No spontaneous immune dysregulation disease in LRBA-deficient mice

In contrast to the morbid immune dysregulation in human LRBA deficiency, LRBA-deficient mice developed at normal frequencies with no obvious morphological abnormalities, normal body weight, normal erythrocyte and platelet counts, and no splenomegaly, whether analysed at 6 weeks of age or at 6 months of age (Figure 2 and Supplementary Figure 2). There was also no lymphadenopathy (not shown). Flow cytometric analysis of the spleen revealed normal numbers of neutrophils, NK cells, B-cell subsets or T-lymphocyte subsets, with no increase in effector/memory T cells or decrease in Tregs as has been observed in LRBA-deficient patients. Based on the low levels of CTLA-4 on Treg and effector T cells, we tested for increased CD86 on the surface of B cells, but found it to be comparable in young mice with or without LRBA (Figure 2d). We also investigated the expression of BTLA, a homologue of CTLA-4, on total B cells and GC B cells. However unlike CTLA-4, BTLA expression was not decreased on lymphocytes from LRBA-deficient mice (Supplementary Figure 2E). Developing subsets of B and T cells in the bone marrow and thymus were present in normal numbers (Figures 2e and f), despite the low expression of CTLA-4 on thymic Tregs.

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Figure 2

Absence of immune dysregulation disease in LRBA-deficient mice. (a–f) Analysis of sex-matched WT (red) and Lrba^−/− (blue) mice at 6 weeks of age, showing individual results and means for each group. (a–c) Flow cytometric analysis of spleen, showing the numbers of: (a) CD4⁺ or CD8⁺ T-cell subsets of central memory (CD62L⁺ CD44⁺), effector memory (CD62L⁻ CD44⁺), naïve (CD62L⁺ CD44⁻) or Tregs (CD4⁺ FOXP3⁺). (b) B cells (B220⁺) and subsets of transitional 1 (T1, B220⁺ CD93⁺ CD23⁻), transitional 2 and 3 (B220⁺ CD93⁺ CD23⁺), marginal zone (B220⁺ CD21⁺ CD23⁻), mature follicular (B220⁺ CD93⁻ CD23⁺) and B-1 (B220⁻ CD19⁺) B cells per spleen. (c) neutrophils (B220⁻ MHC II⁻ Ly6G⁺ CD11b⁺) and NK lymphocytes (B220⁻ CD8⁻MHC II⁻ NK1.1⁺). (d) CD86 mean fluorescence intensity (MFI) on marginal zone and follicular B cells. (e) Flow cytometric analysis of bone marrow, showing % of lymphocytes that are pro-B cells (B220⁺ CD43^int), pre-B cells (B220⁺ CD43⁻ IgM⁻), immature B cells (B220^int CD43⁻ IgM⁺ IgD⁻), transitional B cells (B220^hi CD43 ⁻IgM⁺ CD24⁺ IgD^lo) and mature B cells (B220^hi CD43⁻ IgM⁺ CD24⁻ IgD⁺). (f) Flow cytometric analysis of thymus, showing number of CD8⁻ CD4⁻ double-negative (DN), double-positive (DP), CD8⁺ or CD4⁺ single-positive, or CD4⁺ FoxP3⁺ Treg cells. N=6 per group. Data are representative of one experiment. (g) Titres of IgG2b, IgA, IgG1, IgM, IgG3 Ig2Ga(b) and IgE antibodies in the serum of unimmunised mice, determined by enzyme-linked immunosorbent assay (ELISA). N=10 per group. Representative of two comparable experiments. Statistical analysis was carried out using t-test: **P<0.01.

To investigate potential disease progression over time, mice were also analysed at 6 months of age (Supplementary Figure 2). Older LRBA-deficient mice appeared normal in regards to their total body weight and frequencies of red blood cells, platelets, neutrophils, NK cells and B- and T-cell subsets and development. Interestingly, we did notice a slight but significant decrease in B220^lo CD19⁺ B-1 cells within the spleen that was not evident in the younger mice (Supplementary Figures 2L and M).

LRBA mice show altered frequencies in the serum of antibody isotypes

Given hypogammaglobulinemia is a frequent clinical feature of human Lrba deficiency, we measured basal Ig levels in the serum of Lrba^−/− and WT mice. This revealed the serum of unimmunised Lrba^−/− mice to contain significantly higher levels of IgG2b than age- and sex-matched WT mice (Figure 2g). There was also a trend towards an increase in IgM levels, although this was not significant (P=0.056).

Peritoneal B-1 cells are reduced in Lrba^−/− mice

Given the finding that LRBA-deficient mice aged >6 months of age have a reduced frequency of splenic B-1 cells and in light of the chronic diarrhoea and IBD occurring in 60% of LRBA-deficient patients, we tested for cellular abnormalities in the peritoneal fluid of LRBA-deficient mice. This revealed a significant relative decrease in B-1 cells (CD19⁺ B220^lo IgM⁺ CD23⁻) with no corresponding decrease in B-2 cells (CD19⁻ B220^hi) in both 6-week-old and 6-month-old LRBA-deficient mice (Figure 3). This decrease was largely due to decreased B-1a cells (CD19⁺ B220^lo IgM⁺ CD23⁻ CD5⁺). Interestingly, T cells were increased relative to other cell types within the peritoneal compartment in young and aged LRBA-deficient mice, although this could have been secondary to decreased B-1 cells.

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Figure 3

Decreased peritoneal B-1 B cells in LRBA-deficient mice. Flow cytometric analysis of peritoneal cavity lymphocytes from sex-matched WT (red) and Lrba^−/− (blue) mice at 6 (n=6) or 26 (n=4) weeks of age, showing individual results and means for each group. (a and b) Percentage of lymphocytes that are T cells (B220⁻ IgM⁻ CD23⁻ CD5⁺), B-2 cells (CD19⁻B220^hi), B-1 cells (CD19⁺ B220^lo IgM⁺ CD23⁻), B-1a cells (CD19⁺ B220^lo IgM⁺ CD23⁻ CD5⁺) and B-1b cells (CD19⁺ B220^lo IgM⁺ CD23⁻ CD5⁻). (c and d) Representative flow cytometric plots showing gating IgM, CD23, B220 and CD19 staining for B-1 and B-2 cells and representative CD5 histograms of B-1 cells showing gates on CD5+ B-1a and CD5- B-1b cells from mice at 6 (c) and 26 (d) weeks of age. Statistical analysis was carried out using t-test: *P<0.05; **P<0.01. Data are representative of one experiment.

Primary effects of LRBA deficiency within different subsets of lymphocytes after bone marrow transplantation

To investigate the possibility that homeostatic compensation mechanisms mask cell-autonomous deficits in LRBA-deficient T or B cells, mixtures of Lrba^−/− and WT bone marrow distinguished by CD45.2 and CD45.1 congenic markers were transplanted to lymphocyte-deficient Rag1^−/− mice. The marrow transplant competitively reconstituted the immune system with a mixture of cells with and without Lrba expression. A control group of recipients were transplanted with marrow mixtures where both CD45.2 and CD45.1 cells had normal Lrba expression. A third group of recipients were reconstituted with 100% CD45.2 marrow that was either Lrba^−/− or Lrba^+/+. The resulting chimeric animals were immunised repeatedly with sheep red blood cells (SRBCs), and blood lymphocytes were analysed by flow cytometry. Despite competing with CD45.1 WT cells, Lrba^−/− CD45.2⁺ cells accounted for a high percentage of B cells, CD4⁺ T cells, CD8⁺ T cells, effector CD4⁺ T cells, effector CD8⁺ T cells and Treg cells (Figure 4a). The contribution of LRBA-deficient CD45.2⁺ cells to each subset was equivalent to the contribution of WT CD45.2⁺ cells in control mixed chimera animals analysed at the same time (Figure 4a), and this was true for blood samples collected at multiple timepoints before and after repeated SRBC immunisation (Supplementary Figures 3 and 4).

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Figure 4

Cell-autonomous loss of CTLA-4 on LRBA-deficient T cells in bone marrow chimeras. Irradiated Rag1^−/− recipient mice were transplanted with a bone marrow mixture comprising 50% CD45.1⁺ Lrba^+/+ cells and 50% CD45.2⁺ cells that were either Lrba^−/− or Lrba^+/+ (WT). One hundred percent CD45.2⁺ Lrba^−/− or Lrba^+/+ (WT) marrow was transplanted into a parallel group of Rag1^−/− recipients. At 8 weeks after transplantation, the chimeric mice were immunised with SRBCs three times 3 days apart and blood was analysed by flow cytometry 62 days after the first immunisation. Lines connect paired values for CD45.1 and CD45.2 cells in individual chimeric mice. (a) Percentage of CD45.2⁺ Lrba^−/− (blue) or Lrba^+/+ (WT, red) cells among the indicated lymphocyte subsets in individual mixed chimeric mice and arithmetic means: B cells (B220⁺), T cells (CD3⁺), CD4⁺ T cells, effector CD4⁺ T cells (CD25⁺, CD44^hi), CD8⁺ T cells, effector CD8⁺ T cells (CD25⁺, CD44^hi) and Tregs (CD4⁺FOXP3⁺). (b–e) Analysis of CD4⁺ FOXP3⁻ cells. (b) Representative plots of intracellular CTLA-4 and CD44 gated on CD45.1⁺ (WT) or CD45.2⁺ (WT or Lrba^−/−) CD4⁺ FOXP3⁻ cells. (c–e) Analysis of CTLA4⁺ CD44⁺ CD4⁺ FOXP3⁻ cells, showing: (c) percentage among the CD45.1⁺ or CD45.2⁺ subsets of total T cells; (d) representative CTLA-4 histograms; (e) CTLA-4 MFI values. Dotted lines represent CD45.2+ (Lrba^+/+ or Lrba^−/−) cells in mixed chimeras; solid lines represent equivalent cells in 100% Lrba^+/+ or Lrba^−/− chimeras. (f–j) Analysis of CD4⁺ FOXP3⁺ cells. (f) Representative flow cytometric plots of CD45.2⁺ CD4⁺ cells, showing % Tregs. (g) CTLA-4 MFI values for each chimeric mouse. (h) Representative intracellular CTLA-4 histograms. (i) Representative intracellular FOXP3 histograms. (j) Representative CD25 histograms. Statistical analysis was carried out using t-test between mice or paired t-test when cells were from the same chimeric mouse: *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001. N=5 per group. Data are representative of one experiment.

Despite normal contribution of LRBA-deficient CD45.2⁺ cells to the CD4⁺ memory/effector and Treg cell populations, they exhibited a selective and marked decrease in the total (intracellular and surface) cellular pool of CTLA-4. Within FOXP3⁻ CD4⁺ cells, CTLA-4 was highly expressed in a subset of CD44^hi effector/memory cells in Lrba^+/+ T-cell populations, but CTLA-4⁺ cells were 80% less frequent among CD45.2⁺ Lrba^−/− T cells (Figures 4b and c). Furthermore, in the CTLA-4⁺ CD44^hi cells that were present, these expressed CTLA-4 at mean intensities that were only 40% of CD45.1⁺ internal control cells and of WT CD45.2⁺ cells in control mixed chimeras (Figures 4d and e). Among FOXP3⁺ CD4⁺ Treg cells, CD45.2⁺ Lrba^−/− Tregs expressed CTLA-4 at mean intensities that were only 10% of CD45.1⁺ internal control cells or of WT CD45.2⁺ cells in control mixed chimeras (Figures 4g and h). By contrast, there was no decrease in FOXP3 or CD25 in CD45.2⁺ Lrba^−/− Tregs (Figures 4i and j). These results demonstrate that the loss of CTLA-4 protein caused by LRBA deficiency is cell autonomous within Treg and helper T cells. The loss of CTLA-4 was similar in chimeras where only half the Tregs were LRBA deficient and in chimeric mice where all cells were defective (Figures 4e and h).

Potent immune activation by xenoantigens or virus infection fails to precipitate immune dysregulation in LRBA-deficient mice

The experiments above showed loss of normal CTLA-4 protein caused by LRBA deficiency, but it was surprisingly not accompanied by the morbid immune dysregulation observed in CTLA-4-knockout mice.^{21, 22} We hypothesised that the residual CTLA-4 in LRBA-deficient Tregs might become insufficient in the face of potent inducers of CD80 and CD86 on dendritic cells and B cells. We first tested this possibility by repeatedly injecting SRBCs, which induces potent dendritic cell activation, CD86 induction, follicular helper T-cell and GC B-cell proliferation because they carry numerous xenoantigens and sheep CD47 does not bind mouse SIRPα and fails to deliver the inhibitory signal carried by healthy self-erythrocytes.²³ In the experiments above, a parallel set of Rag1^−/− recipients were transplanted with 100% Lrba^−/− or Lrba^+/+ CD45.2 bone marrow, so that all T cells were LRBA deficient or sufficient. CTLA-4 was decreased on the Lrba^−/− Tregs to 30% of the levels on Lrba^+/+ Tregs. Despite the loss of normal CTLA-4 on 100% of the Tregs, there was no evidence of immune dysregulation syndrome and no increased formation of CD44^hi CD4 or CD8 effector/memory T cells either before or following three sequential immunisations with SRBCs (Supplementary Figure 4B).

To test if a systemic viral infection would precipitate immune dysregulation, Lrba^−/− mice and age- and sex-matched WT control mice were infected with lymphocytic choriomeningitis virus (LCMV) clone 13, which multiplies to high viral titres throughout the body peaking within 7 days, establishes a chronic viral infection and induces powerful innate and adaptive immune activation during the acute and chronic phase of the infection. Following infection, Lrba^−/− mice exhibited no visible evidence of morbidity and were indistinguishable from their WT counterparts. Measured by flow cytometry, the immune response to LCMV infection 20 days after infection was remarkably normal, with no significant difference in the percentage of CD4⁺ T cells that were effector/memory cells (Figure 5a) nor in the percentage of CD4⁺ or CD8⁺ T cells that bound major histocompatibility complex (MHC) tetramers containing the dominant viral peptides (Figures 5b and c). Rather than developing autoimmunity or exaggerated formation of effector T cells, we found that Lrba^−/− mice actually showed subtle signs of increased T-cell exhaustion compared with their WT counterparts. This included a slight but significant decrease in effector CD8⁺ T cells, and changes in markers of T-cell exhaustion²⁴ on viral peptide/MHC tetramer-binding T cells including increased TIM3 and decreased Ly6C (Figures 5b and c), slightly decreased mean interferon-γ per cell and decreased the proportion of tumour necrosis factor-α-positive cells (Figures 5d and e). Nevertheless, the key finding from these experiments is that LRBA-deficient mice mount a remarkably normal immune response to a chronic, systemic viral infection.

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Figure 5

Response of LRBA-deficient mice to chronic systemic viral infection. C57BL/6 Lrba^+/+ (WT) (red) or Lrba^−/− mice (blue) were infected intravenously with 2 × 10⁶ plaque-forming unit (PFU) LCMV clone 13 to induce chronic viral infection. At 20 days after infection, spleen cells were analysed by flow cytometry. (a) Analysis plots of spleens, showing % of lymphocytes that are CD44^hi effector/memory cells in individual mice with arithmetic means. (b and c) Representative flow cytometric plots (b) and anlaysis plots (c) showing % of CD8⁺ T cells binding MHC I tetramers loaded with LCMV peptides GP_33–41 or NP_396–404, or the % of CD4⁺ cells binding MHC II tetramers fused with LCMV peptide GP_66–77 in individual mice with arithmetic means. (d) Representative histograms showing TIM3 staining on NP_396–404 MHC I tetramer-binding CD8⁺ T cells in WT and Lrba^−/− mice, and MFIs in individual animals for TIM3, CD160 and PD-1. Similar trends were seen with the GP_33–41 tetramer. (e) Representative LY6C staining on GP_66–77 MHC II tetramer-binding CD4⁺ T cells, and MFI’s in individual animals for LY6C, PSGL1 and PD-1. (f) IFNγ⁺ MFI of the IFNγ⁺ T cells (top) and % TNFα⁺ within the IFNγ⁺ cells. Representative IFNγ and TNFα histograms are shown for the NP396 peptide-stimulated cells. Statistical analysis was carried out using t-test: *P<0.05; **P<0.01. N=5 per group. Data are representative of one experiment. IFNγ, interferon-γ.

LRBA-deficient B cells undergo normal affinity maturation

Given the decreased frequency of switched memory B cells in the majority of LRBA-deficient patients, we investigated whether or not LRBA-deficient B cells would respond normally to antigenic stimulation in vivo. To do this, Lrba^−/− C57BL/6 mice were crossed to SW_HEL IgH knock-in transgenic mice where many B cells express the Hy10 antibody against hen egg lysozyme (HEL).^{25, 26} CD45.1 congenic mice with normal LRBA were injected intravenously with 30000 HEL-binding spleen cells from CD45.2 SW_HEL donor mice, where the donors were either Lrba^−/− or Lrba^+/+. On days 0 and 4 after receiving the donor SW_HEL B cells, the recipient mice were immunised with SRBCs that had been covalently decorated with HEL^3X, a triply mutant HEL protein that binds with low affinity to HyHEL10 on the SW_HEL B cells.^{25, 26} Flow cytometric analysis of the spleen on day 15 revealed total GC B-cell numbers to be equivalent in recipients of LRBA-deficient or WT SW_HEL B cells (Figure 6a). Unexpectedly, given the human phenotype of defective B-cell responses, we found a slight but significant increase in the total number of donor-derived, antigen-specific SW_HEL GC B cells when these cells lacked LRBA (Figure 6b). In other respects, there was no significant difference between WT and LRBA-deficient SW_HEL GC B cells in the fraction that bound low concentrations of HEL^3X antigen (Figure 6c), had switched to IgG1 (Figure 6d) or exhibited a CD86⁺ CXCR4⁻ light-zone phenotype (Figure 6e). The mean CD86 on the donor-derived SW_HEL GC B cells was also equivalent in the two groups (Figure 6f).

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Figure 6

Normal GC formation and affinity maturation by LRBA-deficient B cells. CD45.1 congenic C57BL/6 recipient mice, with WT LRBA, were injected intravenously with 30000 HyHEL10⁺ SW_HEL spleen B cells from Lrba^−/− or Lrba^+/+ C57BL/6 donor mice. The recipient mice were immunised two times on days 0 and 4 after B-cell transfer with HEL^3X-SRBC, or unconjugated SRBC for a control group of recipients, and spleen cells were analysed by flow cytometry, sorting and single-cell Igh sequencing on day 15. (a) Total Fas^hi CD38⁻ B220⁺ GC B cells per spleen of individual mice, and arithmetic mean for each group. (b) Donor-derived HyHEL10⁺ CD45.2⁺ CD45.1⁻ GC B cells per spleen. (c) Affinity-matured cells, measured as % donor-derived GC B cells stained brightly with 200ng/ml HEL^3X. (d) IgG1-switched cells, measured as % of donor-derived GC B cells. (e) Light-zone CD86⁺ CXCR4⁻ GC B cells, measured as % of donor-derived GC B cells. (f) CD86 MFI on donor-derived GC B cells. (g) Number of VDJ_H amino-acid changing or silent nucleotide substitutions per donor-derived GC B cell. (h) Percentage of donor-derived GC B cells with affinity-increasing VDJ_H mutations S31R, Y53D or Y58F. (i) Percentage of donor-derived GC B cells with substitutions at each VDJ_H amino-acid position. (j) Co-occurrence of S31R, Y53D and Y58F mutations (rows) in individual cells (columns) sorted from separate recipient mice (boxes). Data are pooled from two independent experiments with comparable results. N=9 mice per HEL^3X-immunised group and four unconjugated controls. Statistical analysis was carried out using t-test: *P<0.05.

To investigate affinity maturation of LRBA-deficient cells in more detail, donor-derived SW_HEL GC B cells from both groups of mice were individually sorted and the Hy10 VDJ_H exon sequenced. There was a slight but significant increase in the mean number of amino-acid substitutions per cell in LRBA-deficient GC cells with no corresponding increase in silent nucleotide changes (Figure 6g). The pattern of amino-acid substitutions was equivalent between the two groups, including comparable percentages of cells that had acquired known HEL^3X affinity-increasing mutations²⁷ (Figures 6h–j). Thus, LRBA-deficient B cells have no discernable deficit in T-cell-dependent activation, GC accumulation, isotype switching or affinity maturation when transplanted into mice with normal LRBA.

Bone marrow chimeras and SRBC immunisation

Recipient C57BL/6 Rag1^−/− mice 8–12 weeks old were irradiated with 425cGy using an X-RAD 320 Biological Irradiator (Precision X-Ray, North Branford, CT, USA). Donor bone marrow was aspirated from femurs, humeri and tibia into B-cell medium comprising RPMI (Gibco, Carlsbad, CA, USA) with 10% heat-inactivated foetal calf serum (Gibco), 2mM l-glutamine and 100U/ml penicillin RPMI media (Gibco). At 15h after irradiation, recipient mice were transplanted with an intravenous injection of 5–10 × 10⁶ bone marrow cells, comprising a mixture of 50% from CD45.1 congenic C57BL/6 mice and 50% from C57BL6 (CD45.2) mice that were either Lrba^−/− or Lrba^+/+.

Unmanipulated mice 8 weeks old, and bone marrow chimeras 8 weeks after marrow transplantation, were immunised with 2 × 10⁸ SRBCs given intravenously.

Recombinant HEL proteins

Recombinant HEL^3X were made as secreted proteins in Pichia pastoris yeast (Invitrogen, Carlsbad, CA, USA) and purified from culture supernatants by ion exchange chromatography as described previously.^{26, 27, 45} Proteins were stored in phosphate-buffered saline at 1–2.5mgml⁻¹ at −80°C. Before use samples were thawed and stored at 4°C for a maximum of 8 months. Upon thawing, protein concentrations were determined by spectrophotometry at 280nm.

SRBC conjugation and transfer

HEL proteins were desalted into Conjugation buffer (distilled water with 0.35m d-mannitol (Sigma, St Louis, MO, USA) and 0.01m sodium chloride (Sigma)). For this process, PD-10 columns (Amersham, Piscataway, NJ, USA) were equilibrated with 30ml Conjugation buffer. One hundred micrograms of protein was loaded onto each column and pushed through the column using 2.5ml Conjugation buffer. For elution of the protein 3.5ml Conjugation buffer was added and the HEL protein was collected as fractions in the following volumes: 250, 1000, 250, 250 and 250μl. Protein concentrations of each fraction were determined by spectrophotometry.

For conjugation, SRBCs were washed in 30ml of phosphate-buffered saline per 6–8 × 10⁹ cells and then once in the Conjugation buffer. SRBCs were then resuspended in a final volume of 1000μl conjugation buffer in a 50ml Falcon tube containing 10μgml⁻¹ of HEL^3X. The solution was mixed on a platform rocker on ice for 10min. One hundred microliters of 100mgml⁻¹ N-(3-dimethylaminopropyl)-N-ethylcarbodimide hydrochloride (Sigma) was then added and the solution was mixed for a further 30min on ice. Confirmation of successful conjugation was performed by flow cytometric analysis of SRBC using AlexaFluor 647-conjugated HyHEL9 antibody (generated in-house).

Intravenous transfers of 3 × 10⁴ SW_HEL B cells per recipient mouse together with 2 × 10⁸ HEL^3X SRBC as described previously.²⁶

Haematology and flow cytometry

Red blood cells and platelet numbers were determined in blood collected from the tail-vein into EDTA (Sarstedt, North Rhine-Westphalia, Nümbrecht, Germany) tubes and analysed on a Sysmex XT-2000iV automated haematology analyser, Kobe, Hyōgo Prefecture, Japan.

On the day of harvest organs were collected into B-cell medium, cell suspensions passed through a 70μm cell strainer (Falcon, Corning, NY, USA). Fc receptors were blocked with unlabelled anti-CD16/32 (eBioscience, San Diego, CA, USA) before staining. To detect HEL^3X-binding cells, cells were stained with 200ngml⁻¹ HEL^3X, followed by AlexaFluor 647-conjugated HyHEL9.

Anti-IgG1-FITC (BD Pharminigen, San Diego, CA, USA) stains were followed by 5% mouse serum before staining for other surface molecules. CD4-BV786 (BD Pharminigen), CD8-APCCy7 (BD Pharminigen), CD62L-PerCPCy5.5 (BD Pharminigen) CD44-FITC (BD Pharminigen) and CD25-PE (BD Pharminigen) were used as surface stains. For the intracellular stains, CTLA-4-APC (eBioscience) and FOXP3-EF450 (eBioscience) cells were first permeabilised using FOXP3 Staining Permeabilisation Kit (eBioscience) according to the manufacturer's instructions. Cells were filtered using 35μm filter round-bottom FACS tubes (BD Pharminigen) immediately before data acquisition on an LSR II analyser (BD Pharminigen).

Cytometer files were analysed with the FlowJo Software (FlowJo LLC, Ashland, OR, USA).

Single-cell FACS sorting

Cell suspensions were prepared and GC B cells were identified using flow cytometry. Single-cell sorting into 96-well plates (Thermo Fisher Scientific, Boston, MA, USA) was performed on the FACSAria or FACSAriaIII (BD Pharminigen). B cells from each mouse were analysed individually to ensure that over-representation of one particular clone did not affect the mutation analysis. The VDJ_H exon of the Hy10 heavy-chain gene was amplified from genomic DNA by PCR, sequenced and analysed as described previously.²⁷

Enzyme-linked immunosorbent assay

High-binding plates (Corning, NY, USA) were coated with the indicated Ig isotypes at 5μgml⁻¹ (IgG2b (BD Pharminigen), clone: R9-91; IgA (BD Pharminigen), clone: C10-3; IgG1 (BD Pharminigen), clone: A85-1; IgM (BD Pharminigen), clone: 11/41; IgG3 (BD Pharminigen), clone: R2-38; IgG2a(b) (BD Pharminigen), clone: R11-89; IgE (BD Pharminigen), clone: R35-72). Bound serum antibody was quantified using Igκ (BD Pharminigen). Antibody levels were quantified against isotype-specific standards (IgG2b BD, IgA BD, IgG1 BD, IgM BD, IgG3 BD, IgG2a(b) (Southern Biotech, Birmingham, AL, USA) and IgE (BioLegend, San Diego, CA, USA)).

Viral infection

Mice were infected intravenously with 2 × 10⁶ plaque-forming units of LCMV clone 13. T-cell stimulation with LCMV peptide, tetramer staining, surface staining and intracellular cytokine staining were performed as described previously.^{47, 48, 49} MHC I LCMV tetramers were purchased from the Biomolecular Resource Facility, JCSMR, ANU, ACT, Australia, while the MHC II LCMV GP_66–77 tetramer was obtained from the NIH Tetramer Core Facility (Emory University, Atlanta, GA, USA).

We thank the Garvan Institute ABR, GMG and Flow Cytometry facilities for expert animal husbandry, genotyping and cell sorting. This work was supported by NHMRC Project Grant 1108800, NHMRC Program Grants 1016953 and 1113904, NIH Grant U19 AI100627 and NHMRC Fellowship 1081858, and by the Ritchie Family Foundation.