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Galectin 1 Modulates Plasma Cell Homeostasis and Regulates the Humoral Immune Response

Adrienne Anginot, Marion Espeli, Lionel Chasson, Stéphane J. C. Mancini and Claudine Schiff
J Immunol June 1, 2013, 190 (11) 5526-5533; DOI: https://doi.org/10.4049/jimmunol.1201885
Adrienne Anginot
*Centre d'Immunologie de Marseille-Luminy, Faculté des Sciences de Luminy, Aix Marseille University, UM2, Marseille F-13288, France;
†INSERM, U1104, Marseille F-13288, France;
‡Centre National de la Recherche Scientifique, UMR7280, Marseille F-13288, France; and
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Marion Espeli
§Department of Medicine, Cambridge Institute for Medical Research, University of Cambridge, United Kingdom
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Lionel Chasson
*Centre d'Immunologie de Marseille-Luminy, Faculté des Sciences de Luminy, Aix Marseille University, UM2, Marseille F-13288, France;
†INSERM, U1104, Marseille F-13288, France;
‡Centre National de la Recherche Scientifique, UMR7280, Marseille F-13288, France; and
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Stéphane J. C. Mancini
*Centre d'Immunologie de Marseille-Luminy, Faculté des Sciences de Luminy, Aix Marseille University, UM2, Marseille F-13288, France;
†INSERM, U1104, Marseille F-13288, France;
‡Centre National de la Recherche Scientifique, UMR7280, Marseille F-13288, France; and
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Claudine Schiff
*Centre d'Immunologie de Marseille-Luminy, Faculté des Sciences de Luminy, Aix Marseille University, UM2, Marseille F-13288, France;
†INSERM, U1104, Marseille F-13288, France;
‡Centre National de la Recherche Scientifique, UMR7280, Marseille F-13288, France; and
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Abstract

Galectin-1 (GAL1) is an S-type lectin with multiple functions, including the control of B cell homeostasis. GAL1 expression was reported to be under the control of the plasma cell master regulator BLIMP-1. GAL1 was detected at the protein level in LPS-stimulated B cells and was shown to promote Ig secretion in vitro. However, the pattern of GAL1 expression and function of GAL1 in B cells in vivo are still unclear. In this study, we show that, among B cells, GAL1 is only expressed by differentiating plasma cells following T-dependent or T-independent immunization. Using GAL1-deficient mice we demonstrate that GAL1 expression is required for the maintenance of Ag-specific Ig titers and Ab-secreting cell numbers. Using an in vitro differentiation assay we find that GAL1-deficient plasmablasts can develop normally but die rapidly, through caspase 8 activation, under serum starvation–induced death conditions. TUNEL assays show that in vivo–generated GAL1-deficient plasma cells exhibit an increased sensitivity to apoptosis. Taken together, our data indicate that endogenous GAL1 supports plasma cell survival and participates in the regulation of the humoral immune response.

Introduction

Galectin-1 (GAL1) is a small lectin belonging to a well-conserved family consisting of 15 members sharing a high affinity for β-galactosides (1). GAL1 consists of a single carbohydrate recognition domain with a short NH2 sequence and is active as a noncovalently bonded homodimer (2). GAL1 can be found in several cellular compartments, including the cytoplasm, where it can play multiple roles, and in the nucleus, where it acts as a splicing factor (3, 4). Although galectins do not have the signal sequence required for protein secretion through the usual secretory pathway (5), some galectins, such as GAL1, can be secreted. Although the intracellular functions of GAL1 are generally independent of carbohydrate binding, its extracellular activity mostly requires its lectin activity (6).

GAL1 was shown to be involved in the regulation of immune functions. It acts as a homeostatic agent by modulating innate and adaptive immune responses. Notably, GAL1 was shown to control T cell homeostasis, as well as cancer progression (6–8). Under physiological conditions, extracellular GAL1 promotes apoptosis of activated, but not resting, immune cells (9, 10), with the notable exception of resting T cells, which are sensitized to CD95/Fas-mediated cell death by GAL1 (11). GAL1 also induces phosphatidyl-serine externalization without associated apoptosis (12, 13). Moreover, galectins can cross-link glycosylated proteins, leading to signal transduction and direct cell death or activation of other signals regulating cell fate (14). Finally, GAL1 secretion was shown to induce the death of antitumor T cells and, thus, might contribute to immune escape by tumors (6, 15).

Although the roles of GAL1 in the control of immune responses, inflammation, and tumor progression have been well characterized, very little is known about its expression and function in B lymphocytes. We demonstrated that GAL1 is important for early B cell differentiation in the bone marrow (BM) (16). In this case, GAL1 secreted by BM stromal cells supports B cell differentiation through pre-BCR activation and signaling (16, 17). Further, using an in vitro model of LPS-activated B cells, Tsai et al. (18) reported that splenic plasmablasts (PBs) expressed GAL1 in a Blimp-1–dependent manner. They also showed that ectopic expression of GAL1 in mature B cells increased Ig transcripts and secretion.

To further study the role of GAL1 in vivo, we assessed GAL1 expression and the immune responses of wild type (WT) and GAL1-knockout mice (Lgals1−/−). We showed that, among B cells, GAL1 is specifically expressed by plasma cells (PCs) and more precisely by Blimp1-GFPlow plasma cells that are not fully differentiated. Moreover, Lgals1−/− mice can initiate a normal immune response but fail to maintain serum Ig levels and Ab-secreting cell (ASC) numbers. Finally, we report that GAL1-deficient PBs are more susceptible to apoptosis than are their normal counterparts.

These findings provide new insight into how GAL1 modulates the immune response and emphasize the regulatory role of GAL1 in finely tuning biological processes.

Materials and Methods

Mice

GAL1-deficient mice (19), backcrossed for six generations onto a 129SV background, and 129SV control mice were housed under specific pathogen–free conditions and handled in accordance with European directives, with the approval of the Institutional Review Board of INSERM/Centre National de la Recherche Scientifique. Mice were immunized with two i.p. injections (separated by 2 wk) of 100 μg 4-hydroxy-3-nitrophenylacetyl–keyhole limpet hemocyanin [NP(30)-KLH] (Biosearch Technologies) with Alum 1:1 (v/v) (Pierce) for T-dependent responses and with one i.p. injection of 1 μg NP(40)-Ficoll (Biosearch Technologies) in PBS for T-independent responses. Blood samples were harvested, and Ab responses were analyzed by ELISA at the indicated time points after immunizations.

C57BL/6 Blimpgfp mice were bred and maintained under specific pathogen-free conditions. They were immunized with 100 μg NP(30)-KLH in Alum i.p. and analyzed 2 wk later. All experiments were performed according to the regulations of the U.K. Home Office Scientific Procedures Act (1986).

Flow cytometry

Single-cell suspensions were stained using standard protocols for flow cytometry and the Abs listed in Table I. Intracellular staining was performed after fixation and cell permeabilization using the Cytofix/Cytoperm kit (BD Biosciences, Pont de Claix, France). Alternatively, cells were fixed with 4% paraformaldehyde for 10 min and permeabilized with PBS/0.2% saponin to allow codetection of GFP and intracellular GAL1. FACS analysis was performed on a FACSCanto II, LSR II (BD Biosciences), or CyAn (DAKO) apparatus. Data were analyzed with FlowJo (TreeStar) or DiVa (BD Biosciences) software.

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Table I. Abs used for flow cytometry

In vitro PC differentiation

Splenic B cells from WT and Lgals1−/− mice were isolated using B220 MicroBeads and the autoMACS Separator (Miltenyi Biotec, Paris, France). Cells (1 × 106) were cultivated in IMDM, 10% FCS, 100 U/ml penicillin, 100 μg/ml streptomycin, and 50 μM 2-ME. When specified, cultures were also performed in 5% FCS. Cells were stimulated with 1 μg/ml LPS for 4 d (Sigma). PC differentiation was also tested using 1 μg/ml NP-Ficoll (Biosearch Technologies) or 5 μg anti-IgM (Sigma Aldrich) and 1 ng/ml CD40L (ABCYS) and 25 ng/ml IL-4 (R&D Systems).

ELISA

Serum of immunized or control mice was obtained after blood coagulation and kept at −20°C. Ninety-six–well plates (Nunc) were coated with 5 μg/ml NP(23)-BSA to detect low- and high-affinity Abs or NP(3)-BSA (Biosearch Technologies) to detect only high-affinity Abs overnight at 4°C. Ig concentrations in culture supernatants were quantified using the Clonotyping System-AP (SouthernBiotech). In brief, plates were coated with 5 μg/ml anti-Ig (H+L) overnight at 4°C and then saturated with PBS-BSA 2%. Standard mouse Ig (SouthernBiotech) or samples were then added to the wells at different dilutions. After extensive washing in PBS and 0.05% Tween 20 (PBST), anti-mouse Ig subclasses coupled with alkaline phosphatase (SouthernBiotech) were added to the wells and revealed by alkaline phosphatase substrate (Sigma). NaOH 3 M was used to quench color development, and OD was measured at 405 nm and referred to plate background at 620 nm. Ig concentrations were calculated by comparison with each Ig subclass’s standard curves.

ELISPOT

MultiScreen HTS-IP filter plates (Millipore) were coated with 5 μg/ml NP(23)-BSA to detect low- and high-affinity Abs or with 5 μg/ml NP(3)-BSA (Biosearch Technologies) to detect only high-affinity Abs overnight at 4°C. Alternatively, plates were coated with 5 μg/ml goat anti-mouse IgG(H+L) for detection of in vitro–generated ASCs. Plates were then washed with double-distilled water and saturated with MEMα medium supplemented with 10% FCS. The number of viable cells was evaluated by trypan blue exclusion, and the indicated number of live cells (105 splenic and 106 BM cells, and 103–104 cells from in vitro cultures) was plated carefully in each well. The number of ASCs was analyzed after 16 h at 37°C. Cells were harvested, and plates were washed three times in PBS, three times in PBST, and twice in double-distilled water. Goat anti-mouse Ig subclasses Abs labeled to alkaline phosphatase were added to the wells for 1 h (SBA-Clonotyping System-AP; SouthernBiotech). After extensive washing in PBST, alkaline phosphatase conjugate substrate (Bio-Rad kit) was added and incubated at 37°C for 30 min. Reaction was stopped by PBS and then water and plates were air-dried. Plates were read using an AID ELISPOT reader, according to the manufacturer’s instructions.

Immunohistofluorescence

Spleens were snap-frozen in Tissue-Tek OCT compound (Sakura Finetek). Frozen sections (7 μm) were fixed with cold acetone for 2 min. Unspecific binding site blockade was performed with 2% BSA and 1% donkey serum in PBS for 1 h, and permeabilization was achieved by incubation with 0.1% Triton X-100 and 0.05% Tween 20 in PBS at room temperature for 1 h. Femurs were fixed in formalin for 48 h and then decalcified in PBS/10% EDTA for 20 d. Following decalcification, bones were dehydrated in PBS/30% sucrose for 48 h, embedded in OCT (Microm Microtech, Francheville, France), and snap-frozen in isopentane on dry ice before 20-μm sections were cut using a cryostat. Sections were fixed and permeabilized in saturation buffer (2% BSA, 1% horse serum, 0.05% Tween 20, 0.1% Triton X-100 in PBS) for 1 h at room temperature. Sections were labeled overnight at 4°C with an anti-GAL1 Ab (clone 201002; R&D Systems), incubated for 1 h at room temperature with an anti-rat biotinylated Ab, and revealed with streptavidin coupled to Alexa Fluor 488 or Alexa Fluor 647 (Invitrogen, Carlsbad, CA). Sections were also labeled using anti-mouse IgM coupled to PE, anti-mouse IgG coupled to FITC, or Alexa Fluor 568 and NP-PE for 1 h at room temperature. TO-PRO-3 (Molecular Probes, Invitrogen) was used to stain nuclei. Confocal microscopy was performed with a Zeiss LSM510 microscope, and images were processed with Zeiss LSM software.

RNA extraction and real-time PCR

RNA was extracted from total cells in culture with the RNeasy mini kit (QIAGEN). One microgram of RNA was converted into cDNA using Superscript II reverse transcriptase (Invitrogen). Real-time PCR was performed with 100 ng cDNA using SYBR Green PCR Master Mix (Applied Biosystems). Each sample was run in triplicate in a PCR System (ABI PRISM 7500; Applied Biosystems). Primers were designed on two different exons for each gene of interest, and the basic local alignment search tool from the National Center for Biotechnology Information (Bethesda, MD) was used to confirm amplicon specificity. At the end of the PCR, specificity of the amplification was controlled by generating a melting curve of the PCR product. Expression values were normalized using the HPRT housekeeping gene, run on the same plate. The results were analyzed according to the ΔΔCt comparative method, normalizing to 1 for WT expression at day 0.

The following primers were used: HPRT forward: 5′-GGCCCTCTGTGTGCTCAAG-3′ and reverse: 5′-CTGATAAAATCTACAGTCATAGGAATGGA-3′ and GAL1 forward: 5′-CCTGGTCCATCTTCACTTCCAT-3′ and reverse 5′-CTTTGGCCTGGAAAGCACAA-3′.

Apoptosis assays

Membrane permeabilization was evaluated using 7-aminoactinomycin D (7-AAD) (Molecular Probes, Invitrogen). Mitochondrial potential and phosphatidyl serine externalization were evaluated using a Mitochondrial Membrane Potential Apoptosis Kit with MitoTracker Red and Annexin V, Alexa Fluor 488 (Molecular Probes, Invitrogen), according to the manufacturer’s instructions. Caspase 8 activation was quantified with a CaspGLOW Fluorescein Active Caspase-8 Staining kit (Biovision, CliniSciences), according to the manufacturer’s instructions.

TUNEL assays

TUNEL assays were performed using the DeadEnd Fluorometric TUNEL System (Promega), according to the manufacturer’s instructions. Briefly, frozen sections were fixed with 4% paraformaldehyde and rinsed twice in PBS. Samples were then permeabilized for 5 min with 0.2% Triton X-100 and washed with PBS. The slides were covered with equilibration buffer for 5 min and incubated with the reaction buffer (45 μl equilibration buffer + 5 μl nucleotide mix + 1 μl rTDT enzyme) for 1 h at 37°C. Reactions were stopped by immersing the slides in 2× saline sodium citrate buffer for 15 min at room temperature. The number of TUNEL+ PCs/field was enumerated with Volocity 3D Image analysis software (Perkin Elmer).

Proliferation assays

Splenic B cells from WT and Lgals1−/− mice were isolated using B220 MicroBeads and the autoMACS Separator (Miltenyi Biotec). Cells (1.5 × 105) were plated in complete IMDM in the presence of 5 or 10% FCS in 96-well plates and then stimulated with 1 μg/ml LPS. Two and three days after plating, 1 μl [3H]thymidine was added to each well for 12 h. Plates were directly frozen and analyzed using a beta counter (Wallac Trilux).

Statistical analysis

Data were evaluated using the Mann–Whitney unpaired test with a risk of 5%. The p value < 0.05 were considered statistically significant. Error bars represent SEM.

Results

GAL1 is expressed by PBs

We first analyzed GAL1 expression in splenic B cell subpopulations in nonimmunized and immunized mice by flow cytometry (see Abs in Table I). At steady state in nonimmunized mice, we observed that follicular (B220+CD21+CD23high), marginal zone (B220+CD21highCD23low), and early transitional (B220+CD21−CD23−) B cells did not express detectable levels of GAL1 (Fig. 1A). After T-dependent immunization with NP-KLH, follicular, marginal zone, transitional, and germinal center (B220+Gl7+) B cells were negative for GAL1 expression (Fig. 1B, data not shown). In contrast, B220lowCD138+ PCs were GAL1+ (Fig. 1B), and GAL1 expression was also detected in B220lowCD138+ PCs after T-independent responses against NP-Ficoll (Supplemental Fig. 1). Among splenic B cell subpopulations, only CD138+ PCs express GAL1; this expression is detected after primary and secondary immune responses and at least until day 40 postimmunization (Supplemental Fig. 2).

FIGURE 1.
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FIGURE 1.

GAL1 is expressed by PBs. Spleen cells from nonimmunized mice (A) and mice immunized with a T-dependent Ag (NP-KLH) at day 21 (B) were stained for cell surface expression of B220, CD21, CD23, GL7, and CD138, as well as for intracytoplasmic expression of GAL1 (Table I), and analyzed by flow cytometry. B220+CD21+CD23high follicular (FO), B220+CD21highCD23low marginal zone (MZ), B220+CD21−CD23− early transitional and B220+GL7+ germinal center (GC) B cells, and B220lowCD138+ PCs are indicated. Cells from Lgals1−/− mice were similarly stained and used as negative controls for GAL1 expression. Graphs for Lgals1−/− and WT mice are shown in gray and white, respectively. Data are representative of more than five independent experiments with at least four mice/group. (C) Immunohistofluorescence was performed on spleen sections of mice immunized with a T-independent Ag (NP-Ficoll) 7 d after injection. Anti-GAL1, anti-IgM, anti IgG, and NP-PE staining were also performed. A representative extrafollicular zone (low nuclei density) of the spleen is shown. Images are representative of more than five different fields from five immunized mice. GAL1 is in green; IgM, IgG, and NP are in red; and nuclei are in blue. (D) Immunohistofluorescence was performed on BM sections of mice immunized with a T-dependent Ag (NP-KLH) 8 d after the secondary injection. Anti-GAL1 and anti-IgG staining were performed, and a representative field of the BM is shown. GAL1 is in green, and IgG is in red. Asterisks show double-stained cells. The image is representative of three fields from two immunized mice. (E) Spleen (upper left panels) and BM (upper right panels) cells from NP-KLH–immunized Blimp1gfp mice were stained at day 15 with anti-GAL1, anti-CD138, and anti-B220. MFI of GAL1 staining for B cells (B220+), PBs (CD138+GFPlow), and fully differentiated PCs (CD138+GFPhigh) in the spleen (left lower panel) and BM (lower right panel). Data are representative of two independent experiments with four mice/group. *p < 0.05.

To visualize GAL1-expressing cells in situ, immunohistofluorescence was performed on spleen sections from immunized mice. As shown in Fig. 1C, after a T-independent response, a fraction of IgM+, IgG1+, and NP-specific cells expresses GAL1. These cells are localized in the extrafollicular regions of the spleen, similar to conventional PCs. We also observed expression of GAL1 by some, but not all, PCs in the BM 7 d after secondary immunization with NP-KLH (Fig. 1D).

We used Blimp1gfp reporter mice to assess whether GAL1 was expressed at all stages of PC differentiation (20). These mice express the GFP reporter under the control of the Blimp1 promoter and allow the discrimination of CD138+GFPlow PBs from CD138+GFPhigh fully differentiated PCs. Fourteen days after NP-KLH T-dependent immunization, we observed that CD138+GFPlow PBs in the spleen and BM expressed higher levels of GAL1 compared with B220+ cells (Fig. 1E). Interestingly, the mean fluorescence intensity (MFI) values for GAL1 expression were reduced on splenic and BM CD138+GFPhigh PCs, suggesting that GAL1 is transiently expressed by PBs before being downregulated in fully differentiated PCs.

Serum Ig levels are decreased in GAL1-deficient mice

We then tested the impact of GAL1 expression on humoral immune responses. Following T-independent immunization, we observed a significant reduction in the serum titer of NP-specific IgG3 at days 8 and 15 in Lgals1−/− mice compared with WT mice (Fig. 2A). Although not statistically significant, we also observed a trend toward a lower NP-specific IgM titer in the GAL1-deficient mice compared with WT mice. After T-dependent immunization, the kinetic of NP-specific IgM and IgG1 formation in Lgals1−/− mice parallels that of WT mice (Fig. 2B, 2C). However, in the absence of GAL1, IgM titers at days 15 and 21 (Fig. 2B) and low- and high-affinity IgG1 titers at days 21 and 40 (Fig. 2C) were significantly reduced. The NP3 (high)/NP23 (low) IgG1 affinity ratio was the same for WT and Lgals1−/− immunized mice, except at day 21, where it was slightly increased in Lgals1−/− mice (Fig. 2D). This increased ratio at day 21 could be explained by the fact that the low-affinity Ab titer would be affected before that of high-affinity Abs. Thus, GAL1 does not seem to play a role in the initiation of the immune response, but it seems to be important for the maintenance of Ag-specific Ig titers after boost, without affecting Ig affinity maturation.

FIGURE 2.
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FIGURE 2.

Serum Ig levels are decreased in immunized Lgals1−/− mice. WT and Lgals1−/− mice were immunized with NP-Ficoll (T-independent Ag) (n = 7 mice/group) (A) and NP-KLH in Alum (T-dependent Ag) (n = 5 mice/group) (B, C), and Ig secretion in serum was quantified by ELISA. (A) NP-specific IgM and IgG3 levels were determined at days 2, 5, 8, and 15 for the T-independent responses. NP-specific IgM (B) and low (NP23) and high (NP3) NP-specific IgG1 serum titers (C) were analyzed at days 2, 5, 15, 18, 21, and 40 for T-dependent responses. (D) Affinity maturation of T-dependent immune responses at days 18, 21, and 40 was measured by the NP3 (high)/NP23 (low) affinity IgG1 ratio. Open and black squares correspond to WT and Lgals1−/− mice, respectively. *p < 0.05 versus WT, **p < 0.005 versus WT.

Altogether, our results indicate that Lgals1−/− mice normally initiate immune responses after T-independent and T-dependent immunizations but failed to maintain high titers of NP-specific Igs at later time points.

Ag-specific PC numbers are altered in GAL1-deficient mice

To determine whether the reduction in NP-specific Ig titers in Lgals1−/− mice was due to a defect in PB/PC formation, we used ELISPOT to monitor the number of NP-specific ASCs generated following T-dependent immunization. Total splenocytes and BM cells from WT and Lgals1−/− mice were plated and cultured for 16 h on NP-coated ELISPOT plates, and the number of IgM, low-, and high-affinity IgG1 ASCs was determined. As shown in Fig. 3A, the numbers of IgM and IgG1 NP-specific ASCs generated in the spleens of Lgals1−/− mice were initially normal, but they decreased significantly at day 40 compared with WT mice. At day 21, low-affinity IgG1 ASCs also were decreased significantly in GAL1-deficient mice. For Lgals1−/− BM–derived ASCs, decreases in IgM and low- and high-affinity IgG1 were noted at day 40, as well as at earlier time points for IgM and high-affinity IgG1 ASCs (Fig. 3B). However, affinity maturation, as measured by the NP3 (high)/NP23 (low) ratio, was not modified in the absence of GAL1 in the spleen (Fig. 3C) and BM (Fig. 3D) at days 21 and 40. Interestingly, spot intensities were similar in WT and Lgals1−/− cultures at all times tested (data not shown), suggesting a correct secretion process for Lgals1−/− PCs.

FIGURE 3.
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FIGURE 3.

In vivo–generated PCs are altered in Lgals1−/− mice. WT and Lgals1−/− mice were immunized with NP-KLH in Alum. The numbers of NP-specific IgM and IgG1 ASCs in the spleen (A) and BM (B) of WT and Lgals1−/− mice were evaluated by ELISPOT. Studies were performed in triplicate with at least five mice/group. Affinity maturation was measured by the NP3 (high)/NP23 (low) affinity ASC ratio in spleen (C) and BM (D) at days 21 and 40. Data are means of two independent experiments with four mice/group. Open and black squares correspond to WT and Lgals1−/− mice, respectively. *p < 0.05 versus WT, **p < 0.005 versus WT.

Altogether, these data indicate that PCs are correctly generated in the spleen of Lgals1−/− mice after T-dependent immunization, but their numbers decrease in spleen and BM at late time points of the immune response. The impairment in sustaining PCs following immunization could explain the defect in NP-specific Ig titers and ASCs observed in these mice.

In vitro generation of ASCs is altered in the absence of GAL1

To dissect the role of GAL1 in PC numbers and Ig secretion, we developed an in vitro assay in which PC differentiation was challenged. Splenic B cells from WT and Lgals1−/− mice were activated using various stimuli for 4 d, and the generation of CD138+ PBs was assessed by flow cytometry. After LPS stimulation, the same percentage (25–30%) and total number (2–3 × 105) of CD138+ cells were generated from WT and Lgals1−/− B cells (Fig. 4A, 4B). Similar results were obtained after NP-Ficoll or anti-IgM plus CD40L plus IL-4 stimulation (data not shown). We showed by real-time PCR that GAL1 mRNA was faintly expressed in nonstimulated WT B cells but was upregulated in the first days of LPS-induced differentiation (Fig. 4C). Ig secretion in LPS-stimulated B cell supernatants was monitored by ELISA at different time points. During differentiation, an increase in IgM (from day 2) and IgG3 (from day 3) levels was observed for both WT and Lgals1−/− B cell cultures (Fig. 4D). However, when the same experiments were performed in limiting culture conditions (i.e., by decreasing FCS concentration from 10 to 5%), IgM and IgG3 concentrations were reduced in Lgals1−/− B cell culture supernatant compared with WT B cell culture supernatant (Fig. 4E). The number of ASCs generated from LPS-stimulated B220+ splenic B was quantified by ELISPOT. Stimulated B cells from WT and Lgals1−/− mice were cultured in medium containing 10 or 5% FCS; after 3 d, viable cells were counted and plated with the same FCS concentration on ELISPOT plates. When culture media were supplemented with 10% FCS, no difference in the number of ASCs was observed between WT and Lgals1−/− cultures, whereas a reduced number of ASCs was noted for Lgals1−/−-derived PBs using 5% FCS (Fig. 4F). This decreased number of ASCs generated from Lgals1−/− cells was not due to compromised cell differentiation or proliferation at low serum concentration. Indeed, the number of CD138+ cells generated after 4 d of LPS stimulation in the presence of 5% FCS was the same in WT and Lgals1−/− cultures (Fig. 4G). Moreover, there was no difference in the proliferation rate, assessed by [3H]thymidine incorporation at days 2 and 3, for WT and Lgals1−/− cells cultured in the presence of 10 or 5% FCS (Fig. 4H). Thus, decreasing the FCS concentration in culture medium after stimulation of Lgals1−/− B cells reveals a defect in ASC number, which does not affect cell differentiation/proliferation, and a decrease in Ig concentration in culture supernatants.

FIGURE 4.
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FIGURE 4.

Ig levels are decreased in in vitro–generated Lgals1−/− PBs. B220+ splenic B cells from WT and Lgals1−/− mice were enriched by magnetic selection and cultured for 4 d in the presence of 1 μg/ml of LPS in complete RPMI 1640 medium with 10% FCS. The percentage (A) and the absolute cell number (B) of B220lowCD138+ PBs generated from 106 B220+ spleen cells 4 d after stimulation were evaluated by flow cytometry. (C) GAL1 expression in nonstimulated and LPS-stimulated B cells was determined by Q-PCR at days 1 and 2 after stimulation. (D and E) IgM and IgG3 secretion in culture supernatants from WT and Lgals1−/− cells were quantified by ELISA. Cultures were performed in culture medium supplemented with 10% (D) or 5% (E) FCS, and cells were allowed to secrete Ig for 16 h. (F) The number of ASCs obtained from 1000 viable plated cells after WT and Lgas1−/− cultures performed in 10 and 5% FCS was determined by ELISPOT. Data are representative of more than five independent experiments. (G) Absolute number of B220low CD138+ PBs generated from 106 B220+ spleen cells in the presence of 5% FCS and after 4 d of LPS stimulation. (H) Incorporation of [3H]thymidine at days 2 and 3 (evaluated in cpm) by LPS-stimulated B220+ cells in the presence of 10 and 5% FCS. Open and black squares correspond to WT and Lgals1−/− mice, respectively. *p < 0.05 versus WT.

PBs from GAL1-deficient mice are more susceptible to apoptosis

We first evaluated the sensitivity of the in vitro–generated Lgals1−/− PBs to apoptosis. When cultures contained 10% FCS, the percentage of CD138+7-AAD+ cells was the same in WT and Lgals1−/−-derived PBs (i.e., 3–4% of the cells) (Fig. 5A). In contrast, decreasing FCS to 5% increased the frequency of 7-AAD+ PBs to 7% in WT cultures and to 18% in Lgals1−/− cultures (i.e., 2.5-fold more than in WT cultures) (Fig. 5A). Thus, in vitro–generated PBs deficient for GAL1 display a higher susceptibility to cell death under serum starvation–induced death conditions. To further dissect this phenomenon, we tested different mechanisms of apoptosis, such as mitochondrial depolarization, phosphatidyl serine externalization, and caspase 8 activation. We used MitoTracker and annexin V staining to assess MitoTrackerhigh/annexinV+ cells, which have externalized phosphatidyl serine, and MitoTrackerlow/annexinV+ cells, which exhibit a decreased mitochondrial potential. As shown in Fig. 5B, we did not observe any difference in the depolarization potential of the mitochondria or the phosphatidyl serine externalization of WT and Lgals1−/− PBs generated in normal or low FCS concentrations. In contrast, we observed increased caspase 8 activation in Lgals1−/− PBs compared with WT cells (Fig. 5C). This phenomenon seems to be PB specific, because WT and Lgals1−/− B220+CD138− B cells exhibit similar rates of cell death mainly due to mitochondrial depolarization and phosphatidyl serine externalization rather than to caspase 8 activation (Supplemental Fig. 3). Thus, using an in vitro–differentiation assay, we showed that GAL1-deficient PBs formed normally but died rapidly by apoptosis through caspase 8 activation in serum starvation conditions.

FIGURE 5.
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FIGURE 5.

In vitro– and in vivo–generated Lgals1−/− PBs are more susceptible to apoptosis than are WT PBs. (A–C) B220+ splenic B cells from WT and Lgals1−/− mice were cultured with 1 μg/ml LPS for 3 d in complete medium containing 10 or 5% FCS. (A) Quantification of apoptotic B220lowCD138+ PBs was determined by 7-AAD incorporation. (B) Quantification of mitochondrial depolarization and phosphatidyl serine externalization on B220lowCD138+ PBs using MitoTracker and annexin V staining. Live cells are MitoTrackerhigh/annexinV−, cells with externalized phosphatidyl serines are MitoTrackerhigh/annexinV+, and cells with a decreased mitochondrial potential are MitoTrackerlow/annexinV+. (C) Quantification of caspase 8 activation. Fold changes in active caspase 8 MFI in B220lowCD138+ PBs generated in culture medium supplemented with 10 or 5% FCS. Data in (A) and (B) are representative of three independent experiments. (D) WT and Lgals1−/− mice (n = 6 mice/group) were immunized with NP-KLH in alum (T-dependent Ag), and the percentage of splenic apoptotic cells within IgM+CD138+ cells was quantified using TUNEL assays at day 21. For WT and Lgas1−/− mice, 3893 and 2154 cells were counted, respectively. *p < 0.05 versus WT, **p < 0.005 versus WT.

To confirm the in vitro results, TUNEL assays were performed to measure the sensitivity of in vivo–generated GAL1-deficient PCs to apoptosis. WT and Lgals1−/− mice were immunized with the NP-KLH T-dependent Ag, and TUNEL assays were performed on spleen sections at days 21 and 40 after immunization. The percentage of TUNEL+ cells was higher in IgM+CD138+ PCs cells in Lgals1−/− mice compared with WT mice at day 21 (Fig. 5D) and day 40 (data not shown).

Altogether, our results show that GAL1-deficient PCs are more susceptible to apoptosis than are their normal counterparts.

Discussion

Many reports highlighted the crucial role of GAL1 in immunity, inflammation, and tumor progression, but little was known about its effect on B cells. In this study, we showed that GAL1 is specifically expressed by splenic and BM PBs during PC differentiation following T-independent and T-dependent immune responses. This is in accordance with the detection of GAL1 transcripts after Blimp1 transfection of human and mouse B cell lines or after in vitro LPS stimulation of mouse B cells (18, 21). We also revealed a new role for GAL1 in the maintenance of ASC numbers and Ig titers during T-dependent and T-independent immune responses. Using an in vitro–differentiation assay, we showed that Lgals1−/− PBs are more susceptible than are their WT counterparts to serum starvation–induced death conditions. Actually, TUNEL assays show that in vivo–generated GAL1-deficient PCs present an increased sensitivity to apoptosis. GAL1 does not seem to interfere with affinity maturation and Ig class switching, because we observed normal levels of the different Ig isotypes and of high-affinity NP-specific IgG1 in Lgals1−/− mice after immunization. Thus, our results indicate that GAL1 controls the number of PBs during the immune response by regulation of caspase 8–mediated apoptosis.

In contrast to our results, Tsai et al. (18) reported that immune responses were not altered in Lgals1−/− mice. Because they refer to data not shown, we could not compare the Ag, the doses, and the immunization protocol used in the two studies. Moreover, it is well known that some phenotypes are strongly dependent on the mouse genetic background (22). Thus, the differences observed could reflect the fact that our experiments were performed on Lgals1−/− mice backcrossed with 129SV mice for a minimum of six generations, whereas Tsai et al. (18) used Lgals1−/− mice on a mixed C57Bl6/129SV genetic background, with chimerism markers still present.

Although we observed a significant defect in Ig serum titers and PC numbers in Lgals1−/− mice on pure genetic backgrounds (129SV in this study and C57Bl6, data not shown), we cannot completely rule out that other galectins may be important for PC homeostasis. Indeed, galectin redundancy was reported in other biological systems. In line with this, Tsai et al. (23) observed the expression of GAL8 in in vitro LPS-activated B cells. In our in vivo system, we used PCR to confirm that GAL8 was also expressed in sorted CD138+ B220low cells obtained from WT mice after T-dependent immunization (data not shown).

In contrast to the well-established proapoptotic effect of GAL1 on T cells (8), it appears to have a protective effect during PC differentiation. Antagonist roles for GAL1 were described in the lymphoid lineage. For example, in the thymus, GAL1 is expressed by epithelial cells (24), and it induces cell-cycle arrest and/or apoptosis of human and murine thymocytes (9, 25–27). In contrast, in the BM, GAL1 expressed by stromal cells supports B cell differentiation through pre-BCR activation and signaling (16, 17, 28). These activities are both dependent on extracellular GAL1. Recently, secreted GAL1 was found to bind to splenic mature B220+ B cells but not to CD138+ B220low cells (18). This is due to the differential expression of glycosyl transferases by the two cell types, which results in the expression of specific cell surface glycans (23). Taking into account these results, we postulate that the antiapoptotic effect of GAL1 on CD138+B220low cells is not dependent on GAL1 secretion by PCs that do not possess the mandatory glycosylations to bind it, rather it is due to intracellular mechanisms. Nevertheless, in vivo, an extrinsic role for GAL1 cannot be excluded, because GAL1+ dendritic cells and resident and inflammatory macrophages are present in the spleen and lymph nodes (A. Anginot, unpublished observations); they may influence immune responses, especially PC differentiation. Moreover, GAL1 was shown to be expressed by BM stromal cells and could influence PC homing and survival in this organ (17). However, our in vitro experiments were performed using purified B220+ B cells in the absence of exogenous recombinant GAL1 in the cultures, supporting a cell autonomous effect of GAL1 on the newly generated PBs.

Because few CD138+ cells are generated in vivo and because these cells are extremely sensitive to apoptosis during cell sorting, we used in vitro studies to determine whether the proapoptotic signaling pathways were overstimulated in GAL1-deficient mice. Although the apoptosis was not associated with mitochondria depolarization, we observed an increase in caspase 8 activation. Caspase 8 was shown to be activated by FADD, which is the main adaptor protein transmitting apoptotic signals mediated by death receptors, such as FAS, TNFRs, or TRAIL receptors (29). It remains to be determined which of these death receptors could be implicated and how GAL1 can influence this apoptotic pathway.

In pathological situations, such as multiple myeloma (MM), malignant PCs, which accumulate in the BM, express high levels of GAL1 (21). The bad prognosis associated with GAL1 expression by tumor cells or by the microenvironment has mainly been related to GAL1-mediated antitumor T cell apoptosis (6). However, taking into account our results, it is conceivable that the high ectopic level of intracellular GAL1 protects MM cells from apoptosis. Thus, GAL1 could be considered a new therapeutic target in MM.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank F. Poirier for providing GAL1-deficient mice; F. Mallet and A. Bole for helping with ELISPOT and ELISA techniques, respectively; and the flow cytometry facility of the Centre d'Immunologie de Marseille-Luminy (Marseille, France) for expert technical assistance. We gratefully acknowledge Lee Leserman for critical reading of the manuscript.

Footnotes

  • This work was supported by grants from the Agence Nationale de la Recherche (05-BLAN-0035-01) and Association pour la Recherche contre le Cancer (contract no. 1089) and institutional grants from INSERM and Centre National de la Recherche Scientifique. A.A. was funded by the Agence Nationale de la Recherche, and M.E. was funded by a Federation of European Biochemical Societies long-term fellowship and by The Wellcome Trust (Programme Grant 083650/Z/07/Z).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    7-AAD
    7-aminoactinomycin D
    ASC
    Ab-secreting cell
    BM
    bone marrow
    GAL1
    galectin-1
    KLH
    keyhole limpet hemocyanin
    MFI
    mean fluorescence intensity
    MM
    multiple myeloma
    NP
    4-hydroxy-3-nitrophenylacetyl
    PB
    plasmablast
    PBST
    PBS and 0.05% Tween 20
    PC
    plasma cell
    WT
    wild type.

  • Received July 9, 2012.
  • Accepted March 20, 2013.
  • Copyright © 2013 by The American Association of Immunologists, Inc.

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The Journal of Immunology: 190 (11)
The Journal of Immunology
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Galectin 1 Modulates Plasma Cell Homeostasis and Regulates the Humoral Immune Response
Adrienne Anginot, Marion Espeli, Lionel Chasson, Stéphane J. C. Mancini, Claudine Schiff
The Journal of Immunology June 1, 2013, 190 (11) 5526-5533; DOI: 10.4049/jimmunol.1201885

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Galectin 1 Modulates Plasma Cell Homeostasis and Regulates the Humoral Immune Response
Adrienne Anginot, Marion Espeli, Lionel Chasson, Stéphane J. C. Mancini, Claudine Schiff
The Journal of Immunology June 1, 2013, 190 (11) 5526-5533; DOI: 10.4049/jimmunol.1201885
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