B cell depletion reduces T cell activation in pancreatic islets in a murine autoimmune diabetes model

Human CD20 (hCD20) transgenic mice on a BALB/c background (in which the human gene MS4A1, encoding hCD20, is expressed) [17] were backcrossed to the NOD genetic background more than ten generations, and designated hCD20/NOD mice [7]. Mice were maintained at Cardiff University in specific pathogen-free isolators or scantainers. Mice received water and food ad libitum and were housed in a 12 h dark–light cycle. Animal experiments were conducted in accordance with United Kingdom Animals (Scientific Procedures) Act, 1986 and associated guidelines.

Mice were monitored weekly for glycosuria (Bayer Diastix) from 12 weeks of age. Diabetes was confirmed by blood glucose levels >13.9 mmol/l.

Female hCD20/NOD mice, aged 6–8 or 12–15 weeks were chosen at random to receive anti-hCD20 antibody (clone 2H7; Bio-XCell [West Lebanon, NH, USA]) or control IgG2b antibody (clone MPC-11; Bio-XCell [7, 14, 15]). An i.v. injection of 500 μg of antibody in 200 μl of saline solution (154 mmol/l NaCl) was followed at 3 day intervals by three i.p. injections (modified from [7]).

Pancreatic lymph nodes (PLNs) were disrupted mechanically with a 30G needle. Bone-marrow cells were flushed out from the hind legs (femur and tibia). Spleens were homogenised and erythrocytes were lysed. Pancreases were inflated with collagenase P solution (Roche, Burgess Hill, UK) in Hanks’ Balanced Salt Solution (HBSS) via the common bile duct, followed by collagenase digestion with shaking at 37°C for 10 min. Islets were isolated by Histopaque density centrifugation (Sigma-Aldrich, Gillingham, UK), hand-picked under a dissecting microscope and trypsinised to generate single-cell suspensions. Islet cells were rested at 37°C, 5% CO₂ in complete Iscove’s Modified Dulbecco’s Medium (IMDM) overnight, before stimulation for intracellular staining.

Single-cell suspensions were incubated with TruStain (anti-mouse CD16/32; Biolegend [London, UK]) for 10 min at 4°C, followed by fluorochrome-conjugated monoclonal antibodies against cell surface markers for 30 min at 4°C. Multivariable flow cytometry was carried out using monoclonal antibodies: CD4-FITC (GK1.5 [1:200]), CD8-PE-594 (53-6.7 [1:800]), CD103-BV510 (2E7 [1:100]), PD-1-BV785 (29F.1A12 [1:200]), IFN-γ-BV711 (XMG1.2 [1:100]), CD107a-PeCy7 (1D4B [1:200]), CD69-AF700 (H1.2F3 [1:100]), TGF-β-BV421 (TW7-16B4 [1:100]), CD38-FITC (90 [1:200]) and CD86-PeCy7 (PO3 [1:200]) (all from Biolegend); IL-10-APC (JES5-16E3 [1:200]), CD1d-BV510 (1B1 [1:200]), CD21-PE-594 (7G6 [1:1000]), CD23-BV711(B3B4 [1:800]), CD24-BV650 (M1/69 [1:400]), CD3-BV786 (145-2C11 [1:100]) and CD80-BV650 (16-10A1 [1:100]) (all from BD Biosciences, Reading, UK); CD19-efluor780 (eBio1D3 [1:800]), CD5-PeCy7 (53-7.3 [1:200]), IL-6-PerCP-Cy5.5 (MP5-20F3 [1:200]) and CD44-PerCP-Cy5.5 (IM7 [1:1600]) (all from eBiosciences, Waltham, MA, USA). All antibodies were titrated before use. Dead cells were excluded from the analysis by Live/Dead exclusion (Invitrogen, Paisley, UK). For intracellular cytokine analysis, splenocytes were either unstimulated or stimulated for 24 h with 5 μg/ml lipopolysaccharide (LPS) (Sigma-Aldrich) or 5 μg/ml anti-CD40 (Bio-XCell) and washed. After an overnight resting period, 3 h before antibody staining, phorbal 12-myristrate-13-acetate (PMA) (50 ng/ml), ionomycin (500 ng/ml) and monensin (3 μg/ml) (all from Sigma-Aldrich) were added to the cells. CD107a antibody was added prior to stimulation, as previously described [18]. Fc receptors were blocked using TruStain and, after extracellular staining, cells were fixed using fixation/permeabilisation kit (BD Biosciences) according to the manufacturer’s instructions and then stained for intracellular cytokines or appropriate isotype controls. Cell suspensions were acquired on LSRFortessa (FACSDIVA software; BD Biosciences). All analysis was performed using Flowjo software (Tree Star, Ashland, OR, USA).

No data were excluded from the analysis. Statistical analysis was performed using GraphPad Prism version 5 (GraphPad Software, San Diego, CA, USA). For islet T cells, multivariable flow cytometric analysis was performed using SPICE (Simplified Presentation of Incredibly Complex Evaluations) version 5.1 (http://​exon.​niaid.​nih.​gov). Comparison of distributions was performed using the Mann–Whitney U test and a partial permutation test [19]. For diabetes incidence, the Gehan–Breslow–Wilcoxon test was used. All other data were analysed by the Mann–Whitney U test.

hCD20/NOD mice were treated with 2H7 or isotype control antibodies at 6–8 weeks (little insulitis) or 12–15 weeks of age (established insulitis) (Fig. 1a). We monitored disease progression in mice that were B cell-depleted at 6–8 weeks old. Diabetes was first seen in the treated mice at 29 weeks of age, delayed by 10 weeks, and incidence was reduced in the 2H7-treated groups (ESM Fig. 1). At the time of diabetes onset in the experimental group, 63% of the control mice that ultimately developed disease were diabetic, although the difference at the termination of the experiment was not statistically significant (p = 0.07). This delayed incidence was in keeping with our previously published observation [7]. B cells were successfully depleted in both age groups and had fully repopulated the spleen by 12 weeks post depletion (Fig. 1b, e). No change was observed in the numbers of CD4 or CD8 T cells during depletion or repopulation (Fig. 1c, d, f, g). B cell depletion in PLNs (Fig. 1i, l), vs spleen (Fig. 1h, k), was prolonged. B cells were not fully depleted in the bone marrow, compared with other lymphoid tissues (Fig. 1j, m).

The various B cell depletion methods target different B cell zones in the spleen [7, 20]. Splenic B cell populations were mostly depleted 24 h after 2H7 treatment (Fig. 2a) and B cell numbers were significantly reduced (Fig. 2b–g). The marginal zone (Fig. 2h, k) and T2 (Fig. 2i, l), enriched in regulatory B cells (Bregs), were more successfully depleted than the follicular zone after anti-CD20 treatment (Fig. 2j, m), indicating that Bregs were not spared during depletion. Follicular zone and T2 B cells repopulated before the marginal zone. At 12 or 30 weeks after B cell depletion, there was no increase in T2 cell numbers (data not shown). This contrasts with our previous findings of increased numbers of T2 cells in older diabetic mice, which became normoglycaemic after B cell depletion with anti-CD20 antibody [7], or in normoglycaemic 30-week-old mice treated with anti-CD22 depleting antibody [8], indicating that Bregs with T2 phenotype were not enriched after B cell repopulation. These differences may be due to the use of younger and non-diabetic mice in our current study.

There were significantly fewer IL-10⁺ B cells in spleens from mice treated with 2H7 vs control antibody, unstimulated or following stimulation with LPS or anti-CD40, at either 8 or 12 weeks post depletion, (Fig. 3b, d). This difference was more marked at 12 weeks, when the B cells were repopulated. Moreover, at 30 weeks post depletion, there were still fewer IL-10⁺ B cells in 2H7-treated mice, unstimulated or stimulated with anti-CD40, than in age-matched control antibody-treated mice (ESM Fig. 2). We observed no enrichment of IL-10⁺ B cells in the marginal zone or T2 compartments (data not shown) or in CD1d^hiCD5⁺ or CD24^hiCD38^hi Breg cells during or after depletion (ESM Fig. 3).

We analysed the TGF-β response, as B cell regulation can occur via TGF-β [21, 22]. In mice aged 6–8 weeks, the number of TGF-β⁺ B cells was significantly increased 8 weeks post depletion, albeit at small percentages overall (Fig. 3c, e). Differences were not maintained at 12 weeks post treatment in either of the age groups. Therefore, anti-CD20 treatment did not promote a sustained Breg phenotype either during or long after repopulation of the spleen, in either of the age groups of mice studied. We examined the proinflammatory cytokine-producing B cell populations. In the younger mice, 2H7 and control antibody treatment produced little change in the number of IL-6⁺ B cells, with or without LPS or anti-CD40 stimulation; these mice had significantly fewer IL-6⁺ B cells at 8 weeks post depletion but at 12 weeks this effect was lost and stimulation with anti-CD40 caused an increase in IL-6⁺ B cells (ESM Fig. 4).

To test the repopulating B cells for altered co-stimulatory potential, we studied the expression of CD86 and CD80 on B cells following stimulation with LPS or anti-CD40 (Fig. 4). 2H7-treated mice in both age groups expressed significantly fewer CD19⁺CD80⁺ B cells when stimulated with anti-CD40 during regeneration at 8 weeks (Fig. 4b, d); this difference was less marked at 12 weeks. More strikingly, significantly fewer B cells expressed CD86 during regeneration at 8 weeks, following the different stimuli used in both age groups of mice (Fig. 4c, e). This reduction was maintained at 12 weeks (when cells were fully repopulated), particularly following stimulation with anti-CD40, in mice aged 6–8 weeks. Unstimulated B cells from 2H7- and control antibody-treated mice at repopulation (12 weeks) were used as antigen-presenting cells in a proliferation assay to stimulate insulin-peptide-reactive CD8⁺ T cells as responders. We observed no difference in carboxyfluorescein succinimidyl ester (CFSE) dilution as a measure of proliferation of these CD8⁺ T cells (data not shown). At 30 weeks of age, whether unstimulated or stimulated, B cells expressing CD86 in both age groups were comparable, although fewer B cells expressed CD80 when stimulated with anti-CD40 in 2H7-treated mice (ESM Fig. 5).

We next investigated the effects of anti-CD20 on local pancreatic islet B cells during B cell depletion and repopulation in mice aged 6–8 and 12–15 weeks. Significantly fewer B cells were isolated from the pancreatic islets of anti-CD20-treated mice in both groups, as early as 24 h post antibody administration (Fig. 5a, d and ESM Fig. 6a). At 12 weeks post treatment, B cells in the islets of both groups had returned to the levels found in the control antibody-treated mice. Consistent with B cells in the spleen, a downregulation of IL-10 was observed (Fig. 5b, e). There were fewer IL-10⁺ B cells in 2H7-treated mice aged 12–15 weeks, at both 8 and 12 weeks post depletion (p < 0.01 at 12 weeks). Unlike splenocytes, we observed no difference in islet TGF-β⁺ B cells following 2H7 vs control antibody treatment (Fig. 5c, f). We further phenotyped islet B cells at 12 and 30 weeks after depletion in mice aged 6–8 weeks (Fig. 5g–l; for representative staining, see ESM Fig. 6b). Multivariable analysis demonstrated that, at both 12 and 30 weeks post depletion, there was no overall change in B cell phenotype or difference between the proportions of the B cell subsets. However, individual populations of IL-10⁺ and TGF-β⁺ B cells were significantly reduced 12 weeks post depletion, confirming our earlier analysis of peripheral cells. At 30 weeks post depletion, no significant difference was seen in the frequencies of the B cells within the islets, confirming that B cells had fully repopulated. However, less CD44 expression was noted, indicating lower activation.

B cell depletion has been shown to induce peripheral regulatory T cells (Tregs) after B cell regeneration. We observed no differences in CD4⁺ or CD8⁺ T cell numbers in mouse islets following anti-CD20 vs control antibody treatment (Fig. 6a–d), at any point during depletion (24 h, 8 weeks and 12 weeks; for representative staining see ESM Fig. 7a) or at 30 weeks post depletion (data not shown). Thus, B cell depletion did not affect T cell homing to the pancreas. However, the effect of anti-CD20 treatment on CD4⁺ and CD8⁺ T cells was significantly greater in islets from mice aged 6–8 weeks vs 12–15 weeks (Fig. 6e–l). In the younger mice, during B cell depletion and regeneration (8 and 12 weeks, respectively) both IFN-γ⁺ (Fig. 6e, g) and, surprisingly, IL-10⁺ (Fig. 6f, h) CD4⁺ T cells were significantly reduced. Islet-infiltrating CD8⁺ T cells also significantly downregulated IFN-γ production during B cell regeneration at 12 weeks (Fig. 6i, k), accompanied by a decrease in CD107a expression (Fig. 6j, l) representing cytotoxic degranulation on stimulation, although this was not statistically significant. Both IFN-γ and CD107a expression in CD8⁺ T cells from 12- to 15-week-old treated mice were comparable.

We examined activation (CD44, CD69, PD-1) and cytokine production (IFN-γ, TGF-β, IL-10) of pancreatic CD4⁺ T cells during B cell repopulation (for representative staining, see ESM Fig. 7b). Islet CD4⁺ T cells from anti-CD20 antibody-treated mice aged 6–8 weeks were examined at 12 weeks and 30 weeks after B cell depletion, when complete B cell regeneration had occurred (Fig. 7). We observed significant differences in CD4⁺ T cell composition at 12 weeks post B cell depletion (p < 0.05) (Fig. 7a, c), although CD4⁺ T cell subsets were comparable at 30 weeks (Fig. 7b, d). The CD4⁺ T cell subsets at 12 weeks post treatment had a less-activated phenotype, characterised by fewer CD44⁺CD4⁺ T cells, which lacked functional expression of IFN-γ; however, there was no enrichment of IL-10⁺ or TGF-β⁺CD4⁺ T cells (Fig. 7e). At 30 weeks post depletion, there were still significantly fewer IFN-γ⁺CD4⁺ T cells in 2H7-treated mice than in control IgG-treated mice (Fig. 7f). Here, the CD69 increase may be a result of altered kinetics and an increase in new CD4⁺ T cells that home to the tissue and encounter a proinflammatory environment.

We next analysed the impact of B cell depletion on local CD8⁺ T cells in the islets of hCD20/NOD mice aged 6–8 weeks, focusing on the islet environment during repopulation (Fig. 8; see ESM Fig. 8 for representative staining). Particularly striking, was the high influx of heterogeneous CD8⁺ T cell subsets into the pancreatic islets after initial damage has occurred [23, 24]. At 12 weeks post depletion, islet-infiltrating CD8⁺ T cells significantly differed in their composition (Fig. 8a, c) and were characterised by a lack of effector function typified by less CD44 expression coupled with decreased IFN-γ and CD107a production, confirming our earlier observations (Fig. 8e) (p < 0.05). As with islet CD4⁺ T cells, we observed an increase in CD69⁺CD8⁺ T cells in the 2H7-treated mice, which may represent naive CD8⁺ T cell subsets being activated locally.

At 30 weeks post treatment, islet CD8⁺ T cell subsets were comparable; although there were fewer activated CD8⁺ T cells, this was not statistically significant (Fig. 8b, d). Interestingly, CD8⁺ T cells expressing CD103 and CD69, which indicate a tissue-resident memory (T_RM) CD8 T cell phenotype and are associated with controlling tissue immunity after infection [25], were enriched in the islets of 2H7-treated mice (Fig. 8f).