The physiological environment which hosts the conformational conversion of the cellular prion protein (PrPC) to disease-associated isoforms has remained enigmatic. A quantitative investigation of the PrPC interactome was conducted in a cell culture model permissive to prion replication. To facilitate recognition of relevant interactors, the study was extended to Doppel (Prnd) and Shadoo (Sprn), two mammalian PrPC paralogs. Interestingly, this work not only established a similar physiological environment for the three prion protein family members in neuroblastoma cells, but also suggested direct interactions amongst them. Furthermore, multiple interactions between PrPC and the neural cell adhesion molecule, the laminin receptor precursor, Na/K ATPases and protein disulfide isomerases (PDI) were confirmed, thereby reconciling previously separate findings. Subsequent validation experiments established that interactions of PrPC with PDIs may extend beyond the endoplasmic reticulum and may play a hitherto unrecognized role in the accumulation of PrPSc. A simple hypothesis is presented which accounts for the majority of interactions observed in uninfected cells and suggests that PrPC organizes its molecular environment on account of its ability to bind to adhesion molecules harboring immunoglobulin-like domains, which in turn recognize oligomannose-bearing membrane proteins.
Prions are the causative agents underlying a range of rare but invariably fatal neurodegenerative diseases in humans and other mammals. In disease, the cellular prion protein (Prnp; herein referred to as PrPC) converts to a disease-associated conformer (PrPSc) with different physicochemical properties [1]. PrPC is a relatively small protein which assembles into an unstructured N-terminal domain and a globular C-terminal half characterized by the presence of an internal disulfide bridge, up to two N-linked glycans and a glycosylphosphatidylinositol (GPI) anchor for insertion into the cellular plasma membrane. Multiple lines of investigation have led to the conclusion that mature PrPC is embedded in specialized membrane domains, so-called raft-like domains, rich in cholesterol and sphingolipids [2]. It has been suggested that these raft-like domains host the self-perpetuated accumulation of PrPSc which subsequently triggers a poorly-understood cascade of events that ultimately leads to cell death. Whereas significant progress has been made in the past few years in defining the minimal requirements for PrPC conversion in vitro [3], the molecular environment which hosts the earliest steps in prion disease manifestation in neuronal cells remains enigmatic. This shortcoming is not due to a lack of proteins proposed to interact directly with PrPC. In fact, more than three dozen proteins have been suggested to reside in spatial proximity to PrPC using multiple experimental paradigms [4]. Additional candidate interactors have been proposed to bind preferentially to PrPSc [5], [6], [7]. In surveying this body of literature, however, it is apparent that very few of the candidate interactors have been independently verified by multiple investigators and, overall, little agreement exists as to their relative importance for prion protein biology. Notable exceptions may represent the 37-kDa/67-kDa laminin receptor precursor (herein referred to as LRP; also known as ribosomal protein SA (Rpsa), and not to be confused with Lrp1, the low density lipoprotein receptor-related protein 1) [8], one of the first proteins identified to bind to the prion protein in a yeast two-hybrid (Y2H) screen [9]; the neural cell adhesion molecule (Ncam1; herein referred to as NCAM), which was initially identified in a cellular crosslinking study based on a co-immunoprecipitation methodology [10]; and heparin sulfate proteoglycans (HSPGs) [11], [12]. Since its original discovery, LRP has been proposed to act as a cell surface receptor for PrPC that may play a role in prion propagation [13]. In the case of NCAM, binding to PrP has been shown to play a role in neuritic outgrowth possibly mediated through an interaction with Fyn tyrosine kinase [14], [15]. In studying the molecular environment of membrane proteins, a limitation exists in that the proteins must be solubilized by the addition of detergents in a manner that does not disrupt the protein-protein interactions under investigation. A solution to this obstacle constitutes the covalent stabilization of interactions by chemical crosslinking prior to the disruption of cellular integrity. In this regard, we previously reported on a large-scale investigation of the molecular PrPC neighborhood in mice following limited in vivo crosslinking by time-controlled transcardiac perfusion (tcTPC) -based delivery of formaldehyde (FA) to the brain [16]. A conspicuous feature of this PrPC interactome dataset was the relative abundance of membrane proteins which harbor immunoglobulin (Ig) -like folding motifs in their extracellular domains confirming previous data describing a binding domain of PrPC within these folds [10]. Given the large diversity of cell types present in the brain, we questioned whether an equivalent concentration of Ig-like domain-harboring proteins would have been found if the investigation of the PrPC interactome had been restricted to a single cell type. A particular concern was that the relative abundance of Ig-like domain-harboring proteins in the previous dataset may have masked the ability to identify biologically important interactions of PrPC that are more transient in nature or involve less abundant proteins. We therefore chose to reinvestigate the PrPC interactome in mouse neuroblastoma cells (N2a), which are by far the best-characterized cell model for the study of the biology and conversion of PrPC to PrPSc [17], [18]. To facilitate the discrimination of unspecific binders from specific interactors in this study, we incorporated quantitative mass spectrometry based on isotopic labeling and extended our investigation to the mammalian PrPC paralogs Doppel (encoded by the Prnd gene; protein product herein referred to as Dpl) and Shadoo (encoded by the Sprn gene; protein product herein referred to as Sho). To avoid the use of non-identical affinity chromatography steps, we equipped the three bait proteins with the same N-terminal FLAG epitope [19], with the awareness that a similar PrPC expression construct had in prior investigations neither interfered with the posttranslational processing nor the conversion of PrPC in cell or animal models [18]. We now present data which suggest that members of the mammalian prion protein family may populate highly similar molecular environments when expressed in the neuroblastoma cell model system. We further document that a subset of endoplasmic reticulum (ER) chaperones which interact strongly with PrPC escape from the ER to reside in spatial proximity to PrPC at the plasma membrane. Pharmacological inhibition of these chaperones could increase PrPSc levels in prion-infected N2a sublines, suggesting a protective role for the chaperones in this paradigm. Finally, our data consolidate multiple previously controversial interactions and suggest a scenario whereby PrPC may organize its molecular environment by its ability to recognize a specialized subset of cell adhesion molecules which recruit membrane proteins carrying high-mannose glycans into spatial proximity with PrPC.
In preparation for this study, mouse neuroblastoma cells (N2a) stably expressing N-terminally FLAG-tagged full-length versions of Dpl, PrP and Sho were generated (Fig. 1). Small-scale expression tests followed by diagnostic N-glycosidase F or phosphatidylinositol-specific phospholipase C digestions confirmed that all three bait proteins mimicked their untagged parent molecules in (i) the presence of N-glycans, and (ii) membrane attachment by means of a GPI anchor [20] (and data not shown). To generate the biological source material for a large-scale comparative interactome investigation, the three cell lines and an “empty-vector” stably-transfected N2a cell line serving as a negative control were expanded to 109 cells each using cell culture conditions which promote adherent growth. To covalently stabilize protein-protein interactions prior to the disruption of cellular integrity, cells were subjected to a 15-min treatment with FA [10]. Subsequently, cells were lysed by the addition of detergents and the four extracts (each containing approximately 500 mg of cellular protein) purified side-by-side on anti-FLAG affinity agarose matrices. The presence of covalent linkages between proteins permitted the use of highly stringent washing conditions to minimize the presence of unspecific binders. Following elution from the affinity matrix (with a yield of approximately 100 µg of protein material per sample), protein complexes were denatured and trypsinized. Finally, peptide mixtures were tagged with isobaric tags for relative and absolute quantitation (iTRAQ) [21], and the samples were combined and subjected to a comprehensive analysis by tandem mass spectrometry (MS/MS) (Fig. 2). A query of mouse protein databases led to the identification of more than 100 proteins. All identifications can be considered confident by multiple measures: (i) identical identifications were made by two matching algorithms; (ii) scores assigned by the algorithms exceeded significance thresholds (P<0. 05) for all identifications and were based on a minimum of two peptides; and (iii) a search of a decoy database generated by the inversion of sequences for all mouse protein entries resulted in no identifications which shared any of the above features [22]. A reduction of the total list of identified proteins to the subset of proteins whose identification correlated with the presence of at least one of the three bait proteins was based on peak intensities of iTRAQ reporter ions found in the low mass range of individual collision induced dissociation (CID) spectra. The relative intensity of these ions is indicative of the relative contribution of each of the iTRAQ-labeled samples to the generation of a given CID spectrum. Thus, by calculating the ratio of reporter ion intensities for each of the three bait-specific reporter ions (iTRAQ 115: Dpl; iTRAQ 116: PrP; iTRAQ 117: Sho) and the negative control ion (iTRAQ 114 reporter), peptides purifying with at least one of the three baits were recognized by iTRAQ reporter ion ratios greater than 1. The above analysis revealed that more than 50 proteins co-purified with at least one of the three bait proteins (Table 1). Interestingly, the majority of these proteins co-purified with all three bait proteins, suggesting that the molecular environment of the three members of the mammalian prion protein family assessed in the context of mouse neuroblastoma cells is highly similar. We next explored the known cellular localization of candidate interactors based on bioinformatic methods and literature mining. This investigation revealed that multiple proteins in the dataset were likely to encounter the three bait proteins during their early passage through the secretory pathway, since they constitute (i) classical ER chaperones (heat shock protein 5, Hspa5; calnexin, Canx; calreticulin, Calr; endoplasmin, Hsp90b1), (ii) isomerases which facilitate disulfide or proline cis-trans rearrangements (protein disulfide isomerase associated 3, Pdia3; protein disulfide isomerase associated 4, Pdia4; prolyl 4-hydroxylase beta polypeptide, P4hb; peptidylprolyl isomerase B, Ppib), or (iii) proteins involved in the trafficking between ER and Golgi compartments (transmembrane emp24 transport domain containing 9, Tmed9; and transmembrane emp24-like trafficking protein 10, Tmed10). Other proteins in the dataset were likely to reside in spatial proximity to the mature bait proteins at the plasma membrane, because they are themselves known to be either (iv) embedded in the plasma membrane through transmembrane (TM) domains (NCAM; transferrin receptor, Tfrc; integrins; neuropilin 1, Nrp1; L1 cell adhesion molecule, L1cam; basigin, Bsg), or constitute (v) secreted proteins (galectin-1, Lgals1; family with sequence similarity 3, member C, Fam3c; ectonucleotide pyrophosphatase/phosphodiesterase 1 and 3, Enpp1/3) or (vi) cytosolic proteins (14-3-3 proteins zeta/beta/gamma, Ywhaz/Ywhab/Ywhag; guanine nucleotide binding proteins, Gnao1/Gnai2/Gna13; and Fyn proto-oncogene) that attach to the plasma membrane from outside or within the cell, respectively (Fig. 3). In contrast to the above proteins that were designated as specific interactors by their iTRAQ reporter ion ratios, proteins which were equally represented in control and specific interactome cells and therefore constituted unspecific binders could be mapped to a wide range of cellular compartments including the nucleus and mitochondria. This result was expected based on the known cellular biology of PrPC and therefore served as strong biological validation of the iTRAQ-based discrimination of specific interactors from unspecific contaminants. It was anticipated that iTRAQ reporter ion signals in CID spectra derived from tryptic peptides of the three bait proteins would reflect the logic of the iTRAQ conjugation step. For example, CID spectra which supported the identification of Dpl had to be mainly contributed by the sample which was labeled with the iTRAQ 115 reagent. A close inspection of the low mass range of CID spectra confirmed that this expectation was met for all three bait proteins and thereby served as an internal control for the reliability with which relative iTRAQ reporter ion intensities could be employed to trace the contribution of samples to the identification of proteins (Table 1). Interestingly, however, a closer inspection of the above spectra also revealed that approximately one-third of the peptides which supported the identification of PrP were contributed by the FLAG-Dpl and FLAG-Sho affinity purified samples. This observation was supported by five CID spectra consistent with the interpretation that the purification of FLAG-Dpl and FLAG-Sho caused a significant co-purification of endogenous wild-type (wt) PrPC (P<0. 05). A similar co-purification of endogenous wt Sho in N2a cells was observed with FLAG-Dpl and FLAG-PrP. A co-purification of wt Dpl with FLAG-PrP and FLAG-Sho baits was neither observed nor expected because no endogenous expression of Dpl can be reliably detected in N2a cells (or indeed in the adult mouse central nervous system) [23]. Taken together, these data suggest that PrP, Dpl and Sho are capable of residing in close spatial proximity at the cell membrane and may engage in reciprocal contacts with each other. It is unlikely that members of the mammalian prion protein family engage in direct interactions with all candidate interactors we identified. For example, some retrieved proteins may represent complexes of the type PrPC- crosslink - interactor 1 - crosslink - interactor 2. Such indirect interactors are often less strongly represented in such datasets, due to both the non-stoichiometric crosslinking that can be achieved and the stringent washing conditions applied. A good correlate for the relative abundance of a protein in proteomics investigations of this nature is the sequence coverage with which it was identified. Based on this rationale, the three bait proteins may bind directly to lactate dehydrogenase (Ldha), protein disulfide isomerases Pdia3 and P4hb, galectin-1, NCAM, Gm9234 (also known as EG668548), neuromodulin (Gap43), LRP and its ribosomal protein partner Rps21, heat shock proteins Hspa5 and Hsp90ab1, histocompatibility antigens H2-K1 and H2-D1, Fam3c, Ppib, calnexin, calreticulin, 14-3-3 zeta/delta and basigin, as these proteins were identified with the highest sequence coverage (25–75%). Due to their relatively small size, members of the mammalian prion protein family would be expected to contain a small number of distinct surface domains that can engage in direct protein-protein interactions. Consequently, at least a subset of binders may bind to the same face within the bait proteins and may even resemble each other in their primary structure or structural folds. When candidate interactors were subjected to scrutiny from this angle, similarities between NCAM and basigin, two of the proteins represented with the highest sequence coverage in the dataset, became apparent. Not only do both proteins harbor Ig-like domains, but a subset of these domains bear a striking resemblance in their sequence, an observation made previously [24], [25]. Interestingly, these very same Ig-like domains are capable of binding to membrane proteins carrying oligomannosidic N-glycans [26], [27], [28], [29]. The latter modification is only found on a relatively small number of mature N-glycosylated proteins expressed mainly in the brain, because oligomannose structures typically represent transient intermediates subject to additional posttranslational modification during their passage through the secretory pathway [26], [29], [30]. Amongst the few known proteins carrying oligomannosidic N-glycan structures are the cell adhesion molecule L1, the β-subunit of the Na/K ATPase, the transferrin receptor, integrins [31] and members of the ectonucleotide pyrophosphatase/phosphodiesterase protein family [30]. Remarkably, not only were all the above proteins present in our dataset, but their iTRAQ ratios also documented their specific co-purification with members of the mammalian prion protein family. In light of the relatively low sequence coverage observed for proteins carrying oligomannosidic N-glycans, their presence in the dataset likely reflects indirect binding to the bait proteins mediated by the mentioned molecules that harbor Ig-like domains with an affinity for oligomannosidic N-glycans. To investigate whether PrPC also resides in spatial proximity to membrane proteins equipped with oligomannosidic N-glycans in the mouse brain, we subjected anaesthetized wild-type mice to brief crosslinking with formaldehyde to stabilize protein-protein interactions in vivo prior to the disruption of cellular integrity [16]. Perfused mouse brains were then rapidly dissected and homogenized in the presence of a quenching reagent. To verify the success of the crosslinking reaction and characterize the size distribution of crosslinked protein complexes containing PrPC an aliquot of extracts was passed through a pre-calibrated size-exclusion chromatography column. This pilot analysis documented that PrPC was found in protein complexes co-eluting with calibration markers which span a range of 160 kDa to 669 kDa. Uncrosslinked PrPC could be recovered from the crosslinked complexes by a crosslink reversal procedure based on the administration of heat and reducing agents (Fig. S1). Subsequently, the crosslinked extracts were passed over snowdrop lectin (Galanthus nivalis agglutinin, GNA) agarose, a lectin which specifically recognizes oligomannosidic N-glycans [30]. Eluate fractions, when subjected to Western blotting analysis, documented the presence of a subset of PrPC in high-molecular weight protein complexes that were specifically enriched by snowdrop lectin agarose (Fig. 4). The binding to the affinity matrix was determined to be specific by (i) the PrPC-specific band pattern in the eluate fraction demonstrating the selective capture of PrPC in high-molecular weight crosslinked complexes, rather than in its uncrosslinked form; and (ii) the observation that pre-incubation of the extract with soluble snowdrop lectin prevented affinity capture. A crosslink reversal time series revealed that the high-molecular weight complexes preferentially contain fully-glycosylated PrPC consistent with the notion that this interaction involves mature PrPC present at the cell surface. A conspicuous feature of the Dpl/PrP/Sho-interactome dataset constituted the identification of multiple PDIs. In particular, Pdia3 (also known as ERp57) was represented with strong sequence coverage (61%) and its identification was further corroborated by similarly strong identifications for calnexin and calreticulin, known interactors of this PDI. Whereas binding of PDIs to both Dpl and PrP was expected due to the presence of internal disulfide bridges in these proteins, co-segregation with Sho (Pdia3 117∶114 iTRAQ reporter ion ratio of 2∶1), which does not carry a disulfide bridge, was an unexpected finding. PDIs are known to harbor both isomerase and chaperone functions. In light of previous calls for an involvement of chaperone-like activities in the conversion of PrPC to PrPSc, we asked whether a subset of PDIs may play a role in this process. It is currently thought that the conversion of PrPC occurs only after it has reached the cell surface [32], [33], i. e. well-removed from its presence within the ER where it would be expected to encounter PDIs. Thus, if PDIs were involved in PrPC conversion, these proteins would also have to be present outside the ER, preferably at the plasma membrane. Intriguingly, a small number of reports suggest that, at least in some cells, this might indeed be the case (please refer to Discussion section). To investigate whether Pdia3, P4hb and calreticulin are also present at the cell surface in mouse neuroblastoma cells, a cell surface biotinylation experiment was undertaken. Cells were incubated for a brief duration with a cell-impermeable biotin-labeling reagent, which conjugates to primary amines through an N-hydroxysuccinimide ester group, or were subjected to mock treatment. To determine the degree to which broken cells were present at the time of cell surface biotinylation, the downstream analysis was extended to histone 2B. If a large amount of broken cells had been available, histone 2B would have been present at appreciable levels in the cell culture medium resulting in an unequal signal, absent in the mock-treated sample and strong in the biotinylated sample. The observation that histone 2B signals in streptavidin agarose eluates were equal documented that low levels of non-biotinylated histone 2B had been captured unspecifically and served as evidence that levels of extract employed in the side-by-side streptavidin agarose capture steps had indeed been equal. Western blot analysis of extract and streptavidin eluate fractions probed with a PrPC-specific antibody served as a positive control (Fig. 5). Taken together this experiment revealed that a subset of Pdia3, P4hb and calreticulin was enriched in streptavidin eluate fractions derived from in vivo biotinylated cells compared to the corresponding samples derived from mock-treated control cells. The possibility that this distribution may have been the result of an unspecific binding to the affinity matrix was excluded based on a comparison with histone 2B signals observed evenly across all streptavidin eluate fractions, a distribution expected for a protein which binds to the affinity matrix unspecifically. A calculation based on a semi-quantitative titration analysis of Western blot signal intensities revealed that the relative amounts of PDIs and calreticulin at the plasma membrane are small compared to their respective quantities in the ER, but constitute at least 1∶1000 of the total amounts in these cells. Actual amounts may exceed these estimates as the calculation did not account for the non-stoichiometric labeling achieved during the biotin conjugation step and losses of biotin-derivatized PDIs incurred during the streptavidin affinity purification step. We next addressed whether PDI-inherent chaperone activities influence PrP conversion in prion-infected mouse neuroblastoma cells (ScN2a). The identification of four different PDIs in the dataset suggested that the knockdown of just one PDI may be uninformative. We therefore employed inhibitors known to interfere broadly with the activity of PDIs. One possible outcome of PDI inhibition in vivo would include a depression of PrPSc levels by inhibiting PDI-mediated unfolding of an obligate precursor form of PrPC. Interestingly, both the addition of the dodecapeptide antibiotic bacitracin or 5,5′-dithiobis (2-nitrobenzoic acid) (DTNB), membrane-impermeable reagents known to inhibit PDIs through noncovalent and covalent modes of binding, respectively [34], caused an increase in the level of proteinase K-resistant PrPSc (Fig. 6). The effect sizes correlated with inhibitor concentrations applied, and in the presence of up to 1 mM of either of the inhibitors, the viability of ScN2a cells (assessed by the trypan blue dye exclusion assay) was not impaired during the course of the experiment. A direct inhibitory effect of these reagents on the proteinase K digestion itself can be ruled out, since levels of the faster migrating PrPSc-derived signals, characteristic for the presence of PrPSc in this disease model (attributed to the action of endogenous proteases that produce a “trimming” of PrP N-terminal sequences [35], [36]), were similarly elevated even prior to the proteinase K digestion step. This observation was made with two clones of ScN2a cells obtained from independent sources; surprisingly, however, it could not be observed with an isolate of chronically prion-infected GT1 cells or two additional ScN2a clones derived from yet other sources (Fig. S2), possibly indicating the existence of cell line- or subclone-specific PDI family member expression profiles. We note that variations in the behavior of different ScN2a cell clones are well-documented, as is the occurrence of heteroploidy. While variations in our experimental paradigm preclude a generalized observation, they nonetheless put a spotlight on ER-derived chaperones that may engage in physiological interactions with PrPC even at the cell surface, as discussed below.
Taken together, this work suggests that members of the prion protein family may be embedded in specialized membrane domains characterized by an extended molecular network which supports functions in adhesion control, lactate metabolism and cell fusion events. Our investigation leads to the conclusion that biochemical interactions among the members of the mammalian prion protein family exist. It confirms multiple candidate interactors of PrPC in a unified paradigm including an independent verification of the laminin receptor precursor (LRP) as an authentic PrP interactor. It thereby not only reconciles a fragmented literature, but should facilitate future efforts to further define the relative contribution of these interactors to the biology of PrPC. Novel interactors uncovered in this study paint a coherent picture based on which an argument for the spatial proximity of NCAM, basigin, Na/K ATPase and LRP can be made, even independent of their ability to bind to PrPC. Moreover, data presented here raise many new questions as to (i) whether biological outputs of PrPC implied by the “guilt-by-association” logic will be confirmed, (ii) the importance of oligomannosidic N-glycans for the structural integrity and assembly of the molecular environment of PrPC, (iii) the role which a subset of cellular chaperones may play in PrP biology and disease, and (iv) the exact environment which hosts the conversion of PrPC. It is hoped that this work will stimulate investigations which will not only resolve these questions, but lead to insights from which a therapeutic strategy for addressing prion diseases can be derived.
All procedures which required the handling of mice were conducted in accordance with an animal use protocol (No. 20006633) approved by the University of Toronto Animal Care Committee. The mouse monoclonal antibody directed against PrP (clone 7A12) was a generous gift from Dr. Man-Sun Sy (Case Western Reserve University School of Medicine, Cleveland, OH, USA). The anti-FLAG M2 antibody (F3165) was obtained from Sigma-Aldrich (Oakville, ON, Canada). Anti-ERp57 (Pdia3) antiserum was raised against glutathione S-transferase-fused mouse ERp57. Anti-P4hb antiserum (SPA-891) was purchased from StressGen Biotechnologies (Victoria, BC, Canada). FLAG affinity tags (N-DYKDDDDK-C) were inserted before residue 29 of mouse PrP, residue 27 of mouse Dpl, and residue 26 of mouse Sho (all in the pcDNA3 mammalian expression vector) using standard PCR-based mutagenesis techniques. The identity of all constructs was verified by DNA sequencing. Cell culture and formaldehyde crosslinking of N2a cells followed a protocol described before [10]. For bulk selection of stably transfected cells, cultures were expanded in the presence of 1 mg/mL G418 and maintained at a concentration of 0. 2 mg/mL G418. RML-infected N2a cells (ScN2a) were maintained as above. For PDI inhibition experiments, DTNB or bacitracin (Sigma-Aldrich) was added to the cell culture medium at indicated concentrations followed by a two-day incubation at 37°C. The effect of inhibitor treatments on cell viability was assessed by the Trypan blue dye exclusion assay. Briefly, 0. 4% Trypan blue was gently mixed with trypsinized cells and incubated for 5 minutes at room temperature. Subsequently, the percentage of viable cells per volume equivalent was determined with the use of a hemocytometer. Approximately 109 in vivo FA-crosslinked cells each of control and FLAG-prion expressing N2a cell lines were lysed in homogenization buffer (50 mM NH4Cl, 80 mM Tris, pH 8. 0) supplemented with 1× Complete Protease Inhibitor Cocktail (Roche, Palo Alto, CA, USA). To ensure near quantitative extraction of membrane proteins, an equal volume of extraction buffer (20 mM NaCl, 1% sodium deoxycholate, 1% NP-40,20 mM Tris, pH 8. 0) was added, followed by a 30-min incubation and 5-min sonication in a water bath sonicator. Insoluble cellular debris was removed by high-speed centrifugation (100,000×g, 1 h). Subsequently, the crosslinked bait protein complexes were immunoaffinity-captured on anti-FLAG-agarose (Sigma-Aldrich). During this step samples were gently agitated on a turning wheel for 12 h, then washed extensively with 0. 5 M NaCl, 0. 05% SDS, 1% NP-40,20 mM HEPES, pH 7. 3, and detergents removed by a pre-elution wash with 10 mM NH4HCO3, pH 8. 0. Proteins were eluted by acidification with 0. 2% trifluoroacetic acid, 20% acetonitrile, pH 2. 0. Protein-containing fractions were denatured in the presence of 6 M urea, 20 mM NH4HCO3, pH 8. 0, followed by reduction with 1 mM tris- (2-carboxyethyl) -phosphine for 30 min at 60°C and alkylation with 2. 5 mM 4-vinylpyridine for 1 h at room temperature in the dark. Samples were diluted four-fold to ensure that the concentration of urea did not exceed 1. 5 M. Tryptic digestion was initiated by the addition of 1% (wt/wt) of side chain-modified, TPCK-treated porcine trypsin and allowed to proceed at 37°C for 6 h. Following trypsinization, equal quantities of tryptic peptide mixtures were spiked with 1 pmol of synthetic (Glu1) -Fibrinopeptide B (GluFib) (Sigma-Aldrich) to serve in the downstream analysis as an internal control for the efficiency of individual labeling reactions. Equal labeling with all four reagents was confirmed by equal intensities of 114∶115∶116∶117 signature peaks upon forced fragmentation of the GluFib [M+2H]2+ parent ion at m/z 785. 85. Any strong deviation from this ratio would have indicated problems with the labeling reaction or recovery of individual samples prior to the sample mixing step. Individual iTRAQ labeling reagents (Applied Biosystems, Foster City, CA, USA) were reconstituted in ethanol, added to peptide mixtures derived from the tryptic digestion of IP eluates (Control: iTRAQ 114, Dpl: iTRAQ 115, PrP: iTRAQ 116, Sho: iTRAQ 117) and incubated at room temperature in the dark for 3 h. Strong cation exchange (SCX) chromatography was used to achieve peptide fractionation of the complex digest mixture. Samples digested with trypsin were adjusted to 25% acetonitrile and acidified (pH 3. 0) by 20-fold dilution in 25% acetonitrile, 20 mM KH2PO4, pH 3. 0. High performance liquid chromatography (HPLC) was carried out using the Ultimate System (Dionex, Sunnyvale, CA, USA) equipped with a microflow calibration cartridge, a Valco injection port and a 180 nL volume UV cell. Separation was achieved on a self-packed 0. 5 mm×110 mm Luna SCX column (Phenomenex, Torrance, CA, USA) at a flow rate of 18 µl/min with a steep salt gradient from 0–400 mM NH4Cl in 25% acetonitrile, 20 mM KH2PO4, pH 3. 0. Fractions eluted from the SCX column were desalted with C18 Empore (3 M, Minneapolis, MN, USA) stop and go extraction (STAGE) tips and subsequently subjected to nano-flow RP-HPLC using the Ultimate LC system (Dionex, Sunnyvale, CA, USA) equipped with a nanoflow calibration cartridge at a flow rate of 250 nL/min. Peptides were separated on a 75-µm ID self-packed column containing Proteo C12 reverse-phase matrix (Phenomenex) using a 100-min gradient from 2%–34% acetonitrile in water, with 0. 1% (wt/vol) formic acid as the ion-pairing agent. The column effluent was coupled directly via a fused silica capillary transfer line to a QSTAR XL hybrid quadrupole/time-of-flight tandem mass spectrometer (Applied Biosystems; MDS Sciex, Concord, ON, Canada) equipped with a MicroIonSpray source. The progress of each LC/MS run was monitored by recording the total ion current (TIC) as a function of time for ions in the m/z range 300 to 1800. At 5-s intervals through the gradient, a mass spectrum was acquired for 1 s, followed by one collision-induced dissociation (CID) acquisition of 4 s each on ions selected by preset parameters of the information-dependent acquisition (IDA) method, using nitrogen as the collision gas. Singly-charged ions were excluded from CID selection. The collision energy was adjusted automatically for each CID spectrum using an empirically-optimized formula which considers the charge state and m/z value of the precursor ion. Peak lists for database searching were created using Mascot Distiller (Version 1; MatrixScience, London, UK). Searches were performed using designated MS/MS data interpretation algorithms within Protein Prospector (Version 4. 21. 3; University of California, San Francisco, CA, USA) [69] and Mascot (Version 2. 2; MatrixScience). Searches considered up to one missed cleavage and charge states ranging from +2 to +4. The analysis of iTRAQ data was assisted by the software program ProteinPilot (Version 2. 0; Applied Biosystems; MDS Sciex). For a protein to be listed in the data tables, it had to be identified by all three search algorithms. All proteins listed in the table were identified with high confidence by the application of the following filters: (i) CID spectra with individual confidence scores of less than 80% were not included (the determination of these confidence scores is documented in the Protein Pilot 2. 0 Software Help); (ii) assignments to non-iTRAQ-labeled peptides were not considered; (iii) all identifications of proteins had to be based on at least two CID spectra. Raw iTRAQ ratios were corrected for impurity levels of individual reagent lots determined by the manufacturer. In instances where only two peptides supported the identification of a protein, we required the underlying CID spectra to generate a Mascot score indicating a <5% probability that the match could be considered a random event and further confirmed matches by manual interpretation of spectra. As searches were carried out without species restriction, the correct assignment of matches to mouse entries served as an additional internal control. Identifications were confirmed in repeat experiments performed at a two-fold lower scale. It should be noted that the vast majority of proteins were identified with Mascot scores exceeding thresholds conventionally applied for confident identifications. The mass tolerance range between expected and observed masses used for database searches was ±150 ppm for MS peaks, and ±0. 15 Da for MS/MS fragment ions. These relatively large thresholds were used to capture more of the low intense peaks that frequently display broader distribution and are therefore assigned with lower mass accuracy. Threshold levels were optimized based on LC/MS/MS datasets of tryptic digests of standard proteins. All samples were searched against the National Center for Biotechnology Information nonredundant database (nrNCBI, release: June 2008) and a ‘decoy’ database in which all entries of the above NCBI database had been inverted. The interactome diagram was constructed using Cytoscape 2. 6. 0 [70]. Mice were subjected to time-controlled transcardiac perfusion crosslinking and brain extracts generated as described before [16]. Brain extracts which had been cleared of all insoluble content by high speed centrifugation (100,000×g, 1 hr) were added to snowdrop (Galanthus nivalis) lectin agarose (Sigma-Aldrich), which had been pre-equilibrated in Lysis buffer (50 mM Tris, pH 8. 3,150 mM NaCl, 0. 5% NP-40,0. 5% sodium deoxycholate, 1× Complete Protease Inhibitor), and incubated on an end-over-end turning wheel for 12 h at 4°C. Following extensive washes with lysis buffer, proteins were eluted by incubation of lectin agarose beads with 1× Laemmli sample buffer containing 100 mM DTT. To generate the crosslink reversal series, snowdrop lectin agarose eluate fractions were subjected to 90°C heat treatment for the indicated durations. N2a and ScN2a cells, grown to 90% confluency, were washed three times with 10 ml of ice-cold PBS and subsequently subjected to biotinylation with 0. 5 mg/ml HOOK-Sulfo-NHS-SS-Biotin (Pierce Biotechnology, Rockford, IL, USA) in PBS for a duration of 30 min at 4°C. Residual biotin was quenched by the addition of 100 mM glycine in PBS and further incubation for 15 min. Following cell lysis in detergent containing buffer (0. 5% NP40,0. 5% DOC, 150 mM NaCl, 1× Complete Protease Inhibitor Cocktail) the capture of biotinylated proteins on streptavidin agarose (Sigma-Aldrich) occurred during a 12 h incubation. Following extensive washing with lysis buffer, captured proteins were eluted by reducing the disulfide bond within the HOOK-Sulfo-NHS-SS Biotin conjugation reagent in the presence of 100 mM DTT, 62. 5 mM Tris, pH 6. 8,2% SDS. Confluent cells were lysed in 10 mM Tris-HCl, pH 8. 0; 100 mM NaCl; 0. 5% NP-40; 0. 5% sodium deoxycholate (DOC). Equal amounts of protein determined using the bicinchoninic acid (BCA) reagent were digested with 20 µg/ml PK (GIBCO, Carlsbad, CA, USA) at a ratio of 1∶25 (w/w protease to protein) for 1 h at 37°C. Reactions were stopped by the addition of 2 mM phenylmethylsulfonyl fluoride (PMSF). Following ultracentrifugation, pellets were resuspended in SDS-loading buffer and subjected to immunoblot analysis.
Prions underlie rare but invariably fatal neurodegenerative diseases in humans and other mammals. Public awareness of these diseases has grown with the occurrence of cases of BSE (also known as “Mad Cow Disease”) and the realization that this disease can, in rare instances, be transmitted to humans. The normal cellular prion protein is found in most cell types within the body. In disease, this protein acquires a different shape which tends to aggregate and poison nearby cells. The disease-associated conversion of the prion protein appears to require its localization to specialized cellular membrane regions rich in cholesterol; however, the precise molecular environment which hosts this event has remained elusive. We used a cell-based disease model to identify proteins which reside in close proximity to the prion protein and two closely related mammalian proteins. We demonstrate that these three proteins may not only populate highly similar environments but may also interact with each other. We further identified an extended molecular network in proximity of these proteins which supports functions in adhesion control, lactate metabolism and cell fusion events. It is anticipated that these insights will contribute to efforts to rationally interfere with aberrant protein-protein interactions underlying these diseases.
infectious diseases/prion diseases
Interactome Analyses Identify Ties of PrPC and Its Mammalian Paralogs to Oligomannosidic N-Glycans and Endoplasmic Reticulum-Derived Chaperones