Biochemical and proteomic approaches for the study of membrane microdomains

J O U RN A L OF P ROT EO M IC S 7 2 (2 0 0 9) 1 2–2 2

a v a i l a b l e a t w w w. s c i e n c e d i r e c t . c o m

w w w. e l s e v i e r. c o m / l o c a t e / j p r o t

Review

Biochemical and proteomic approaches for the study of membrane microdomains Yu Zi Zheng, Leonard J. Foster⁎ Centre for High-Throughput Biology and Department of Biochemistry and Molecular Biology, 435-2125 East Mall, University of British Columbia, Vancouver, BC, Canada V6T 1Z4

AR TIC LE D ATA

ABSTR ACT

Keywords:

Many cellular signaling and communication events take place at the plasma membrane and thus

Plasma membrane proteomics

the characterization of the plasma membrane proteome has been a hot research area in the

Membrane microdomain

hopes of learning more about these processes. Membrane microdomains are large protein and

Quantitative proteomics

lipid complexes found on the cell surface membrane, able to concentrate or recruit signaling molecules or factors. The first step of any organelle proteomics study is to get a pure and enriched protein sample yet this has always been problematic in membrane proteomics as it is virtually impossible to purify a specific membrane type to homogeneity. In this review, we summarize the biochemical and proteomic approaches that have been used recently in the isolation and identification of several membrane microdomains and non-typical membrane proteins. © 2008 Elsevier B.V. All rights reserved.

Contents 1. 2.

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5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lipid rafts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Detergent-resistant membranes (DRMs) and lipid rafts. . . . . . . . . . . . . . . . . . . 2.2. Detergent-based methods to isolate DRMs/rafts . . . . . . . . . . . . . . . . . . . . . 2.3. A detergent-free method to isolate DRMs/rafts . . . . . . . . . . . . . . . . . . . . . . 2.4. A method for improving the specificity of raft proteomic studies . . . . . . . . . . . . . Caveolae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Caveolae and structural protein—caveolins . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Detergent-based and detergent-free methods to isolate caveolae . . . . . . . . . . . . . 3.3. Silica coating and immunoisolation for the isolation of caveolae . . . . . . . . . . . . 3.4. Comparative proteomics of caveolae-containing and -deficient cells . . . . . . . . . . Tetraspanin-enriched microdomains (TEMs) . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Tetraspanins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. The use of milder detergent combined with co-immunoprecipitation to isolate TEMs. GPI-anchored proteins (GPI-APs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Proteins anchored to the plasma membrane by glycosylphosphatidylinositol (GPI) . . 5.2. The isolation of GPI-APs by GPI-specific phospholipases followed by phase separation. .

⁎ Corresponding author. Tel.: +1 604 822 8311. E-mail address: [email protected] (L.J. Foster). 1874-3919/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.jprot.2008.09.003

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6. Downstream processing of isolated microdomain samples before MS analysis . 7. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1.

Introduction

Biological membranes, in addition to their paramount role in maintaining cell structure, are also involved in many essential cellular processes such as cell signaling and trafficking due to their unique interaction with both the inside and the outside of cells. Whole genome analysis predicts that integral membrane proteins comprise one-third of all the proteins encoded by the human genome [1]. Generally, membrane-associated proteins include not only proteins that are physically embedded in lipid bilayers, such as ion channels, but also proteins anchored to the membrane (e.g., glycosylphosphatidylinositol (GPI)-anchored proteins (GPI-APs) in the outer leaflet of the plasma membrane (PM), acylated proteins in the inner leaflet of the PM), as well as otherwise soluble proteins that interact with the above two classes of proteins. Thus, the number of proteins that should actually be considered as ‘membrane proteins’ is even much larger than the one-third of the proteome that is integral to the membrane. The old fluid mosaic model [2], where proteins and lipids are evenly distributed across a two dimensional surface, has been replaced by the membrane compartmentalization model [3] that states that membranes are compartmentalized, or non-uniform, as a result of an uneven distribution of specific lipids and/or proteins into various microdomains and these microdomains can take up to 10–30% of the total membrane area [4]. These membrane microdomains often act as reaction platforms, harboring enzymes, substrates and adaptor/scaffold proteins [5] at a scale much beyond simple protein complexes, up to microns in diameter [6,7]. The challenge of correctly defining the proteome of a whole membrane or a membrane microdomain has been intensely investigated for many years, in part because it is more tractable than more complex targets such as a whole cell. Of greater significance, however, is the potential to provide a deeper understanding of the biological processes happening on the cell membrane and to identify new targets for drug development as membrane proteins represent more than two-third of the known protein targets for existing and likely future drugs [8]. Several properties of membrane proteins make them difficult to detect in proteomic studies. In liquid chromatography–tandem mass spectrometry (LC–MS/MS) experiments the primary challenges to identifying membrane proteins are: 1) abundance—as a fraction of the volume of a cell membranes occupy a very small portion so membrane proteins have an intrinsically lower potential concentration than soluble proteins, 2) hydrophobicity—there are two aspects to this: a) few tryptic cleavage sites occur within the hydrophobic domains—Arg and Lys residues are very hydrophilic and so occur infrequently in hydrophobic domains, meaning that the favorite protease for proteomic studies often does not generate peptides of an appropriate size, b) hydrophobic peptides don't elute from reversed phase—even when peptides of an appropriate mass are generated from a hydrophobic domain, they can often be so hydrophobic that they are not eluted from

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the standard C18 reversed phase material that most groups use in LC–MS/MS and thus are never detected. Organelle proteomics, within which we consider the study of membrane microdomains, presents an additional challenge in the form of sample purity. If one is interested in studying the proteome of an organelle a typical first step involves biochemical enrichment of that organelle [9]. However, it is essentially impossible to purify an organelle to homogeneity so there can often be some uncertainty surrounding any given protein seen in a proteomics study: is the protein really a resident of the organelle of interest or is it a contaminant? Different methods have been developed to overcome this problem in the case of membrane microdomains: various detergents have been used to solubilize hydrophobic membrane proteins [10–16], membrane surface labeling and affinity purifications have been used to isolate particular protein populations [17–20], and quantitative proteomics approaches have been used to define, without bias, lipid raft components [21]. This review focuses on methods used in different biochemical and/or proteomic approaches for the analysis of membrane microdomains, going beyond the classic microdomains, lipid rafts, to touch on caveolae, tetraspanin-enriched microdomains and proteins anchored in membrane by a specific chemical linker, in particular GPI-anchored proteins.

2.

Lipid rafts

2.1.

Detergent-resistant membranes (DRMs) and lipid rafts

Lipid rafts are viewed as platforms for many cell signaling factors to integrate and to interact. Lipid raft theory [22] proposes that certain proteins preferentially cluster into this unique environment, forming membrane reaction centers essential for many cellular processes, such as cell signaling and trafficking [22–24]. Because of their plasma membrane localization, rafts are also considered to be the entry points of certain pathogens to host cells [25–28]. In fact, lipid rafts are one class of membrane microdomains that were originally defined biochemically as the low-buoyant density DRM fraction of cells but that are now recognized as a subset of DRMs enriched in cholesterol and sphingolipids [29–33]. Cholesterol is thought to intercalate between the rigid hydrophobic tails of sphingolipids and saturated phospholipids, allowing a very tightly packed structure with unique biophysical characteristics compared with surrounding membranes. Lipid–lipid and lipid–protein interactions are the likely biophysical basis behind the formation of rafts as cholesterol inserts into the lipid bilayer, clustering together and then recruiting proteins with a high likelihood to partition into an environment rich in cholesterol and/or sphingolipids. The tight lipid interactions create a lipidordered phase that allows for phase separation by certain nonionic detergents at low temperature, followed by subsequent

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floating by density gradient centrifugation. Protein–protein interactions are also an important aspect of raft biology, as more proteins come to rafts either in the plane of the membrane (i.e., membrane proteins diffusing into rafts) or from the cytosol or lumen of the cell to form, in some cases, very large protein complexes [34,35]. Flotillins, tyrosine kinases and GPI-anchored proteins are among the most well studied protein markers of rafts. The raft field has been dogged by biophysical arguments that rafts are only an artifact of detergent extraction [36–38] but their recent visualization by several methods [39–44] has finally laid such uncertainties to rest.

2.2.

Detergent-based methods to isolate DRMs/rafts

Most lipid raft proteomics studies performed so far have taken advantage of the early observations that rafts or cholesterol and sphingolipids enriched membranes are insoluble in cold, non-ionic detergents (e.g., Triton X-100) and that their lowbuoyant density allows them to float easily by density gradient centrifugation [32]. Typically, DRMs/rafts are separated from total cell lysate or membrane fractions solubilized with 1% Triton X-100 and further purified through a density gradient fractionation procedure comprised of three layers of sucrose at 45% (which contains the DRMs initially), 35% and 5%. After centrifugation at or near 200,000 relative centrifugal force (rcf), a purified DRM fraction can be isolated as a band from the 35 and 5% sucrose interference [23]. Although rafts are only a subset of DRMs [21], the two terms are often confused. Triton X-100 is the most widely used non-ionic detergent for extracting DRMs. Several studies have the extraction properties of other detergents, including NP-40, CHAPS, Tween, Lubrol WX and several of the Brij series [45–48]. All of these can be used to enrich raft marker proteins but the DRM proteome extracted with each are not consistent, with some proteins being extracted quite easily with one detergent and not with another. Other factors that contribute to the proteome variability include the abilities of detergents to promote

domain formation and to break up some protein–protein interactions. A DRM proteomics study by Blonder et al. compared Brij-96 and Triton X-100 and concluded that Triton X-100 extracted more DRM material than Brij-96. Furthermore, Triton X-100 increases the yield of DRM extraction but at the same time more non-DRMs were also observed, such as ribosomal proteins [46].

2.3.

A detergent-free method to isolate DRMs/rafts

Another method for enriching rafts employs extraction of the cells in high salt, for example 500 mM sodium carbonate, and high pH (≥11), followed by ultracentrifugation on a sucrose density gradient [49]. This approach also enriches some raft markers proteins but, surprisingly, fails to enrich GPIanchored proteins, suggesting that this method may preferentially enrich caveolae, another related membrane microdomain that are also a subset of DRMs. To compound this finding, the specificity of the high pH approach seems to be very poor, with as many as 75% of the proteins enriched in this way being non-raft proteins [21]. A related but likely even less stringent method for enriching rafts involves an initial total membrane enrichment step, followed by membrane fragmentation using sonication and then floatation of raft/DRM membranes on sucrose or Optiprep density gradients [50,51]. This method seems to preserve more proteins, although a large fraction of those proteins have been otherwise demonstrated to be non-raft localized.

2.4. A method for improving the specificity of raft proteomic studies Biochemical purification of an organelle or subcellular domain to homogeneity is essentially impossible [9]. Thus, since both detergent-based and detergent-free methods fail to yield biochemically pure rafts, other properties of rafts have to be considered in order to correctly define their proteome.

Table 1 – Different biochemical and proteomic methods used for isolating or enriching membrane microdomains and comparison of their relative purity Membrane microdomains Lipid rafts

Caveolae

Tetraspaninenriched microdomains GPI-anchored proteins

a

Not applied.

Methods Detergent Triton X-100, NP-40, CHAPS, Tween, Lubrol WX and Brij-96 [21,32,45–48] Triton X-100 [73,87,92]

Brij-99, CHAPS [104] Brij-58, NP-40, CHAPS, Triton X-114 [47,125,127–131]

Detergent free

Immunoisolation

Purity Phase transfer

High salt and alkaline pH, membrane fragmentation using sonication [49–51]

a

NA

Cholesterol disruption drug [21]N detergentN detergent free

High salt and alkaline pH, membrane fragmentation using sonication or Dounce [50,56,89] NA

Caveolin antibodies [93,94]

NA

2D gel comparison [95] N immunoisolation N detergentN detergent free

Tetraspanin antibodies [108–110]

NA

Immunoisolation N detergent

NA

Enzymatic cleavage by GPI-specific phospholipases [127–131]

Phase transfer N detergent N detergent free

High salt and alkaline pH [47,130,131]

NA

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We have previously reported the use of a quantitative proteomics approach employing Stable Isotope Labeling by Amino acids in Cell culture (SILAC) [52] as an unbiased method for distinguishing the true components of rafts out of a messy DRM proteome [21]. This was done in the epithelium-derived HeLa cell line through the use of SILAC in combination with cholesterol disrupting drugs; one half of the cells was labeled with heavy stable isotopes and left untreated while the other, unlabelled half was treated with the cholesterol disrupting drug methyl-β-cyclodextrin (MβCD) [53,54]. The two cell populations were then being combined together and Triton X-100 was used to enrich rafts. Proteins displaying sensitivity to MβCD (the hallmark of a raft protein) would present a large heavy:light SILAC ratio while proteins insensitive to MβCD, and thus likely contaminants or co-purifying proteins, would present a heavy:light ratio near 1. Previously known or characterized raft marker proteins all fell into the former, high ratio group, whereas mitochondrial and ER proteins that had been considered as components of rafts [47,55,56] could be assigned as contaminants. Besides MβCD, several other cholesterol disrupting agents have been reported but they were not as efficient as MβCD for these purposes [21]. This approach allowed us to overcome the inability to biochemically purify (as opposed to enrich) rafts (Table 1).

suppression of enzymatic activities [75,76]. However, for cases like insulin receptor signaling, binding to caveolae has an activating function instead [77–80]. Besides signal transduction, caveolae are also involved in vesicle transport through caveolae-mediated endocytosis; several pathogens, such as simian virus 40 (SV40) and human immunodeficiency virus (HIV) interact with caveolae [81,82]. Distinct from clathrin-mediated endocytosis, the use of caveolae for cellular intake allows the pathogens to bypass the classical endosome–lysosome trafficking pathway and, as a consequence, avoid lysosomal degradation. Caveolae are also associated with non-infectious diseases; certain mutations in caveolin proteins mislocalize them, often resulting in cancer or muscle degeneration diseases [61,83]. As one example, genetic analysis of human breast cancer samples revealed that up to 16% of these patients have a single Cav-1 gene point mutation (P132L) [84,85]. Caveolae and lipid rafts have been shown to co-exist in cells. One school of thought proposes that caveolae are specialized lipid rafts [86]; however, the association of caveolin with cholesterol has been questioned [21]. Proteomic studies have reported the overlap of these two compartments at the proteome level but whether the two entities are completely separate microdomains or connected in some way still requires further study.

3.

Caveolae

3.2. Detergent-based and detergent-free methods to isolate caveolae

3.1.

Caveolae and structural protein—caveolins

Caveolae are membrane microdomains found in many cells and they are classically pictured as flask-shaped, stable invaginations of the plasma membrane. Because of their unique morphology, caveolae can be easily distinguished by electron microscopy, appearing when looking onto a plasma membrane from outside the cell as small caves with diameters in the range of 50 to 100 nm [57,58]. When numerous, caveolae can have a substantial impact on the surface area of cells; ultrastructural analysis has shown that as much as 20% of adipocyte plasma membrane can be caveolae [59]. The genesis of caveolae depends on the presence of caveolins, small, structural proteins that have also been observed at low levels in DRMs [60]. Caveolae formation is inhibited when caveolin expression is knocked out [61,62] and the human genome contains three caveolin proteins, Cav-1, -2 and -3 [63–65]. All three are integral but not trans-membrane proteins targeted to the inner leaflet of the plasma membrane (PM); Cav-1 and -2 are expressed in most cells whereas Cav-3 is muscle specific [49,66–68]. Caveolins themselves do not have enzymatic activities, rather they homo- or hetero-oligomerize with other caveolins to form a coat that stabilizes caveolae and forces its concave structure; at the same time, they interact with other membrane proteins and recruit other cytosolic proteins to caveolae through their scaffolding domains (CSD) [69–72]. Therefore, unlike lipid rafts, caveolae are formed by protein–protein interactions. Caveolae have been suggested to function in signaling events through the compartmentalization of signaling molecules interacting with caveolin proteins [73,74]: the interactions are normally thought to have negative effects through the

The first caveolae isolation experiment was based on the detergent-insolubility and the low-buoyant density of these membrane compartments [60]. The non-ionic detergent Triton X-100 was used to extract crude membranes from murine lung tissue, following which sucrose density ultracentrifugation was applied to enrich caveolae-rich membrane domains [73]. The purified membrane fractions, or DRMs, retain approximately 85% of caveolin and approximately 55% of a GPI-linked marker protein, while they exclude ≥98% of integral plasma membrane protein markers and ≥99.6% of other organelle-specific membrane markers tested. However, the assumption here was that the purified DRMs were equivalent to caveolae, which seems highly unlikely given more recent results [21]. This technique has also been used to isolate caveolae from chicken smooth muscle cells where plasma membranes were subjected to several purification steps before detergent treatment [87]. To avoid artifacts with using detergents to extract membrane domains, detergent-free methods were developed to extract caveolae [51]. The initial step in all these processes is always mechanical disruption (e.g., a Dounce homogenizer or sonication) of cells, sometimes in high salt or alkaline pH conditions. Then the low-buoyant density of caveolae is enriched by floatation on a density gradient, similar to the detergent extraction method. The hope is that weakly-associating proteins are more readily retained by avoiding detergents and these methods have been used to study caveolae in human heart tissue [88], rat ventricular myocytes [89] and several cell lines [56,90]. However, the downside of detergent-free methods is that the resulting low-density membrane fraction is probably even less enriched in caveolae, making their use as a caveolae model questionable.

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3.3. Silica coating and immunoisolation for the isolation of caveolae In order to improve the purity of isolated caveolae, silica coating and immunoisolation were introduced more recently. Silica coating of plasma membranes was first applied by Caney and Jacobson in the early 80s to enhance the yield and purity of isolated PM of cells in culture [91]. The procedure consists of coating intact cells with dense, positively charged silica particles, which functions to increase the density of the PM for easier isolation and also to prevent vesiculation or lateral reorientation. More recently, Schnitzer et al. applied this silica-coating approach to purified caveolae from luminal endothelial plasma membranes of rat lung. Cell surface membranes were isolated by differential centrifugation after coated with silica; then the PM pellet was homogenized in 1% Triton X-100 followed by sucrose density gradient to yield a DRM preparation containing caveolae [92]. A modified version of this procedure was also published later where caveolae were immunoisolated with the aid of a monoclonal anti-Cav-1 antibody [93]. DRMs were isolated from sonicated plasma membranes with or without silica coating and then subjected to immunoprecipitation by anti-Cav-1-coated magnetic beads. Similar immunoisolation procedures were also employed by another group to enrich caveolae from rat lung vasculature where the membrane fraction was ultracentrifuged in the presence of high salt prior to immunoenrichment [94].

3.4. Comparative proteomics of caveolae-containing and -deficient cells Without question the use of a DRM extraction enriches caveolae versus a whole cell lysate but even an immunoisolation step does not produce pure caveolae. Therefore, to get more to the question of specificity of caveolar components, Hill et al. used 2D gels to distinguish differences in the DRM proteome between mouse embryonic fibroblasts from both wild-type and Cav-1-deficient mice [95]. Since Cav1−/− cells are devoid of identifiable caveolae by microscopy, proteins that are absent in Cav-1−/− DRMs compared to WT DRMs are expected to be components of caveolae. The 2D gel approach used allowed the unbiased assignment of seven proteins unique to caveolae; the use of such tools in the future is expected to yield even more caveolae-specific components through the use of other technologies.

4.

Tetraspanin-enriched microdomains (TEMs)

4.1.

Tetraspanins

Tetraspanins are a class of integral membrane proteins containing, as their name suggests, four trans-membrane domains and two extracellular loops [96]. They are expressed in all cells and tissues in almost all animal species [97]. In humans, the tetraspanin family comprises 33 proteins that are defined by their unique structural features. Tetraspanins can interact with each other and also with other proteins or factors on cell membranes, particularly in the plasma membrane, to form so called “tetraspanin web” or tetraspanin-enriched microdomains (TEMs) [98].

Some tetraspanins are well characterized, such as CD9, CD81, CD82 and CD151; they do not have any known enzymatic activities in cells but rather they act as “organizers” to recruit other factors to TEMs. Tetraspanins have been reported to interact with integrins, growth factor receptors and many intracellular signaling molecules [99,100]. Therefore, tetraspanin-associated microdomains are involved in a variety of physiological processes, such as immune cell activation, and cell migration, as well as cellular differentiation [98,101]. Tetraspanin proteins are usually underrepresented in biochemical assays because they are small, hydrophobic and low abundance membrane proteins; however, due to their functional significance (i.e., mutations in some tetraspanins are associated with human diseases like mental retardation [102]), TEMs are gaining more attention. There is one report that tetraspanins are in lipid rafts because they can be associated with cholesterol through palmitoylated resides [103], but others believe that TEMs and rafts are two distinct membrane microdomains because tetraspanins appear to be resistant to cholesterol depletion [104]. Another possibility is that TEMs might reside next to lipid rafts, which would also explain why the two microdomains sometimes appear to share similar cell functions and facilitating signal transduction [105].

4.2. The use of milder detergent combined with co-immunoprecipitation to isolate TEMs Like lipid rafts, TEMs or tetraspanin complexes can be separated from other membranes based on their biophysical properties. These complexes are insoluble in milder detergents (less hydrophobic) such as in 1% Brij-99 or CHAPS [104]: Firstly, cells are lysed in these detergents and then tetraspanins and their associated proteins are immuno-enriched using antibodies against tetraspanins [106,107]. Stronger non-ionic detergents like Triton X-100 are not used to study TEMs because tetraspanins were not observed in DRMs after Triton X-100 extraction. Furthermore, studies have reported that Triton X-100 can affect homodimeric tetraspanin interactions, effectively solubilizing whole tetraspanin complexes. Studies in B-cells using Brij-98 to enrich DRMs prior to coimmunoprecipitating CD9, a tetraspanin protein, identified several co-purifying proteins that are likely components of TEMs [108], as did similar studies in other cell types such as T cell and epithelial cells [108–110]. One elegant approach used in such studies to elute proteins from the immune complex is to use a stronger detergent such as Triton X-100 to disrupt the tetraspanin–tetraspanin interactions, leading to the release of the complexes.

5.

GPI-anchored proteins (GPI-APs)

5.1. Proteins anchored to the plasma membrane by glycosylphosphatidylinositol (GPI) GPI-anchored proteins (GPI-APs) are otherwise soluble proteins localized on the outer leaflet of plasma membranes via the attachment of a glycosylphosphatidylinositol (GPI)

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Fig. 1 – Cartoon representation of three membrane microdomains: lipid rafts, caveolae and tetraspanin-enriched microdomains, as well as GPI-anchored proteins. Lipid rafts are depicted as cholesterol and sphingolipid-dependent complexes in the plane of the membrane. Caveolae are formed by protein–protein interactions and are invaginations, coming out of the plane of the membrane. Tetraspanin-enriched microdomains have a less well-envisioned structure, with the requirement for tetraspanin the only commonality. Finally, GPI-anchored proteins are soluble proteins anchored to the plasma membrane by a glycosylphosphatidylinositol (GPI) moiety.

moiety [111]. This is a classic example of a post-translational modification determining the targeting, localization and function of proteins within cells. GPI-APs share some common sequence features as well, such as two hydrophobic regions on the N and C-terminus of the sequence, a hydrophilic spacer region and a ω-site located in the middle of the sequence, corresponding to the cleavage site where the phosphoglycan anchors are attached to the carboxy-termini. Moreover, the attachment takes place in the ER where the nascent protein is transferred to the pre-synthesized GPI anchor by a transamidase [112]. One interesting biophysical property of GPI-APs, conferred upon them by the GPI anchor itself, is their insolubility in non-ionic detergents such as Triton X-100 [43]. GPI-APs' membrane localization places them exactly in the interface where cells communicate with their extracellular surfaces and thus GPI-APs mediate many cellular processes including cell–cell interactions, receptor recognition and cell signaling [113–115]. It is now quite clear that GPI-APs play important roles in human diseases and disorders; for example, one causative agent of neural degeneration, misfolded prion protein (PrP), is a GPI-AP [116]. On the other hand, many of the surface proteins of the malaria parasite Plasmodium falciparium are GPI-APs [117] and they are associated with hostpathogen recognition. GPI-APs have also been reported as candidate biomarkers of human hepatocellular carcinoma, as well as pancreatic and biliary carcinomas [118,119]. Another potential biomarker, folate receptor, is not only a marker for myeloid leukemia and chronic inflammatory diseases but also shows potential use in targeted drug delivery [120]. Transamidase recognizes certain constraints of the ω-site of native GPI-APs and thus bioinformatics approaches have been developed to exploit this, including the Big-Pi, DGPI and GPI-SOM predictors [121–123]. Another more sensitive, and certainly more specific, approach has integrated computational and experimental components [124]. However, while the available bioinformatics tools are very useful for predicting GPI-APs, which typically comprise 1–2% of eukaryotic genomes, very few have been confirmed biochemically since their initial description in the 1980s; the main difficulties is that GPI-APs are insoluble and typically present at very low levels in cells (Fig. 1).

5.2. The isolation of GPI-APs by GPI-specific phospholipases followed by phase separation Since GPI-APs are insoluble membrane-anchored proteins, detergents were normally used to isolate or separate them from rest of the soluble proteins in cells; however, detergent free methods for studying GPI-APs have also emerged. In a study by Man et al., both detergent and detergent-free methods were used to extract crude membrane from rat natural killer cells followed by purification of the extracted membrane fraction on a sucrose gradient to obtain the low-buoyant density membrane fraction containing GPI-APs [47]. The two detergents used were Brij-58, a milder detergent that could preserve weakly associated proteins, and NP-40, a much more stringent detergent. The detergent-free method involves using hypertonic sodium carbonate at pH 11. Each of the methods has its advantages and disadvantages, mild conditions could result in more proteins but at the same time introduces more non-specific proteins. Another two dimensional (2D) gel study compared the ability of several detergents, including DHPC/diC7PC (1,2-Diheptanoyl-snglycero-3-phosphocholine), ASB-14 (Amidosulfobetain-14) and CHAPS, to resolve GPI-APs and other trans-membrane proteins of myelin in 2D gels; DHPC/diC7PC and ASB-14 were most effective [125]. Most recently, GPI-AP proteomics studies have used nonionic detergents like the Triton series, since the GPI moiety targets GPI-APs to the cholesterol-rich, detergent-insoluble membrane domains or lipid rafts. GPI-APs are only a subset of the DRM proteome. The GPI moiety itself is generally quite stable in the context of the cell but it can be released by enzymatic treatment of intact cells, membrane fractions like DRMs or a single protein. The enzymes used are bacterial GPIspecific phospholipases [126]. After the enzymatic cleavage, GPI-APs are released from membrane, making them soluble and thus they partition into the aqueous phase [127], allowing subsequent separation from rest of the membrane proteins. This strategy has been termed “modification specific proteomics” and has been used in several studies involving diverse higher eukaryotes, from plants to animals. In an early study by Parkin et al., a two-phase separation with Triton X-114 combined with phosphatidyl inositol-specific phospholipase C (PI-PLC) was used to determine the GPI anchorage of alkaline

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phosphatase, 5′-nucleotidase and aminopeptidase P [128]. Essentially, DRMs were extracted in Triton X-114; by addition of PI-PLC, GPI-APs were released from their membranes and thereby moved from the detergent phase to the aqueous phase. In related studies, McNall et al. used enzymatic cleavage and phase separation of brush border membrane vesicles (BBMV) to isolate GPI-APs [129] while Elortza et al. used the same method, with either PI-PLC or phospholipase D, to characterize the GPI-AP proteome of human and Arabidopsis thaliana plasma membranes [130,131].

6. Downstream processing of isolated microdomain samples before MS analysis Biochemically enriched membrane microdomains necessarily go through several downstream sample preparation steps prior to MS analysis including: solubilizing membrane proteins in detergents, organic solvents and/or chaotropic reagents; separation of proteins by gel electrophoresis and ion exchange chromatography; reduction, alkylation and digestion of proteins into peptides using either in-solution or in-gel proteolytic digestion. Dozens of detergents, organic solvents and chaotropes have been used to solubilize hydrophobic proteins, although the solubilizing ability of organic solvents like ethanol is not as high as detergents and chaotropes. These reagents are imperfect as they can often interfere with the enzymatic activities of the proteases (proteases are also proteins) or they can precipitate in or otherwise gum up the analytical column used in LC–MS/MS. Some of our favorite reagents for solubilizing membrane microdomain samples include: a buffered solution of 6 M urea and 2 M thiourea—easily removed by a solid phase clean-up step—or deoxycholate (quantitatively precipitated by low pH prior to peptide chromatography [16]). One or two dimensional gel electrophoresis is probably the most used protein separation method for fractionating complex protein samples or to specifically separate one or several proteins from a mixture. However, neither method is particularly suited to membrane proteins; hydrophobic membrane proteins tend to aggregate and precipitate so that they are often underrepresented in gels. To compensate for this a “tube-gel” protocol was developed for application in a lipid raft study [132]. DRMs from neonatal mouse brain were directly incorporated into a polyacrylamide tube-gel matrix without prior protein separation. The gel was then cut into slices and subjected to in-gel digestion. Overall, around 60% of all proteins identified were predicted to be membrane or membrane-associated proteins, thus the solubility of membrane proteins seems improved by this protocol.

7.

Conclusions

Any biochemical study, proteomic or otherwise, of a membrane microdomain depends a great deal on the approach used to enrich the domain away from the remaining material of the cell. As an example we have reviewed here, the use of different detergents can have an enormous impact on the

proteins recovered in the DRM preparation, to the extent that the conclusions reached with different detergents could be polar opposites. Since it seems unlikely that the raft field will agree on one detergent to use for such preparations it is important that results obtained with one detergent be confirmed with others or at least not over-interpreted. Likewise, the unbiased nature of proteomic experiments requires that much greater attention or care be given to determining the specificity of proteins to the compartment being studied. For example, assuming that a caveolae preparation obtained by density gradient separation of sonicated membrane fractions does not contain non-caveolar membranes is naïve, just as is the assumption that all proteins in DRMs are lipid raft proteins. With the increasing availability and decreasing cost of quantitative proteomic approaches, the challenge of obtaining homogenous biochemical preparations can be neatly sidestepped, as described in several examples presented here. In addition, quantitative proteomic technologies are [45,133–137] and will allow more biologically-oriented questions to be asked, such as how the composition of microdomains change upon agonist stimulation. In the next five years we expect that several more elegant approaches coupling quantitative proteomics and membrane microdomains will be reported, driving the biological knowledge of these intriguing entities.

Acknowledgements The authors thank Ivan Robert Nabi and Masayuki Numata for helpful discussions about membrane microdomains. LJF is the Canada Research Chair in Organelle Proteomics and a Michael Smith Foundation Scholar. Microdomain research in the Cell Biology Proteomics laboratory is supported by a Canadian Institutes of Health Research grant to LJF.

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