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. Author manuscript; available in PMC: 2014 Jun 11.
Published in final edited form as: J Immunol. 2009 Aug 26;183(6):3690–3699. doi: 10.4049/jimmunol.0900953

XBP-1-deficient plasmablasts show normal protein folding but altered glycosylation and lipid synthesis

Annette M McGehee 1, Stephanie K Dougan 1, Elizabeth J Klemm 1, Guanghou Shui 2, Boyoun Park 1, You-Me Kim 1, Nicki Watson 1, Markus R Wenk 2,3, Hidde L Ploegh 1, Chih-Chi Andrew Hu 1
PMCID: PMC4053221  NIHMSID: NIHMS591872  PMID: 19710472

Abstract

The accumulation of misfolded secreted IgM in the endoplasmic reticulum (ER) of XBP-1-deficient B cells has been held responsible for the inability of such cells to yield plasma cells, through the failure to mount a proper unfolded protein response. Lipopolysaccharide (LPS)-stimulated B cells incapable of secreting IgM still activate the XBP-1 axis normally: XBP-1 is turned on by cues that trigger differentiation and not in response to accumulation of unfolded IgM, but the impact of XBP-1 deficiency on glycoprotein folding and assembly has not been explored. The lack of XBP-1 compromised neither the formation of functional hen egg lysozyme-specific IgM nor the secretion of free κ chains. Although XBP-1 deficiency affects the synthesis of some ER chaperones, including protein disulfide isomerase, their steady state levels do not drop below the threshold required for proper assembly and maturation of the Igα/Igβ heterodimer and MHC molecules. Intracellular transport and surface display of integral membrane proteins are unaffected by XBP-1 deficiency. Given the fact that we failed to observe any defects in folding of a variety of glycoproteins, we looked for other means to explain the requirement for XBP-1 in plasma cell development. We observed significantly reduced levels of phosphatidylcholine, sphingomyelin, and phosphatidylinositol in total membranes of XBP-1-deficient B cells, and reduced ER content. Terminal N-linked glycosylation of IgM and class I MHC was altered in these cells. XBP-1 hence has important roles beyond folding proteins in the ER.

Introduction

Plasma cells produce large amounts of secreted immunoglobulins, which is their primary task in the adaptive immune response. In contrast, naïve B cells express the membrane form of IgM (mIgM) but do not secrete IgM until they are activated. B cell differentiation to plasma cells begins when a B cell is activated by an encounter with its cognate antigen or in conjunction with ligands for Toll-like receptors. This leads to the expansion of the B cell’s endoplasmic reticulum (ER) in preparation for the increase in synthesis of the secreted form of IgM (sIgM) (1). Eventually such B cells fully differentiate into immunoglobulin-secreting plasma cells (2), a process proposed to depend critically on the unfolded protein response (UPR) (3, 4).

XBP-1 is a transcription factor that drives this UPR. Its expression is ultimately controlled by the transmembrane kinase/endoribonuclease IRE-1 (3, 5), the activation of which occurs in response to pharmacologically-induced ER stress. IRE-1 modulates XBP-1 activity by catalyzing an unusual reaction that generates spliced XBP-1 mRNA, encoding a 54-kDa protein (XBP-1s) with transcriptional activity. XBP-1s translocates to the nucleus and regulates the synthesis of chaperones and other proteins believed to contribute to the proper function of the secretory pathway (4, 6, 7).

XBP-1 plays an important role in B cell differentiation: when XBP-1 is absent from B cells, the number of plasma cells is dramatically reduced (8). It has been argued that the action of XBP-1 in B cell differentiation ensures expression of proteins equipped to deal with an excess of unfolded sIgM; this excess is thought to be an unavoidable byproduct of the increased synthesis of sIgM (3, 4). In this model, the increase in synthesis of sIgM subsequent to B cell activation exceeds the folding capacity of the ER and causes an accumulation of excess unfolded proteins that activate IRE-1, which in turn triggers XBP-1 activation. Activation of XBP-1 by IRE-1 serves to increase the size of the ER and enhances its folding capacity to handle the increased levels of sIgM. This model predicts that, in the absence of XBP-1, differentiating B cells are unable to deal with the increased load of sIgM in the ER and thus misfolded sIgM will accumulate in the ER, rather than be secreted. As a correlate, other proteins destined for surface display or secretion may be misfolded, and operation of the secretory pathway in its entirety could be compromised (9). This model would further predict that B cells that do not manufacture sIgM should fail to activate XBP-1 if misfolded sIgM is the exclusive driver of the UPR.

In earlier experiments we have produced evidence that XBP-1 deficiency leads to activation of XBP-1 even in B cells that do not synthesize massive quantities of sIgM (10). Here we set out to examine whether the presence or absence of XBP-1, through its impact on ER homeostasis, affects glycoprotein folding in B cells. To that end we generated mice in which XBP-1 is conditionally deleted in B cells by crossing the XBP-1flox/flox mouse line (XBP-1WT) (11) to mice in which expression of Cre recombinase is under the control of the B cell-specific CD19 promoter (12), henceforth referred to as XBP-1KO mice. In addition to the XBP-1KO mice, we used mice unable to synthesize sIgM (XBP-1KO/μS−/−) (13) and thus expressing only the membrane-bound form of IgM (mIgM). Finally, we used transgenic MD4 mice (XBP-1WT/MD4 and XBP-1KO/MD4) in which both the heavy and light chains of IgM specifically recognize hen egg lysozyme (HEL) (14); this mouse model allowed us to investigate the antigen reactivity of IgM produced in the absence of XBP-1, a rigorous test of successful glycoprotein synthesis, folding and assembly.

Lipids are not template-encoded and cannot be manipulated by genetic mutation as readily as proteins, but they are important in physiology. Enforced expression of XBP-1s in NIH3T3 fibroblasts established the role of XBP-1 in controlling synthesis of phosphatidylcholine and phosphatidylethanolamine (15, 16). Although a liver-specific knockout of XBP-1 leads to hypocholesterolemia and hypotriglyceridemia, a lipomics analysis shows that lipid composition in hepatocytes is not affected by XBP-1 deficiency (17). These results suggested that XBP-1 can regulate expression of a tissue-specific lipid profile in individual organs. The changes in lipid composition have not been systematically examined in the context of B cells with a deficiency in XBP-1. A change in lipid synthesis can alter lipid environments (such as lipid rafts), making B cells respond differently to stimulation with antigen.

Here we show that XBP-1-deficient B cells are fully capable of correctly folding both sIgM and mIgM. Our investigation of the biogenesis of a number of essential membrane proteins in XBP-1-deficient B cells revealed no defects in their synthesis, folding, assembly, or intracellular transport. XBP-1-deficient B cells also secrete free κ chains at rates comparable to those seen in XBP-1-proficient cells. Examination of the lipid contents in total membranes of XBP-1-deficient B cells shows decreased levels of a select group of lipids. XBP-1-deficient B cells expand their ER in response to LPS stimulation, but do so to a lesser extent than wild-type B cells. A subtly altered pattern of terminal N-linked glycosylation was observed for IgM as well as class I MHC products. We conclude that the operation of the secretory pathway shows no obvious abnormalities in XBP-1-deficient B cells and that XBP-1 is required in differentiating B cells for processes other than the upregulation of ER folding capacity.

Materials and Methods

Mice

XBP-1flox/flox mice have been described (11). To obtain B cell-specific deletion, XBP-1flox/flox mice were crossed with CD19-Cre mice (12) obtained from Dr. Yang Shi (Harvard Medical School, Boston). Mice with a B cell-specific XBP-1 deletion were additionally crossed to μS−/− mice (13) or to MD4 mice (14), which harbor a transgene for hen egg lysozyme (HEL)-specific IgM.

Antibodies and Reagents

Polyclonal antibodies against Igα, Igβ and PDI were generated in rabbits. Class I MHC was detected using p8 antiserum directed against the cytoplasmic tail, and class II MHC was detected using JV1 antiserum directed against the Class II α chain. Reagents purchased from commercial sources include antibodies to actin (Sigma), p97 (Fitzgerald), XBP-1 (Santa Cruz), μ (SouthernBiotech), and κ (SouthernBiotech). FACS antibodies were purchased from BD Pharmingen and the following clones were used: CD40 (3/23), CD80 (16-10A1), CD1d (1B1). Ligands to Toll-like receptors were procured from commercial sources: Pam3CSK4 from Alexis Biochemicals; poly(I:C) and lipopolysaccharide (E. coli O26:B6) from Sigma; imiquimod from Invivogen; and CpG DNA from TIB-MOLBIOL.

Cell Culture

B lymphocytes were purified from mouse spleens by negative selection using anti-CD43 magnetic beads (Miltenyi Biotech). B cells were cultured in the RPMI 1640 media (Gibco) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM L-glutamine, 100 U/ml penicillin G sodium, 100 μg/ml streptomycin sulfate, 1 mM sodium pyruvate, 0.1 mM non-essential amino acids, and 0.1 mM β-mercaptoethanol (β-ME). For differentiation of B cells into plasmablasts, LPS (20 μg/ml) was added to the culture media. The NKT cell hybridoma 24.7 (18) was a kind gift from Dr. Samuel Behar (Brigham and Women’s Hospital, Boston). IL-2 and IL-6 concentrations were measured by ELISA (BD Pharmingen).

Protein Isolation and Immunoblotting

Cells were lysed in NP-40 lysis buffer (50 mM Tris, pH 7.4, 0.5% NP-40, 5 mM MgCl2, 150 mM NaCl,) or RIPA buffer (10 mM Tris-HCl, pH 7.4; 150 mM NaCl; 1% NP-40; 0.5% sodium deoxycholate; 0.1% SDS; 1 mM EDTA) supplemented with protease inhibitor cocktail (Roche). The protein concentrations of the supernatants were determined by BCA assay (Pierce). Samples were boiled in SDS-PAGE sample buffer (62.5 mM Tris-HCl, pH 6.8; 2% SDS; 10% glycerol; 0.1% bromophenol blue) with β-ME (or N-ethylmaleimide where indicated) and separated by SDS-PAGE. Proteins were transferred to nitrocellulose or polyvinylidene difluoride membranes, blocked in 5% (wt/vol) milk, and immunoblotted with the indicated antibodies and appropriate horseradish peroxidase-conjugated secondary antibodies. Following three washes in PBSTween 20 (0.1%), the blots were developed using Western Lighting Chemiluminescence Reagent (Perkin-Elmer).

Pulse Chase Labeling

Pulse-chase experiments were performed as described (19). Briefly, plasmablasts were starved in methionine- and cysteine-free media, then pulse-labeled with [35S]-methionine/cysteine (Perkin-Elmer). After labeling, cells were incubated in chase medium containing unlabeled methionine (2.5 mM) and cysteine (0.5 mM). At the end of each chase interval, cells were lysed in RIPA buffer containing protease inhibitors. Lysates were then analyzed by immunoprecipitation, SDS-PAGE, and fluorography. Band intensity was determined using a phosphorimager (Fujifilm BAS 2500), and quantitation was done using the MultiGauge software (Fujifilm).

FACS Analysis

Live plasmablasts were stained with the indicated antibodies and analyzed by a FACS Calibur flow cytometer (BD Biosciences). Data were analyzed using CellQuest (BD Biosciences).

Triton X-114 Phase Separation

Triton X-114 phase separation was performed as described (20). Briefly, cells were lysed in Triton X-114 lysis buffer (10 mM Tris-HCl, pH 7.4; 150 mM NaCl; 1% Triton X-114) containing protease inhibitors. Lysates were placed onto a sucrose cushion (6% sucrose; 10 mM Tris-HCl, pH 7.4; 150 mM NaCl; 0.06% Triton X-114), and incubated at 30°C until the lysates turned cloudy. The detergent phase was recovered by centrifugation for 5 min at 300g. The aqueous supernatant was removed and re-extracted with 0.5% Triton X-114, and overlaid onto the original sucrose cushion. After a subsequent round of separation, both the detergent (sediment) and the soluble (supernatant) fractions were brought to the same buffer and detergent concentrations for further analysis.

Biotinylated-HEL Affinity Purification of HEL-specific Immunoglobulins

Cells were metabolically labeled and subjected to Triton X-114 phase separation as described above. Samples were incubated with either anti-μ, biotinylated HEL, or unconjugated biotin. Proteins were then recovered with either Protein G-agarose beads or anti-biotin beads (Sigma), washed, eluted with SDS-PAGE sample buffer, and analyzed by SDS-PAGE. For serial depletions using biotinylated HEL, supernatants from the initial purification were subjected to five sequential rounds of retrieval by biotinylated HEL, and a final round of immunoprecipitation using the anti-μ antibody.

Enzymatic Deglycosylation

Total lysates or immunoprecipitates from lysates were denatured in glycoprotein denaturing buffer (0.5% SDS, 1% β-ME) at 95°C for 5 min, followed by addition of sodium citrate (pH 5.5) to a final concentration of 50 mM, and incubated with Endo H (New England Biolabs) at 37°C for 2 h. Alternatively, sodium phosphate (pH 7.5) and NP-40 were added to the denatured cell lysates to a final concentration of 50 mM and 1%, respectively, and the mixture was incubated with PNGase F (New England Biolabs) at 37°C for 2 h.

Liquid Chromatography-Mass Spectrometry analysis of lipids

Lipid extract were prepared using a modified version of Bligh and Dyer method (21), with two sequential extraction steps to maximize yield. 0.9 ml of a chloroform:methanol (1:2) mix was added to 10 million cells (in 0.1 ml of PBS) and the mixture was vortexed vigorously for 3×1 min with a 5 min interval in between. Next, 0.3 ml of chloroform and 0.3 ml of 1M KCl was added to the tube and the mixture was again vortexed. The mixture was then centrifuged for 2 min at 9,000 rpm to separate the phases. The lower organic layer was transferred to a clean microfuge tube. Residual aqueous phase and cell remnants are re-extracted with 0.5 ml chloroform as described above, and the organic phase combined with extract 1. The combined extract is dried in a Speedvac. Prior to analysis lipids are dissolved in chloroform/methanol (1:1, v/v).

An Agilent high performance liquid chromatography (HPLC) system coupled with an Applied Biosystem Triple Quadrupole/Ion Trap mass spectrometer (4000Qtrap, Foster City, California, USA) was used for quantification of individual polar lipids. Based on product ion and precursor ion analyses of head groups, two comprehensive sets of multiple reaction monitoring (MRM) transitions were set up for quantitative analysis of various lipids including phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI), phosphatidylglycerol (PG), sphingomyelin (SM) and ceramide (Cer) (22, 23). The signal intensity obtained for each lipid species was calculated by comparing to corresponding internal standards including di14:0-PC, di14:0-PE, di14:0-PS, di14:0-PG, di8:0-PI, Cer d18:1/17:0 and SM18/14:0.

Electron Microscopy

B cells were fixed for electron microscopy either immediately after isolation, or after 3 days in culture in the presence of LPS. The cells were fixed in 2.5% glutaraldehyde, 3% paraformaldehyde with 5% sucrose in 0.1 M sodium cacodylate buffer (pH 7.4). Cells were then post-fixed in 1% OsO4 in veronal-acetate buffer. The cells were stained in block overnight with 0.5% uranyl acetate in veronal-acetate buffer (pH6.0), dehydrated and embedded in Spurr’s resin. Sections were cut on a Reichert Ultracut E microtome with a Diatome diamond knife at a thickness setting of 50nm, and stained with 2% uranyl acetate followed by 0.1% lead citrate. Samples were examined using an FEI Tecnai Spirit TEM at 80 KV and imaged with an AMT camera.

Results

XBP-1 activation occurs in the absence of secreted IgM

We set out to test the connection between XBP-1 and glycoprotein quality control during B cell differentiation. B cells were analyzed over the course of a four-day in vitro differentiation scheme in which naïve splenic B cells were cultured in the presence of LPS, to induce differentiation into immunoglobulin-secreting plasmablasts (24).

We assessed the expression of XBP-1s in B cells from μS−/− mice over four days of in vitro LPS-stimulated differentiation. XBP-1s was induced as early as day 1, and its expression persisted for the remainder of the time in culture (Fig. 1A; (10)). This expression pattern is similar to what is observed in wild-type mice (10, 25). XBP-1WT/μS−/− but not XBP-1KO/μS−/− B cells showed upregulation of protein disulfide isomerase (PDI) following LPS stimulation (Fig. 1A), indicating that upregulation of PDI requires both LPS and XBP-1s.

Figure 1.

Figure 1

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(A-B) Naïve B cells from XBP-1WT/μS−/− and XBP-1KO/μS−/− mice were cultured in vitro in the presence of LPS for the indicated times. Cell lysates were prepared and analyzed by SDS-PAGE and immunoblotting for the indicated proteins. Immunoblots shown in both (A) and (B) were performed using the same set of lysates.

Since μS−/− B cells do not express sIgM but are readily induced by LPS to express XBP-1, activation of XBP-1 in these B cells cannot be due to an accumulation of unfolded sIgM. We therefore investigated the expression of membrane μ (μM), kappa (κ), Igα, and class I and II MHC molecules in μS−/− B cells to determine if there was an aberration in the expression of proteins other than IgM that could result in an accumulation of unfolded proteins and activation of XBP-1. Membrane μ was only modestly upregulated over the four-day time course of differentiation in both control and XBP-1-deficient B cells (Fig. 1B). This increase is insignificant when compared to the increase of sIgM upon LPS stimulation in normal B cells; in XBP-1-proficient cells, sIgM levels increase by at least 15 fold (25). In both XBP-1-deficient and -proficient B cells, expression of κ chain increased over the course of LPS stimulation and was increased in XBP-1-deficient B cells (Fig. 1B). However, free κ chains are unlikely to be the trigger for XBP-1 activation, because pulse chase experiments showed that these κ chains are secreted normally (see below). Expression of Igα remained constant over the 4-day course of LPS stimulation, and MHC molecules were slightly upregulated after LPS stimulation (Fig. 1B).

The absence of XBP-1 does not alter the efficiency of IgM folding

The defect attributable to the absence of XBP-1 is expected to yield systemic folding defects in glycoproteins due to dysregulation of chaperone synthesis. Such a defect could manifest itself in several ways: an increase in misfolded proteins, a defect in protein assembly, or a decrease in trafficking from the ER to the targeted locations.

We performed a stringent test for the presence of misfolded IgM, using MD4 transgenic mice that express IgM specific for hen egg lysozyme (HEL). Antigen binding represents a robust test of correct folding and assembly of IgM, since antigen-antibody interaction requires both correct folding and assembly of the heavy and light chains. To isolate correctly folded IgM from MD4 B cells, the antigen HEL was conjugated to biotin, which was then used to recover IgM capable of recognizing HEL by adsorption to anti-biotin agarose beads. We performed these experiments following separation of mIgM and sIgM using the Triton X-114 phase separation method (20). By retrieval of IgM using biotinylated-HEL, we found that the majority of IgM recovered from MD4 B cells is indeed specific for HEL, since immobilized biotinylated HEL led to retrieval of IgM only from MD4 but not from wild-type B cells (Fig. 2A). There was no obvious difference in the percentage of HEL-reactive IgM recovered from XBP-1WT/MD4 and XBP-1KO/MD4 B cells (Figs. 2B and 2C). After 5 rounds of sequential retrieval with immobilized biotinylated-HEL, we recovered the remaining unfolded IgM by immunoprecipitation using an anti-μ antibody. Had the ability of IgM to fold properly been compromised by the XBP-1 deficiency, we should have detected a surplus of free, unbound μ chains in XBP-1-deficient MD4 B cells, but this was not the case. We observed no significant differences in the percentage of misfolded IgM between XBP-1WT/MD4 and XBP-1KO/MD4 B cells, suggesting a similar ability to correctly fold both sIgM and mIgM (Figs. 2B and 2C).

Figure 2.

Figure 2

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(A) Naïve B cells from XBP-1WT, XBP-1WT/MD4 and XBP-1KO/MD4 mice were cultured in the presence of LPS for 4 days and then labeled with [35S]-methionine/cysteine for 4 h. Triton X-114 lysis and separation were performed. Both the pellet and soluble fractions were precipitated with an antibody against μ, or with biotin or biotinylated-HEL. The pellet fraction containing μM is shown. (B-C) Cell lysates were prepared as in panel A. Lysates were split into 2 fractions, one of which was immunoprecipitated with an anti-μ antibody, and the other of which was subjected to five sequential rounds of precipitation with biotinylated-HEL, followed by a subsequent immunoprecipitation using the anti-μ antibody. For each cell type (XBP-1WT/MD4 and XBP-1KO/MD4), the amount of μ recovered from the first fraction by immunoprecipitation using the anti-μ antibody was designated as the total μ. By comparing to the total μ, the percentage of μ recovered from the second fraction was designated for each of the five sequential biotinylated-HEL recovery steps and for the final recovery with the anti-μ antibody. The quantitation of μS (B) was performed on fractions taken from the Triton X-114 supernatant, and the μM quantitation (C) was performed on the Triton X-114 pellet fractions. The results shown are an average of two separate experiments.

Igα/Igβ heterodimer formation proceeds normally in XBP-1-deficient B cells

Igα must assemble with Igβ before the heterodimer can leave the ER. To assess the association of Igα with Igβ, we radiolabeled both naïve and 3-day LPS-stimulated B cells, and performed immunoprecipitations using antibodies against Igα or Igβ, either of which should allow recovery of both Igα and Igβ due to the fact that these proteins remain disulfide-linked under non-reducing conditions. In both naïve and 3-day LPS-stimulated B cells, there was no detectable difference in the association/stoichiometry of Igα and Igβ as a result of XBP-1 deficiency (Fig. 3). The synthesis of Igα and Igβ was apparently not affected by the absence of XBP-1 because we detected similar amounts of Igα and Igβ in XBP-1WT and XBP-1KO B cells (Fig. 3). Since both Igα and Igβ acquire complex-type glycans in the Golgi apparatus, the exit of Igα and Igβ from the ER is normal in the absence of XBP-1 (Fig. 3). Similar results were found in either XBP-1KO or XBP-1KO/μS−/− B cells (Fig. 3).

Figure 3.

Figure 3

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Naïve B cells or 3-day LPS-stimulated plasmablasts from indicated mice were labeled with [35S]-methionine/cysteine for 4 h. Cell lysates were immunoprecipitated using antibodies to Igα (A) and Igβ (B). The immunoprecipitates were treated with either Endo H (H) or PNGase F (F) before analyses by SDS-PAGE and fluorography. CHO, CHO*, and NAG represent high mannose-type glycans, complex-type glycans, and N-acetylglucosamines, respectively. Note that the secreted IgM was precipitated from the 3-day LPS-stimulated XBP-1WT B cell lysates via non-specific binding of secreted IgM to protein G-conjugated agarose beads.

Disulfide formation proceeds normally in the absence of XBP-1

XBP-1-deficient B cells did not upregulate PDI in response to LPS stimulation (Fig. 1A), consistent with the known mechanism of transcriptional control of the PDI gene by XBP-1 (6, 7). Several components of the B cell receptor (BCR) associate through disulfide bonds: Igα and Igβ form a heterodimer held together by a disulfide bond, as do Ig heavy (μ) and light (κ) chains to yield IgM. Lysates from XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells cultured in the presence of LPS for 2 or 4 days were treated with N-ethylmaleimide (NEM) to preserve disulfide bond arrangements and to prevent de novo formation of disulfide bonds. We found no difference in the levels of non-reduced Igα/Igβ heterodimers in XBP-1KO/μS−/− B cells (Fig. 4A), suggesting that the disulfide bond between them formed normally. Similarly, μ and κ chains assembled correctly into higher-order dimers (μ+κ) and tetramers (μ2κ2) (Fig. 4B). Thus, the prevailingly low levels of PDI in XBP-1-deficient B cells are sufficient to sustain normal disulfide bond formation.

Figure 4.

Figure 4

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(A) XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells were cultured in the presence of LPS for 2 or 4 days and lysed in NP40, and lysates were treated with either N-ethylmaleimide (NEM) to retain disulfide bonds or dithiothreitol (DTT) to reduce disulfide bonds. Cell lysates were then analyzed by SDS-PAGE and immunoblotting for Igα. (B) Four-day LPS-stimulated XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells were lysed in NP40, and lysates were treated with either NEM or DTT. Lysates were then immunoblotted using antibodies to μ or κ.

Normal kinetics of synthesis and ER exit are observed for proteins that enter the secretory pathway in XBP-1-deficient B cells

We investigated protein synthesis and trafficking in XBP-1-deficient B cells by pulse chase analyses on IgM, κ chains, class I MHC, Igα/Igβ and class II MHC molecules. We detected no defects in the trafficking of mIgM to the cell surface, as levels of the complex oligosaccharide-bearing μ chain (indicated by μM(+)CHO*) were not altered in XBP-1-deficient B cells after 120 minutes of chase (Figs. 5A and 5B). Although XBP-1-deficient B cells synthesized more κ chains (Fig. 1) than their wild-type counterparts, all of the excess κ chains were effectively secreted (Fig. 5B).

Figure 5.

Figure 5

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(A) Naïve XBP-1WT/MD4 and XBP-1KO/MD4 B cells were cultured in the presence of LPS for 3 days. Cells were then labeled with [35S]-methionine/cysteine for 10 minutes and chased for the indicated times. Immunoprecipitations were performed using antibodies against κ or μ, and were analyzed by SDS-PAGE and fluorography. (B) Three-day LPS-stimulated XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells were radiolabeled for 10 minutes and chased for the indicated times. Immunoprecipitations were performed on both cell lysates and the culture media using an antibody against κ.

We observed a barely detectable delay in the exit of class I MHC from the ER in XBP-1-deficient B cells as compared to XBP-1-proficient B cells, regardless of whether this was measured for XBP-1KO/μS−/− or XBP-1KO/MD4 mice (Figs. 6A and 6B). At later time points, all class I MHC molecules exit the ER. There was no detectable difference in the synthesis or trafficking of Igα and Igβ when comparing XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells (Fig. 6C). We also did not detect any difference in the rate of trafficking of class II MHC molecules as a result of XBP-1 deficiency (Fig. 6D).

Figure 6.

Figure 6

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Naïve B cells from indicated mouse lines were cultured in the presence of LPS for 3 days. Cells were labeled with [35S]-methionine/cysteine for 10 minutes and chased for the indicated times. Immunoprecipitations were performed using antibodies to class I MHC heavy chain (A-B), Igα (C) or class II MHC α chain (D), and were analyzed by SDS-PAGE and fluorography. Additionally in panel B, lysates from the 90- and 120-min chase points were combined, immunoprecipitated with the antibody to class I MHC heavy chain, and subsequently treated with either Endo H (H) or PNGaseF (F) prior to SDS-PAGE analysis.

XBP-1 deficiency does not alter cell-surface display of glycoproteins or compromise their functions

No differences were found in the cell surface levels of CD1d, CD80, or CD40 (Fig. 7, A-C) as measured by cytofluorimetry. Signaling through various Toll-like receptors (TLRs) by distinct TLR ligands was indistinguishable for XBP-1WT/MD4 and XBP-1KO/MD4 B cells, as measured by the secretion of IL-6 in response to stimulation (Fig. 7D). Consistently, B cells were found irresponsive to poly(I:C) (26). The ability of CD1d to present lipid antigens to NKT hybridoma cells was also tested and no apparent defect was detected in XBP-1-deficient B cells (Fig. 7E). Thus all proteins examined function normally in XBP-1-deficient B cells, implying that they do not obviously suffer from a folding or transport defect.

Figure 7.

Figure 7

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(A-C) XBP-1WT/MD4 and XBP-1KO/MD4 B cells that had been cultured with LPS for 3 days were analyzed for the cell surface display of indicated markers by FACS. The black lines are from XBP-1WT/MD4 cells; and the grey lines, from XBP-1KO/MD4 cells. (D) XBP-1WT/MD4 and XBP-1KO/MD4 B cells were cultured with Pam3CSK4 (100 ng/ml), poly(I:C) (100 μg/ml), LPS (20 μg/ml), imiquimod (25 μM), and CpG DNA (1 μM) for 3 days. Culture supernatants were analyzed for the presence of IL-6 by ELISA. (E) XBP-1WT/MD4 and XBP-1KO/MD4 B cells were cultured in the presence of LPS for 4 days. Varying numbers of these LPS-stimulated B cells were then incubated with a CD1d-responsive NKT cell line (NKT 24.7) overnight, and the NKT cell activation was assessed by secretion of IL-2, measured by ELISA.

XBP-1-deficient B cells synthesize significantly less phosphatidylcholine, sphingomyelin, and phosphatidylinositol

We examined changes in lipid composition of total membranes extracted from both naïve and LPS-stimulated XBP-1-deficient B cells using liquid chromatography-mass spectrometry. Data from four independent experiments suggest that the lipid composition in total membranes of the naïve XBP-1-deficient B cells is similar to that of the naïve XBP-1-proficient B cells. However, significantly lower levels of phosphatidylcholine (PC), sphingomyelin (SM), and phosphatidylinositol (PI) were found in the membranes of 4-day LPS-stimulated XBP-1-deficient B cells (Figs. 8 and S1), suggesting that the presence of XBP-1 is required for production of these lipids in LPS-stimulated plasmablasts.

Figure 8.

Figure 8

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XBP-1-deficient plasmablasts synthesize significantly less phosphatidylcholine (PC), sphingomyelin (SM), and phosphatidylinositol (PI). Naïve B cells purified from the spleens of either XBP-1WT or XBP-1KO mice were cultured with LPS for 4 days. Lipids were extracted from at least 100 million cells. Quantitation of each lipid was carried out using liquid chromatography/mass spectrometry. Lipid abundance was calculated by comparing to internal standards. Experiments were repeated for four times using B cells pooled from 9 mice of each genotype. Data are presented as mean±SD. Lipids analyzed include phosphatidylcholine (PC), ceramide (Cer), sphingomyelin (SM), phosphatidylethanolamide (PE), phosphatidylinositol (PI), phosphatidylserine (PS) and phosphatidylglycerol (PG).

XBP-1-deficient B cells show a reduced expansion of the ER

We investigated expansion of the ER in XBP-1-deficient B cells in response to LPS stimulation. Naïve B cells contained relatively little ER, and no differences between the XBP-1-deficient B cells and their wild-type counterparts were detectable by electron microscopy (Fig. 9). In both XBP-1-proficient and XBP-1-deficient B cells, similar percentages of the cell population respond to LPS stimulation by increasing ER content. The expansion was more pronounced in XBP-1-proficient B cells (Fig. 9).

Figure 9.

Figure 9

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Naïve (Day 0) or three-day LPS-stimulated XBP-1WT and XBP-1KO B cells were fixed and prepared for analysis by electron microscopy. Approximately a third of the purified B cells respond to LPS with increased ER content and show the represented morphology. Scale bar = 500 nm.

XBP-1-deficient B cells have subtly altered patterns of terminal glycosylation

We noticed subtle differences in the patterns of glycosylation of IgM and class I MHC when comparing LPS-stimulated XBP-1-proficient and XBP-1-deficient B cells. XBP-1WT/MD4 B cells contained more of the ER form of both mIgM and sIgM than XBP-1KO/MD4 B cells (Fig. 10A). By carefully investigating the glycosylation status of mIgM, we observed a difference in the mobility of the Endo H-resistant fraction of mIgM between XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells (Fig. 10B), and an increased heterogeneity in the acquisition of Endo H-resistant complex-type glycans in XBP-1KO/μS−/− B cells (Fig. 10B, indicated by <). Because each μ-chain has five potential N-linked glycosylation sites, heterogeneity can arise from any of these sites due to differences in terminal glycosylation. Similar defects in glycosylation also occurred to class I MHC molecules (Figs. 6A and 6B). The XBP-1-proficient B cells synthesized a population of class I MHC with Endo H-resistant glycan modifications that migrated more slowly in SDS-PAGE, which was absent from XBP-1-deficient B cells (Figs. 6A and 6B).

Figure 10.

Figure 10

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(A) Four-day LPS-stimulated XBP-1WT/MD4 and XBP-1KO/MD4 B cells were lysed in Triton X-114, and secreted and membrane IgM were separated as described. Lysates were then immunoblotted using the anti-μ antibody. (B) Naïve and 3-day LPS-stimulated XBP-1WT/μS−/− and XBP-1KO/μS−/− B cells were labeled with [35S]-methionine/cysteine for 4 h. Immunoprecipitations were performed using an antibody to μ. Immunoprecipitates were subsequently treated with either Endo H (H) or PNGaseF (F), and analyzed by SDS-PAGE and fluorography. The arrowhead (<) indicates the partially Endo H-resistant IgM.

Discussion

The unfolded protein response (UPR) is commonly viewed as a stereotypic set of changes in gene expression, triggered by the accumulation of misfolded proteins. Its regulation involves the activation of IRE-1, which executes a splicing reaction to yield XBP-1s mRNA, the translation of which starts a transcriptional program that targets a variety of genes encoding chaperones and enzymes involved in protein folding and lipid synthesis. The literature frequently equates the activation of XBP-1 with the activation of the UPR, a response commonly evoked experimentally through the application of toxic drugs such as tunicamycin, thapsigargin or dithiothreitol. The extent to which the UPR functions in a more physiological manner is not immediately obvious, and none of the above treatments used to induce the UPR can be considered subtle. Tunicamycin treatment interferes with glycoprotein folding through inhibition of glycosylation, but the extent of folding damage inflicted is difficult to gauge. At the same time, tunicamycin treatment also leads to accumulation of the dolichol precursor and presumably affects the lipid environment in which ER membrane proteins function. Thapsigargin depletes the ER of its calcium stores, compromises the function of calcium-dependent ER-resident chaperones such as calnexin and calreticulin, and is therefore expected to increase the failure rate of protein folding in the ER. Imposition of stress by generating a strongly reducing environment using dithiothreitol (DTT) is also a common method to unleash the UPR. All of these triggers activate XBP-1 (3). More rarely the UPR can be induced by overexpression of genetically-engineered misfolded proteins (27, 28), but it is interesting to note that these two examples concern a polytopic (multi-spanning) ER membrane protein and a surfactant-binding protein, respectively, both of which might act not only through their misfolded state per se, but also through sequestration of lipids and their metabolites at inappropriate locations. Experiments in yeast suggest that massive quantities of a misfolded type I membrane protein are far less effective at induction of the UPR than even mild tunicamycin treatment (29).

Development of B cells into plasma cells is an example of a physiological process that triggers the UPR, as defined by activation of XBP-1. Although misfolded sIgM has been speculated to provide the impetus for XBP-1 activation in differentiating plasma cells (3, 4), our data provide no support for this suggestion, because XBP-1 activation occurs also in B cells that cannot make sIgM, with kinetics that mimic those in normal B cells (Fig. 1; (10)). Since an excess of unfolded sIgM cannot be the cause of XBP-1 activation in these cells, we conclude that there must be an as yet unidentified (differentiation-dependent) trigger that leads to XBP-1 activation through IRE-1. IRE-1 is usually activated by the presence of unfolded proteins in the lumen of the ER, either through a direct interaction with unfolded proteins (30), or through its dissociation from BiP as a result of competition for unfolded proteins (31, 32). Unfolded ER proteins alone may not be sufficient for full IRE-1 activation. Protein-tyrosine phosphatase 1B (PTP-1B) has been implicated in assisting complete function of IRE-1, since cells that lack PTP-1B are less efficient in turning on downstream effectors of IRE-1 in response to UPR triggers (33), thus linking PTP-1B to IRE-1. Full activation of the IRE-1 pathway can thus be controlled by multiple inputs, and not merely by the presence of unfolded proteins.

LPS-induced differentiation activates XBP-1 (4, 34), leading to upregulation of chaperones like PDI (Fig. 1A). Assuming that the failure to increase PDI levels in XBP-1-deficient plasmablasts would cause misassembly and/or misfolding of proteins, we investigated the fate of IgM heavy and light chains, as well as the ability of IgM to bind hen egg lysozyme (HEL) in the absence of XBP-1 (Fig. 2). We saw no difference between XBP-1WT/MD4 and XBP-1KO/MD4 plasmablasts in terms of their ability to correctly assemble sIgM and mIgM capable of HEL binding. Free κ chains were synthesized at increased levels and were readily secreted by XBP-1-deficient B cells (Figs. 1B and 5). XBP-1 deficiency also did not affect synthesis, assembly and trafficking of Igα and Igβ, or class I and class II MHC molecules (Figs. 3, 4A, and 6), with the possible exception of a barely perceptible delay in the exit of class I MHC molecules from the ER in XBP-1-deficient B cells (Figs. 6A and 6B). XBP-1-deficient B cells obtained from chimeric XBP-1−/−/RAG2−/− mice showed no obvious delay in class I MHC trafficking, a difference we attribute to the slight difference in the pulse-chase protocols used (25). We hence suggest that glycoproteins fold and function normally in XBP-1-deficient B cells. This conclusion is inconsistent with the proposal that an XBP-1-deficient B cell produces an ER that is defective in protein folding.

CD40, CD80 and CD1d were all present at the cell surface of XBP-1-deficient B cells in normal amounts (Fig. 7, A-C), and CD1d was fully functional in the presentation of lipid antigens to NKT cells (Fig. 7E). Production of IL-6 triggered by various Toll-like receptor (TLR) ligands was not significantly different for XBP-1-proficient and XBP-1-deficient plasmablasts (Fig. 7D), suggesting that the examined TLRs function properly in their distinct cellular compartments. Taken together, these results show that the ER of XBP-1-deficient plasmablasts sustains folding and assembly of all proteins we have examined thus far. The activation of stress responses triggered by prion replication was not influenced by XBP-1 deficiency (11), further supporting the notion that XBP-1 is not required to handle misfolded proteins, in the form of cytotoxic aggregates.

It is possible that we do not detect a defect in protein folding in XBP-1-deficient B cells due to a compensatory upregulation of other branches of the UPR. In addition to the responses initiated by the IRE-1/XBP-1 axis, PERK and ATF6 contribute as well. The presence of unfolded proteins in the ER stimulates phosphorylation by PERK of eukaryotic initiation factor 2α. This attenuates translation initiation, thus reducing the number of proteins that enter the ER (31). However, there is no indication that initiation of translation is somehow compromised through engagement of the PERK axis in XBP-1-deficient B cells. ATF6 is a membrane-bound transcription factor. In response to ER stress, ATF6 traffics to the Golgi apparatus where its transmembrane segment is cleaved to release its transcription factor domain, which then translocates to the nucleus to initiate the transcription of ER chaperones that can aid in protein folding (35, 36). The PERK branch of the UPR is not required for B cell differentiation (37) and XBP-1 deficiency does not lead to the upregulation of eIF2α and phospho-eIF2α, important indictors for PERK activation ((10) and data not shown). The ATF6 pathway is induced during plasma cell differentiation (38) but whether it is essential for plasma cell differentiation remains to be established. Finally, XBP-1 deficiency alone is sufficient to cause a block in plasma cell differentiation, with no obvious compensation by either PERK or ATF6 activation.

The exact role of XBP-1 in plasma cell differentiation remains to be defined. Phosphatidylcholine, sphingomyelin and phosphatidylinositol decrease in response to XBP-1 deficiency in total membranes of LPS-stimulated plasmablasts cells but not unstimulated naïve B cells (Fig. 8), suggesting that XBP-1 is required for lipid synthesis in support of plasma cell differentiation. Phosphatidylcholine is most drastically affected by XBP-1 deficiency in plasmablasts because it is the primary phospholipid of the ER membranes. Although no lipid was found altered in hepatocytes lacking XBP-1 (17), phosphatidylcholine and phosphatidylethanolamine increase significantly in NIH3T3 fibroblasts overexpressing XBP-1s (15, 16). Unlike in fibroblasts, we do not find phosphatidylethanolamine affected by XBP-1 deficiency in B cells; however, sphingomyelin and phosphatidylinositol decrease significantly in XBP-1-deficient plasmablasts although the levels of these two lipids are lower than that of phosphatidylcholine. To find reduced levels of sphingomyelin and phosphatidylinositol in plasmablasts is interesting, because both lipids play important roles in signal transduction. Sphingomyelin is a crucial lipid in rafts and phosphatidylinositol is an essential intermediate in the phosphatidylinositol-3-phosphate (PI3P) signaling pathway. These lipid defects may compromise the recruitment of IgM, Igα/Igβ and other B cell coreceptors (CD19, CD20, CD21, and CD81) into lipid rafts, and contribute to the observed defective signal transduction in XBP-deficient B cells (10).

Activated XBP-1-deficient plasmablasts had decreased ER content compared to XBP-1-proficient plasmablasts, as visualized by transmission electron microscopy (Fig. 9), whereas unstimulated controls showed no such difference. While XBP-1 is not required for B cells to increase their ER content in response to differentiation, the extent of the ER expansion does depend on XBP-1. This is consistent with XBP-1’s role in setting the levels of phosphatidylcholine (Figs. 8 and S1A). Thus it appears that expansion of the ER, but not necessarily the ER folding capacity per se, is under the control of XBP-1.

We observed minor differences in the pattern of terminal glycosylation of both IgM and class I MHC in XBP-1-deficient cells (Figs. 6A, 6B and 10). Though these changes are slight and require a more detailed analysis by glycan sequencing, such differences could certainly contribute to the defect in plasma cell differentiation in XBP-1-deficient mice. Whether alterations in glycosyltransferase levels, nucleotide sugars and their transporters or the environment in which these enzymes function are responsible for the observed glycosylation defects is not known at present. Proper glycosylation is likely critical to B cell differentiation, given that the blockade of high mannose to complex type N-linked glycans imposed by the ER mannosidase inhibitor 1-deoxymannojirimycin can completely block the formation of immunoglobulin-secreting B cell blasts from human peripheral blood B cells (39), although mannosidase inhibition does not inhibit B cell proliferation or immunoglobulin secretion per se. Some factors required for B cell differentiation may well be sensitive to changes in terminal glycan modifications.

XBP-1WT/MD4 and XBP-1KO/MD4 B cells differ in signaling through the B cell receptor (BCR), and regulation of the transcription factors IRF4 and Blimp-1 (10). In addition, XBP-1-deficient B cells after antigen stimulation fail to migrate to the bone marrow (10). These, together with altered lipid synthesis and glycosylation in XBP-1-deficient B cells, could account for failure of B cells to fully differentiate into plasma cells, but accumulation of misfolded proteins does not appear to be a key contributing factor. Altogether, our results suggest that XBP-1 activation in B cells is a differentiation-dependent event unlinked to accumulation of misfolded IgM. Furthermore, protein folding occurs normally in XBP-1-deficient B cells, indicating an essential role for XBP-1 beyond the UPR.

Supplementary Material

01

Supplemental Figure Legend Figure S1. The levels of the different classes of lipids presented in Fig. 8 are the summation of specific lipid species analyzed by mass spectrometry. The abundance of the specific lipids analyzed is presented here according to class: (A) phosphatidylcholine, (B) ceramide, (C) sphingomyelin, (D) phosphatidylethanolamide, (E) phosphatidylinositol, (F) phosphatidylserine and (G) phosphatidylglycerol. Data were acquired as described for Fig. 8 and are presented as mean±SD.

Acknowledgements

We thank J. Antos and M. Brinkmann for critical reading of the manuscript, and members in the Ploegh Lab for their support. These studies were supported by grants from the National Institutes of Health (to H.L.P.). A.M.M. was supported by a National Science Foundation (NSF) Graduate Research Fellowship. S.K.D. is supported by a CRI Fellowship. E.J.K was supported by a NSF East Asia and Pacific Summer Institutes fellowship grant and the Singapore National Research Foundation. National Research Foundation under CRP Award No. 2007-04, the Academic Research Fund (R-183-000-160-112), the Biomedical Research Council of Singapore (R-183-000-211-305) and the National Medical Research Council (R-183-000-224-213) to M.R.W are gratefully acknowledged.

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Supplementary Materials

01

Supplemental Figure Legend Figure S1. The levels of the different classes of lipids presented in Fig. 8 are the summation of specific lipid species analyzed by mass spectrometry. The abundance of the specific lipids analyzed is presented here according to class: (A) phosphatidylcholine, (B) ceramide, (C) sphingomyelin, (D) phosphatidylethanolamide, (E) phosphatidylinositol, (F) phosphatidylserine and (G) phosphatidylglycerol. Data were acquired as described for Fig. 8 and are presented as mean±SD.

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