1968).
81 Zaman Z, Verwilghen RL. Quantitation of proteins solubilized in sodium dodecyl sulfate mercaptoethanol tris electrophoresis buffer. Anal. Biochem. 100(1), 64–69 (1979).
82 Carraro U, Rizzi C, Sandri M. Effective recovery by KCl precipitation of highly diluted muscle proteins solubilized with sodium dodecyl-sulfate. Electrophoresis 12(12), 1005–1010 (1991).
83 Wessel D, Flugge UI. A method for the quantitative recovery of protein in dilute-solution in the presence of detergents and lipids. Anal. Biochem. 138(1), 141–143 (1984).
84 Puchades M, Westman A, Blennow K, Davidsson P. Removal of sodium dodecyl sulfate from protein samples prior to matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 13(5), 344–349 (1999).
85 Amons R, Schrier PI. Removal of sodium dodecyl-sulfate from proteins and peptides by gel-filtration. Anal. Biochem. 116(2), 439–443 (1981).
86 Vissers JP, Hulst WP, Chervet JP, Snijders HM, Cramers CA. Automated on-line ionic detergent removal from minute protein/peptide samples prior to liquid chromatography electrospray mass spectrometry. J. Chromatogr. B 686(2), 119–128 (1996).
87 Kapp OH, Vinogradov SN. Removal of sodium dodecyl-sulfate from proteins. Anal. Biochem. 91(1), 230–235 (1978).
88 Li F, Dong MQ, Miller LJ, Naylor S. Efficient removal of sodium dodecyl sulfate (SDS) enhances analysis of proteins by SDS-polyacrylamide gel electrophoresis coupled with matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Commun. Mass Spectrom. 13(5), 464–466 (1999).
Intact proteome fractionation strategies compatible with mass spectrometry
Expert Rev. Proteomics 8(6), (2011)800
Review
89 Dong MQ, Baggetto LG, Falson P, LeMaire M, Penin F. Complete removal and exchange of sodium dodecyl sulfate bound to soluble and membrane proteins and restoration of their activities, using ceramic hydroxyapatite chromatography. Anal. Biochem. 247(2), 333–341 (1997).
90 Fox JL, Stevens SE, Taylor CP, Poulsen LL. SDS removal from protein by polystyrene beads. Anal. Biochem. 87(1), 253–256 (1978).
91 Jeno P, Scherer PE, Manningkrieg U, Horst M. Desalting electroeluted proteins with hydrophilic interaction chromatography. Anal. Biochem. 215(2), 292–298 (1993).
92 Phillips HM. Method for rapid removal of sodium dodecyl-sulfate from polyacrylamide gels. Anal. Biochem. 117(2), 398–401 (1981).
93 Zhou J, Li J, Li J, Chen P, Wang X, Liang S. Dried polyacrylamide gel absorption: a method for efficient elimination of the interferences from SDS-solubilized protein samples in mass spectrometry-based proteome analysis. Electrophoresis 31(23–24), 3816–3822 (2010).
94 Lu XN, Zhu HN. Tube-gel digestion – a novel proteomic approach for high throughput analysis of membrane proteins. Mol. Cell. Proteomics 4(12), 1948–1958 (2005).
95 Wisniewski JR, Zougman A, Nagaraj N, Mann M. Universal sample preparation method for proteome analysis. Nat. Methods 6(5), 359–362 (2009).
96 Zhou J, Zhou T, Cao R et al. Evaluation of the application of sodium deoxycholate to proteomic ana lysis of rat hippocampal plasma membrane. J. Proteome Res. 5(10), 2547–2553 (2006).
97 Masuda T, Saito N, Tomita M, Ishihama Y. Unbiased quantitation of Escherichia coli membrane proteome using phase transfer surfactants. Mol. Cell Proteomics 8(12), 2770–2777 (2009).
98 Ishihama Y, Katayama H, Asakawa N. Surfactants usable for electrospray ionization mass spectrometry. Anal. Biochem. 287(1), 45–54 (2000).
99 Suder P, Bierczynska A, Konig S, Silberring J. Acid-labile surfactant assists in-solution digestion of proteins resistant to enzymatic attack. Rapid Commun. Mass Spectrom. 18(7), 822–824 (2004).
100 Chen EI, Cociorva D, Norris JL, Yates JR 3rd. Optimization of mass spectrometry-compatible surfactants for shotgun proteomics. J. Proteome Res. 6(7), 2529–2538 (2007).
101 Kadiyala CSR, Tomechko SE, Miyagi M. Perfluorooctanoic acid for shotgun proteomics. PLoS One 5(12), e15332 (2010).
102 Lin Y, Zhou J, Bi D, Chen P, Wang X, Liang S. Sodium-deoxycholate-assisted tryptic digestion and identification of proteolytically resistant proteins. Anal. Biochem. 377(2), 259–266 (2008).
103 Zhang N, Li L. Effects of common surfactants on protein digestion and matrix-assisted laser desorption/ionization mass spectrometric analysis of the digested peptides using two-layer sample preparation. Rapid Commun. Mass Spectrom. 18(8), 889–896 (2004).
104 Wall MJ, Crowell AM, Simms GH, Liu F, Doucette AA. Implications of partial tryptic digestion in organic-aqueous solvent systems for bottom-up proteome analysis. Anal. Chim. Acta 703(2), 194–203 (2011).
106 Tabert AM, Goodwin MP, Duncan JS, Fico CD, Cooks RG. Multiplexed rectilinear ion trap mass spectrometer for high-throughput analysis. Anal. Chem. 78(14), 4830–4838 (2006).
107 Lee JE, Kellie JF, Tran JC et al. A robust two-dimensional separation for top-down tandem mass spectrometry of the low-mass proteome. J. Am. Soc. Mass Spectrom. 20(12), 2183–2191 (2009).
