BCR signal strength upon antigen binding is defined by a host of co-receptors that fine-tune and co-ordinate signal transduction. Increased signal strength leads to productive B cell activation when antigen affinity or concentrations are limiting (Zikherman and Lowell, 2017). Therefore, BCR co-receptors modulate BCR signaling and define the threshold of antigen required for B cell activation (Tedder et al., 1997). We have shown that Gal9 regulates BCR signaling by binding directly to IgM-BCR, altering its membrane organization with respect to the inhibitory co-receptors CD45 and CD22, thereby dampening BCR signal transduction (Cao et al., 2018). To investigate whether Gal9 impacts the threshold of B cell activation, we first asked if Gal9 regulates the intensity of signal transduction downstream of BCR ligation. To address this, we stimulated primary naïve murine wild-type (WT) and Lgals9^−/− (Gal9KO) B cells with defined concentrations of anti-IgM F(ab’)₂ as surrogate antigen and measured total tyrosine phosphorylation at 5 min by flow cytometry. Modulating BCR signal strength through limiting the concentration of stimulatory antigen leads to a decrease in the magnitude of the signaling response at 5 min, as observed by the decrease in geometric mean fluorescence intensity (gMFI) of total-phosphotyrosine (Figure 1A,B). In the absence of Gal9, however, BCR signaling is enhanced at limiting antigen concentrations compared to WT B cells (Figure 1A,B).

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Following antigen binding and initiation of the BCR signaling cascade, the BCR-antigen complex becomes internalized and brought to major histocompatibility complex class II (MHCII) containing lysosomes, which facilitate antigen processing and subsequent presentation to T cells, a crucial step to achieve full B cell activation (Avalos and Ploegh, 2014). To assess whether altered BCR signaling in Gal9-deficient B cells impacts BCR internalization, we stimulated WT and Gal9KO primary murine B cells with defined concentrations of anti-IgM F(ab’)₂ and measured surface expression of IgM-BCR over 20 min. Following stimulation with high concentrations of anti-IgM F(ab’)₂, we see a sharp decrease in surface IgM that plateaus by 20 min, whereas stimulation with decreasing concentrations leads to less IgM internalized (Figure 1C). Notably, in the context of reduced antigen concentrations, IgM internalization is increased in Gal9KO compared to WT B cells (Figure 1C). To quantify the rate of IgM internalization, we plotted the rate of change (k) of the fitted decay curve, which revealed Gal9KO B cells internalize IgM at a faster rate than WT B cells (Figure 1D). Furthermore, this results in an increase in total BCR internalization at 20 min to limiting antigen concentrations in Gal9KO B cells (Figure 1E). Taken together, these data suggest that increased BCR signaling in Gal9-deficient B cells leads to an increase in internalization of IgM-BCR.

To assess the relationship between BCR signal strength and BCR internalization, we used a Nur77-eGFP reporter system, in which GFP expression is driven by Nur77 and is rapidly upregulated upon BCR signaling (Zikherman et al., 2012). Following stimulation for 20 min with defined concentrations of anti-IgM F(ab’)₂, WT and Gal9KO Nur77-eGFP B cells were washed of excess antigen and allowed to incubate for 16 hr to express GFP proportional to the extent of BCR signal transduction. Consistent with our previous findings, BCR signal transduction to low-dose stimuli, as measured by the gMFI of the GFP reporter, is enhanced in the absence of Gal9 (Figure 1—figure supplement 1A,B). Furthermore, the extent of BCR signaling (gMFI of GFP) correlates with the rate of BCR internalization over the same interval (k, over 20 min) (Figure 1—figure supplement 1C). Taken together, these data demonstrate that Gal9 regulates BCR internalization through modulating BCR signal strength.

BCR signal transduction leads to a B cell activation program, marked by upregulation of activation markers and a change in transcriptional and metabolic profiles. We then asked if enhanced BCR signaling in response to limiting antigen concentrations in Gal9KO B cells affects the threshold of B cell activation as assessed by the upregulation of the co-receptor CD86. To address this, we stimulated primary naïve B cells from WT or Gal9 deficient mice with titrated concentrations of anti-IgM F(ab’)₂ for 16 hr and assessed upregulation of CD86 by flow cytometry. Following 16 hr of stimulation we see increased B cell activation proportional to increasing concentration of anti-IgM; however, in the absence of Gal9 B cells respond to lower concentrations of antigen (Figure 1F), quantified by the decreased EC₅₀ value (Figure 1G). Taken together, these data demonstrate that Gal9 restrains B cell activation when antigen concentrations are limiting.

Gal9 is a pleiotropic protein with many described roles in a variety of immune cells. Because of its secreted nature, Gal9 can be produced by one cell and act on another. This enhanced layer of complexity prompted us to ask which cell types within the spleen can produce Gal9 and contribute to extrinsic sources of Gal9 in the tissue? To address this, we compared Gal9 expression on the surface of a variety of splenic immune cell populations by flow cytometry (Yu et al., 2016; Figure 1—figure supplement 2A). Additionally, to delineate cells which may be excreting Gal9, we quenched the surface Gal9 with an unlabeled monoclonal antibody, then permeabilized cells, and stained for intracellular Gal9 using a fluorescent conjugation of the same monoclonal antibody (Figure 1—figure supplement 2B). Importantly, following quenching with unlabeled antibody, unpermeabilized cells did not stain with fluorescent antibody, demonstrating effective saturation of the quenching antibody in this assay (data not shown). Interestingly, we see high levels of intracellular and extracellular Gal9 expression from many myeloid cell populations, with little detectable Gal9 from the T cell compartment. The contribution of subset-derived Gal9 is a factor of both the expression of Gal9 by a given subset and the overall abundance of that subset within the splenic environment. As such, we normalized the total Gal9 expression (intra- and extracellular) against the relative abundance of each cell subset in the spleen, providing a metric of Gal9 expression that considers the size of specific populations (Figure 1—figure supplement 2C). We see that when we factor in subset abundance, B cells appear to have the highest Gal9 expression relative to abundance within the spleen.

We next asked which cell types within the spleen contribute to the Gal9 detected on the surface of B cells. To investigate this, we performed adoptive transfer experiments using Gal9KO B cells transferred into WT hosts, µMT hosts (which lack B cells), and clondronate liposome-treated WT hosts (which results in selective depletion of phagocytic cells, predominantly macrophages/monocytes). We then stained for Gal9 expression on transferred cells to assess the cell type-specific contribution to surface Gal9 expression on B cells. Following treatment with clondronate liposomes, we see approximately 50% reduction in splenic macrophage and monocyte populations compared to control liposome-treated mice (Figure 1—figure supplement 2D). Interestingly, when we compare the expression of Gal9 on transferred Gal9KO B cells with endogenous WT B cells in the same host, we see full reconstitution of Gal9 expression on follicular B cells when transferred into WT hosts (Figure 1—figure supplement 2E). Notably, however, Gal9 expression is only partially restored when gating on marginal zone B cells (Figure 1—figure supplement 2F), suggesting that B cells may have subset-specific requirements for intrinsic vs extrinsic sources of Gal9. Additionally, Gal9 expression was only partially restored when Gal9KO B cells were transferred into macrophage depleted or B cell-deficient hosts (Figure 1—figure supplement 2E,F), demonstrating that both B cell-derived and macrophage-derived Gal9 contribute to extrinsic sources of Gal9 within the splenic environment. Lastly, we asked if these perturbations in Gal9 expression affect the threshold of B cell activation. To address this, we stimulated cells with titrated concentrations of anti-IgM F(ab’)₂ for 16 hr and assessed upregulation of CD86 by flow cytometry. With reconstitution of Gal9 expression by transfer into WT hosts, we see that Gal9KO B cells have similar threshold of activation compared to endogenous WT cells in the same host (Figure 1—figure supplement 2G,H). In contrast, we see a decrease in the threshold of activation of Gal9KO B cells transferred into macrophage-depleted or B cell-deficient hosts, which is proportional to Gal9 expression. Taken together, these data demonstrate that macrophage populations and B cells act as a source of Gal9 within the spleen and that these sources of extrinsic Gal9 are sufficient to alter the threshold of B cell activation.

B cell activation is defined by factors such as the abundance of antigen as well as the affinity of that antigen for cognate BCR (Zikherman and Lowell, 2017). To assess the role of Gal9 in modulating B cell activation to antigens with reduced affinity, we used the transgenic MD4 system. In this model, B cells express an IgM-BCR specific for model antigen hen egg lysozyme, HEL (HyHEL10, K_a = 4.5 × 10¹⁰ M⁻¹). MD4 B cells bind to lysozymes from other avian species with reduced affinity, such as Bobwhite Quail (QEL, K_a = 3 × 10⁸ M⁻¹) and Duck (DEL, K_a = 1.7 × 10⁷ M⁻¹) (Lavoie et al., 1992). To investigate a role for Gal9 in antigen affinity discrimination, we first settled WT or Gal9KO MD4 B cells on artificial planar lipid bilayers displaying HEL, QEL, or DEL antigens, allowing us to simulate antigen presentation in a similar fashion to how B cells interact with antigen in vivo (Harwood and Batista, 2008). At 90 s post-interaction, the point of maximal B cell spreading, cells were fixed and imaged using total internal reflection fluorescence (TIRF) microscopy (Figure 2A). Gal9KO B cells appear to spread more and accumulate more antigen in response to lower affinity antigens compared to WT B cells. To quantify this, we measured the contact area and total amount of antigen accumulated (Figure 2B,C). Indeed, Gal9KO B cells spread more and accumulate more antigen in response to low-affinity antigens presented on lipid bilayers compared to WT B cells. It is well established that the B cell spreading response, and thus the amount of antigen accumulated is regulated by BCR signaling (Fleire et al., 2006; Weber et al., 2008). Thus, to assess the impact of Gal9 on BCR signaling in the context of an antigen presenting cell, we labeled OP9 stromal cells and deposited lysozyme containing immune complexes onto their surface at three different densities, covering a fourfold range, corresponding to low, medium, and high density. OP9 stromal cells were incubated with WT or Gal9KO MD4 B cells for 5 min, and cell conjugates were then permeabilized, stained for total tyrosine phosphorylation, and analyzed by flow cytometry. In the absence of Gal9, BCR signaling is increased in response to low-affinity and low-density membrane-bound antigens compared to WT B cells (Figure 2D,E).

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We then asked if this altered signaling in response to low-affinity and low-density membrane-bound antigens in Gal9KO B cells impacts the threshold of downstream B cell activation. To investigate this, we stimulated WT or Gal9KO MD4 B cells with titrated concentrations of these antigens and assessed B cell activation by upregulation of CD86. In the absence of Gal9, upregulation of CD86 is increased in response to lower concentrations of antigen in all three affinities of lysozyme (Figure 2F). Furthermore, in the absence of Gal9, B cells internalize IgM at a faster rate in response to low-affinity antigens compared to WT B cells (Figure 2G, Figure 2—figure supplement 1A). This results in a greater amount of IgM and antigen internalized in Gal9KO B cells when stimulated with low-affinity antigens (Figure 2H,I, Figure 2—figure supplement 1B). Taken together, these data demonstrate that Gal9 regulates B cell responsiveness to low-affinity and low-density antigens.

BCR signaling is tightly regulated to mitigate inappropriate activation to otherwise innocuous autoantigens. Many central and peripheral tolerance mechanisms are in place to eliminate B cells with high affinity for autoantigens from the B cell repertoire, resulting in a pool of B cells that should have little to no affinity toward autoantigens (Basten and Silveira, 2010). Given our observation of enhanced B cell activation to low-affinity antigens in Gal9-deficient B cells, we then asked, does loss of Gal9 result in spontaneous autoreactivity and the development of autoimmunity in mice? To investigate this, we examined aged (>8 months) WT and Gal9KO mice for evidence of spontaneous autoimmunity. We find that aged Gal9KO mice have enlarged spleens (Figure 3A), with detectable germinal center (GC) B cells in the absence of immunization, suggesting they are driven toward autoantigens (Figure 3B,C). Furthermore, we see enhanced antibody secreting cell (ASC; defined as FSC^hi, surface IgM^neg, and CD138^hi expressing cells) (Pracht et al., 2017) development in the spleen of Gal9KO mice compared with WT littermate controls (Figure 3D,E). Further characterization of the ASC compartment revealed a clear skewing toward B220-negative long-lived ASCs in the absence of Gal9 (Pracht et al., 2017; Figure 3F). Consistent with this, we observe increased frequency of B220^- ASCs in the bone marrow the absence of Gal9 (Figure 3—figure supplement 1A,B). We next asked if these spontaneous germinal centers and increased ASCs are driving autoimmunity. To address this, we looked for circulating autoantibodies in the serum of aged Gal9KO mice. We see increased autoreactivity to HEp-2 cells with serum from Gal9KO mice compared to WT controls, having circulating IgG antibodies that are largely nuclear-reactive and circulating IgM that are largely cytoplasmically reactive (Figure 3G,H). Furthermore, antigen-specific anti-dsDNA and anti-GME (glomerular membrane extract) autoantibodies are increased in Gal9KO mice (Figure 3I,J). In lupus-like disease, autoantigen containing immune complexes (ICs) are a source of inflammation as they deposit in the kidneys and other organs and impair filtration and kidney function. Thus, we next assessed circulating immune complexes and kidney function in aged WT and Gal9KO mice. We observe an increase in circulating ICs in the serum (Figure 3K) as well as increased proteinuria (Figure 3L), indicative of nephritis, in aged Gal9KO mice compared to WT controls. Taken together, these data indicate that Gal9KO mice develop spontaneous autoimmunity with age.

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T follicular helper (Tfh) cells are essential for the development of autoimmune disease as they are required for the formation and maintenance of GCs (Stebegg et al., 2018). A population of regulatory Tfh cells expressing FoxP3 (termed Tfr cells) have been described, and these cells play a critical role in regulating B cell responses in peripheral tissues. Loss of Tfr cells leads to activation of autoreactive B cells and unchecked autoimmunity (Wollenberg, 2011). Thus, we measured the frequency of Tfh and Tfr cells in the spleen of aged Gal9KO mice compared to WT controls. We see that aged Gal9KO mice have an increase in Tfh cell frequency (Figure 3—figure supplement 1C and D), mirroring the increase in GC B cells (Figure 3B,C). Additionally, we observe a similar increase in the frequency of FoxP3+ Tfr cells in the spleen of Gal9KO mice; however, the ratio of regulatory Tfr cells to Tfh cells is similar in both WT and Gal9KO mice (Figure 3—figure supplement 1E–G). These data suggest that loss of Gal9 does not lead to spontaneous autoimmunity through impaired generation of regulatory Tfh cells.

BCR signal strength and T cell help are key regulators of tolerance in peripheral tissues. To assess whether there is a breakdown in peripheral tolerance in Gal9KO B cells, we crossed Gal9KO mice onto the MD4/ML5 mouse model (Goodnow, 2009). In this system, B cells express a transgenic BCR specific for HEL, as well as soluble HEL secreted by somatic cells. Upon binding antigen in this context, B cells undergo a process of peripheral tolerance called anergy where they become refractory to BCR stimulation, downregulate surface IgM, and have a decreased half-life (Goodnow et al., 1989). Therefore, in this model of anergy, there is little to no secretion of antibodies toward the pseudo-autoantigen HEL. We observed a distinct population of cells in the spleen of Gal9KO MD4/ML5 mice that retain high levels of surface IgM (Figure 3—figure supplement 1H and I). Furthermore, following IgM stimulation, this population induces robust signaling compared to both anergic WT cells and IgM-low anergic-like cells in Gal9KO mice (Figure 3—figure supplement 1J,K). Additionally, we detect higher titers of HEL-specific antibodies in the serum of Gal9KO MD4/ML5 mice compared to MD4/ML5 WT controls (Figure 3—figure supplement 1L). These data suggest there is a breakdown in peripheral tolerance in the absence of Gal9, leading to a population of cells that escape tolerance mechanisms.

B-1a cells are a population of B cells derived from fetal progenitors that undergo a positive selection process during development, providing B-1a cells with an autoreactive repertoire. It is thought that this is beneficial for aiding in the clearance of cellular debris and senescent red blood cells (Duan and Morel, 2006). B-1a cells, defined by their expression of the T cell regulatory protein CD5, along with CD5-negative B-1b cells are typically found at barrier sites such as the lining of the gastrointestinal tract and peritoneal (PerC)/pleural cavities. B-1 cells rapidly respond to both BCR and TLR stimulation and can readily secrete antibodies in the absence of T cell help. Intriguingly, B-1a cells are expanded in several models of autoimmunity, though their specific role remains incompletely defined (Duan and Morel, 2006). We therefore asked if B-1a cells were similarly expanded in aged Gal9KO mice. Indeed, we find an increase in the proportion of B-1a cells in the PerC of Gal9KO mice compared to WT mice (Figure 4A,B). Additionally, we see an increase in cell number for B-1a cells in the PerC of Gal9KO mice (data not shown). Furthermore, B-1a cells from Gal9KO mice have higher expression of activation markers at steady-state (Figure 4C,D; Figure 4—figure supplement 1A–D), suggesting that Gal9 restrains B-1a activation.

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Activation of B-1 cells readily drives development into ASCs. Consistent with our previous results, we find an expanded population of ASCs in the PerC of aged Gal9KO mice compared to WT littermate controls (Figure 4E,F). We then asked if these ASCs were contributing to circulating antibody titers in the serum. A large proportion of B-1a cells share a common specificity for phosphoryl-choline (PC) (Baumgarth, 2016). We performed PC-specific ELISAs and found that Gal9KO mice have an increase in PC-specific antibodies, consistent with their increased ASCs (Figure 4G). Upon activation at barrier sites, B-1 cells migrate to secondary lymphoid organs (Ha et al., 2006). Consistent with this, we see an expanded population of B-1a cells in the spleen of Gal9KO mice compared to WT littermates (Figure 4H–K). Additionally, in the NZB/W F1 model of lupus-like autoimmunity, B-1a cells migrate to the thymus and mediate selection of autoreactive T cell clones (Sato et al., 2004). Similarly, we see an increase in B-1a cells in the thymus of aged Gal9KO mice (Figure 4—figure supplement 1E–G). Taken together, these data demonstrate that Gal9 regulates B-1a cell accumulation and activation at steady-state and implicate B-1a cells in the autoimmune phenotype observed in Gal9-deficient mice.

We next asked if the expansion of B-1a cells and their increased expression of activation markers at steady-state in Gal9KO mice is due to altered BCR signaling. To address this, we stimulated PerC B-1 cells with titrated concentrations of anti-IgM F(ab’)₂ for 16 hr and measured B cell activation by upregulation of CD86. We find that loss of Gal9 leads to enhanced activation of B-1a cells in response to lower concentrations of agonist (Figure 5A–C). Notably, this increased activation upon loss of Gal9 is specific to B-1a cells, as we did not observe any difference in B-1b cell responses (Figure 5—figure supplement 1A–C). We then asked if Gal9 interactions directly modulate BCR signal transduction in B-1a cells. We stimulated PerC B-1 cells with anti-IgM F(ab’)₂ for 5 min, followed by fixation and staining for phospho-signaling molecules by flow cytometry. We see that Gal9KO B-1a cells have enhanced phosphorylation of signaling machinery following IgM stimulation (Figure 5D). To assess whether enhanced activation is specifically due to Gal9 at the cell surface, we treated WT PerC cells with the pan-galectin inhibitor lactose to remove all galectins from the cell surface and, alternatively, treated Gal9KO PerC cells with recombinant Gal9 (rGal9) to reconstitute Gal9 expression (Figure 5—figure supplement 2). Following IgM stimulation, pan-galectin inhibition in WT B cells results in increased signal transduction and importantly, reconstitution of Gal9 in Gal9KO cells directly inhibits signaling in B-1a cells (Figure 5D). In contrast, but consistent with our earlier observations, these treatments have little effect on B-1b cell signaling (Figure 5—figure supplement 1D). We wondered if differences in Gal9 expression may explain some of the subset-specific discrepancies in B cell activation. Indeed, B-1a cells have higher expression of Gal9 at steady-state compared to B-1b cells (Figure 5E). Taken together, our results demonstrate a specific role for Gal9 in the regulation of B-1a, but not B-1b cells.

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We have shown that Gal9 regulates B-2 B cell activation through mediating interaction between IgM-BCR and negative co-receptors CD22 and CD45 (Cao et al., 2018); however, B-1a cells are known to express a unique surface profile lacking these negative regulators. Therefore, Gal9 regulation of B-1a cells must occur through a unique mechanism. To investigate this, we performed pull-down assays to identify Gal9 ligands in B-1a B cells by incubating protein lysates from sorted B-1a and B-1b PerC B cells or conventional splenic B-2 cells with rGal9-coated beads. We then washed and stained these beads with antibodies targeting various co-receptors and assessed protein enrichment by flow cytometry. We find that Gal9 interacts with IgM-BCR in B-1a, B-1b, and B-2 B cells, but uniquely interacts with the inhibitory glyco-protein CD5 on the surface of B-1a cells (Figure 5F, Figure 5—figure supplement 1E). Interestingly, we observed only slight enrichment of other known B-1 cell regulators such as Siglec-G or FcγRIIB on Gal9 coated beads, suggesting that Gal9 preferentially interacts with other protein species such as CD5 (Figure 5—figure supplement 3). Taken together, these data demonstrate that Gal9 binds to B-1a cells and regulates their activation, likely through the inhibitory co-receptor CD5.

The nanoscale organization of molecules on the cell surface allow for interactions between receptors and important regulatory co-receptors that enhance or restrain signal transduction (Treanor, 2012). We have shown previously that Gal9 modulates BCR responses in B-2 B cells by altering the nano-scale distribution of the BCR and its association with inhibitory co-receptors (Cao et al., 2018; Wasim et al., 2019). To assess the impact of Gal9 on the nanoscale organization of IgM-BCR and inhibitory co-receptor CD5, we performed dual-color direct stochastic optical reconstruction microscopy (dSTORM) to achieve single-molecule resolution of the cell surface. Upon visual inspection of dSTORM images, IgM appears to be more clustered on B-1a cells in the absence of Gal9 (Figure 5G). Quantification of cluster area demonstrated that IgM clusters are larger in the absence of Gal9 (Figure 5H). To quantify clustering tendency, we calculated the Hopkins index, which assesses clustering tendency relative to a random distribution (0.5). We find that IgM on the surface of Gal9KO B-1a cells has a higher clustering tendency compared to WT B-1a cells (Figure 5I). Upon visual inspection of dSTORM images of CD5, we note that it is more highly clustered compared to IgM; however, we do not observe striking differences between WT and Gal9KO B-1a cells. However, quantification of CD5 cluster area and degree of clustering revealed an increase in Gal9KO B-1a cells (Figure 5J,K). We hypothesized that Gal9 crosslinks IgM and CD5 to regulate BCR signaling in B-1a cells. Dual-dSTORM provides the advantage of single-molecule coordinate-based colocalization analysis to examine whether there are differences in the colocalization of these molecules on the surface of B-1a cells in the absence of Gal9. Visually, we can discern a loss of colocalized IgM and CD5 clusters in the absence of Gal9 (Figure 5G). To quantify this, we performed coordinate-based colocalization (CBC) analysis, which ranges from −1 (perfectly segregated) to +1 (perfectly colocalized). The frequency distribution of CBC values shifts to the left (i.e. toward negative values) in the absence of Gal9, indicating an increase in the segregation of IgM-BCR and CD5, compared to WT B-1a cells (Figure 5L). This results in a decrease in the mean CBC value per cell (Figure 5M) and a corresponding increase in the mean nearest neighbor (NN; Figure 5N). In contrast, but consistent with the lack of effect of Gal9 deficiency on B-1b B cell activation, we find no difference in the organization of IgM on the surface of B-1b cells in the absence of Gal9 (Figure 5—figure supplement 1F–G). Taken together, these findings identify a unique and specific mechanism of Gal9 regulation of B-1a BCR signaling.

B-1 cells respond to T cell-independent antigens at barrier sites and rapidly differentiate into effector cells such as ASCs through BCR or TLR stimulation (Baumgarth, 2016). To assess whether Gal9 influences innate-like responses of B-1 cells, and to identify TLRs potentially regulated by Gal9, we stimulated PerC cells from WT or Gal9KO mice with titrated concentrations of various TLR ligands and measured the proportion of cells activated by upregulation of CD86. Interestingly, we find that B-1a cells from Gal9KO mice are more sensitive to LPS and CpG stimulation, but not imiquimod or zymosan stimulation (Figure 6A–D). In contrast, but consistent with BCR stimulation, Gal9 appears to be dispensable for TLR activation of B-1b cells (Figure 6—figure supplement 1A–D). We then asked if enhanced CD86 upregulation in the absence of Gal9 was due to a similar enhancement of TLR signaling in B-1a cells. To address this, we stimulated WT or Gal9KO PerC cells with LPS and assessed total-tyrosine phosphorylation at 10 min by phospho-flow cytometry. We find that Gal9KO B-1a cells, but not B-1b cells, have increased total tyrosine phosphorylation at 10 min post-LPS stimulation, which can be reduced to levels comparable to WT cells by pre-treatment with rGal9 (Figure 6E, Figure 6—figure supplement 1E). Similarly, pre-treatment of WT B-1a, but not B-1b, cells with lactose to displace galectins, enhanced total tyrosine phosphorylation upon LPS stimulation (Figure 6E, Figure 6—figure supplement 1E). To identify Gal9 ligands in B-1a cells that mediate this phenotype, we performed pull down assays with rGal9-coated beads incubated with B-1a or B-1b cell lysates, as previously. We find that Gal9 binds TLR4 and the inhibitory molecule CD180 on B-1 cells, and to a lesser extent on B-2 B cells (Figure 6F,G), consistent with previous mass spectrometry analysis of Gal9 ligands in B2 B cells (Cao et al., 2018).

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To assess the impact of Gal9 on TLR4 nano-scale organization, we again performed dual-color dSTORM to identify single molecule localizations of TLR4 and CD180 on the surface of B-1 cells from WT and Gal9KO mice (Figure 6H). In the absence of Gal9 the mean cluster area and clustering tendency of TLR4 is decreased on B-1a cells (Figure 6I–J), but interestingly, Gal9 has no affect on the organization of TLR4 in B-1b cells (Figure 6—figure supplement 1F-H). Similarly, the clustering tendency of CD180 is decreased in the absence of Gal9 in B-1a cells; however, the area of CD180 clusters is larger (Figure 6H,K–L), whereas deficiency of Gal9 has no effect on the organization of CD180 in B-1b cells (Figure 6—figure supplement 1F,I–J). To assess whether Gal9 mediates close association of TLR4 with the inhibitory co-receptor CD180, we performed CBC analysis as previously described. The mean CBC value of TLR4 and CD180 is decreased, and mean nearest-neighbor increased, in B-1a cells from Gal9KO mice compared to WT mice (Figure 6M–O), however there is no difference in B-1b cells (Figure 6—figure supplement 1K,M). Taken together, these data demonstrate that Gal9 increases the association of TLR4 and CD180, specifically on the surface of B-1a cells, but not B-1b cells, to modulate TLR4 signaling.

TLR stimulation of B-1 cells leads to rapid ASC differentiation (Ha et al., 2006). So, we asked if Gal9-mediated regulation of TLR signaling influences ASC development. To address this, we isolated PerC B-1 cells from WT and Gal9KO mice and sorted B-1a cells. We stimulated sorted cells ex vivo with 0.5 µg/mL LPS and 0.5 µg/mL anti-IgM and measured ASC differentiation by CD138 expression 3 days post-stimulation. We find that B-1a cells from Gal9KO mice more readily adopt an ASC phenotype following stimulation, which results in more secreted antibody in the culture supernatant (Figure 7A–D).

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As B-1a-derived autoantibodies are thought to aid in clearance of cellular debris (Duan and Morel, 2006), we therefore asked if enhanced sensitivity of B-1a cells to LPS could have an impact in vivo. To address this, we injected mice with a low (0.05 µg/mL) or high dose (1 µg/mL) of LPS by intraperitoneal (IP) injection and assessed B cell activation 24 hr post-injection. Following low-dose LPS injection, upregulation of CD86 is enhanced in Gal9-deficient B-1a cells, but not B-1b cells (Figure 7—figure supplement 1A). We then asked if these activated B-1a cells are secreting autoantibodies into circulation. Indeed, we find a greater fold increase in both PC-specific IgM and IgG in Gal9KO mice 24 hr post low-dose LPS injection (Figure 7—figure supplement 1B).

We then asked if B-1a cell-derived immune complexes that form following LPS injection carry autoantigens into secondary lymphoid organs. To address this, we IP injected mice with low-dose LPS followed by CFSE-labeled apoptotic bodies (ApoBs) by intravenous injection 24 hr later. At 12 hr post-transfer, we looked for ApoB capture by subcapsular sinus (SCS) macrophages in the spleen of WT or Gal9KO mice. We find that ICs formed following LPS injection are capable of capturing CFSE-labeled ApoB and recruiting them to SCS macrophages in the spleen (Figure 7—figure supplement 1C–E). To determine whether autoantigen delivery to SCS macrophages was directly driven by increased antibody secretion by B-1a cells, we isolated secreted antibodies from sorted B-1a and B-1b cultures stimulated ex vivo and injected these antibodies into mice every 3 days over a 2-week period (Figure 7E). Following 2 weeks of antibody transfer, we detect ICs in circulation as well as in the spleens of mice receiving B-1a-derived antibodies in both WT and Gal9KO recipient mice (Figure 7F–K).

To assess whether enhanced delivery of ICs bearing autoantigens facilitates autoimmunity, we measured splenic GC B cells 4 weeks following final antibody transfer. Following transfer of B-1a-derived antibodies, GCs formed in the spleen and autoantibodies are detected in circulation (Figure 7L–O). Additionally, we detect low levels of protein secretion into the urine at this time point, suggesting that these mice begin to develop nephritis (Figure 7P). These observations are further exacerbated in Gal9KO mice, which have a reduced threshold for B cell activation, and subsequently develop a more exaggerated autoimmune response (Figure 7L–P). Collectively, these data demonstrate that Gal9 regulates BCR-driven and unique TLR-driven B-1a responses by regulating the association of these receptors with negative co-receptors, loss of which enhances B-1a-derived antibody production and consequently autoantigen delivery to splenic B cells to drive autoimmunity.

Loss of Gal9 leads to enhanced TLR4 signal transduction in B-1a cells, allowing for enhanced activation and systemic antibody secretion. Similarly, loss of Gal9 decreases the threshold of B cell activation, allowing for B cell responses against low-affinity antigens. We therefore asked if loss of Gal9 would lead to more rapid disease onset in the NZB/W F1 model system for lupus-like disease in mice. To address this, we backcrossed the Gal9KO allele onto the NZB and NZW backgrounds for five generations. We subsequently crossed heterozygous NZB and NZW mice to generate WT and Gal9-deficient NZB/W F1 littermate controls, which develop spontaneous autoimmunity with age. Indeed, we find that Gal9-deficient mice produce anti-dsDNA IgG antibodies earlier in life compared to WT littermate controls (Figure 8A). Furthermore, mice lacking Gal9 expression develop accelerated proteinuria compared to WT controls (Figure 8B). Consistent with these observations, Gal9-deficient mice have a larger population of splenic GC B cells early in disease progression compared to WT littermates (Figure 8C,D). We see a similar increase in the proportion of splenic ASCs in Gal9-deficient mice early in disease onset (Figure 8E,F). These ASCs skew toward B220⁻ long-lived ASCs, and consistent with this, we see enhanced immune complexes in circulation in mice lacking Gal9 (Figure 8G,H). We then asked if loss of Gal9 would lead to an expanded pool of B-1a cells in the peritoneal cavity, as we observed in the C57BL/6 background. Indeed, NZB/W F1 mice deficient in Gal9 have a larger population of peritoneal B-1a cells (Figure 8I). If these expanded B-1a cells are contributing to disease progression, we would expect to see signs of their activation such as migration to secondary lymphoid tissues. Indeed, there is an increased proportion of B-1 cells in the spleen in Gal9-deficient mice (Figure 8J), and this population is enriched for B-1a cells (Figure 8K). Furthermore, loss of Gal9 leads to enhanced production of B-1a-derived PC-specific IgM antibodies detectable in the serum (Figure 8L). Together, these data demonstrate that Gal9 regulates the onset and progression of lupus-like disease in mice. Loss of Gal9 leads to accelerated B cell-driven autoimmunity, marked by GC B cell and ASC formation as well as B-1a cell expansion, migration, and antibody secretion.

Figure 8

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C57BL/6 (WT) mice were obtained from Charles River, Lgals9^–/– (Gal9KO) mice were obtained from Steven Beverley (Washington University) on behalf of The Scripps Research Institute, Nr4a1-eGFP (Nur77-eGFP; C57BL/6-Tg(Nr4a1-EGFP/cre)820Khog/J; Stock No. 016617), NZB/BlNJ (Stock No. 000684), NZW/LacJ (Stock No. 001058), MD4 (C57BL/6-Tg(IghelMD4)4Ccg/J; Stock No. 002595), and µMT (B6.129S2-Ighm^tm1Cgn/J; Stock No. 002288) mice were obtained from The Jackson Laboratory. MD4/ML5 (B6.Ig/sHEL) mice were obtained from Joan Wither (University of Toronto, Krembil Research Institute, University Health Network). Mice were used at 2–6 months of age for all functional and biochemical experiments, and mice were age- and sex-matched within experiments. Mice were used at 8–14 months of age for characterization of spontaneous autoimmunity. Mice were housed in specific pathogen-free animal facility at University of Toronto Scarborough, Toronto, Canada. All procedures were approved by the Local Animal Care Committee at the University of Toronto Scarborough, Animal Use Protocols 20012282 and 20011481.

HEp-2 cells, verified to be contaminated with HeLa cells, were obtained from ATCC and were certified mycoplasma-negative and used acutely. OP9-R7FS cells were kindly provided by Juan Carlos Zúñiga-Pflücker (University of Toronto, Sunnybrook Research Institute) and authenticated by mouse STR profiling by ATCC and mycoplasma tested via MycoAlert Mycoplasma Detection Kit (Lonza). Cells were maintained at 37°C with 5% CO₂ in DMEM containing 10% heat-inactivated fetal bovine serum (FBS, Wisent), 100 U/mL penicillin and streptomycin (Gibco), and 50 µM 2-mercaptoethanol (Amresco).

Lysozymes were isolated from egg whites by ion exchange. Egg whites were diluted in PBS and filtered through 30 kDa filter to restrict protein size (Amicon). Fraction <30 kDa was added to 100 mM ammonium acetate buffer (pH 9.0) and incubated with carboxymethylcellulose (CMC, Sigma-Aldrich). Unbound proteins were washed with ammonium acetate buffer, and lysozymes were then eluted with 400 mM ammonium carbonate buffer (pH 9.2). Isolated lysozymes were diluted in PBS and concentrated using a centrifugation column (Amicon). Purified lysozymes were then mono-biotinylated (Fleire and Batista, 2009) using EZ-Link NHS-LC-LC-Biotin (Thermo Fisher Scientific) according to the manufacturer’s protocol. Mono-biotinylation was confirmed by flow cytometry as described (Fleire and Batista, 2009).

Splenocytes were isolated from mice using a 70 µm cell strainer and phosphate-buffered saline (PBS, pH 7.4, Wisent) to form a single-cell suspension. Bone marrow was isolated by centrifugation of dissected long bones. Peritoneal cells were isolated by peritoneal lavage. B cells were purified using negative magnetic isolation kit (Miltenyi Biotec Inc or Stem Cell Technologies) according to the manufacturer’s protocol.

Primary B cells were cultured in RPMI1640 (Gibco) containing 10% heat-inactivated fetal bovine serum (FBS, Wisent), 100 U/mL penicillin and streptomycin (Gibco), and 50 µM 2-mercaptoethanol (Amresco) at 37°C with 5% CO₂. For activation experiments: cells were stimulated with titrated concentrations of agonists (as indicated) for 16 hr, then stained with fluorescently conjugated antibodies targeting CD86, CD69, and MHCII (BioLegend), and analyzed by flow cytometry. For BCR internalization experiments: primary B cells were stimulated with agonists (as indicated) in RPMI1640 at 37°C. Cells were then fixed at indicated time points in 2% paraformaldehyde stained with fluorescently conjugated antibody Fab fragments targeting IgM (Jackson Immunoresearch) and analyzed by flow cytometry. For signaling experiments: primary B cells were stimulated with agonists (as indicated) in RPMI1640 at 37°C. Cells were fixed at indicated time points in 2% paraformaldehyde and permeabilized on ice in 100% methanol at −20°C. Cells were washed 3× in PBS and then stained for total tyrosine phosphorylation (clone 4G10, Sunnybrook), and cells were subsequently stained with secondary antibodies (Invitrogen) as well as phenotyping markers (BioLegend). Signaling was measured by flow cytometry. To detect internalization of lysozyme, primary MD4 B cells were stimulated with mono-biotinylated lysozyme in RPMI 1640 at 37°C. Cells were fixed in 2% paraformaldehyde, stained with PE-conjugated streptavidin (BioLegend) to occupy mono-biotinylated lysozyme on the cell surface. Cells were then permeabilized in 0.1% Saponin permeabilization buffer (0.1% saponin, 3% FBS, 0.1% sodium azide, in PBS) and stained with Alexa Fluor-633-conjugated streptavidin (Invitrogen) to detect intracellular lysozyme by flow cytometry. For experiments using OP9 stromal cells: OP9 cells were labeled with 5 µM CFSE (Invitrogen). Immune complexes containing mono-biotinylated lysozyme and anti-sera generated in mice were formed in vitro, then incubated with OP9 stromal cells in complete media in suspension, then washed with PBS, and stained with Alexa Fluor-633-conjugated streptavidin (Invitrogen). Amount of lysozyme was determined by flow cytometry using fluorescence quantitation beads according to the manufacturer’s protocol (Bangs Laboratories Inc). Density of molecules was then determined using the average surface area of a cell determined by confocal imaging. OP9 cells were then seeded into wells and centrifuged to draw cells to the bottom. Primary MD4 B cells were added to the top of the wells and allowed to settle onto OP9 cells. Cells were fixed using 2% paraformaldehyde and permeabilized on ice in 100% methanol at −20°C. Cells were washed 3× in PBS and then stained for total tyrosine phosphorylation (clone 4G10, Sunnybrook), followed by secondary antibodies (Invitrogen) as well as anti-B220-Pacific Blue (BioLegend).

Artificial planar lipid bilayers were prepared by spreading liposomes in FCS2 chambers (Bioptechs). 1,2-Dioleoyl-sn-glycero-3-phosphocholine liposomes (Avanti Polar Lipids, Inc) were mixed with 2.5 × 10⁻⁴% biotinylated liposomes (N-Biotinylated Cap PE: Avanti Polar Lipids, Inc). Alexa Fluor 633-streptavidin (1:1000 v/v; Life technologies) was used to tether monobiotinylated lysozymes. Cells were injected into FCS chambers in chamber buffer (0.5% FBS, 2 mM MgCl₂, 0.5 mM CaCl₂, and 1 g/L d-glucose in PBS). Freshly isolated splenocytes from WT and Gal9KO MD4 mice were allowed to spread and interact with the bilayer for 90 s and then fixed with 2% PFA for 15 min at 37°C. Cells were imaged by TIRF microscopy and analyzed using ImageJ. Total contact area and total fluorescence intensities were calculated for each cell. Fluorescence intensity was converted into molecular count using a fluorescence standard (Fleire and Batista, 2009) (Quantitation set, Bangs Laboratories).

HEp-2 cells were grown on glass bottom dishes and fixed with 2% paraformaldehyde. Cells were then stained with sera isolated from mice (1:50) at 4°C. Cells were then washed, and autoantibody binding was detected using secondary anti-mouse IgM or IgG (Jackson ImmunoResearch). Cells were imaged by spinning disc confocal microscopy (Quorum Technologies) consisting of an inverted fluorescence microscope (DM16000B; Leica) equipped with a 63×/1.4 NA oil-immersion objective and an electron-multiplying charge-coupled device (EMCCD) camera (ImagEM; Hamamatsu). Z-section images were collapsed into a Z-projection, and mean fluorescence intensity (MFI) was determined by dividing the total intensity of the field of view by the number of nuclei using ImageJ.

Antibodies were quantified using IgM and IgG quantitation sets (Bethyl Laboratories Inc) according to the manufacturer’s protocol with some adjustments: Thermo Scientific Nunc MaxiSorp 96-well ELISA plates were coated with 50 µg/mL calf thymus DNA (Thermo Fisher Scientific), phosphorylcholine (Sigma-Aldrich), hen egg lysozyme (Sigma Aldrich), glomerulus extract isolated from primary mouse glomeruli lysed in NP40 lysis buffer, or anti-C3 (clone K13/16, BioLegend for immune complex determination) in sodium carbonate coating buffer. Sera was diluted at 1:50 for quantification. Signal was quantified against IgG/IgM standard and reported as absorbance units (AU).

Primary murine B cells were incubated in 20 mM β-lactose or 0.1 µM recombinant Gal9 (R&D Systems) in RPMI1640 at 37°C for 30 min. Cells were washed with PBS and resuspended in RPMI1640. Cells were then stimulated as described above.

Isolated B cells from Gal9KO mice were lysed in NP40 lysis buffer (50 mM HEPES pH 8.0, 150 mM NaCl, 5 mM dithiothreitol, 5 mM EDTA, 0.1% Nonidet-P40, Roche cOmplete mini EDTA-free protease inhibitor, and 1 mM phenylmethylsulfonylfluoride). Lysates were cleared by centrifugation at 21,000 × g for 10 min at 4°C. Cell lysates were incubated with anti-FLAG beads (Sigma-Aldrich) conjugated with 10 µg FLAG-tagged rGal9, kindly provided by the Ostrowski Lab (University of Toronto) for 3 hr at 4°C. Beads were washed with lysis buffer and blocked in 5% horse serum in PBS. Bound proteins were detected by staining with fluorescently conjugated antibodies (BioLegend) and measured by flow cytometry. Anti-FLAG beads alone (without rGal9) were incubated with lysates and served as a negative control of background adsorption.

Primary murine peritoneal cells were sorted into B-1a and B-1b cells using MACS B-1a selection kit (Miltenyi Biotec Inc). Sorted cells were cultured in RPMI1640 containing 10% heat-inactivated fetal bovine serum (FBS, Wisent), 100 U/mL penicillin and streptomycin (Gibco), and 50 µM 2-mercaptoethanol (Amresco) for 3 days with 0.5 µg/mL LPS (Invivogen) and 0.5 µg/mL anti-IgM F(ab’)₂ (Jackson ImmunoResearch). Cultures were collected, and cells were pelleted by centrifugation. Culture supernatants containing secreted antibodies were collected, and antibodies were purified by filtration using a 100 kDa centrifugation filter (retaining the > 100 kDa fraction). Antibody composition was determined by ELISA using mouse IgM/IgG quantitation set (Bethyl Laboratories), according to the manufacturer’s protocol. Total concentration of antibody was determined by BCA assay (Thermo Fisher Scientific).

Mice were transferred 100 µg of total antibody by intraperitoneal injection every 3 days over a 2-week period. At day, 14 mice were euthanized and a single cell suspension was prepared from the spleen using enzymatic digestion (DNAse/Collagenase). Cells were stained with fluorescently labeled antibodies and measured by flow cytometry. Additionally, mice were assessed 4 weeks following the last transfer for signs of autoimmunity as described above.

Primary mouse thymocytes were isolated and formed into a single-cell suspension using a 70 µm cell strainer. Cells were labeled with 10 µm of CFSE and washed as described above. Cells were then cultured for 2 days in RPMI1640 containing 100 U/mL penicillin and streptomycin (Gibco) and 50 µM 2-mercaptoethanol (Amresco) and 5 µg/mL puromycin (Thermo Fisher Scientific) to induce apoptosis. Cultures were collected and washed with PBS; intact cells were pelleted by gentle centrifugation at 300× g for 5 min. Cell supernatants containing ApoB were transferred into mice by intravenous injection.

Mice were injected with 1 µg/mL or 0.05 µg/mL of LPS by intraperitoneal injection to activate B-1 cells in the peritoneal cavity. Mice were euthanized 24 hr after injection, and B-1 cells were analyzed by flow cytometry. Additionally, mice received CFSE-labeled ApoB derived from 2 million initial thymocytes at 12 hr after injection. Mice were then analyzed for ApoB staining on subcapsular sinus macrophages in the spleen, as described above, by flow cytometry.

Protein concentration was determined in the urine by BCA assay (Thermo Fisher Scientific) according to the manufacturer’s protocol.

WT mice received 200 µL of Clophosome-A Anionic Clodronate Liposomes (FormuMax) or control liposomes by intravenous injection to deplete splenic phagocyte populations. Twenty-four hours after injection, splenic B cells were isolated from Gal9KO mice by negative selection (EasySep, Stemcell) and labeled with eFluor 450 dye, according to the manufacturer’s instructions (ThermoFisher). 2 × 10⁶ Gal9KO B cells were transferred by IV injection into control or depleted hosts. Twenty-four hours after adoptive transfer spleens were collected, and Gal9 expression was measured on endogenous and transferred B cells by flow cytometry. Cells were stimulated with titrated concentrations of anti-IgM F(ab’)₂, as above. Depletion of phagocyte populations was determined by flow cytometry.

Gal9KO mice were backcrossed onto NZB and NZW strains for five generations. NZB and NZW mice heterozygous for the Gal9KO allele were crossed to generate NZB/W KO and WT littermate controls. Blood and urine collection was performed biweekly to monitor disease progression. Autoantibody production and proteinuria was quantified as described above.

Thunderstorm plugin for ImageJ is available at http://zitmen.github.io/thunderstorm/ (Ovesny, 2021). DBSCAN algorithm is available in the SMoLR package, and the Hopkins index in the Spatstat package in R, available through Github.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Joan Wither (Krembil Research Institute, University Health Network) for MD4/ML5 mice, Juan Carlos Zúñiga-Pflücker (Sunnybrook Research Institute) for OP9- R7FS cells, and Mario Ostrowski (University of Toronto) for rGal9. This work was supported by funding from the Canadian Institutes of Health Research (CIHR; MOP-136808 and PJT-165938), and Canada Research Chair (CRC; 905–231134) to BT.

Animal experimentation: All procedures were approved by the Local Animal Care Committee at the University of Toronto Scarborough, Animal Use Protocols 20012282 and 20011481.

1. Received: November 2, 2020
2. Preprint posted: November 4, 2020 (view preprint)
3. Accepted: July 5, 2021
4. Version of Record published: August 9, 2021 (version 1)

© 2021, Smith et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.