Inefficient Peptide Binding by Cell-Surface Class II MHC Molecules

CELLULAR IMMUNOLOGY ARTICLE NO.

182, 1–11 (1997)

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Inefficient Peptide Binding by Cell-Surface Class II MHC Molecules Melanie A. Sherman, Dominique A. Weber, Ellen A. Spotts, Joseph C. Moore, and Peter E. Jensen Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Atlanta, Georgia 30322 Received August 1, 1997; accepted October 17, 1997

cules bind peptide antigens through an exchange mechanism involving the release of CLIP and the newly formed class II–peptide complexes progress to the cell surface. Peptide loading predominantly occurs in specialized endosomal compartments (MIIC or CIIV) found in antigen-presenting cells (5–8). However, it is likely that class II molecules can bind peptides in multiple endocytic compartments (9) and there is good evidence that some peptide complexes are generated in early endosomes where class II molecules recycling from the cell surface may undergo further rounds of peptide exchange (10–14). APC require only a few hours to internalize and present protein antigens to T cells (15, 16). Many laboratories have reported that specific class II–peptide complexes can form in less than 1 h (17–20). In addition, a major fraction of class II molecules become stable in SDS within a few hours of biosynthesis (6, 9), which is indicative of stable association with peptide antigens (21). These observations confirm that live APC normally require less than 4 h to load peptides derived from protein antigens onto newly synthesized class II MHC molecules for presentation to T cells. Peptide binding by newly synthesized class II molecules is catalyzed by the MHC-encoded membrane protein, HLADM (H-2M in mice) (22–24). DM facilitates peptideexchange reactions by increasing the rate of dissociation of previously bound peptides, including CLIP (22– 27). In the absence of DM, purified class II molecules bind peptides very slowly, requiring days to reach apparent saturation (28–30). In contrast to protein antigens, short peptides with flexible conformation do not require internalization for presentation to T cells. It appears that such peptides bind almost exclusively to cell surface and not to intracellular class II molecules (18, 31, 32), probably because of rapid degradation after pinocytosis. Importantly, several previous kinetics studies with aldehydefixed (33, 34) and viable APC (17, 35) have suggested that peptides may bind rapidly (less than 4 h for apparent saturation) to cell-surface class II molecules. These reports raise the interesting possibility that cofactors present in the plasma membrane may facilitate peptide loading. DM is selectively localized in specialized anti-

The efficiency of peptide loading onto surface class II MHC molecules in intact APC was investigated, using a previously defined europium immunoassay as well as a simplified Western blot procedure. Conditions normally employed for peptide loading in T cell stimulation assays were suboptimal for peptide binding, which is enhanced at low pH, in the presence of protease inhibitors, and the absence of competing serum proteins. In contrast to some earlier reports, our results indicate that the rate of peptide loading by class II molecules is not enhanced in the environment of the plasma membrane. Peptide association rates were similar for purified and cell-surface class II molecules. As previously reported, rapid peptide binding can be achieved by reconstituting class II molecules into total cellular membranes. We report that this activity is due solely to HLA-DM (which is not present at the cell surface), since it can be specifically removed by immunodepletion with an anti-DM mAb. Thus, we find no evidence for additional cellular cofactors capable of catalyzing peptide binding to class II molecules. q 1997 Academic Press

INTRODUCTION CD4/ T cells recognize antigenic peptides stably associated with class II MHC molecules expressed on the surface of APC.1 Following biosynthesis, the class II a and b chains associate with the nonpolymorphic invariant chain in the endoplasmic reticulum. After traveling through the golgi, the complexes are directed to the endosomal pathway by a sorting signal in the cytoplasmic domain of invariant chain (1). After reaching endosomal compartments, proteases cleave the invariant chain, leaving residual fragments, CLIP, associated with the peptide-binding site (2–4). Class II mole1 Abbreviations used: APC, antigen-presenting cell(s); CIIV, class II vesicles; CLIP, class II-associated invariant chain peptides; DM, HLA-DM or H-2M; ER, endoplasmic reticulum; HA, influenza hemagglutinin peptide; HEL, hen egg lysozyme; MIIC, MHC class II compartment; MAT, influenza matrix peptide; PCC, pigeon cytochrome c.

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gen-processing compartments and it is not expressed in the plasma membrane (36–38). Thus, DM cannot enhance peptide binding by cell-surface class II molecules. In addition, analysis of mice heterozygous for a targeted mutation in DM (H-2Ma) as well as transfected cell lines expressing different levels of DM indicate that intracellular concentrations of DM are critical for efficient peptide exchange (39–41). DM is largely concentrated in dense endosomal compartments (36, 37) and thus it is unlikely to catalyze peptide exchange reactions involving recycling class II molecules (10–13) which access low-density early endosomes but not dense prelysosomal compartments. The present study was initiated to determine whether peptide binding by class II molecules is, indeed, accelerated in the environment of the plasma membrane and to determine whether additional membrane-associated cofactors, other than DM, can catalyze peptide exchange. MATERIALS AND METHODS Cell Culture T1 and T2 human (.174 1 CEM.T1 and .174 1 CEM.T2) hybrids (42) were cultured in DMEM containing 10% FCS, 2 mM L-glutamine, 2 mM sodium pyruvate, and the antibiotics penicillin–streptomycin and gentamicin. CH27 (43) and TH2.2 (44) murine B cell hybridomas, A20 murine B cells (45), and human LG2 B cells were cultured in RPMI containing the same additives and 50 mM 2-mercaptoethanol (R10). LG2 expresses DR1, CH27 expresses IAk and IEk, TH2.2 expresses IAb, IAd, and IEd, and A20 expresses IAd and IEd. Cells were maintained at 377C in 7% CO2 .

leimide, 10 mM leupeptin, 270 mM TLCK, and 10 mM EDTA as previously described (all purchased from Sigma Chemical Co., St. Louis, MO) (22). Suspensions were cleared of debris and nuclei by centrifugation at 3500g for 10 min at 47C, and membranes were pelleted from supernatants by centrifugation at 22,000g for 30 min at 47C. Membranes (1 mg/ml total protein, 108 cell equivalents/ml) were solubilized in 0.5% 1-S-octyl-bD-thioglucopyranoside (SOG, Pfanstiehl Laboratories, Inc., Waukegan, IL) in phosphate-buffered saline (PBS) with protease inhibitors in the presence or absence of 0.4 mM purified DR1 or IAk. SOG was used to solubilize membranes because of the high critical micelle concentration of the detergent which can be easily removed by dialysis against PBS, allowing liposome formation (34). In some experiments 0.4 mM class II protein was introduced into detergent-solubilized purified lipid containing 500 mM L-a-dipalmitoylphosphatidylcholine (DP-PC) and 140 mM cholesterol (both from Sigma Chemical Co.) before dialysis. The liposomes were used directly in binding assays. Protein content was measured by BCA assay (Pierce) with BSA as a standard. Peptide Binding Assay

DR1 was purified from the EBV-transformed homozygous LG2 B cell line (DRB1*0101) and IAk was purified from CH27 using a monoclonal antibody LB3.1 (anti-DRa) or 10-2.16 (anti-IAk) immunoaffinity column as previously described (34). Membranes were prepared from TH2.2, T1, or T2 cells by hypotonic lysis (10 mM Tris, pH 8.0) in the presence of protease inhibitors: 5 mM phenanthroline, 4 mM PMSF, 15 mM pepstatin, 568 mM TPCK, 1 mM benzamidine, 1 mM iodoacetamide, 6 mM N-ethylma-

The class II–peptide binding immunoassay has been previously described (46, 47). In brief, binding reactions using purified or reconstituted class II molecules were performed with 2 pmol DR1 or IAk (40–67 nM final concentration) and 1 mM biotinylated peptide at 377C in microfuge tubes in a 30–50 ml volume of binding buffer with (purified class II) or without (membrane-associated class II) detergent. Binding buffer contained 100 mM citrate/phosphate, pH 5, 0.2% NP-40 (for purified class II samples) and protease inhibitors (detailed above). After the indicated time period, the pH was then neutralized by addition of an equal volume of neutralization buffer (3.5% skim milk, 0.7% BSA, 335 mM Tris, pH 7.5, 0.07% sodium azide, 0.07% Tween 20, and 0.35% NP-40) and the samples were plated on precoated microtiter wells (see below). Peptide binding to cell-surface class II was measured using 1 or 2 1 106 cells/sample. LG2 (expressing DR1) or CH27 (expressing IAk) were washed twice in HBSS and fixed with 0.5% paraformaldehyde for 10 min at 247C. After extensive washing in RPMI 1640 media with 10% serum followed by PBS, the cells were incubated with biotinylated peptide in 50–100 ml binding buffer lacking detergent. After incubation at 377C for the indicated period of time, cells were washed twice with HBSS and lysed for 40 min on ice in 60 ml lysis buffer (0.5% NP-40, 0.15 M NaCl, 50 mM Tris, pH 8.0, and protease inhibitors). The lysates were cleared by centrifugation for 10 min at 20,000g and added to precoated assay plates. Plates were coated with 100 ml of 20 mg/ml L243 mAb (DRa specific) or 10-2.16 (IAk specific) in borate-buffered saline, pH 8, overnight at 47C and blocked for 30 min at 247C.

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Peptides Peptides were synthesized as previously described (34). MAT(17-31), sequence: SGPLKAEIAQRLEDV; HEL(46-61), sequence: NTDGSTDYGILQINSR; and HA(306-318), sequence: PKYVKQNTLKLAT. Peptides were labeled with biotin at the a amino group by reaction with biotinamidocaproate N-hydroxysuccinimide (34). Reconstitution of Class II in Membranes and Liposomes

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Samples were allowed to bind to the antibody-coated plates for 2 h at 47C. After washing, excess europiumstreptavidin was added and fluorescence was measured at 612 nm as described (47). The data points represent the mean fluorescent counts per second/1000 (cps 1 1003) of duplicate or triplicate samples. In some experiments, data were normalized to maximum observed binding. Actual fluorescence signals are reported in the figure legend. The results shown in this communication are representative of at least three separate experiments.

by electrophoresis on SDS–PAGE. After the proteins were transferred to nitrocellulose, DM was detected with anti-DMa rabbit sera (22), donkey anti-rabbit horseradish peroxidase (Amersham), and chemiluminescent substrate (Renaissance kit, Dupont). Quantification of bands in immunoblots was accomplished by optical densitometry using a Hewlett scanner (Scan Jet 4C) and the MacIntosh-based software NIH Image1.55f. The black level was adjusted to ensure linearity of the densitometry analysis. The concentration of DM in each fraction is represented in arbitrary units.

Detection of Peptide–Class II Complexes by Western Blot Analysis

Immunodepletion

Class II–peptide complexes were formed at pH 5 as described above using 2 1 106 fixed B cells or 2 pmol (67 nM) purified class II protein. No attempt was made to use exactly the same concentrations of IAk or DR1 molecules in samples containing purified versus cellassociated class II molecules. SDS-stable complexes were detected by neutralization of the samples with 0.4 M Tris, pH 7.5 (final concentration), and incubation with SDS sample buffer (1.25% final concentration of SDS) for 15 min. The samples were separated on linear 12% SDS polyacrylamide gels by electrophoresis (120 V for 90 min). Protein was transferred to nitrocellulose membranes for 40 min at 12 V in 25 mM Tris, 192 mM glycine, pH 8.3. Class II–biotin–peptide complexes were detected with avidin–horseradish peroxidase (Sigma) and chemiluminescent substrate (Renaissance kit, DuPont). Film exposures were chosen to optimize linearity and increase our capacity to detect changes in peptide binding over the time course of the experiments.

Membranes of T1 and T2 were prepared as described above and solubilized in 0.5% NP40 in the presence of protease inhibitors. Protein A–Sepharose beads (Pharmacia Biotech Inc., Piscataway, NJ) were precoated with mAb anti-DMb cytoplasmic domain (a kind gift from Dr. S. K. Pierce) or MK-D6 (anti-I-Ad) as control, by incubating 200 ml 1:1 bead suspension in PBS with 30 mg mAb for 2 h at 47C. Coated beads were washed extensively in PBS. One hundred fifty-microliter membranes (1 mg/ml total protein, Ç108 cell equivalents/ ml) were added to 50 ml of washed beads and incubated overnight at 47C. The depletion was repeated with freshly coated beads for an additional 2 h. Fifty-microliter depleted membranes were incubated with 2 pmol DR1 and 1 mM biotinylated HA(306-318) for 4 h at 377C. Class II–peptide complexes were quantified using the europium–streptavidin assay. RESULTS

T1 cells (2 1 108) were washed with PBS and disrupted in 5 ml cold homogenization buffer (0.25 M sucrose, 10 mM Hepes, pH 7.2, protease inhibitors) with a Dounce homogenizer. Nuclei were pelleted by lowspeed centrifugation (2000g for 10 min at 47C) and the cleared supernatant was layered onto a 17% Percoll gradient in homogenization buffer. The gradient was centrifuged at 35,000g for 1 h, 47C, in a Beckman SW28 rotor. Fractions were collected from the bottom of the gradient. Each fraction was tested for b-hexoseaminidase (lysosomal marker) and alkaline phosphodiesterase (plasma membrane marker) as described (48). To assess functional activity, 200 ml of each fraction was incubated in 0.5% NP-40 overnight at 47C to solubilize the membranes. DR1 (2 pmol) and 1 mM biotinylated MAT(17-31) were added to each fraction and incubated for 4 h at 377C. Class II–peptide complexes were detected with the europium fluorescence assay. To measure DM protein, 20 ml of each fraction was mixed with reducing SDS sample buffer and separated

Several previous studies have shown rapid binding of synthetic peptides to cell-associated class II molecules (17, 33–35). Experimental conditions, however, can substantially affect peptide binding by cell-surface class II glycoproteins. The conclusion that peptide binding occurs predominantly at the cell surface rather than in endosomal compartments (18) is further supported by the observation that chloroquine does not affect peptide binding to class II molecules in unfixed cells ((32) and Fig. 1A), although it is known to interfere with presentation of protein antigens by raising the endosomal pH (49). In order to compare the association rates for peptides binding to purified or cell-associated class II molecules, similar binding conditions had to be used. Although, in functional assays with T cells, APC are typically exposed to peptide antigens in serum-containing media at neutral pH, peptide binding to purified class II is generally enhanced at low pH (34, 50 – 53). A similar enhancement can be measured at low pH for the binding of peptide by class II proteins on the surface of fixed or unfixed cells, such as HEL(4661) with IAk (Fig. 1A), and MAT(17-31) with DR1

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Subcellular Fractionation

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FIG. 1. Optimal conditions for peptide binding by cell-surface class II molecules. (A) APC bind peptide optimally at low pH with protease inhibitors. 2 1 106 unfixed CH27 cells were pulsed with 2 mM biotin–HEL(46-61) for 4 h at 377C in pH 5 or 7 citrate–phosphate buffer with (solid bars) and without (hatched bars) a cocktail of protease inhibitors. Some samples contained 60 mM chloroquine as indicated. (B) Serum inhibits peptide binding to class II. Unfixed LG2 cells were incubated with or without 2 mM biotin–MAT(17-31) for 4 h at pH 7.4 in RPMI (open bars) or RPMI containing 10% FCS (hatched bars). Protease inhibitors were included as indicated. Peptide binding was quantified by using the europium fluorescence immunoassay.

(47). Peptide binding by cell surface class II molecules is enhanced in low pH binding buffer compared with cell culture media at neutral pH. Proteases present in serum or on the cell surface may also complicate peptide loading. We generally observe increased class II–peptide complex formation after incubating cells with peptide in the presence of a cocktail of protease inhibitors (Figs. 1A and B). It has been shown that, in the presence of proteases, peptide antigens may have a very short half-life (54). Our data also show that serum inhibits peptide loading (Fig. 1B). Serum may contain peptides or partially unfolded proteins that can directly compete with peptide for binding to class II molecules (55). Thus, class II–peptide binding is generally suboptimal in the experimental conditions typically used to measure the relative potency of peptide antigens in stimulating CD4/ T cells. A simple streptavidin Western blot assay was initially used to compare peptide binding by cell-surface and purified class II molecules. This assay relies on the unusual stability of class II–peptide complexes in SDS detergent. Fixed CH27 B cells (expressing IAk and IEk) were incubated with biotinylated HEL(46-61), washed, solubilized in NP-40 lysis buffer, and unheated samples were resolved by SDS–PAGE. Western blots probed with streptavidin–HRP showed a band migrating at Ç65 kDa, consistent with SDS-stable class II–peptide complexes (Fig. 2A). The band was absent from boiled samples, indicating noncovalent association of biotin– peptide with class II molecules. Excess HEL(46-61) but not a control peptide, PCC(91-104), inhibited formation of the band. PCC(91-104) binds with high affinity to

IEk but not to IAk. No band was observed with A20 B cells, which express IAd and IEd but not IAk. The band was removed from lysates precleared with anti-IAk but not anti-IEk mAb (Fig. 2B), confirming the identity of the band as biotin–HEL(46-61)–IAk complexes. We conclude that this simplified assay can be used to specifically measure peptide binding by cell-surface class II molecules. The streptavidin Western blot assay may be useful in laboratories where more complex assays for class II–peptide binding are unavailable. It should be emphasized, however, that this assay will not be useful for measuring class II–peptide complexes that are unstable or partially stable in SDS. This assay was used to directly compare peptide binding to purified and cell-associated IAk. Fixed CH27 B cells or purified IAk were incubated with an optimal concentration (1 mM) of biotin–HEL(46-61) at pH 5 for various time periods and complexes were detected by Western blotting. The quantity of purified IAk or DR1 (2 pmol) used in our experiments was chosen to roughly correspond to the quantity of these proteins expressed in 2 1 106 CH27 or L243 cells (data not shown). At least 19 h were required to approach a plateau in peptide binding by IAk in either situation (Fig. 2C, top). Similar results were obtained by comparing the rate of peptide binding by cell-surface and purified DR1 molecules (Fig. 2C, bottom). These results suggested that the relative rate of peptide binding by cell-surface class II molecules does not greatly differ from that observed with purified class II proteins. These results were confirmed using a europium fluorescence immunoassay to quantify peptide binding.

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FIG. 2. Detection of class II–peptide complexes using a streptavidin Western blot assay. (A) Fixed CH27 or A20 B cells (2 1 106) were incubated with or without 1 mM biotin–HEL(46-61) for 2 h at 377C in 50 ml pH 5 buffer in the presence or absence of 100 mM competitor peptide as indicated. (B) Fixed CH27 (5 1 106) were incubated with 5 mM biotin–HEL(46-61) for 3 h at 377C in 100 ml pH 5 buffer. The cells were washed and solubilized in 0.5% NP-40 buffer. The cell lysates were precleared with two rounds of Protein A–Sepharose preloaded with 10-2.16 (anti-IAk) or 14-4-4 (anti-IE) mAb before separation of unheated samples on SDS–PAGE. (C) Fixed LG2 or CH27 (2 1 106) or purified DR1 or IAk (2 or 5 pmol) were incubated with 1 mM biotin–MAT(17-31) (DR1) or biotin–HEL(46-61) (IAk) for the indicated period of time at 377C in 50 ml pH 5 buffer. Each sample was pH neutralized and samples containing cells were solubilized in NP-40 detergent before SDS–PAGE. Samples were not boiled unless otherwise indicated. Biotin–peptide–class II complexes were detected by Western blotting with streptavidin–HRP as described under Materials and Methods.

This assay is very sensitive with a wide range of linearity and the results are not complicated by the potential effect of SDS on the stability of particular class II– peptide complexes (46). Purified IAk and DR1, or fixed APC that express these proteins, were incubated with excess biotinylated peptide for various periods of time. The membranes were solubilized in NP-40 detergent and specific class II proteins were captured on assay plates precoated with appropriate mAb. Class II–biotin–peptide complexes were quantified by incubation with excess europium-labeled streptavidin and the time-resolved fluorescence of chelated europium was

measured. Again, the kinetics of peptide binding by cell-associated class II molecules was similar to that observed with purified proteins (Figs. 3A and 3B). The results were normalized to the maximum observed fluorescence signal to facilitate comparison. In the experiment shown, a somewhat greater quantity of peptide complexes were formed in samples containing CH27 cells compared to samples containing purified IAk (see legend, Fig. 3). By contrast, more peptide complexes were formed in samples containing purified DR1 than those containing DR1-expressing LG2 cells. Despite these differences, which result from differences in the

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FIG. 3. Kinetics of peptide loading using the europium fluorescence immunoassay. (A) Fixed CH27 (1 1 106, open circles) or purified IAk (2 pmol, closed circles) were incubated with 1 mM biotin–HEL(46-61) for the indicated time period at 377C in pH 5 buffer. The samples were pH neutralized and the cells were solubilized before addition to a 10-2.16 (anti-IAk) precoated assay plate. Class II–peptide complexes were detected as described under Materials and Methods. (B) Fixed LG2 (2 1 106, open circles) or purified DR1 (2 pmol, closed circles) were incubated with 1 mM biotin–MAT(17-31) as described above. Class II–peptide complexes were measured by europium fluorescence as described under Materials and Methods. Maximum binding for LG2: 371; CH27: 527; DR1: 1839; IAk: 264 (cps 1 1003).

concentrations of functional class II molecules available for peptide binding in the samples, the relative kinetics of peptide binding to purified and cell-surface class II molecules are strikingly similar. The fraction of cell-surface or purified class II molecules that bind peptide after 4 or 5 h of incubation is relatively small compared to the binding observed after 1 or 2 days of incubation. Two-fold changes in the concentration of purified class II protein or class II-expressing cells did not change the general kinetics profile (data not shown). These biochemical experiments do not support the conclusion, reached in previous functional studies measuring T cell activation (33, 34), that peptide binding is enhanced in the environment of the plasma membrane. Fixed APC were used in the experiments described above. Ceppellini et al. (35) reported rapid binding of radioiodinated peptide to HLA-DR molecules on living B lymphoblastoid cells. The kinetics of binding were strikingly delayed in fixed cells (35). We considered the possibility that aldehyde fixation inhibits the activity of a cell-surface cofactor that facilitates peptide binding to class II molecules. To address this possibility, experiments were done to compare the rate of peptide binding to class II molecules on fixed and unfixed cells. Incubations at pH 5 resulted in a considerable loss of viability as measured by trypan blue exclusion in unfixed APC; however, the cells remained morphologically intact. As shown in Figs. 4A and 4B, the fixation procedure does not greatly alter the rate of peptide binding by cellsurface class II molecules. The small decrease in peptide binding observed in fixed compared to unfixed APC may in part reflect direct damage to class II molecules by the fixation procedure. The degree of peptide binding

was substantially reduced under conditions in which viable B cells were incubated with peptide at neutral pH in RPMI 1640 with or without serum compared to binding at pH 5 (Fig. 1 and data not shown). These results clearly demonstrate that, under a variety of experimental conditions, cell-surface molecules do not bind peptide faster than purified class II molecules. Further experiments were done to explore the possibility that intracellular membrane components, other than DM, may facilitate peptide loading by class II molecules recycling through endosomes. Murine B cell membrane preparations containing endosomal, ER, Golgi, and plasma membranes were solubilized in detergent. Purified IAk or DR1 was added to the solubilized membranes and the preparations were dialyzed to reconstitute membranes in the form of unilamellar liposomes. As a control, class II was also reconstituted into purified phosphatidylcholine/cholesterol vesicles. There was no significant loss of class II during dialysis and the liposome preparations contained similar concentrations of class II protein (data not shown). After reconstitution into B cell membranes, DR1 and IAk bound peptide rapidly, showing an initial plateau after only 4 h (Fig. 5). Liposomes containing purified lipids and the same quantity of class II molecules bound peptide much more slowly (Fig. 5), at a rate similar to peptide binding to purified class II in detergent (data not shown). Interestingly, the human DR1 protein showed a greater enhancement of peptide binding compared with the murine IAk molecule, even though the membranes were derived from a murine B lymphoblastoid line. Reconstituted liposomes without IAk or DR1 did not bind peptides (data not shown). These results confirm previous experiments with reconstituted human B cell membranes (22).

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FIG. 4. Peptide binds to cell-surface class II molecules on fixed and unfixed cells with similar kinetics. 1 1 106 fixed (closed circles) or unfixed (open circles) CH27 (A) or LG2 (B) cells were incubated with 2 mM biotin–HEL(46-61) or biotin–MAT(17-31) for the indicated time periods at 377C in 0.15 M citrate–phosphate buffer, pH 5, with protease inhibitors. The cells were washed and solubilized in NP-40 lysis buffer before measuring peptide complexes with the europium fluorescence assay.

Total cellular membranes were fractionated on Percoll density gradients to determine whether a cofactor capable of catalyzing peptide binding was associated with low-density early endosomes or plasma membranes. Detergent-solubilized membrane fractions were tested for the ability to enhance peptide binding to purified DR1 (22). This activity was mostly localized to dense membrane fractions, probably containing MIIC (8, 56, 57), and was almost completely

segregated from the plasma membrane marker, alkaline phosphodiesterase (Fig. 6A). This is consistent with our results demonstrating inefficient peptide binding by cell-surface class II molecules. Some activity is present in intermediate-density fractions, which could reflect smaller quantities of DM distributed through a variety of endosomal compartments (37, 58, 59). Alternatively, we considered the possibility that a distinct cofactor present in low-density

FIG. 5. Rapid peptide binding by class II molecules reconstituted in mouse B cell membranes. (A) Five picomoles purified DR1 reconstituted into vesicles containing TH2.2 cell membranes (12.5 mg total membrane protein, open circles) or DP-PC and cholesterol (see Materials and Methods) (closed circles) were incubated with 2 mM biotin–MAT(17-31) for the indicated period of time at 377C in pH 5 buffer. (B) Five picomoles IAk reconstituted into TH2.2 membranes (12.5 mg protein, open circles) or DP-PC/cholesterol vesicles (closed circles) were incubated with 2 mM biotin-HEL(46-61) at 377C in pH 5 buffer for the indicated time period. The liposomes were pH neutralized and solubilized in NP-40 before measuring class II–peptide complexes using the europium fluorescence assay.

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FIG. 6. Intracellular DM fully accounts for the cofactor activity of B cell membranes. (A) Cofactor activity segregates from plasma membrane fractions. T1 cells were lysed and fractionated on a Percoll density gradient as described under Materials and Methods. Aliquots of each fraction were tested for alkaline phosphodiesterase activity, DM concentration, and for enhancing activity in a class II–peptide binding assay as described under Materials and Methods. The level of peptide binding in control samples containing unfractionated T1 or T2 membranes is indicated. T2 cells are derived from T1, but contain a large homozygous deletion in the class II region of the MHC and therefore do not express DM. (B) Immunodepletion of cofactor activity with anti-DM mAb. Detergent solubilized T1 (solid bars) or T2 (hatched bars) membranes were incubated for 18 h at 47C with protein A–Sepharose precoated with control mAb or anti-DM mAb. Treated samples were tested for the capacity to enhance biotin-MAT(17-31) binding to DR1 as described under Materials and Methods.

endosomal compartments may catalyze peptide exchange reactions. Western blot analysis demonstrated DM largely distributed in dense fractions 2–4 and in the light membrane fractions 20–23 (Fig. 6A). The latter peak probably represents DM localized in the ER, which comigrates with plasma membranes on Percoll gradients. It is interesting that the light membrane fractions had minimal activity in the functional assay despite the presence of DM. It is possible that DM requires some posttranslational modification for activity, that a large fraction of the ER-associated DM is improperly folded, or that an inhibitor is present in light membrane fractions. Small quantities of DM were detected by Western

blotting in intermediate-density fractions containing endosomes. The potential presence of endosome components, other than DM, with the capacity to facilitate peptide exchange reactions was evaluated in immunodepletion experiments. The catalytic activity present in solubilized B cell membranes was completely removed after incubation with protein A Sepharose-bound anti-DM mAb but not control mAb (Fig. 6B). These results support the conclusion that DM can fully account for the accelerated peptide binding observed with cell-associated class II molecules and that other cofactors with similar activity are not present in the plasma membranes or endosomes of B cells.

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The results presented here demonstrate that class II molecules on the surface of B cells bind exogenous peptide at a rate similar to that observed with purified class II molecules. However, the rate of peptide binding by class II molecules is markedly enhanced after reconstitution into liposomes generated from total cellular membranes isolated from murine or human B cells. The activity responsible for accelerated peptide binding is not present in plasma membrane fractions separated by Percoll density gradients. Immunodepletion experiments demonstrate that this activity can be fully accounted for by DM, with no need to postulate the existence of a second cofactor capable of catalyzing peptide loading at the cell surface or in endosomes containing recycling class II molecules. Previous reports from our own laboratory (33, 34) and others (17, 35) have provided support for the hypothesis that components of the plasma membrane may facilitate rapid peptide binding by cell-surface class II molecules. Roosneck et al. (17) demonstrated that unfixed B cells were able to optimally stimulate specific T cells after incubation with peptide for less than 1 h. We also reported apparent saturation of peptide-binding sites on fixed B cells within 4 h of incubation at low pH (33, 34). In these studies, peptide binding was determined indirectly by measuring T cell activation. It is probable that the apparent saturation was a result of maximal T cell stimulation which can be reached with the presentation of few class II–peptide complexes (60, 61), and therefore does not represent maximal occupancy of peptide-binding sites. This explanation cannot account for the observations of Ceppellini et al. (35), who reported maximal binding of radioiodinated peptide to DR1 on living B cells within 45 min of incubation. The kinetics of binding were markedly delayed in aldehyde-fixed B cells (35). By contrast, the present studies indicate that the rate of peptide binding by cell-surface class II molecules is equal to or less than that observed with purified class II molecules at the same pH. This was observed for both living and fixed APC. In the study of Ceppellini et al. (35), binding was done in medium with no attempt to inhibit protease activity. Thus, it is possible that proteolytic degradation of peptide may account for the inability to detect increased binding of peptide at later time points. Our results lead us to suggest that DM is the only membrane-associated cofactor that can catalyze peptide loading by class II molecules. This conclusion is supported by the findings that catalytic activity segregates from plasma membrane fractions on Percoll density gradients (Fig. 6A) and that this activity is fully depleted upon removal of DM (Fig. 6B). Under physiological conditions, the bulk of class II–peptide complexes appears to be generated in specialized antigenprocessing compartments through DM-catalyzed pep-

tide binding to newly synthesized class II molecules. However, strong evidence has recently been provided indicating that class II molecules recycling between the plasma membrane and endosomes can also bind selected epitopes presumably generated by the unfolding and fragmentation of proteins in early endosomes (10– 14). The role of DM in these peptide-exchange reactions is unclear. DM accumulates in dense endosomal compartments in human B lymphoblastoid cells with relatively small amounts distributed in lower density endosomes that have access to class II molecules recycling from the cell surface (Fig. 6A and (37, 58, 59, 62)). The range of concentrations of DM expressed in vivo or in transfected cell lines critically affects the efficiency of peptide exchange indicating that DM concentration is a limiting factor and that DM is not present in large excess in dense endosomal compartments (39–41). It is therefore likely that there is only a very limited capacity for DM-catalyzed peptide exchange in early endosomes. Pinet et al. have demonstrated that presentation of an epitope from influenza hemagglutinin depends on recycling of cell-surface DR1 molecules with no requirement for newly synthesized molecules (10, 11). Presentation was not dependent on expression of invariant chain or DM (10). The findings presented in the current study indicate that undefined additional cofactors present in low-density endosomes do not compensate for the low concentration of DM found in compartments with access to recycling class II molecules. Given that class II molecules do not recycle through DM-enriched dense endosomal compartments, peptide exchange by recycling class II molecules should be a relatively inefficient process. In T1 cells, DM was present in a bimodal distribution on Percoll density fractionation with most of the protein localized in lysosomal and ER/plasma membrane fractions (Fig. 6A). It is interesting that the DM-containing ER/plasma membrane fractions had little or no catalytic activity. DM (H-2M) has been reported to associate with invariant chain (36, 63) as well as the nonpolymorphic class II protein, HLA-DO (64), in the ER. It is possible that interaction with either of these proteins or a presently unidentified inhibitor blocks DM activity. Alternatively, DM may require posttranslational modification for catalytic activity. Finally, we cannot exclude the possibility that the ER-associated molecules are improperly folded, since DM was measured by Western blotting with antisera that react with denatured protein. These alternative possibilities are presently under investigation. The effect of DM on peptide binding by IAk was somewhat less than that observed for DR1 in experiments with either murine H-2M (Fig. 5) or human HLA-DM (data not shown). This may be due to a relatively fast spontaneous dissociation of CLIP (or other previously bound peptides with similar properties) from IAk as compared to DR1, and it has been speculated that IAk

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DISCUSSION

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SHERMAN ET AL.

is less dependent upon DM because it binds CLIP with low affinity (65). Interestingly, DM transfection only partially restores antigen presentation capability to mutant T2 cells expressing IAk, suggesting that a factor, in addition to class II and DM, is required for full APC potential (66). The results of the present study indicate that DM is the only vesicle-associated factor capable of catalyzing class II–peptide binding in T1 B cells. Thus, any additional cofactors must act through distinct mechanisms to facilitate antigen presentation. A common approach used to evaluate the antigenicity of peptide antigens and substitution analogs involves direct measurement of their potency in T cell stimulation assays. Under these conditions, peptides bind to cell-surface class II molecules through an uncatalyzed exchange mechanism that is subject to a number of complications. Unfolded protein or peptides released by the cells or derived from serum may inhibit peptide binding to class II molecules through direct competition (55). Proteases may substantially reduce the half-life of peptide, making it difficult to know actual concentrations. They may also differentially reduce the observed potency of peptides with different sequences based on substrate specificity. We have found that the best general approach for evaluating peptide antigens in assays with T cell hybridomas is to pulse lightly-fixed APC with peptide at low pH in serum-free buffer containing a cocktail of protease inhibitors. This approach has its own set of limitations, including the potential for inactivation of costimulatory and adhesion molecules and the loss of APC viability. Data from the present study indicate that even under optimal conditions, peptide loading and exchange by cell-surface class II molecules remains relatively inefficient. There is considerable interest in the characterization of naturally processed peptides eluted from purified class II proteins which are purified in very limiting amounts. In the light of this report pointing to the slow kinetics of peptide binding at the cell surface of APC, we are considering the possibility that transfected APC, expressing DM molecules on the cell surface by truncation of the endosomal targeting signal in the b-chain cytoplasmic domain (63, 67, 68), may provide a more sensitive system for the functional evaluation of limiting quantities of peptide. ACKNOWLEDGMENTS We are indebted to Dr. Susan K. Pierce for providing a mAb directed against the cytoplasmic tail of DMb. This work was supported by research grants from the National Institute of Health. M.A.S. was supported by a Howard Hughes Predoctoral Fellowship.

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