Enhanced detergent extraction for analysis of membrane proteomes by two-dimensional gel electrophoresis
© Churchward et al; licensee BioMed Central Ltd. 2005
Received: 23 January 2005
Accepted: 07 June 2005
Published: 07 June 2005
The analysis of hydrophobic membrane proteins by two-dimensional gel electrophoresis has long been hampered by the concept of inherent difficulty due to solubility issues. We have optimized extraction protocols by varying the detergent composition of the solubilization buffer with a variety of commercially available non-ionic and zwitterionic detergents and detergent-like phospholipids.
After initial analyses by one-dimensional SDS-PAGE, quantitative two-dimensional analyses of human erythrocyte membranes, mouse liver membranes, and mouse brain membranes, extracted with buffers that included the zwitterionic detergent MEGA 10 (decanoyl-N-methylglucamide) and the zwitterionic lipid LPC (1-lauroyl lysophosphatidylcholine), showed selective improvement over extraction with the common 2-DE detergent CHAPS (3 [(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate). Mixtures of the three detergents showed additive improvements in spot number, density, and resolution. Substantial improvements in the analysis of a brain membrane proteome were observed.
This study demonstrates that an optimized detergent mix, coupled with rigorous sample handling and electrophoretic protocols, enables simple and effective analysis of membrane proteomes using two-dimensional electrophoresis.
Historically, the proteomic analysis of hydrophobic membrane proteins has been considered to be difficult within the bounds of conventional protocols for two-dimensional gel electrophoresis (2-DE). The nature of first dimension isoelectric focusing (IEF) requires that proteins be thoroughly solubilized as they are subjected to an electric field in which they migrate to their isoelectric point, by definition the state of lowest possible net charge and thus lowest solubility in aqueous environments. In addition to being highly hydrophobic, many integral membrane proteins tend to be very large: human Ca2+ channels have 24 transmembrane helices and are typically > 200 kDa , and tyrosine kinase receptors are frequently > 100 kDa [2, 3]. This leads to two major problems in the preparation of membrane protein samples for 2-DE. First, effectively extracting membrane proteins into a detergent that is IEF compatible. Second, maintaining protein solubility throughout loading onto IPG strips and the subsequent first dimension IEF separation. Although highly efficient membrane protein extractions are routinely carried out with a detergent such as SDS for one-dimensional PAGE, SDS is incompatible with IEF due to the charged head group. To overcome this, SDS solubilized samples often undergo solvent or acid precipitation to remove or reduce SDS and lipids. Despite these harsh treatments and even subsequent treatment of the precipitate with a strong base , delipidation by solvent extraction is often cited as enhancing protein recovery [5, 6] without discussion of the loss or modification of proteins during precipitation. For example, highly hydrophobic proteins (such as proteolipids) and proteins with particular post-translational modifications (such as palmitoylation) are capable of partitioning into the solvent phase [7–9], and TCA treatment can cause acid hydrolysis of proteins or alter post-translational modifications. Additionally, some early general problems with effectively separating hydrophobic proteins by 2-DE have led to widespread general disregard for the analysis of membrane proteins, particularly in the development of alternate proteomic approaches [4, 5, 10, 11].
Since membrane proteins comprise approximately 30% of human proteins, and may account for substantially more cellular functions, the focus on soluble proteins in so-called 'full' proteomic analyses is somewhat concerning. There is evidence that optimization of extraction conditions by alteration of buffers, chaotropes, and detergents is sufficient to reliably achieve high-resolution maps of membrane proteins [13–16]. To this end we have sought simple alternatives to optimize the detergent conditions used to extract proteins from native membranes by systematic analysis of the solubilization properties of a wide range of commercially available non-ionic and zwitterionic detergents and a range of natural and synthetic detergent-like lipids [17–19]. Using proven synthetic detergents, together with more native lipophilic agents, we find that combinations of these reagents generally improve the resolution of membrane proteomes analyzed by 2-DE, providing for select improvements in the yields of specific proteins. Optimization of conditions for particular samples remains a key to any successful analysis [20–22].
Results & Discussion
1D SDS-PAGE of RBC membrane
Summary of detergents tested using systematic 1D SDS-PAGE analysis. Overall extraction efficacy analyzed by 1D SDS-PAGE or 2-DE separation is expressed qualitatively relative to SDS extraction (for 1D analysis) or CHAPS extraction (for 2-DE). + indicates compatibility but poor perfomance, ++ indicates similar or slightly worse than CHAPS extraction, +++ indicates performance equal to or better than CHAPS, – indicates incompatibility.
Comments & Rationale
Poor extraction of hydrophobic and high molecular weight proteins
Group of nonionic detergents commonly used for protein purification [35,36]
Sulfobetaine-based detergents reported to improve membrane protein extraction [24-26]
Zwittergent® 3–10/SB 3–10d
LPC (synthetic, lauroyl chain)e
LPC (egg, mixed chain)e
LPS (bovine brain)f
Anionic lysophospholipid, incompatible with IEF
LPE (egg, mixed chain)g
Zwitterionic lysophospholipid; low solubility in high urea buffer
LPG (egg, mixed chain)h
Anionic lysophospholipid, incompatible with IEF
LPA (egg, mixed chain)i
Anionic lysophospholipid, incompatible with IEF
cardiolipin (bovine heart)
Anionic lipid, incompatible with IEF, low solubility in high urea buffer
Synthetic fatty acid; low solubility in high urea buffer
Medium chain fatty acid; low solubility in high urea buffer
free fatty acids (mixed)
Mixed natural fatty acids; low solubility in high urea buffer
Cationic lipid used as a transfection reagent , IEF incompatible
Uncharged monoacylated lipid, very low solubility in high urea buffer
Nonionic detergent used to study membrane proteins 
Amphipathic cyclic glycerol conjugate
Amphipathic heterocyclic metabolite of tryptophan; ionizable at low pH
2-DE analysis of Red Blood Cell membrane
RBC membranes extracted with the 4% total detergent mixtures of CHAPS and LPC showed general evidence of improved spot densities (Fig. 2B), as well as specific improvements in terms of reduced horizontal streaking in the intermediate molecular weight region (Fig. 2A). Automated spot detection and quantitative comparative analysis using Progenesis Workstation software identified specific changes in the protein pattern. Specific areas of the gels showed improvement relative to parallel 4% CHAPS control gels (Fig. 2Bi–iv). In particular, a prominent spot corresponding to band III was clearly observed [15, 16], as well as a 2.2 ± 0.1 – fold increase in the density of a string of spots relative to the CHAPS extract (Fig. 2Bii). Three additional unique spots were observed when extracted with 3% CHAPS : 1% LPC (Fig 2Biii–iv). RBC membrane samples extracted with 3% CHAPS : 1% MEGA 10 (Fig. 2C) yielded protein maps of generally equivalent resolution to both the 3% CHAPS : 1% LPC and the 4% CHAPS maps, but did not resolve the protein band III as effectively as 3% CHAPS : 1% LPC.
In order to examine the overlapping effects of both these test detergents, but ameliorate the observed losses of protein, 0.5% of each was mixed with 3% CHAPS and tested in the extraction and 2-DE analysis of RBC membrane proteins (Fig. 2D). Extraction with 3% CHAPS : 0.5% LPC : 0.5% MEGA 10 does not yield the extent of differences identified in the 3% CHAPS : 1% LPC extracted condition, although the maps show general improvements over the control CHAPS condition that correlate with the improvements seen in the two individual detergent extractions (Figs. 2B, C). The density of the indicated string of proteins was increased an average of 1.7 ± 0.2 – fold over CHAPS (Fig. 2Dii). In general then, the addition of LPC to the extraction buffer enhances both protein recovery and resolution in the subsequent 2D protein maps.
Our initial findings using the RBC membrane as a model system lead us to expand the analyses to additional tissue types. Mouse brain membranes  and mouse liver membranes were chosen due to their availability, and broad international interest in improved analyses of these tissue proteomes.
2-DE analysis of mouse brain membrane
Additional 2-DE Analyses
Interestingly, extracting mouse liver membranes with the same detergent combinations described above resulted in protein maps that were highly similar, with very limited improvements. Automated analysis indicated almost complete overlap of the resulting 2-DE protein patterns (Fig 5), with the specific and substantial recovery of one additional protein spot. We interpret the marked similarity in these liver protein profiles, relative to the differences seen in the RBC and brain samples, to be due to variability between tissues in terms of relative homogenization/extraction efficiency and compatibility with our current buffer system.
To control for possible differences arising from the changing CHAPS concentration in these test extraction buffers, mouse brain membranes were also extracted with 5% total detergent (5% CHAPS or 4% CHAPS : 0.5% LPC : 0.5% MEGA 10) and analyzed in parallel with membranes extracted with 4% total detergent. No significant difference in overall spot pattern or specific differences as described above was observed between the 5% and the 4% total detergent mixtures (data not shown), indicating that the differences described here are specifically attributable to the addition of LPC and MEGA 10 as solubilizing agents. Indeed, overall, membrane protein patterns were generally of somewhat lower resolution when the CHAPS concentration or total detergent concentration was increased to 5%.
During initial experiments we found total protein load to be the most significant variable confounding quantitative analyses. As such, great care was taken to ensure that the analyses meaningfully tested protein extraction and solubilization efficiency, in isolation from complicating variables. Simply, the goal was to compare reagents and conditions, not to compare different final total protein loads by 2-DE. Initially many protein samples were quantified using a modified Folin total protein assay (RC DC Protein Assay kit, BioRad). Colourimetric assays of this type (eg. Bradford, Lowry, BCA, and so forth) perform acceptably under many circumstances requiring routine normalization of a series of very similar samples. However one of several limitations of such total protein assays is a marked sensitivity to interfering substances, including components of typical IEF solubilization solutions such as detergents, reducing agents, and urea. In our experiments, detergents and detergent concentrations were systematically altered and combined. Not unexpectedly, we observed substantial variability in the results of the total protein assay, depending upon the solubilizing reagents present. The complications of applying systematic corrective controls, or of preparing separate standard curves for each of the solubilization conditions tested, simply increased the potential for error. Regardless, separate standard curves are not even feasible in the case of the RC DC assay, as urea causes a saturating false positive signal.
We have found that the EZQ Protein Quantitation kit (Molecular Probes) is insensitive to the nature and concentrations of detergent in all samples tested. In this assay format, the immobilized protein sample is washed exhaustively with methanol to remove components of the solubilization solution prior to addition of the fluorescent protein detection reagent. Thus, the chemistry of the assay proceeds in the absence of potentially confounding contaminants. In extensive comparisons, there were no significant differences in standard protein assay curves regardless of the type or quantity of detergent included (data not shown). Additionally, the method proved quite sensitive (routine detection of 0.030 μg of total protein/spot, or 15 μg/ml); this is fully 10-fold more sensitive and requires 4-fold less material than the RC DC Assay. Thus, as the chemistry of the assay was not altered under our different experimental conditions, we are confident that the improvements observed in our final protein maps were truly the result of differences in extraction and solubilization efficiency, and not artifacts generated by erroneous total protein assays leading to inconsistent total protein IEF loads between different test conditions. Although the EZQ protein assay certainly has its caveats, not least of which is cost, it does offer distinct benefits that support its utility in these and other ongoing proteomic analyses.
In order to optimize recovery of hydrophobic proteins for 2-DE, we have sought a simple, direct solution to the problem of protein extraction and solubility during IEF. The systematic screening and combination of commercially available detergents offers a direct, inexpensive, and convenient method for optimizing the conditions of IEF without entering into the complexities of a systematic synthesis of new detergents based on specific base molecules, or the potential losses or modification of proteins associated with solvent extraction techniques. Coupled with our ability to effectively analyze membrane proteomes using 2-DE  the resulting findings should also prove of use in defining optimized combinations of extraction reagents for use with alternate protein separation protocols.
Based on the hypothesis that highly lipophilic molecules (albeit at lower total concentrations than can be achieved with the more standard detergents), might better mimic native lipid-membrane protein interactions and thus improve protein solubilization, we found that LPC can substantially augment the extraction of membrane proteins from different sources. This finding does not obviate the need for optimization of extraction and 2-DE conditions for different samples, but does provide a powerful, widely available and reasonably priced alternative that can be readily tested in parallel with more routine solubilization reagents. Rigorous testing of protein assays ensured that these findings reflect a true effect on extraction and protein solubility, rather than an artifact of inconsistent protein loads between different 2-DE analyses. Notably, LPC and MEGA 10 provided particularly marked improvements in the resolution of the mouse brain membrane proteome.
L-α-lysophosphatidylcholine lauroyl, urea, tris acetate, lauric acid, pH 3–10 ampholytes, ammonium persulfate, decyl-N,N-dimethyl-3-ammonio-1-propanesulfonate (SB 3–10), amidosulfobetaine-14 (ASB-14), DL-α-O-benzylglycerol, tributylphosphine (TBP), HEPES, sodium orthovanadate, staurosporine, cantharidin, and components of the broad spectrum protease inhibitor cocktail  were purchased from Sigma (St. Louis, Missouri). IPG strips (pH 3–10), 30% acrylamide/bisacrylamide solution, low melting agarose, Sypro Ruby, 10×TGS running buffer, RC DC Protein Assay Kit, bovine γ-globulin, and SDS were from BioRad (Hercules, California). EZQ Protein Quantitation Kit was from Molecular Probes (Eugene, OR), Zwittergent® 3–10 was from Calbiochem (La Jolla, California), and CHAPS was from Anatrace (Maumee, Ohio). 1,2-dioleoyloxy-3-(dimethylamino)propane, 5,7-docosadiynoic acid, and 1-oleoyl-sn-glycerol were from Toronto Research Chemicals (Toronto, Ontario). Bovine brain L-α-lysophosphatidylserine, egg L-α-lysophosphatidylcholine, egg L-α-lysophosphatidylethanolamine, egg L-α-lysophosphatidylglycerol, egg L-α-lysophosphatidic acid, bovine heart cardiolipin, and free fatty acids were from Doosan Serdary Research (Toronto, Ontario). Thiourea was from Fisher Scientific (Hampton, New Hampshire), and PBS, DTT, octanoyl-N-methylglucamide (MEGA 8), nonanoyl-N-methylglucamide (MEGA 9), decanoyl-N-methylglucamide (MEGA 10), TEMED, glycerol, 40% acrylamide solution, and octaethylene glycol monododecyl ether (C12E8) were from Bio Basic Inc. (Markham, Ontario). Narrow range ampholytes (pH 2.5–4, 3.5–5, 5–7, 7–9, and 8–9.5) were from Fluka (Buchs, Switzerland), and tryptophol and trans, trans- farnesol was from Aldrich (St. Louis, Missouri). All other chemicals were of at least analytical grade.
Red Blood Cell membrane preparation
Packed RBC were obtained from Canadian Blood Services, (Calgary, AB) and washed 3× with isotonic buffer (20 mM sodium phosphate pH 7.4, 0.9% NaCl). RBC ghosts were prepared according to the method of Chernomordik  with slight modifications. Cells were lysed osmotically in hypotonic lysis buffer (5 mM sodium phosphate pH 7.4, protease inhibitor cocktail , 5 mM DTT) for 20 minutes on ice. The lysate was flash frozen in a dry ice / ethanol bath, thawed, and membranes were collected by centrifugation (3000×g, 20 min, 4°C). Pellets were washed with wash buffer (20 mM sodium phosphate pH 8.5, protease inhibitor cocktail, 5 mM DTT) until supernatants were clear, and then subjected to a second round of hypotonic lysis and freeze-thaw. After washing until supernatants were clear, membranes were collected by centrifugation (3000×g, 40 min, 4°C), suspended in a minimal volume of wash buffer, and stored at -80°C. Before extraction, membrane isolates were washed with PBS containing protease inhibitors (PBS-PI) and pelleted (3 hours, 120 000×g, 4°C).
Membrane preparations from mammalian tissues
Membranes were isolated as previously described . Briefly, mouse brains or livers were flash frozen after dissection and stored at -80°C until needed. For the isolation of all cellular membranes, we applied a simple physical separation / fractionation protocol. Briefly, frozen tissues were thawed in hypotonic lysis buffer (20 mM HEPES pH 7.4, protease inhibitor cocktail, 10 mM sodium orthovanadate, 4 μM staurosporin, 4 μM cantharidin) and manually homogenized on ice with a polyethylene pestle in a 1.5 mL microcentrifuge tube. The homogenate was subjected to one round of freeze-thaw (-80°C), before being combined with an equal volume of 2×PBS to restore isotonicity. Membranes were collected by ultracentrifugation (3 hours, 120 000×g, 4°C), and were washed twice; pellets were resuspended in PBS-PI for each wash and collected by ultracentrifugation, as described above.
Detergent extraction buffers were prepared for 1D (7 M urea, 2 M thiourea, 9 mM Tris acetate pH 7.0, protease inhibitor cocktail, and detergent as indicated) or 2-DE (IEF buffer 1 containing 8 M urea, 2 M thiourea, protease inhibitor cocktail, and detergent as indicated ). Membrane pellets were resuspended by pipetting and vortexing. Extractions were incubated for 1 hour on ice, with periodic vortexing. Any insoluble material was separated by ultracentrifugation as previously described. Solubilized samples were assayed for total protein content using the EZQ Protein Quantitation Kit (Molecular Probes, Eugene, OR).
Total protein was assayed using either the EZQ Protein Quantitation Kit or the RC DC Protein Assay Kit (BioRad, Hercules, CA). The RC DC assay was carried out according to manufacturers instructions in 96-well plates and absorbance was measured using the Wallac Victor2 Multilabel HTS Counter (PerkinElmer Life Sciences, Boston, MA). EZQ Protein Quantitation was carried out essentially according to manufacturers instructions except fluorescence was recorded by imaging on the Proexpress multiwavelength fluorescent imager (PerkinElmer, Boston MA) and spot fluorescence was quantified using ImageQuant 5.2 software (Molecular Dynamics, Sunnyvale, CA).
1D SDS-PAGE was performed in mini gel format using the BioRad Protean II Electrophoresis system, essentially as described  with minor modifications . Samples were normalized to 2 mg/ml in the appropriate extraction buffer, and then diluted 1:1 (v/v) with 2 × SDS sample buffer . 10 μg total protein was loaded per well on 12.5%T separating gels with 5%T stacking gels, buffered with 375 mM Tris (pH 8.8) as described . Gels were run at 125 V for 10 min to stack proteins, and then the voltage was reduced to 90 V to completion .
Samples for IEF were normalized to 2 mg/ml with the appropriate IEF buffer, then combined 1:1 (v/v) with an ampholyte-containing IEF buffer (8 M urea, 2 M thiourea, 1% pH 3–10 broad range ampholytes, 0.2% each narrow range ampholytes (pH 2.5–4, 3.5–5, 5–7, 7–9, and 8–9.5) and detergent as indicated ), to introduce a working concentration of ampholytes to the sample.
Samples were sequentially reduced and alkylated essentially according to Herbert et al. [33, 34] with some minor modifications. Briefly, the sample was reduced by the addition of TBP and DTT to final concentrations of 2.3 mM and 45 mM DTT, respectively, and incubated for 1 hour at 25°C. The reduced sample was then alkylated with 230 mM acrylamide monomer for 1 hour at 25°C. Immediately following alkylation, the sample was loaded onto IPG strips for passive hydration at 25°C (12 hours). IEF was carried out at 15°C using the BioRad Protean IEF Cell; voltage was ramped linearly to 4000 V (2 hours) and IEF was carried out at 4000 V (constant) for 37500 Vhours. After focusing, IPG strips were equilibrated essentially according to the manufacturer's instructions by sequential immersion in equilibration buffer (6 M urea, 2% SDS (w/v), 20 % glycerol (w/v), and 375 mM Tris pH 8.8) containing 130 mM DTT for 10 minutes, followed by equilibration buffer with 350 mM acrylamide monomer for 10 minutes. Following equilibration, IPG strips were loaded onto 12.5%T separating gels with 5%T stacking gels (buffered as described for 1D) and sealed in place with an agarose overlay (0.5% low melting agarose, 0.1% SDS and 375 mM Tris pH 8.8). SDS-PAGE was otherwise carried out as described for 1D SDS-PAGE.
After electrophoresis, gels were fixed in 10% methanol, 7% acetic acid for 1 hour, washed thoroughly with water and stained with Sypro Ruby overnight. Gels were visualized using the Proexpress multiwavelength fluorescent imager (PerkinElmer, Boston MA). Quantitative image analysis was performed using Progenesis Workstation 2004 (Nonlinear Dynamics, Cambridge, UK). Parallel sets of gels were warped and matched by automated analysis, and volumes were normalized to a single spot consistent in size, shape, density and location across all gels.
- MEGA 8:
- MEGA 9:
- MEGA 10:
- SB 3–10:
(octaethylene glycol monododecyl ether)
(red blood cell)
The authors would like to thank Tiffany Rice, Tammy Wilson, and Dr. V. Wee Yong for kindly providing mouse tissues. We would also like to thank Jeff Lamb, Dr. Sina Ahmadi Pirshahid, Marlies Ernst, and all the members of the Coorssen lab for helpful discussions and advice. J.R.C. acknowledges support from the Canadian Institutes of Health Research, the Canada Foundation for Innovation, the Alberta Heritage Foundation for Medical Research, the Alberta Network for Proteomics Innovation, and the Heart and Stroke Foundation of Canada.
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