1. Immunology and Inflammation
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TLR induces reorganization of the IgM-BCR complex regulating murine B-1 cell responses to infections

  1. Hannah P Savage
  2. Kathrin Kläsener
  3. Fauna L Smith
  4. Zheng Luo
  5. Michael Reth
  6. Nicole Baumgarth  Is a corresponding author
  1. University of California, Davis, United States
  2. University of Freiburg, Germany
  3. Institute of Biology III at the Faculty of Biology of the University of Freiburg, Germany
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Cite this article as: eLife 2019;8:e46997 doi: 10.7554/eLife.46997

Abstract

In mice, neonatally-developing, self-reactive B-1 cells generate steady levels of natural antibodies throughout life. B-1 cells can, however, also rapidly respond to infections with increased local antibody production. The mechanisms regulating these two seemingly very distinct functions are poorly understood, but have been linked to expression of CD5, an inhibitor of BCR-signaling. Here we demonstrate that TLR-mediated activation of CD5+ B-1 cells induced the rapid reorganization of the IgM-BCR complex, leading to the eventual loss of CD5 expression, and a concomitant increase in BCR-downstream signaling, both in vitro and in vivo after infections of mice with influenza virus and Salmonella typhimurium. Both, initial CD5 expression and TLR-mediated stimulation, were required for the differentiation of B-1 cells to IgM-producing plasmablasts after infections. Thus, TLR-mediated signals support participation of B-1 cells in immune defense via BCR-complex reorganization.

https://doi.org/10.7554/eLife.46997.001

Introduction

During lymphocyte development (self)-antigen binding by the TCR and BCR results in negative selection, leading to the removal of strongly self-reactive lymphocytes from the T and B cell repertoire. Depending on the strengths of these antigen-BCR interactions, self-reactive B cells undergo deletion, receptor-editing, or they become anergic, that is unresponsive to antigen-receptor engagement (Melchers, 2015).

Self-reactive, anergic bone marrow-derived B cells up-regulate expression of the signaling inhibitor CD5 (Hippen et al., 2000). On developing T cells, the levels of CD5 expression correlate with TCR signaling intensity encountered during thymic development, with those most strongly binding to self-antigens expressing the highest levels of CD5 (Punt et al., 1994; Azzam et al., 1998). While several ligands have been proposed for CD5 (Punt et al., 1994; Berland and Wortis, 2002; Biancone et al., 1996; Bikah et al., 1998; Brown and Lacey, 2010; Calvo et al., 1999; Pospisil et al., 1996; Van de Velde et al., 1991), none seem to have significant CD5-dependent biological effects. Instead, CD5 expression seems to directly reduce antigen-receptor signaling (Hippen et al., 2000; Punt et al., 1994; Azzam et al., 1998; Perez-Villar et al., 1999). Thus, CD5 seems to act primarily as a component of the antigen-receptor complex, directly modulating TCR and BCR signaling.

In addition to anergic B cells, most B-1 cells also express CD5. In contrast to lymphocytes developing postnatal, these primarily fetal and neonatal-derived B cells (Yuan et al., 2012; Zhou et al., 2015; Hardy and Hayakawa, 2001) seem to undergo a positive selection step during development, requiring self-antigen recognition and strong BCR signaling. The lack of self-antigen expression, or the deletion of co-stimulatory molecules that enhance BCR signaling, diminished B-1 cell development, while deletion of negative co-stimulatory signals, or enhanced surface expression of the BCR, resulted in enhanced B-1a cell development (Berland and Wortis, 2002; Hardy and Hayakawa, 2001; Casola et al., 2004; Hayakawa and Hardy, 2000; Nguyen et al., 2017). Specificity of CD5+ B-1 cells for self-antigens and self-antigen binding during development is consistent with their known self-reactive BCR repertoire (Hayakawa et al., 1999; Lalor and Morahan, 1990; Mercolino et al., 1988; Yang et al., 2015) and thus a role for CD5 in silencing B-1 cell responses to BCR-engagement in order to avert autoimmune responses.

Yet, not all B-1 cells express CD5, instead depending on their expression or not of CD5, they are typically divided into two subsets, B-1a and B-1b, respectively. In contrast to B-1b cells, and consistent with their expression of CD5, B-1a cells do not proliferate in response to BCR stimulation (Morris and Rothstein, 1993). However, in CD5-/- mice and in mice in which the association of membrane IgM with CD5 was inhibited, mature B-1 cells demonstrated a proliferative response similar to that of conventional B (B-2) cells (Tarakhovsky et al., 1994; Bikah et al., 1996), further confirming that CD5 expression reduces B-1 cell responsiveness to BCR-signaling.

A BCR-signaling independent response of B-1 cells might be inferred from the fact that B-1 cells strongly respond to innate, TLR-mediated signals, such as LPS, and that they are a major source for ‘natural’ self-reactive IgM. Moreover, steady-state secretion of natural IgM does not appear to require external antigenic stimulation, as total serum levels of natural IgM and frequencies of IgM-secreting B-1 cells are similar in mice held under both, SPF and germfree housing conditions (Baumgarth et al., 2015; Ochsenbein et al., 1999). However, natural IgM production is not stochastic, but instead likely driven by expression of self-antigens. This was demonstrated by Hayakawa et al, who showed a lack of anti-Thy-1 self-reactive IgM antibodies in the serum of Thy-1-deficient but not Thy-1 expressing mice (Hayakawa et al., 1999; Hayakawa et al., 2003), as well as repertoire studies by Yang et al, which showed selective and extensive clonal expansion of certain CD5+ B-1 cell clones during the first 6 months of life, including in germfree mice (Yang et al., 2015).

Furthermore, B-1 cells can be also actively involved in immune responses to various pathogens (Haas et al., 2005; Alugupalli et al., 2003; Alugupalli et al., 2004; Gil-Cruz et al., 2009; Choi and Baumgarth, 2008). Given that CD5 is a BCR inhibitor, it was suggested that CD5- B-1b cells, but not B-1a cells, respond to pathogen encounters in an antigen-dependent manner. Haas and colleagues, conducting studies in CD19-deficient mice that lack B-1a development, concluded that B-1a cells are responsible for natural IgM secretion, while only the B-1b cells responded to Streptococcus pneumonia infection (Haas et al., 2005). Similarly, CD5- B-1b cells were shown to expand and secrete protective IgM after infection with Borrelia hermsii and Salmonella typhimurium (Alugupalli et al., 2003; Alugupalli et al., 2004; Gil-Cruz et al., 2009).

This model of a ‘division of labor’ between B-1a and B-1b cells leaves the B-1 cell response to influenza infection as an outlier. Chimeric mice reconstituted with either allotypically-marked CD5+ or CD5- B-1 cells showed that only CD5+ B-1 cells were responding in vivo to influenza infection with migration from the pleural cavity to the draining mediastinal lymph nodes (MedLN) in a Type I IFN-dependent process, where they differentiated into IgM-secreting cells (Choi and Baumgarth, 2008; Waffarn et al., 2015). The reasons for the apparent different behaviors of CD5+ and CD5- B-1 cells in the various infectious disease models are unexplained. Furthermore, it is unclear how B-1 cells expressing CD5 can participate in antigen-specific immune responses.

This study addresses some of these questions and reconciles previous divergent findings on B-1 cell responses to infections by demonstrating that only CD5+ B-1 cells respond to influenza virus as well as Salmonella infections, but that once activated, these B-1 cells lose expression of CD5 and thus become ‘B-1b’ like. Mechanistically, the downregulation of CD5 requires expression of TLR, triggering of which resulted in the reorganization of the IgM-BCR complex. BCR reorganization led to the rapid dissociation, and then eventual loss of CD5 from the complex, and triggered enhanced IgM-CD19 and CD79:Syk interactions, resulting in enhanced down-stream BCR-signaling. Thus, TLR-mediated signals support participation of B-1 cells in immune defense via BCR-complex reorganization, linking innate and adaptive antigen-recognition by B-1 cells.

Results

CD5 negative B-1 cells are responsible for local IgM secretion after influenza infection

We previously identified three populations of cells involved in natural IgM secretion: CD5+ B-1 cells, CD5- B-1 cells, and plasma cells, the latter are CD19- and CD138/Blimp-1+ (Savage et al., 2017) and also B-1-derived (B-1PC) (Savage et al., 2017). This was shown using a ‘neonatal chimera’ model, in which host B-1 cells are replaced in neonatal host mice by congenic but Ig-allotype-disparate donor B-1 cells, while the host B-2 cells remain of the host and thus its allotype (Lalor et al., 1989). After full reconstitution B-1 cells as well as their secreted IgM can be identified and quantified using allotype-specific anti-IgM (and anti-IgD) antibodies. Because B-1-derived IgM is important for protection from lethal influenza infection (Baumgarth et al., 2000), we sought to determine which B-1 cell populations generate IgM in the draining (mediastinal) lymph nodes (MedLN) after influenza infection (Choi and Baumgarth, 2008).

Examination of the MedLN of neonatal chimeras showed that B-1 cells migrated to MedLN and then rapidly differentiated to IgM-secreting B-1PC on day seven after infection with influenza A Puerto Rico 8/34 (A/PR8) (Figure 1A). Neonatal chimeric mice generated with B-1 donor cells from Blimp-1 YFP reporter mice (Fooksman et al., 2010; Rutishauser et al., 2009) confirmed the presence of Blimp-1-YFP+ B-1PC in the MedLN (Figure 1B). The MedLN B-1PC mostly lacked expression of CD5, particularly among the Blimp-1hi cells (Figure 1C). Also, the CD5+ Blimp-1-YFP+ cells expressed less Blimp-1-YFP than the CD5- Blimp-1-YFP+ B-1 cells (Figure 1C, left). The data were unexpected, as we had shown previously that only the CD19+CD43+CD5+but not the CD5- B-1 cells were able to migrate from the pleural cavity to the MedLN after influenza infection, where they differentiated into IgM-secreting cells (Choi and Baumgarth, 2008; Waffarn et al., 2015).

CD5 negative B-1 cells secrete most IgM in the mediastinal lymph nodes (MedLN) after influenza infection.

(A) FACS plot of MedLN cells from day seven influenza-A/PR8-infected neonatal chimeric mice generated with Ighb B-1 donor cells and Igha host cells. Shown is gating to identify IgMb+CD19+B-1 cells and IgMb+CD19 CD138+B-1PC. FMO, ‘fluorescence minus one’ control stains. (B) Mean number ± SD of Blimp YFP+ cells in peripheral LN (PLN) and MedLN of day seven influenza-infected neonatal chimera generated with B-1 donor cells from Blimp-1-YFP mice (n = 4). (C) FACS plot (left) and (right) mean percentage ± SD of CD5+ and CD5- cells among total Blimp-1 YFP+ cells (n = 13). (D) FACS gating strategy for sorting CD19+IgM+IgDloCD43+CD5+ or CD5- cells in the MedLN on days 3, 5, or seven after influenza infection of C57BL/6 mice, pooled from n = 2–3 per time point. (E) Concentration (ng/ml) IgM in supernatant (left) and secreted (ng x 10−4) per cell (right) of sorted cells measured by ELISA. (F) FACS gating strategy and (G) mean percentage ± SD of CD19+CD43+IgMb+IgDlo and CD5+ or CD5- B-1 cells among at indicated times after infection (n = 6–7 per time point). (H–I) Samples from (G) regated to show total B-1 populations. (H) Sample FACS plot and (I) percentage ± SD of CD5+ and CD5- B-1 cells among total (Dump- IgMb+ IgMa-) B-1 cells at indicated times after infection (n = 6–7 per time point). (J) Mean percentage ± SD (left) and total number ± SD (right) of FACS-sorted CD5+ and CD5- B-1 cells (IgMb+IgMa-) that formed IgM antibody-secreting cells (ASC) in each MedLN, as measured by ELISPOT (n = 3–4 per time point). Results are representative of >4 (A), 3 (B), and 2 (F, I), or are combined from 2 (D, E, G, H) or 3 (C) independent experiments. Values in (I) were compared by unpaired Student’s t test (*=p < 0.05, **=p < 0.005).

https://doi.org/10.7554/eLife.46997.002

To investigate the contribution of CD5- B cells to local IgM secretion, we FACS-sorted CD19+IgM+IgDloCD43+ CD5+ and CD5- B cells on days 3, 5, and seven after influenza infection from C57BL/6 mice (Figure 1D), which were then cultured for 2 days to analyze spontaneous IgM secretion by ELISA. Consistent with the presence of CD5- B-1PC, the CD5- cells secreted more IgM compared to the CD5+ cells, measuring total IgM concentrations and calculating IgM production per cell (Figure 1E). Sorted CD5+ cells did not secrete measurable amounts of IgM unless harvested after day 5 of infection.

Because CD5- B-1 cells and IgM-secreting B-2 derived plasmablasts express a similar phenotype (IgM+ IgD- CD5- CD19+ CD43+), the CD5- cells could have contained both B-1 cells and/or B-2-derived IgM-secreting cells. To determine the contribution of CD5- B-1 cells to IgM secretion in the MedLN, we infected allotype-disparate B-1 cell neonatal chimeras, in which B-1 (Ighb) and B-2 (Igha) cells and their secreted antibodies can be distinguished based on Igh-allotype differences (Lalor et al., 1989). The studies confirmed our previous findings that among CD19hiIgMb+IgDloCD43+B-1 cells in the MedLN, about 70% expressed CD5 after influenza infection (Figure 1F–G).

Because we had shown previously that Blimp-1+ B-1PC have reduced or absent CD19-expression (Savage et al., 2017) and found here that these cells are present after influenza infection (Figure 1A–B) and often lacked CD5-expression (Figure 1C), we expanded the analysis to include all IgMb-expressing (B-1 donor-derived) and IgMa negative (recipient-derived) cells, regardless of expression of CD19 or other surface markers (Figure 1H). Of note, a small number of double positive (IgMa+IgMb+) cells were always observed but excluded from the analysis, as these are likely IgMa+ B-2 cells with surface-bound serum IgM (B-1-derived and thus IgMb), attached via surface FcµR (Nguyen et al., 2017) (Figure 1A/H). In contrast to the analysis described above, this expanded analysis of all B-1 donor Igh-b cells revealed that the frequency of CD5 negative MedLN B-1 cells increased after influenza infection (Figure 1H–I), consistent with the development of CD5- B-1PC in this compartment (Figure 1A–C). Furthermore, FACS-sorting and culture of CD5+ and CD5- B-1 cells showed that a higher frequency and total number of CD5- B-1 cells secreted IgM in the MedLN compared to CD5+ B-1 cells on days 3, 5, and seven after infection (Figure 1J). Thus, CD5- B-1 cells increase in the MedLN and are a major source of local IgM production after influenza infection.

CD5+ B-1 cells decrease CD5 expression after LPS stimulation in vitro

To reconcile our previous findings about the role of CD5+ B-1 cells in influenza infection (Choi and Baumgarth, 2008; Waffarn et al., 2015), we considered whether CD5 surface expression may change after B-1 cell activation. Indeed, approximately 40% of highly purified FACS-sorted CD5+ B-1 cells from the peritoneal cavity lacked CD5 expression when cultured for 3 days in the presence but not absence of LPS, a stimuli that is known to induce IgM production by body cavity B-1 cells (Su et al., 1991) (Figure 2A). CD5 surface expression was unaffected during the first 2 cell divisions following stimulation, but was then quickly lost during the next 1–2 divisions (Figure 2B). Both, surface-expressed CD5 and cd5 mRNA, as assessed by qRT-PCR, were decreased among B-1 cells after 3 days of LPS stimulation (Figure 2C–D). Surface CD5 levels were decreased by 1.5 days of culture, while cd5 mRNA was not reduced until 2 days after culture onset (Figure 2C–D). The stimulated cells began secreting IgM before CD5 levels were reduced, but the increase in IgM secretion was more pronounced after 2 days of stimulation compared to the earlier time points (Figure 2E).

Figure 2 with 1 supplement see all
CD5+ B-1 cells decrease CD5 expression after LPS stimulation in vitro.

(A) Representative FACS plots (left) and mean percentage ± SD (right) of CD5+ and CD5- B-1 cells after FACS-purified peritoneal cavity CD19+ CD23- CD5+ B-1 cells were cultured with or without 10 µg/ml LPS for 3 days (n = 18). (B) CD5 expression on FACS-purified Efluor 670-stained proliferating peritoneal cavity CD5+ B-1 cells stimulated with LPS compared to CD5 FMO (fluorescence minus one) control. (C) Mean CD5 MFI ± SD, determined by flow cytometry, (D) mean Log(cd5 mRNA expression) ± SD, determined by qRT-PCR, and (E) mean IgM secretion ± SD (µg/ml), determined by ELISA, after purified peritoneal cavity CD5+ B-1 cells were cultured for indicated times with LPS (n = 3–4 per time and data point). Results are combined from 4 (A), or are representative of >5 (B), and 2 (C-E) independent experiments, respectively. Values in (C–E) were compared using an unpaired Student’s t test (*=p < 0.05, **=p < 0.005, ***=p < 0.0005, ****=p < 0.00005).

https://doi.org/10.7554/eLife.46997.003

A number of control studies were performed to ensure that the reduced frequencies of CD5+ B-1 cells in the cultures were not due to selective expansion of small numbers (<5%) of CD5- cells that might have contaminated the cultures at onset. First we separated CD5+ and CD5- B-1 cells from the body cavities by FACS to very high purities, and then cultured pure (100%) CD5+ B-1 cells, as well as cultures of CD5+ B-1 cells to which we added 1% and 5% CD5- B-1 cells, respectively. The data showed that the frequencies of CD5+ and CD5- cells after 3 days of culture were unaffected by the initial composition of the culture wells (Figure 2—figure supplement 1A). There was no significant difference in either CD5 MFI or in the percent of CD5+ and CD5- cells on day 3 of culture, independent of whether small numbers of CD5- cells were added to the CD5+ B-1 cell cultures (Figure 2—figure supplement 1B). Thus, small percentages of CD5- B-1 cells at culture onset, representative of potential sort impurities, could not explain the lack of CD5 expression by the CD5+ B-1 cells stimulated with LPS for 3 days.

Next, we compared the ability of CD5+ and CD5- B-1 cells to survive and/or proliferate with and without LPS stimulation to ensure that CD5- B-1 cells do not demonstrate a more robust response than CD5+ B-1 cells. To ensure that the two populations were exposed to the same culture conditions, CD5+ and CD5- B-1 cells were sorted, labeled with different proliferation dyes, and cultured together (Figure 2—figure supplement 1C). Compared to B-1 cells that expressed CD5 on day 0, CD5- cells did neither demonstrate better survival (Figure 2—figure supplement 1D) nor enhanced proliferation in response to LPS stimulation compared to the CD5+ cells, in terms of both, the percentage of cells that underwent division, as well as the numbers of divisions each B-1 cell underwent (Figure 2—figure supplement 1E–F). In fact, the CD5+ B-1 cells had better overall survival rates compared to CD5- B-1 cells. Reflecting the similar rates of proliferation and the increased survival of the CD5+ B-1 cells, populations of B-1 cells that expressed CD5 at culture onset were present at higher frequencies of input cells compared to B-1 cells that were CD5 negative (Figure 2—figure supplement 1G). We conclude that CD5+ B-1 cells lose CD5 surface and mRNA expression after in vitro LPS stimulation.

CD5+ B-1 cells differentiate into CD5- IgM secreting cells after stimulation with multiple TLR agonists

Endosomal TLR agonists Imiquimod (TLR7) and ODN CpG 7909 (TLR9) also induced CD5 downregulation on CD5+ B-1 cells after 3 days of culture (Figure 3A), as did stimulation with lipids from Mycobacterium tuberculosis (Mtb lipids), which activate cells primarily through TLR2 (Basu et al., 2012) (Figure 3C). These findings are consistent with the loss of CD5 expression seen after stimulation with various TLR agonists by Kreuk et al. (2019). Similar to LPS stimulation, CD5 expression decreased as the cells divided (Figure 3B). In contrast, stimulation of CD5+ B-1 cells isolated from mice lacking all TLR-signaling due to a deletion of genes encoding Unc93, TLR2 and TLR4 (kind gift of Greg Barton, UC Berkeley), were unable to respond with proliferation (Figure 3D), and failed to lose CD5 (not shown). Thus, B-1 cells were stimulated via TLR-engagement and not via the BCR. In all instances, the loss of CD5 was correlated with the differentiation of CD5+ B-1 cells to IgM-secreting cells, as stimulation of these cells with Imiquimod, CpG, and LPS for 3 days resulted in increased percentages of CD138+ cells (Figure 3E–F) and an increase in IgM concentrations in the culture supernatants (Figure 3G).

CD5+ B-1 cells differentiate into CD5- IgM secreting cells after TLR-mediated activation.

(A) CD5 MFI ± SD and (B) representative FACS plots for CD5+ B-1 cells cultured without stimulation or with Imiquimod (TLR7 agonist), CpG 7909 (TLR9 agonist), or LPS (TLR4 agonist) (n = 3–5). (C) Mean CD5 MFI ± SD of CD5+ B-1 cells cultured without stimulation or with Mycobacterium tuberculosis (Mtb) lipids (TLR2 agonist) or LPS (n = 4–5). (D) Mean percentage ± SD of B-1 cells from wild type (WT) or Tlr2−/−xTlr4−/−xUnc93b13d/3d (TLR KO) mice that underwent at least one division after culture without stimulation or stimulated with Mycobacterium tuberculosis (Mtb) lipids or LPS (n = 6–9 per group). (E) FACS plots (left) and mean percentage ± SD (right) of CD138+ cells, and (F) representative FACS plots for CD138 expression among proliferating cells. (G) Mean IgM concentration ± SD (µg total per culture well) of cultured CD5+ B-1 cells stimulated or not with Imiquimod (TLR7 agonist), CpG 7909 (TLR9 agonist), or LPS (TLR4 agonist) (n = 2 for no stimulation and LPS, n = 5 for Imiquimod and ODN). (H) Sample FACS plot (left) and mean CD5 MFI ± SD (right) of PTC liposome-binding (PTC+) and non-PTC liposome-binding (PTC-) cells for CD5+ B-1 cells cultured without stimulation or with Imiquimod (TLR7 agonist), CpG 7909 (TLR9 agonist), or LPS (TLR4 agonist) (n = 3–5). Results are combined from two (D, E–G), or are representative of three (A) or two (B, C, H) independent experiments, respectively. Values compared in (A, C–D) using an unpaired Student’s t test (*=p < 0.05, **=p < 0.005, ***=p < 0.0005, ****=p < 0.00005).

https://doi.org/10.7554/eLife.46997.005

Finally, we examined whether phosphatidylcholine (PTC)-binding B-1 cells can lose CD5 surface expression after TLR-stimulation. PTC is a well-known specificity of a large subset of peritoneal cavity CD5+ B-1 cells (Mercolino et al., 1988; Arnold and Haughton, 1992). PTC-binding B-1 cells, identified by incubation of cells with a fluorescent PTC-liposome (kind gift of A. Kantor, Stanford University), lost CD5 expression similarly to PTC non-binders (Figure 3H). We conclude that TLR-mediated stimulation of CD5+ B-1 cells (‘B-1a’) causes the loss of CD5 surface expression, making these cells phenotypically indistinguishable from the proposed ‘sister’ B-1 cell population, the CD5- ‘B-1b’ cells.

CD5+ B-1 cells become CD5- IgM ASC in the MedLN after Influenza infection

These results raised the possibility that pleural cavity CD5+ B-1 cells respond to influenza virus infection with down-regulation of CD5 in the MedLN. The data would explain the increases in CD5- B-1 cells in the MedLN after influenza infection (Figure 1). They would also explain how the frequencies of CD5- B-1 cells increased at that site, despite the fact that we had shown previously with neonatal chimeras reconstituted with only CD5- B-1 cells that CD5- B-1 cells cannot enter the MedLN after influenza infection, and that the CD5+ B-1 cells were sufficient to induce the entire B-1 cell response (Choi and Baumgarth, 2008).

To confirm and expand these data and because these previous studies showed that only CD5+ B-1 cells could enter the MedLN, we established neonatal chimeras with varying mixes of CD5+ and CD5- B-1 cells (Figure 4A) and tested whether we could see a correlation between the frequencies of CD5+ and CD5- cells in the MedLN and/or the levels of B-1-derived IgM secretion in MedLN after influenza infection. For clarity we binned the results into chimeras reconstituted with >50% CD5- B-1 cells,>50% CD5+B-1 cells or only CD5+ B-1 cells (Figure 4B). As shown previously (Choi and Baumgarth, 2008), MedLN of mice reconstituted with mostly CD5- B-1 cells had reduced MedLN B-1 cell after infection (Figure 4B). Of note, among the total donor B-1 cells in the MedLN, the frequencies of CD5+ and CD5- cells were similar, regardless of the initial percentage of CD5+ cells (Figure 4C–D), consistent with the CD5+ B-1 cells losing surface CD5 expression. This is further consistent with the fact that in chimeras generated with only CD5+ B-1,>50% of B-1 cells in the MedLN lacked CD5 expression (Figure 4D).

Figure 4 with 1 supplement see all
CD5+ B-1 cells differentiate to CD5- IgM ASC in the MedLN after Influenza infection.

(A) Neonatal chimeric mice were generated with FACS sorted CD19+ CD23- Ighb+ CD5+ (100%, blue), mostly CD5+ (orange), or mostly CD5- (green) peritoneal cavity-derived B-1 cells and infected with influenza A/Puerto Rico 8/34 for 7 days. (B) Mean number ± SD of B-1 cells in the MedLN of mice 7 days after infection. (C) FACS plot and (D, left) mean percentage ± SD of Dump- IgMb+ IgMa CD5+ and CD5- MedLN B-1 cells on day 7. CD5 FMO (fluorescence minus one) control for CD5. (D, right) Mice were grouped by initial percentage of CD5+ and CD5- B-1 cells (left) and % MedLN CD5+ B-1 cells present on days 0 (initial %) and 7 of infection were plotted with a line of best fit. (E) Mean B-1 derived IgM ASC ± SD per MedLN, grouped by initial percentage of CD5+ and CD5- cells (left) and plotted based on initial starting percentage of CD5+ cells (right) with a line of best fit. (F) Mean proliferation rate per day ± SD of CD5+, CD5-, and CD138+ B-1 cells and CD138+ B-2 cells (B-2 PC) in the MedLN of infected chimeras compared to proliferation rate per day of similar populations (B-1 or B-2 cells) in the peritoneal cavity of each mouse as determined by BrDU incorporation. Results for infected mice in (B–F) are combined from four independent experiments (n = 4 for>50% CD5-, n = 7 for>50% CD5+ cells, n = 5 for pure CD5+ cells). Results for uninfected chimeras in (E) are combined from three independent experiments, n = 6. Values in (B, D–F) were compared by unpaired Student’s t test (*=p < 0.05, **=p < 0.005).

https://doi.org/10.7554/eLife.46997.006

Allotype-specific ELISPOTs showed that chimeric mice generated with only CD5+ B-1 cells formed significantly higher frequencies of B-1-derived IgM-secreting cells compared to chimeric mice generated with predominantly CD5- B-1 cells (Figure 4E). In fact, chimeras generated with CD5- B-1 cells showed no more B-1 derived IgM ASC in their MedLN than uninfected chimeras, consistent with their previously reported deficiency in entering the MedLN after infection (Figure 4E, left panel) (Waffarn et al., 2015; Choi and Baumgarth, 2008). There was a significant positive correlation between the frequencies of CD5+ B-1 cells transferred to generate the neonatal chimeras and the ability of the mice to generate IgM ASC following influenza virus infection (Figure 4E, right panel).

CD5+ B-1 cells failed to show signs of clonal expansion following their accumulation in the MedLN (Choi and Baumgarth, 2008), which was confirmed using BrdU injection on day six after infection. However, the CD5- MedLN B-1 cells showed increased proliferation compared to their counterparts in body cavities (Figure 4F). Among B-1 cells the proliferation rate was highest among the CD138+ B-1PC, with rates similar to that of the B-2 CD138+ plasma cell compartment (Figure 4F). The data support the hypothesis that CD5- B-1 cells, and in particular B-1PC in the MedLN, arise from proliferating CD5+ pleural cavity B-1 cells that accumulate in the MedLN and differentiate into CD5- IgM ASC following acute influenza virus infection.

CD5+ B-1 cells become CD5- IgM ASC in the Mesenteric LNs and Peyer’s Patches after Salmonella typhimurium infection

Numerous infection models have reported CD5- ‘B-1b’ cell responses after infection, including studies on mice infected with Streptococcus pneumonia (Haas et al., 2005) and S. typhimurium (Gil-Cruz et al., 2009). This has led to the concept that the CD5- B-1b are a ‘responder’ B-1 cell population, whereas CD5+ B-1 cells generate natural IgM exclusively in the steady state (Haas et al., 2005; Alugupalli and Gerstein, 2005). We therefore aimed to reexamine whether activation and differentiation of CD5+ B-1 cells into CD5- IgM ASC were more universal outcomes of CD5+ B-1 cell activation to infections.

Neonatal chimeric mice generated with varying ratios of FACS-purified CD5+ and/or CD5- B-1 cells, as described above were orally infected with S. typhimurium (Figure 4—figure supplement 1A). Consistent with the studies of MedLN B-1 cell populations after influenza infection, we found an increased number of B-1 cells in the infected Mesenteric LN (MesLN) of mice given CD5+ B-1 cells vs. those given CD5- B-1 cells (Figure 4—figure supplement 1B, left). A similar trend was seen in the Peyer’s Patches, but did not reach statistical significance, likely due to the very small number of total B-1 cells in that tissue (Figure 4—figure supplement 1B, right). Also consistent with studies on the MedLN after influenza infection, we found a similar percentage of CD5+ and CD5- B-1 cells (identified as IgMb+IgMa-) in the MesLN and Peyer’s Patches on day four after infection, regardless of the initial percentage of CD5+ cells used to reconstitute the B-1 compartment of these mice . We again found that > 50% of the B-1 cells in tissues of chimeras established with only CD5+ B-1 cells lacked CD5 surface expression (Figure 4—figure supplement 1C and these chimeras were the most competent at forming IgM secreting cells after infection (Figure 4—figure supplement 1D). In contrast, the MesLN and PP of chimeric mice that were given primarily B-1b cells formed fewer IgM secreting cells, although more than the uninfected chimeras . Although we did see a significant decrease in the number of B-1 cells in the MesLN of mice given 95% CD5+ B-1 cells compared to those receiving 100% CD5+ Figure 4—figure supplement 1B, left), this is unlikely of biological significance, since there is similar IgM secretion in the MesLN between these two groups of mice, Figure 4—figure supplement 1B, left).

The S. typhimurium surface antigen OmpD had been reported previously to stimulate IgM secretion exclusively by CD5- ‘B-1b’ cells (Gil-Cruz et al., 2009). However, in our hands, although total B-1-derived IgM was increased after infection in the MesLN (Figure 4—figure supplement 1D) OmpD-specific B-1-derived IgM ASC in the MesLN did not increase significantly (Figure 4—figure supplement 1E). Instead, we found OmpD-specific IgM secretion only by host-derived, thus B-2 plasmablasts. Of note, the phenotype of activated B-2-derived plasmablasts is indistinguishable from that of the so-called ‘B-1b’ cells (CD19lo CD5- CD45Rlo IgM+ CD43+ (Figure 4—figure supplement 1F) and thus only identifiable using a lineage-marking approach, such as used here.

Together these findings demonstrate that B-1 cells accumulate in draining lymph nodes at the site of both, bacterial and viral infections, where they lose CD5 expression and become the main source of B-1 derived secreted IgM. In vitro this process is recapitulated by stimulation with various TLR-ligands.

Changes in BCR signaling following innate activation of B-1 cells

Surface CD5 expression by B-1 cells has been linked previously to their inability to proliferate in response to BCR-mediated signaling (Bikah et al., 1996). To analyze the association of CD5 with the BCR on B-1 cells in steady state and to determine what changes the stimulation of the BCR may induce on B-1 cells, we analyzed the IgM-BCR-complexes on the cell surface of highly FACS-purified, then rested, peritoneal cavity CD5+ CD45 Rlo CD23- B-1 and splenic CD45Rhi CD23+ CD5 follicular B cells using Proximal Ligation Assay (PLA). On B-1 cells, both CD19 and CD5 were strongly associated with the surface-expressed IgM-BCR, while CD5 was not directly associated with the co-stimulator and signaling molecule CD19 (Figure 5A). Consistent with the lack of stimulation and strong interaction between IgM and CD5, the BCR-signaling chain CD79 only weakly interacted with the adaptor molecule Syk in B-1 cells in the steady-state. B-2 cells lack CD5 expression, and CD19 did not interact with the IgM-BCR prior to stimulation (Figure 5A).

Association of CD5 with IgM-BCR in resting B-1a cells is increased after BCR-stimulation.

(A) Indicated FACS-purified B cell subsets from the peritoneal cavity (PC) and spleen (Spl) of BALB/C mice were analyzed by proximal ligation assay for the following interactions (left to right): IgM:CD19, IgM:CD5, CD19:CD5 and CD79:syk. Left panel shows summarizes data on signal counts for 200 individual cells analyzed. Each symbol represents one cell, horizontal line indicates mean signal count per cell. Right panel show representative fluorescent images. (B) FACS-purified CD19hi CD23- CD5+ CD43+ B-1 cells from the peritoneal cavity and CD19+ CD23+ splenic B-2 cells of BALB/C mice were labeled with efluor670 and then cultured in the absence (top) or presence of 20 ug/ml anti-IgM (middle) or 10 ug/ml CpGs for 72 hr. Left panels show representative histogram plots, middle panel shows the % cells in each culture having undergone at least one cell division and right panel indicates the proliferation index (average number of proliferations undergone per divided cell). (C) Summary of proximal ligation assay results of B-1 cells purified as in (A) and then stimulated for indicated times with anti-IgM(Fab)2. Interactions of the following proteins were analyzed on 200 cells per condition (left to right): IgM:CD19, IgM:CD5, CD19:CD5 and CD79:syk. Right panels shows representative fluorescent images from one experiment of at least two done. Values were compared using an unpaired Student’s t test (*=p < 0.05, **=p < 0.005, ***=p < 0.0005, ****=p < 0.00005).

https://doi.org/10.7554/eLife.46997.007

Stimulation of B-1 and B-2 cells with CpG induced strong proliferation of both cell populations (Figure 5B). As expected, and although anti-IgM induced strong proliferation by B-2 cells, CD5+ B-1 cells failed to respond to the same stimulus (Figure 5B). The lack of responsiveness of the CD5+ B-1 cells to BCR-stimulation was explained by the PLA data, which showed the maintenance and even increase in IgM-BCR:CD5 association and an increase in the association of the inhibitor CD5 with CD19 following anti-IgM treatment. Furthermore, B-1 cells lost the interaction of the IgM-BCR with CD19. Consequently, CD79-Syk interactions remained very low (Figure 5C). Thus, BCR-engagement on B-1 cells inhibits BCR-signaling by reducing steady-state IgM-CD19 interactions and likely also by initiating instead interactions between CD5 and CD19.

In contrast to direct stimulation of the IgM-BCR, CpG stimulation led to changes in the BCR-signaling complex that are consistent with induction of positive BCR-signaling, and/or the ability to signal through the antigen-receptor (Figure 6). CpG stimulation strongly increased the interaction of IgM-BCR:CD19 and reduced CD5:IgM-BCR proximity. The dissociation between the IgM-BCR and CD5 occurred within 24 hr of stimulation (Figure 6A) and thus significantly before the reduction in surface CD5 expression was measurable by flow cytometry (Figure 2C) and before changes to CD5 transcription (Figure 2D). The already weak CD5:CD19 interaction was further reduced (Figure 6A), consistent with the eventual loss in surface CD5 expression noted following stimulation (Figure 3). These rapid changes in the BCR-signaling complex were associated with increases in CD79:Syk interaction, suggesting active BCR-signaling in CpG-stimulated B-1 cells (Figure 6A). This was further supported by sustained increased levels of phosphorylated Akt (pAkt; pS473) following stimulation of CD5+ B-1 cells with CpG, while anti-IgM stimulation reduced pAkt levels below that of unstimulated B-1 cells by 24 hr, after an initial increase (Figure 6B). We also noted increased Nur77 expression in CpG- but not anti-IgM-stimulated B-1 cells, further suggesting BCR-signaling is linked to CpG-stimulation after 24 hr (Figure 6C), but not after 3 hr (not shown). Given that B-1 cell responses following CpG-stimulation were TLR-expression dependent in vitro (Figure 3D) and no external antigen was provided to the cultures, TLR-signaling may directly link to BCR-signaling. Alternatively, TLR-signaling may ‘license’ subsequent self-antigen recognition, by altering the BCR-signaling complex. In support of the latter, we noted a strong increase in the frequency of PTC-binding B-1 cells during culture (Figure 6D), which may be due to the expansion of CpG-activated B-1 cell in response to PTC-antigen present on dead and dying cells in the cultures.

TLR-mediated stimulation of CD5+ B-1 cells alters the BCR-signalosome.

(A) FACS-purified peritoneal cavity CD19hi CD23- CD43+ CD5+B-1 and splenic CD19+ CD23+ CD43 CD5- B-2 cell of BALB/C mice were stimulated for the indicated times with TLR9-agonist ODN7909 prior to analysis by proximal ligation assay, probing for the following interactions (left to right): IgM:CD19, IgM:CD5, CD19:CD5 and CD79:syk. Left panel summarizes data on signal counts for 200 individual cells analyzed. Each symbol represents one cell, horizontal line indicates mean signal count per cell. Right panel show representative fluorescent images. (B) Analysis of the phosphorylation status of Akt by probing for Akt pS473 by flow cytometry on FACS-purified CD19hi CD23- CD5+ CD43+ B-1 cells from the peritoneal cavity and CD19+ CD23+splenic B-2 cells of BALB/C mice. Top panels show representative histogram plots, bottom summarizes the results. (C) Mean fluorescence intensity ± SD of staining for the immediate early activation factor Nur77, in CD5+ B-1 cells isolated as described in (A) and cultured for up to 2 days in the absence and presence of the indicated stimuli. (D) Shown are % frequencies of live PtC-binding B-1 cells among live FACS-purified CD5+ peritoneal cavity B-1 cells cultured with LPS stimulation for the indicated times, as assessed by flow cytometry. Each symbol represents results obtained from one culture well. Results are representative from experiments conducted at least twice with multiple repeats done per experiment (n = 2–5). Results in D are combined from two independent experiments. Values were compared using an unpaired Student’s t test (*=p < 0.05, **=p < 0.005, ***=p < 0.0005).

https://doi.org/10.7554/eLife.46997.009

Initial stimulation through IgM-BCR suppresses subsequent activation of B-1 cells via TLR- stimulation

We further analysed the impact of stimulation through the TLR or IgM-BCR on stimulation of B-1 cells by the other receptor. For that a set of in vitro experiments was conducted, in which FACS-purified, eFluor 670-stained and rested CD5+ body cavity B-1 cells and splenic follicular B-2 cells were stimulated in vitro with anti-IgM and/or CpG for a total of 72 hr at which time proliferation was assessed by flow cytometry. Stimulation with either anti-IgM or CpG for 2 hr followed by wash-out and then stimulation with the other stimulus for 70 hr showed no difference to stimulation with the second stimulus alone (not shown). However, initial stimulation for 24 hr followed by wash-out and restimulation with the second stimulus showed significant effects of initial IgM-BCR stimulation on subsequent CpG responsiveness by B-1 and B-2 cells (Figure 7). While B-1 cell responses to anti-IgM alone for 24 or 72 hr did not result in significant proliferation, when CpG was given after 24 hr anti-IgM stimulation a small but significant increase in proliferation was noted (Figure 7A). However, proliferation rates were greatly lower compared to stimulation with CpG first, followed by anti-IgM stimulation, or stimulation with CpG alone (Figure 7A). In contrast, initial anti-IgM stimulation of B-2 cells followed by stimulation with CpG resulted in the most robust B-2 cell proliferative response (Figure 7B), exceeding that of CpG stimulation alone or CpG stimulation followed by anti-IgM.

TLR but not BCR-stimulation induces CD5+ B-1 cell proliferation FACS-purified.

(A) peritoneal cavity CD19hiCD23- CD43+ CD5+ B-1 and (B) splenic CD19+ CD23+ CD43 CD5- B-2 cell from BALB/C mice were labeled with eFluor 670, rested for 2 hr and then cultured for 24 hr with the indicated stimulus 1 (none/anti-IgM at 10 μg/ml or CpG ODN7909 at 5 μg/ml), washed and recultured for 48 hr with stimulus 2 (none/anti-IgM at 10 μg/ml/ CpG ODN7909 at 5 μg/ml) prior to analysis for efluor 670 staining. Top panels show representative FACS histogram plots and bottom panels shows the % cells in each culture having undergone at least one cell division. Each symbol represents results from one culture well, horizontal line indicates mean for the group. Results are compiled from two independent experiments. Statistical analysis was done by one-way ANOVA, followed by an unpaired Student’s t test with Holm-Sidak correction for multiple comparisons (*p<0.05, **p<0.005, ***p<0.0005, ****p<0.00005).

https://doi.org/10.7554/eLife.46997.010

Thus B-1 and B-2 cells greatly differ in their responses not only to BCR but also to TLR stimulation. While TLR stimulation by B-2 cells induced a strong and synergistic enhancement of proliferation initiated by BCR-signaling, B-1 cells did not respond to BCR-mediated stimulation with proliferation, independent of whether the BCR signal was given first or after a TLR-stimulus. However, only when TLR stimulation was provided first, did B-1 cells show robust proliferation in context of anti-IgM BCR-signaling.

Local IgM production following influenza infection depends on TLR expression

The data suggest that TLR-mediated stimulation alters the BCR-signalosome complex, which may drive B-1 cell responses to pathogens in vivo. Indeed, complete TLR-deficient mice (due to a lack of TLR2, TLR4 and Unc93) showed significant deficits in CD5+ B-1 cell responses following influenza infection (Figure 8). Significant increases in the ratios of CD5+ over CD5- and CD19+ CD43+ B cells were noted (Figure 8A). This suggested that CD5+ B-1 cells in TLR-deficient mice could enter the MedLN, consistent with our previous findings that this step is TLR-independent but Type I IFN-dependent (Waffarn et al., 2015), but once in the MedLN they were not activated via TLR-dependent signals, that is failed to downregulate CD5. Of importance, the lack of TLR-stimulation also resulted in a near complete loss of CD19lo/- IgM+ CD138+ B-1PC in the MedLN at day 5 of infection (Figure 8A/B) and a corresponding drop in IgM ASC in TLR-deficient compared to control mice at that timepoint (Figure 8C), while viral loads were similarly low in the lungs of both mouse strains (not shown). Generation of Ig-allotype chimeras in which only B-1 cells lacked TLR expression confirmed a B-1 cell-intrinsic requirement for TLR-signaling in B-1 cell differentiation to CD138+ ASC after influenza virus infection (Figure 8D–F).

TLR-mediated stimulation is required for maximal IgM responses to influenza virus infection.

(A) C57BL/6 (n = 5) and congenic total TLR-deficient mice (n = 5; lacking TLR2, TLR4 and Unc93) were infected with influenza A/Puerto Rico/8/34 for 5 days. Shown are representative FACS plots from control C57BL/6 (top) and TLR-deficient (bottom) mice FACS analysis of MedLN for the presence of B-1 and B-1PC. (B) Number of B-1PC per MedLN as assessed by FACS and (C) number of IgM-secreting cells in MedLN as assessed by ELISPOT analysis. (D-F) Similar analysis as for A-C but using allotype chimeras generated with wild type recipients and B-1 cells from either C57BL/6 or tlr-/- mice. (D) Representative FACS analysis of CD138+ B-1PC pre-gated for live, dump-, B-1 donor (IgMb+) cells in MedLN on days 5 and 7 after influenza infection. (E) Mean ± SD of data summarized from analysis shown in D. (F) Mean ± SD of B-1 IgM-ASC in MedLN on day 7 after infection, as assessed by ELISPOT. Each symbol represents results from one mouse with female mice shown as open symbols, males as closed symbols. Results are combined from two independent experiments. Values were compared using an unpaired Student’s t test (*=p < 0.05, **=p < 0.005, n.s. not significant). 

https://doi.org/10.7554/eLife.46997.011

Together the data demonstrate a linkage of TLR and BCR-signaling during B-1 cell responses to infections, with intrinsic TLR-mediated signaling triggering a rapid reorganization of the IgM-BCR-signalosome complex, including the removal of the BCR-signaling inhibitor CD5 and increased association of IgM-BCR with the co-receptor CD19, and the TLR-dependent differentiation of CD5+ B-1 to CD5- IgM-secreting B-1 and B-1PC.

Discussion

Self-reactive, fetal and neonatal-developing B-1 cells do not respond to BCR-stimulation with clonal expansion. This was shown previously to be associated with their expression of the BCR-signaling inhibitor CD5, and a lack of fully functional CD19 signaling. It is consistent with the notion that these self-reactive cells must be silenced in order to avert the risk of autoimmune disease induction. Yet B-1 cells do respond rapidly to various infections with migration to secondary lymphoid tissues and with differentiation to IgM secreting cells. The present study resolves this conundrum by providing a mechanism by which B-1 cell can overcome their inherent BCR-signaling block, namely the TLR-mediated activation and reorganization of the BCR-signalosome complex. This non-redundant signal induced the dissociation of CD5 from the IgM-BCR and eventual its loss from the cell surface of the initially CD5-expressing B-1 cells. It also enhanced the association between IgM and the co-stimulatory molecule CD19, and caused strong increases in CD79:Syk interaction and phosphorylation of BCR downstream effectors.

The study also clarifies the respective roles of the CD5+ and CD5- B-1 cells, previously suggested to form two distinct subsets ‘B-1a’ and ‘B-1b’. By demonstrating that CD5+ B-1 cells respond to infections with both, influenza and S. typhimurium, the latter previously identified as an exclusive ‘B-1b’ response (Gil-Cruz et al., 2009; Marshall et al., 2012), with rapid downregulation of CD5, the study suggests that pathogen-induced responses by B-1 cells represent responses of CD5+ B-1 cells (Haas et al., 2005; Alugupalli et al., 2003; Alugupalli et al., 2004; Gil-Cruz et al., 2009; Marshall et al., 2012). The lack of CD5-expression on B-1 cells may thus more broadly identify activated and differentiated B-1 cells, a conclusion supported also by the work in the accompanying manuscript by Kreuk et al. (2019). This is consistent with previous reports on the phenotype of natural IgM-secreting cells as mostly CD5- (Savage et al., 2017; Ohdan et al., 2000), and could explain earlier reports that CD5+, not CD5- B-1 cells form natural IgM secreting cells (Haas et al., 2005; Masmoudi et al., 1990; Hayakawa et al., 1984). In addition, it could explain the findings that CD5- B-1 cells contain CD5- memory-like B-1 cells in the body cavities of previously infected mice (Alugupalli et al., 2004; Foote and Kearney, 2009) which also likely respond to antigen.

Thus, the data presented here and in the accompanying work by Kreuk and colleagues are inconsistent with models that consider B-1 cell responses as a division of labor between two subsets of B-1 cells: B-1a and B-1b cells, where CD5+ B-1a contribute ‘natural’ IgM and CD5- B-1b the induced IgM, proposed previously (Haas et al., 2005; Alugupalli and Gerstein, 2005). Given CD5 expression is dynamically expressed and thus cannot be used to identify B-1 cell subsets, and no other clearly subset-defining differences have been reported to-date, we suggest that the separation of B-1 cells into B-1a and B-1b ‘sister populations’ be revoked, and that instead these cells are simply referred to as B-1 cells. If distinctions in CD5 expression are important, those could be indicated by describing B-1 cells as CD5+/CD5- instead.

Our data do not exclude the possibility that some B-1 cells develop which either express no, or undetectable levels of surface CD5, as described previously (Kantor et al., 1992). Given the known functions of CD5 as an inhibitor of BCR-signaling (Hippen et al., 2000; Punt et al., 1994; Azzam et al., 1998; Perez-Villar et al., 1999) and the fact that CD5-expression levels on thymocytes correlated with the strengths of the positively selecting TCR–MHC-ligand interactions (Azzam et al., 1998), such CD5lo/neg B-1 cells might have lower levels of self-reactivity (Hayakawa et al., 1999; Lalor and Morahan, 1990; Mercolino et al., 1988; Yang et al., 2015) and lack the need for CD5-mediated silencing of BCR-signaling in order to avoid inappropriate hyperactivation of these self-reactive B cells. De novo development of both CD5+ and CD5- B-1 cells has been reported to occur also in stromal cell cultures seeded with B-1 cell precursors (Montecino-Rodriguez et al., 2006). It remains to be explored whether the presence of DAMPS in the in vitro cultures could contribute to the loss of CD5 on otherwise CD5+ B-1 cells, or whether these cells never expressed CD5. Our data are not consistent with early reports suggesting that CD5+ B-1 cells could only reconstitute themselves, but not CD5- B-1 cells (Kantor et al., 1992; Stall et al., 1992), as reconstitution of neonatal mice with even very highly FACS-purified body cavity CD5+ B-1 cells led to significant numbers of CD5- B-1 cells recovered from these mice (Choi and Baumgarth, 2008 and Figure 4—figure supplement 1).

The mechanisms by which the TLR-induced reorganization of the BCR-signalosome drives the differentiation of B-1 cells to IgM-secreting B-1PC will require future in-depth analysis. We favor the concept of a ‘licensing’ step, in which initial TLR-stimulation supports subsequent BCR-stimulation through the here observed reorganization of the BCR-signalosome. This is supported also by the in vitro studies which demonstrated that an initial engagement of the TLR could overcome the block in B-1 cell proliferation after BCR stimulation, while an initial BCR stimulus followed by TLR stimulation failed to induce B-1 cell proliferation (Figure 7). Such a licensing step may explain the recent data by Kreuk and colleagues (see Kreuk et al., 2019), which showed a degree of B-1 response specificity, and thus presumably BCR-responsiveness, that was dependent on the type of TLR stimulus provided to the B-1 cell. However, how such licensing step could induce antigen-specific B-1 cell responses that are dependent on the specificity of the TLR remains to be fully elucidated.

An attractive alternative and perhaps simpler concept would be that the BCR-mediated binding to antigen and then its uptake by BCR-internalization leads to engagement of particular TLR, which then synergize with BCR-mediated stimulation and support the strong proliferation and differentiation of antigen-specific B cells. This concept is very consistent with the outcome of B-2 cell stimulation in vitro (Figure 7), where TLR stimulation strongly and synergistically enhanced the initial BCR-mediated activation of these cells. However, the model cannot explain the complete lack of B-1 cell proliferation in response to BCR-engagement, the strong inhibition of TLR-mediated proliferation when preceded by BCR-engagement, and the BCR-signaling induced enhanced unresponsiveness of the BCR signalosome at the molecular level, as demonstrated by a failure to enhance CD79:syk interaction and upregulation of pAkt.

A potential third model would involve a direct linkage of TLR and BCR-signalosome effects. For example, low-affinity BCR-antigen interactions might be enabled by having DAMPS and PAMPS first bind to TLR on the B cell surface, which then brings these antigens in close proximity to the BCR, triggering antigen-specific BCR activation event. Although in some cultures we did not provide antigens other than the TLR-ligands to the in vitro cultures, dead and dying cells provide ample antigens, including PTC, that could have stimulated these cells. This would potentially explain why the stimulation of B-1 cells with TLR-agonists activates BCR signaling pathways, and why we found such strong enrichment for PTC-binders among cultures of TLR-stimulated B-1 cells. However, not all TLR are expressed on the cell surface, and their initial engagement prior to that of the BCR would necessitate the initial BCR-independent phagocytosis of antigen by B-1 cells, which others have reported (Popi et al., 2016).

The lack of phenotypic differences between CD5- B-1 cells and B-2 cell-derived non-switched plasmablasts, both expressing CD19, CD43, low levels of CD45 and lack IgD, further complicates the identification of responding B cell subsets in vivo, as demonstrated with the analysis to the Salmonella antigen OmpD. Using allotype-marked B cell lineages, we show here that anti-OmpD secreting B cells were derived predominantly from B-2 cells, and not as previously suggested from B-1b cells (Gil-Cruz et al., 2009). When CD5+ B-1 cells lose CD5, some also seem to begin to express CD138, thus becoming indistinguishable from B-2-derived plasma cells and plasmablasts that carry the same phenotype. Some investigators used expression of CD11b to identify B-1 versus B-2 cells in infectious models (Haas et al., 2005; Gil-Cruz et al., 2009), but this marker is also dynamically regulated depending on tissue site and B-1 cell activation status (Waffarn et al., 2015; Ohdan et al., 2000; Yang et al., 2007). Recent lineage-tracing approaches (Yuan et al., 2012; Zhou et al., 2015; Montecino-Rodriguez et al., 2016) may help the development of novel approaches or markers that can unequivocally identify B-1 cells. In the meantime, the use of neonatal B-1 allotype-chimeras remains a valuable tool for such analyzes.

Taken together, our data suggest that the BCR-complex composition on neonatally-derived, self-reactive B-1 cells is controlled by TLR-mediated signals, preventing inappropriate activation and autoimmune disease on the one hand, while facilitating rapid B-1 cell participation in anti-viral and anti-bacterial infections on the other. TLR-signaling thereby influences not only innate B-1 cell activation, but may also affect their antigen-specific responses.

Materials and methods

Key resources table
Reagent type
(species) or
resource
DesignationSource or
reference
IdentifiersAdditional
information
Strain, strain background (mouse) musculus (mouse), C57BL/6J (M + F)C57BL/6,
Control, Ighb
The Jackson LaboratoriesStock 000664Mus musculus (mouse), C57BL/6J (M + F)
Strain, strain background (mouse)IghaThe Jackson LaboratoriesStock 001317Mouse, B6.Cg- Gpi1aThy1aIgha/J (M + F)
Strain, strain background (mouse)BALB/CThe Jackson LaboratoriesStock 000651Mouse, BALB/CJ mice (F)
Strain, strain background (mouse)Blimp-1YFPRutishauser et al., 2009Breeding pairs from Michel Nussenzweig (The Rockefeller University)
Mouse, B6-Cg- Tg(PRDM1- EYFP)^(1Mnz) (M + F)
Strain, strain background (mouse)TLR-/-OtherBreeding pairs from Greg Barton (The University of California, Berkeley)
Mouse, Tlr2-/- Tlr4-/- Unc93b1^(3d/3d) (M + F)
Strain, strain background (mouse)ChimeraLalor et al., 1989Generated in-house
Mouse, Igha/Ighb B-1 Cell Neonatal Chimera
Strain, strain background (mouse)ChimeraLalor et al., 1989Generated in-house
Mouse, Igha/Ighb- YFP B-1 Cell Neonatal Chimera

Mice

8-16 week old male and female C57BL/6J or female BALB/c mice and breeding pairs of B6.Cg-Gpi1aThy1aIgha/J (Igha) mice were purchased from The Jackson Laboratory. Female, 10 weeks old BALB/C mice were purchased from Jackson Laboratory. B6-Cg-Tg(prdmi-EYFP)1Mnz (Blimp-1 YFP) breeders were kindly provided by Michel Nussenzweig (The Rockefeller University, NY) and breeding pairs of Tlr2-/- x Tlr4-/- x Unc93b13d/3d (TLR-deficient) mice by Greg Barton (University of California, Berkeley, CA). Mice were housed under SPF conditions in micro-isolator cages with food and water provided ad libitum. Mice were euthanized by overexposure to carbon dioxide. All procedures were approved by the UC Davis Animal Care and Use Committee.

Chimera generation

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Neonatal chimeric mice were generated as described previously (Lalor et al., 1989; Baumgarth et al., 2000; Baumgarth et al., 1999). Briefly, one-day old Igha C57BL/6 congenic mice were injected intraperitoneally with anti-IgMa (DS-1.1) diluted in PBS. On day 2 or three after birth mice were injected with total peritoneal cavity wash out, or with FACS-purified dump- CD19+ CD43+ CD5+ and/or CD5- B-1 cells from C57BL/6 (Ighb) mice. Over the next 6 weeks, these donor B-1 cells expand to fill all tested B-1 compartments, including the peritoneal and pleural body cavities, bone marrow, spleen, lymph nodes (mediastinal, mesenteric, inguinal, axilaris, cervical), gastrointestinal tract and lung, while host B-1 cells are depleted. Intraperitoneal anti-IgMa injections were continued twice weekly until mice reached 6 weeks of age. Mice were then rested for at least 6 weeks before use, for reconstitution of the conventional B cell populations from the host bone marrow. Due to the lack of significant B-1 cell development from bone marrow precursors after 6 weeks (Dorshkind et al.), the reconstituted mouse has exclusively host-derived B-2 cells (Igha) as well as a B-1 cell compartment that is 80–95% donor-derived (Ighb).

Influenza virus infection

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Influenza A/Puerto Rico/8/34 was grown and harvested as previously described (Doucett et al., 2005). Mice were anesthetized with isoflurane and virus was diluted to a previously titrated sublethal dose of infection and administered intranasally in PBS.

Salmonella typhimurium infection

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Oral infections with S. typhimurium were performed following previously described protocols (O'Donnell et al., 2015). S. typhimurium, strain SL1344, kindly provided by Stephen McSorley (University of California, Davis, CA), was grown overnight at 37°C in Luria-Bertani broth. A known volume of bacteria were centrifuged for 20 min at 6,000–8,000 rcf at 4°C after concentration was determined by spectrophotometer reading at OD600. Bacterial pellets were resuspended in mouse drinking water to a concentration of 109 CFU/ml. Water was provided to mice ad lib.

Flow cytometry and sorting

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Tissues were processed and stained as described previously (Rothaeusler and Baumgarth, 2006). Briefly, single cell suspensions of spleen, lymph node, and Peyer’s patches were obtained by grinding tissues between the frosted ends of two microscope slides, then resuspended in ‘Staining Media’ (Rothaeusler and Baumgarth, 2006). Peritoneal cavity washout was obtained by introducing Staining Media into the peritoneal cavity with a glass pipet and bulb, agitating the abdomen, and then removing the media. Samples were filtered through nylon mesh and treated with ACK lysis buffer as needed. Cell counts were performed using Trypan Blue exclusion to identify live cells.

Fc receptors were blocked using anti-CD16/32 antibody (2.4G2) and cells were stained using fluorochrome conjugates generated in-house unless otherwise specified against the following antigens: CD19 (clone ID3)-Cy5PE, allophycocyanin, FITC, CD4- (GK1.5), CD8a- (53–6.7), CD90.2- (30H12.1), Gr1- (RB68-C5), F4/80- (F4/80), and NK1.1- (PK136) Pacific blue (‘Dump’), CD43- (S7) allophycocyanin or PE, IgM- (331) allophycocyanin, Cy7-allophycocyanin, FITC, Alexa700, IgMa- (DS-1.1) allophycocyanin, biotin, IgMb- (AF6-78.2.5) allophycocyanin, FITC, biotin, CD5- (53–7.8) PE, biotin, IgD- (11-26) Cy7PE, Cy5.5PE, CD138- (281-2) allophycocyanin, PE; CD138-BV605 (BD Bioscience), CD19-BV786, PE-CF594 (BD Bioscience), SA-Qdot605 (Invitrogen), SA- allophycocyanin (eBioscience), BrDU-FITC (BD Bioscience), B220 (CD45R) APC-eFluor 780 (Invitrogen) and CD23-biotin (eBioscience), BV605, BV711 (BD Bioscience). PTC-FITC liposomes were a kind gift of Aaron Kantor (Stanford University, CA). Dead cells were identified using Live/Dead Fixable Aqua or Live/Dead Fixable Violet stain (Invitrogen).

Intracellular staining: Cells were surfaced stained, then fixed (eBiosience IC Fixation Buffer) for 30 min and then permeabilized (eBioscience Permeablization Buffer) for 30 min, followed by staining with anti-Nur77-Alexa Fluor 488 for 30 min all at room temperature.

Phosphoflow: Cells were fixed (BD Cytofix) for 12 min at 37°C. Cells were then permeabilized (BD Perm Buffer III) for 30 min on ice and intracellularly stained with anti-phospho-Akt-Alexa Fluor 488 for 30 min on ice.

FACS analysis was done using either a 4-laser, 22-parameter LSR Fortessa (BD Bioscience) or a 3-laser FACSAria (BD Bioscience). Cells were sorted as previously described (Rothaeusler and Baumgarth, 2006) using the FACSAria and a 100 µm nozzle. Data were analyzed using FlowJo software (FlowJo LLC, kind gift of Adam Treister).

Elisa

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Sandwich ELISA was performed as previously described (Rothaeusler and Baumgarth, 2006). Briefly, MaxiSorp 96 well plates (ThermoFisher) were coated with anti-IgM (Southern Biotech) and nonspecific binding was blocked with 1% NCS/0.1% dried milk powder, 0.05% Tween20 in PBS (‘ELISA Blocking Buffer’). Two-fold serial dilutions in PBS of culture supernatants and an IgM standard (Southern Biotech) were added to the plates at previously optimized starting dilutions. Binding was revealed with biotinylated anti-IgM (Southern Biotech), Streptavidin-Horseradish Peroxidase, both diluted in ELISA Blocking Buffer, and 0.005% 3,3’,5,5’-tetramethylbenzidine (TMB)/0.015% hydrogen peroxide in 0.05 M citric acid. The reaction was stopped with 1N sulfuric acid. Antibody concentrations were determined by measuring sample absorbance on a spectrophotometer (SpectraMax M5, Molecular Devices) at 450 nm (595 nm reference wavelength) and then compared to a standard curve created with a mouse IgM standard (Southern Biotech) of known concentration.

Culture and proliferation dye labeling

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After FACS sorting, cells were labeled with Efluor670 or CFSE at previously determined optimal concentrations, by incubation at 37°C for 10 mins., then washed three times with staining medium containing 10% neonatal calf serum and resuspended into ‘Culture Media’ (RPMI 1640 with 10% heat inactivated fetal bovine serum, 292 µg/ml L-glutamine, 100 Units/ml penicillin, 100 µg/ml streptomycin, and 50 µM 2-mercaptoethanol). Cells were plated at 105 cells/well of 96-well U bottom tissue culture plates (BD Bioscience), and unless otherwise indicated, cultured at 37°C/5% CO2 for 3 days. When indicated, LPS at 10 µg/ml, Mycobacterium TB lipids at 20 µg/ml (BIA), Imiquimod (R837, InvivoGen) at 1 µg/ml, CpG ODN 7909 at 5 µg/ml or anti-IgM (Fab)2 at 10–20 ug/ml were added to the wells. Cell enumeration after culture was performed using Molecular Probes CountBright Beads (Thermo Fisher) by flow cytometry, per manufacturer instructions. After culture, culture plates were spun and supernatant was collected and stored at −20°C, and cells were stained for FACS.

Elispot

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IgM antibody secreting cells were enumerated as previously described (Doucett et al., 2005). Briefly, 96 well ELISPOT plates (Multi-Screen HA Filtration, Millipore) were coated overnight with anti-IgM (331) or recombinant OmpD (MyBioSource) and non-specific binding was blocked with 4% Bovine Serum Albumin (BSA)/PBS. Cell suspensions were processed, counted, and directly plated in culture medium into ELISPOT wells and subsequently serially diluted two-fold, or they were FACS-sorted directly into culture media-containing ELISPOT wells. Cells were incubated overnight at 37°C/5% CO2. Binding was revealed with biotinylated anti-IgM (Southern Biotech), anti-IgMa (BD Bioscience), or anti-IgMb (BD Bioscience), Streptavidin-Horseradish Peroxidase (Vector Labs) both diluted in 2% BSA/PBS, and 3.3 mg 3-amino-9-ethylcarbazole (Sigma Aldrich) dissolved in dimethyl formamide/0.015% hydrogen perioxide/0.1M sodium acetate. The reaction was stopped with water. Spots were enumerated using the AID EliSpot Reader System (Autoimmun Diagnostika, Strassberg, Germany).

qRT-PCR

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mRNA was isolated from cells using the RNeasy mini kit (Qiagen), per manufacturer instructions. cDNA was generated using random hexamer primers and SuperSript II reverse transcriptase (Invitrogen). qRT-PCR was performed using commercially available Taqman primer/probes for cd5 and ubc (Thermo Fisher).

BrDU labeling

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Mice were injected intraperitoneally with 1 mg of BrDU (Sigma-Aldrich) per mouse diluted in 100 µL PBS, 24 hr before tissue collection. Staining for BrDU was performed as described previously (Rothaeusler and Baumgarth, 2006).

Proximity Ligation Assay (PLA)

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After FACS sorting, cells were resuspended in RPMI and rested for at least two hours before designated stimuli were added to culture media. Stimulated and unstimulated cells were cultured for 5 min, and 24 and 48 hr prior to PLA. PLA was performed as previously described (Kläsener et al., 2014). In brief: For PLA-probes against specific targets, the following unlabeled Abs were used: anti-IgM (Biolegend, clone RMM-1), anti-CD79a (Thermo Fisher, clone 24C2.5), anti-CD5 (Biolegend, clone 53–7.3), anti-Syk (Biolegend, clone Syk-01), and anti-CD19 (Biolegend, clone 6D5). Fab fragments against CD79a, Syk, IgM, and CD19 were prepared with Pierce Fab Micro preparation kit (Thermo Scientific) using immobilized papain according to the manufacturer’s protocol. After desalting (Zeba spin desalting columns, Thermo Scientific), all antibodies were coupled with PLA Probemaker Plus or Minus oligonucleotides (Sigma-Aldrich) to generate PLA-probes. For in situ PLA, B cells were settled on polytetrafluoroethylene slides (Thermo Fisher Scientific) for 30 min at 37°C. BCR. Cells were fixed with paraformaldehyde 4%, for 20 min. For intracellular PLA, B cells were permeabilized with 0,5% Saponin for 30 min at room temperature, and blocked for 30 min with Blocking buffer (containing 25 μg/ml sonicated salmon sperm DNA, and 250 μg/ml bovine serum albumin). PLA was performed with the Duolink In-Situ-Orange kit. Resulting samples were directly mounted on slides with DAPI Fluoromount-G (SouthernBiotech) to visualize the PLA signals in relationship to the nuclei. Microscope images were acquired with a Leica DMi8 microscope, 63 oil objective (Leica-microsystems). For each experiment a minimum of 100 B-1a/B-1b/B-2 peritoneal cavity or 1000 splenic B-2 cells from several images were analyzed with CellProfiler-3.0.0 (CellProfiler.org). Raw data were exported to Prism7 (GraphPad, La Jolla, CA). For each sample, the mean PLA signal count per cell was calculated from the corresponding images and the statistical significance with Mann–Whitney test.

Statistical analysis

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Statistical analysis was done using a two-tailed Student t test with help of Prism software (GraphPad Software). For time-course data, an ANOVA was performed with the help of Prism software, and if significant, Student t tests were performed to determine which time points were significant. When multiple comparisons were run on the same sets of data, Holm-Sidak correction was applied, using Prism software. p<0.05 was considered statistically significant.

References

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    The primary B cell response to the O/core region of bacterial lipopolysaccharide is restricted to the Ly-1 lineage
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    Journal of Immunology 146:327–331.
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Decision letter

  1. Andrew J MacPherson
    Reviewing Editor; University of Bern, Switzerland
  2. Wendy S Garrett
    Senior Editor; Harvard T.H. Chan School of Public Health, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "TLR-signaling induces reorganization of the IgM-BCR complex regulating B-1 cell responses to infections" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Wendy Garrett as the Senior Editor. The reviewers have opted to remain anonymous.

Thank you very much for your patience in the reviewing process. Since the paper was submitted with "B cell receptor and TLR signaling coordinate to control distinct B-1 responses to both self and the microbiota" by Dr Barton and colleagues, we aimed to secure reviewers (Nos. 1 and 2) for both papers, so that the complementary nature could be fairly considered. The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

A key point in the discussion was the B cell-intrinsic nature of the TLR signaling in the other paper. We have requested a revision of both papers, and we would be grateful if you could liaise especially on this point to call out the complementarity.

Reviewer #1:

In this manuscript, Baumgarth et al. describe their discovery and characterization of a novel mechanism by which innate signals can license B1 cells to respond to antigen receptor signaling and produce IgM. Specifically, they show that B cell-intrinsic TLR signaling can trigger the downregulation of the inhibitor of BCR signaling CD5, which effectively blocks antigen recognition-based activation of CD5+ B-1 cells. In so doing, the authors resolve multiple long-standing mysteries surrounding the mechanisms by which CD5+ B1 cells respond to antigen-dependent stimuli, overturn previously held views regarding the cellular origins of specific IgM responses, and uncover a defined mechanism by which B cells integrate innate and adaptive stimuli. The experiments that support these main conclusions are both elegant and, in my opinion, performed to an exceptionally high standard-e.g., the extensive use of neonatal B-1 allotype-chimeras enabled detailed and definitive examinations of the roles of B cell subsets in vivo. Overall, I have little to criticize regarding the veracity of the work and believe that this will be an important paper. My one remaining confusion has to do with the order of engagement of the innate versus adaptive stimuli. The authors speculate a bit about this in the discussion, but as far as I can tell don't directly address this experimentally. To me, the logical order of events under in vivo conditions (in response to self-derived debris or infection) would be BCR mediated uptake of complexes that contain innate stimuli. However, the experiments performed by the authors in vitro instead seem to test the opposite scenario-where TLR activation precedes BCR activation and licenses B cells to respond to antigen. I think that a few additional in vitro experiments that address how the timing of exposure to these signals affects B1 responsiveness could at least begin to clarify some of these issues and thus improve the paper. For example, does prior engagement of the BCR in CD5+ B1 cells stabilize the inhibitory complex and prevent future licensing by TLRs? Also, can CD5+ B1 cells acquire antigen through BCR mediated endocytosis despite the blunting of signaling by CD5 (the answer to this may already be known)? If not, then how do they respond to TLR stimuli such as nucleic acids from dying cells that would typically be recognized in endosomes after BCR-mediated endocytosis? Or, is this licensing in vivo restricted mostly to TLR ligands that can either signal on the cell surface or be readily acquired by other mechanisms (pinocytosis)?

Reviewer #2:

In this manuscript, the authors showed that CD5 negative B1 cells were the source of IgM producing cells in the draining lymph nodes upon influenza infection. These CD5 negative B1 cells were originally CD5 positive B1 cells, and down-regulated CD5 expression during cell proliferation upon TLR stimulation. In vitro experiments showed that the down-regulation of CD5 was at the level of gene expression, and was not due to the expansion of contaminated originally CD5 negative B1 cells. The B1 cells secreted significantly higher amount of IgM after the down-regulation of CD5 when stimulated with TLR ligands. An in vivo approach using neonatal chimeric mice showed that CD5 positive B1 cells lose CD5 expression in the draining lymph nodes upon influenza or Salmonella infection, and these CD5 negative B1 cells secrete high amount of IgM. The molecular analysis of purified B1 cells showed that the down-regulation of CD5 enhanced the BCR signaling by releasing from the suppressive effect of CD5. Finally, TLR-/- CD5 positive B1 cells were not able to down-regulate CD5 and thus failed to elicit IgM response upon the influenza infection.

This study addressed a long-standing question how B1 cells secrete a large amount of IgM by rapidly responding to infections. Importantly, the authors demonstrated that IgM-producing CD5 negative B1b cells that were previously thought different origin from CD5 positive B1a cells, were phenotypically indistinguishable from TLR-stimulated B1a cells. This finding is exciting in the field of B1 cell biology, as it requests reconsideration for the interpretation of previous studies.

In general, the experiments were well done, but the data presentation and explanations could be improved. For example, some of the data could be moved to supplementary information to improve readability. In addition, some more elaboration would be required for a few of the data (see below). Although it is not strictly required, the hypothesis that TLR-signaling may "license" BCR-dependent reaction could be more deeply explored, as it is functionally important. For example, in Figure 7, what happens for the BCR-signaling in vitro if B1 cells are stimulated both with TLR-ligands and anti-IgM? Also, in Figure 8, does VDJ repertoire skew upon influenza infection due to the "licensed BCR reaction" in WT B1 cells, which may be disappeared in TLR-/- B1 cells? These data may add further value to this study.

Specific points:

Results section, paragraph four.

B-1(Igha) and B-2(Ighb) cells should read B-1(Ighb) and B-2(Igha) cells.

Results section, paragraph five.

“we expanded the analysis to include all IgMb-expressing (B-1 donor-derived) and IgMa- (recipient-derived) cells, regardless of expression of CD19 or other surface markers.” Please provide the gating strategy together with Figure 1F, as this gating is important to show the increased CD5-negative population.

Figure 2H-I: Please provide explanation for the purpose of this experiment. The authors concluded that "CD5+ B-1 cells lose CD5 surface and mRNA expression after in vitro LPS stimulation" based on this experiment. However, these experiments show that "CD5+B1 cells survive better, and also CD5+ and CD5- B1 cells proliferate similarly upon LPS stimulation". Thus, the data do not support the conclusion in the text. Please provide better explanation.

Results paragraph eight: Figure 3J-K should read Figure 2J-K.

Figure 3C-D: Why did the authors use Mtb lipids? Is this used as the alternative TLR4-ligand, which is in a different form from LPS? Please provide some explanation.

Figure 4F: Which chimeric mice were used for this analysis?

Figure 5: Please carefully check this figure and legend, because there are many typos here. This figure may be moved to supplementary information.

Title of the figure legend: “…and spleen after S. typhimurium infection” No data is shown for spleen.

Figure 5B: Graph label for the Y-axis: #CD5+ B-1 cells MedLN should read MesLN ?

Figure 5B: Why the numbers of CD5+ B-1 cell significantly reduced in 95% CD5+ chimera comparing with CD5+ chimera in mesenteric LN?

Figure 5B contain 2 graphs (MesLN and Peyer's patch), but the legend indicates different presentation: (B) MesLN and (C) Peyer's patch.

Third paragraph of subsection “CD5+ B-1 cells become CD5- IgM ASC in the Mesenteric LNs and Peyer’s Patches after Salmonella typhimurium infection”:

– Figure 5F is not provided.

– “Instead, we found OmpD-specific IgM secretion by B-2 derived plasmablasts.” should read “Instead, we found OmpD-specific IgM secretion by B-2 derived plasmablasts in mesenteric LN” (?).

– “Of note, the phenotype of B-2 derived plasmablasts is indistinguishable from that of "B-1b" cells (CD19lowCD45loIgM+CD43+ (Figure 5F)),…” Please provide the data to support this description (Figure 5F).

Reviewer #3:

In this interesting manuscript, Savage and colleagues provide evidence that innate-like CD5+ B1 cells respond to innate signals by production of IgM and differentiation into CD138+ plasmablasts in secondary lymphoid tissues. This was already known, however Savage and colleagues here demonstrate that CD5+ B1 cells are the actual TLR-responsive cells, and that TLR signaling activates these cells and results in downregulation of CD5. Potentially, this leads to disruption of CD5-IgM-BCR association and allows for subsequent BCR-medicated signaling, also the latter is not demonstrated.

This manuscript proposes a model in which CD5- B1 cells (B1b cells) are derived from CD5+ B1 cells (B1a cells), and reflect a post-activation state. This opposes the theory of 'division of labor' of B1a and B1b cells.

The manuscript is well written and the data are well presented. The overall message is interesting and adds to our knowledge of these innate-like B cells.

Questions/remarks:

1) As a general remark, the authors mostly focus on secondary lymphoid tissues, in which the differentiation into plasma cells probably occurs. Why do the authors not investigate the body cavities, where initial activation of B1 cells occurs? Downregulation of CD5 likely already occurs at this site.

2) What is the rationale of choosing the neonatal chimeric model as described in the Materials and methods for pleural cavity and mediastinal lymph node B1 cells, as the model is peritoneum-oriented (anti-IgM treatment and transfer of cells is i.p.)? This should be better explained.

3) The authors should include the transfer of CD5- cells as a control (now only 100% CD5+, 99% or 95% CD5+). Do CD5-B1 cells also upregulate CD138 upon TLR signaling?

4) Is there plasticity of CD5+ and CD5- B1 cells or is their fate set (in the pre-PC differentiated state)? mRNA (Figure 2) suggests transcriptional regulation of CD5 expression by TLR signaling (is this significant?). Is CD5 mRNA and surface expression restored upon removal of TLR agonists in vitro? This could be easily tested, and would add to the message of this manuscript.

5) Figure 6: the PLA data are difficult to interpret and may be redundant, as CD5 surface expression was already been shown to be lost upon TLR signals. This obviously affects the assay and it is thus questionable as of whether these results are useful. In addition, how can CD5 be detected in the PLA in CD5- B1 cells? (Figure 6A, panel 3).

6) Many plots are gated from IgM+cells (i.e. Figure 1A and Figure 8A): is it possible that the authors miss populations, as the B1 cells may have already undergone CSR, for example to IgA? Did they check for IgA+ or IgG+ B1-like cells? (dump-CD23-CD43+CD19+).

https://doi.org/10.7554/eLife.46997.014

Author response

A key point in the discussion was the B cell-intrinsic nature of the TLR signaling in the other paper. We have requested a revision of both papers, and we would be grateful if you could liaise especially on this point to call out the complementarity.

We have inserted a more extensive discussion on the paper by Barton and colleagues in paragraphs two, three and five of the Discussion section.

Reviewer #1:

In this manuscript, Baumgarth et al. describe their discovery and characterization of a novel mechanism by which innate signals can license B1 cells to respond to antigen receptor signaling and produce IgM. Specifically, they show that B cell-intrinsic TLR signaling can trigger the downregulation of the inhibitor of BCR signaling CD5, which effectively blocks antigen recognition-based activation of CD5+ B-1 cells. In so doing, the authors resolve multiple long-standing mysteries surrounding the mechanisms by which CD5+ B1 cells respond to antigen-dependent stimuli, overturn previously held views regarding the cellular origins of specific IgM responses, and uncover a defined mechanism by which B cells integrate innate and adaptive stimuli. The experiments that support these main conclusions are both elegant and, in my opinion, performed to an exceptionally high standard-e.g., the extensive use of neonatal B-1 allotype-chimeras enabled detailed and definitive examinations of the roles of B cell subsets in vivo. Overall, I have little to criticize regarding the veracity of the work and believe that this will be an important paper. My one remaining confusion has to do with the order of engagement of the innate versus adaptive stimuli. The authors speculate a bit about this in the discussion, but as far as I can tell don't directly address this experimentally. To me, the logical order of events under in vivo conditions (in response to self-derived debris or infection) would be BCR mediated uptake of complexes that contain innate stimuli. However, the experiments performed by the authors in vitro instead seem to test the opposite scenario-where TLR activation precedes BCR activation and licenses B cells to respond to antigen. I think that a few additional in vitro experiments that address how the timing of exposure to these signals affects B1 responsiveness could at least begin to clarify some of these issues and thus improve the paper. For example, does prior engagement of the BCR in CD5+ B1 cells stabilize the inhibitory complex and prevent future licensing by TLRs? Also, can CD5+ B1 cells acquire antigen through BCR mediated endocytosis despite the blunting of signaling by CD5 (the answer to this may already be known)? If not, then how do they respond to TLR stimuli such as nucleic acids from dying cells that would typically be recognized in endosomes after BCR-mediated endocytosis? Or, is this licensing in vivo restricted mostly to TLR ligands that can either signal on the cell surface or be readily acquired by other mechanisms (pinocytosis)?

We thank the reviewers for those comments. The question raised by this reviewer (and also reviewer #2) is very important and one we are currently considering how best to explore in the future. We agree that the order of stimulation (BCR/TLR) is not fully addressed in this manuscript. We expect additional extensive work is required to fully address this critical point which we believe to be outside the scope of this manuscript. However, in response to the comment, we have conducted additional experiments in vitro, now shown in a new Figure 7 and subsection “Initial stimulation through IgM-BCR suppresses subsequent activation of B-1 cells via TLR- stimulation” of the revised manuscript, in which we studied the effects of pulsed stimulation with one stimuli (either BCR or TLR for 2h and 24h followed by wash-out), followed by exposure to the second stimulus (for 70h and 48h, respectively). At 72h after initial stimulation we measured B-1 cell proliferation.

Stimulation for 2h with either ligand, followed by 70h stimulation with the other ligand showed no significant difference compared to 72h stimulation with the second stimulus alone (data not shown). However, when we exposed B-1 cells to the first stimulus for 24h and then the other stimulus for an additional 48h we found significant effects. Stimulation first with anti-IgM (BCR-signaling) and then CpG resulted in significant reductions in B-1 cell proliferation compared to stimulation only with CpG. In contrast, 24h stimulation with CpG followed by 48h stimulation via the BCR resulted in significant enhanced B-1 cell proliferation compared to BCR stimulation alone, however, still less than CpG only (Figure 7).

The data indicate that TLR stimulation must proceed BCR-stimulation for B-1 cells to be able to maximally respond to the innate stimulus. These data are consistent with the PLA experiments, which suggested strong BCR signaling-enhancing effects of TLR stimulation, while initial BCR stimulation enhanced the dissociation of the BCR with positive signaling components of the BCR complex.

How these findings relate to the findings by Barton and colleagues in the companion paper, in which they find effects of TLR-signaling on antigen specificity of the B cell response is now outlined in more detail in paragraphs two, three and five of the Discussion section.

Our current view is that the TLR-signal enhances potential responsiveness of B-1 cells in vivo, followed by a BCR-mediated signal that would then drive antigen-specific expansion of those cells that have already received a TLR signal. We are not aware of studies that have studied antigen uptake via BCR-internalization by B-1 cells. There are reports, however, that B-1 cells can uptake antigen via phagocytosis (Popi, Longo-Maugeri and Mariano, 2016), which would explain engagement of specific endosomal TLRs without first engagement and internalization of the BCR. This clearly requires further exploration, which we believe is outside the scope of this manuscript.

Importantly, for conventional B cells TLR-mediated stimulation may function quite differently, given their ability to vigorously respond to BCR-signaling alone. Here antigen-binding via the BCR would cause antigen/BCR-internalization and then exposure of the antigen to endosomal TLRs, enhancing antigen-specific responses.

Reviewer #2:

In this manuscript, the authors showed that CD5 negative B1 cells were the source of IgM producing cells in the draining lymph nodes upon influenza infection. These CD5 negative B1 cells were originally CD5 positive B1 cells, and down-regulated CD5 expression during cell proliferation upon TLR stimulation. In vitro experiments showed that the down-regulation of CD5 was at the level of gene expression, and was not due to the expansion of contaminated originally CD5 negative B1 cells. The B1 cells secreted significantly higher amount of IgM after the down-regulation of CD5 when stimulated with TLR ligands. An in vivo approach using neonatal chimeric mice showed that CD5 positive B1 cells lose CD5 expression in the draining lymph nodes upon influenza or Salmonella infection, and these CD5 negative B1 cells secrete high amount of IgM. The molecular analysis of purified B1 cells showed that the down-regulation of CD5 enhanced the BCR signaling by releasing from the suppressive effect of CD5. Finally, TLR-/- CD5 positive B1 cells were not able to down-regulate CD5 and thus failed to elicit IgM response upon the influenza infection.

This study addressed a long-standing question how B1 cells secrete a large amount of IgM by rapidly responding to infections. Importantly, the authors demonstrated that IgM-producing CD5 negative B1b cells that were previously thought different origin from CD5 positive B1a cells, were phenotypically indistinguishable from TLR-stimulated B1a cells. This finding is exciting in the field of B1 cell biology, as it requests reconsideration for the interpretation of previous studies.

In general, the experiments were well done, but the data presentation and explanations could be improved. For example, some of the data could be moved to supplementary information to improve readability.

We thank the reviewer for these comments. We have now moved part of our data into two supplemental figures. Figure 2—figure supplement 1 contains data demonstrating that expansion of CD5- B-1 cells is not due to enhanced proliferation or survival by small frequencies of contaminating CD5- cultures at onset. Figure 5—figure supplement 1 now contains the data from the Salmonella infection experiments, which demonstrate that similar to what was observed after influenza infection, it is the CD5+ B-1 cell that responds to infection with downregulation of CD5 and differentiation to an IgM secreting cell. We hope that this together with changes listed below allows for an easier reading of the manuscript.

In addition, some more elaboration would be required for a few of the data (see below). Although it is not strictly required, the hypothesis that TLR-signaling may "license" BCR-dependent reaction could be more deeply explored, as it is functionally important. For example, in Figure 7, what happens for the BCR-signaling in vitro if B1 cells are stimulated both with TLR-ligands and anti-IgM?

As outlined in response to reviewer #1, who had similar questions, we now provide a new Figure 7 with in vitro data analyzing how the order of TLR and BCR signaling affects B-1 cell responses. We refer to the above response for details.

Also, in Figure 8, does VDJ repertoire skew upon influenza infection due to the "licensed BCR reaction" in WT B1 cells, which may be disappeared in TLR-/- B1 cells? These data may add further value to this study.

We thank the reviewer for that question. We agree that assessing the repertoire of B-1 cells in the mediastinal lymph nodes of influenza infected mice would be highly informative. Unfortunately, this is also a very challenging study to do, given the small numbers of B-1 cells that can be obtained from one mouse (about 2,000 – 5,000 total B-1 cells per animal). We hope that recent advances in single-cell RNA sequencing will help us to do this in the future. In terms of overall influenza-binding IgM-secreting cells, we previously published (Choi and Baumgarth, 2008) that the frequency of influenza-binding cells was similar in the MedLN between days 7 and 10 after infection and also similar compared to splenic B-1 cells prior to infection by ELISPOT analysis. Thus, we were unable to demonstrate influenza-specific clonal expansion. Thus, either the ELISPOT analysis lacked the necessary sensitivity to demonstrate expansion of influenza-binding B-1 cells, or the specificity of B-1 cells might be direct against altered self-antigens, rather than influenza virus and thus possibly triggered by DAMPS rather than PAMPS.

Specific points:

Results section, paragraph four.

B-1(Igha) and B-2(Ighb) cells should read B-1(Ighb) and B-2(Igha) cells.

We apologize for this mistake – it has been corrected in the revised manuscript.

Results section, paragraph five.

“we expanded the analysis to include all IgMb-expressing (B-1 donor-derived) and IgMa- (recipient-derived) cells, regardless of expression of CD19 or other surface markers.” Please provide the gating strategy together with Figure 1F, as this gating is important to show the increased CD5-negative population.

In response we expanded the data shown to include all IgMb-expressing (B-1 donor-derived) and IgMa- (recipient-derived) cells, regardless of expression of CD19 or other surface markers. The data are added as a new part to Figure 1H.

Figure 2H-I: Please provide explanation for the purpose of this experiment. The authors concluded that "CD5+ B-1 cells lose CD5 surface and mRNA expression after in vitro LPS stimulation" based on this experiment. However, these experiments show that "CD5+B1 cells survive better, and also CD5+ and CD5- B1 cells proliferate similarly upon LPS stimulation". Thus, the data do not support the conclusion in the text. Please provide better explanation.

We are sorry for the confusion. We have moved subfigures 2F-I into the new Figure 2—figure supplement 1 to deconvolute the data. We have also reworded the text to make the purpose for the experiments clearer (subsection “CD5+ B-1 cells decrease CD5 expression after LPS stimulation in vitro”). In short, the question we attempted to address was whether the expanding population of CD5- cells we see emerge after stimulation of B-1 cells with TLRs could have arisen from small contaminations with CD5- cells that would survive better and/or proliferate stronger in response to the stimulus. We did not find that this was the case. Instead, if anything CD5- cells spiked into the cultures died faster and proliferated less than the CD5+ cells. From that we concluded that the increase in CD5lo/- cells in the culture was due to the loss of CD5 expression of originally CD5+ cells.

Results paragraph eight: Figure 3J-K should read Figure 2J-K.

We have moved this data now into the new Figure 2—figure supplement 1 and reordered the figure.

Figure 3C-D: Why did the authors use Mtb lipids? Is this used as the alternative TLR4-ligand, which is in a different form from LPS? Please provide some explanation.

We now provide information and a reference (Basu, Shin and Jo, 2012) to indicate that the Mtb lipid provides a TLR 2 agonist.

Figure 4F: Which chimeric mice were used for this analysis?

We apologize for this not being clear. The entire figure was done with studies using the same sets of neonatal chimeras as outlined in the first sentence of the figure legend and then infected for 7 days with influenza A/PR8. To ensure this is clear we have now indicated this also in the Figure legend to Figure 4F.

Figure 5: Please carefully check this figure and legend, because there are many typos here. This figure may be moved to supplementary information.

We apologize for these errors. The figure has been carefully edited and moved to the supplemental information (Figure 2).

Title of the figure legend: “…and spleen after S. typhimurium infection” No data is shown for spleen.

We removed the reference to the spleen data.

Figure 5B: Graph label for the Y-axis: #CD5+ B-1 cells MedLN should read MesLN ?

We corrected the name of the lymph node.

Figure 5B: Why the numbers of CD5+ B-1 cell significantly reduced in 95% CD5+ chimera comparing with CD5+ chimera in mesenteric LN?

The chimeras always show a larger degree of variation in the sizes of their lymph nodes compared to that of non-chimeric mice. This is likely due to the fact that are set-up as a mix of males and females (it is difficult to determine the sex of a 1-day old mouse). B-1 cells are often somewhat more prevalent in females than in males. However, overall we found that the lymph nodes had similar levels of IgM secretion (5C). We therefore believe the variation to be not to be of biological significance (albeit reporting statistical significance) for the work done here. We now address this point in paragraph two of subsection “CD5+ B-1 cells become CD5- IgM ASC in the Mesenteric LNs and Peyer’s Patches after Salmonella typhimurium infection”.

Figure 5B contain 2 graphs (MesLN and Peyer's patch), but the legend indicates different presentation: (B) MesLN and (C) Peyer's patch.

We apologize for this mistake. The legend has been corrected.

Third paragraph of subsection “CD5+ B-1 cells become CD5- IgM ASC in the Mesenteric LNs and Peyer’s Patches after Salmonella typhimurium infection”:

– Figure 5F is not provided.

The numbering of the figures has been updated.

– “Instead, we found OmpD-specific IgM secretion by B-2 derived plasmablasts.” should read “Instead, we found OmpD-specific IgM secretion by B-2 derived plasmablasts in mesenteric LN” (?).

We have altered the text as suggested.

– “Of note, the phenotype of B-2 derived plasmablasts is indistinguishable from that of "B-1b" cells (CD19lowCD45loIgM+CD43+ (Figure 5F)),…” Please provide the data to support this description (Figure 5F).

We have added a subfigure (Figure 5—figure supplement 1F) showing both IgMb+ and IgMa+ cells among the CD19lowCD43+CD138+ (IgM+) compartment in the MesLN.

Reviewer #3:

In this interesting manuscript, Savage and colleagues provide evidence that innate-like CD5+ B1 cells respond to innate signals by production of IgM and differentiation into CD138+ plasmablasts in secondary lymphoid tissues. This was already known, however Savage and colleagues here demonstrate that CD5+ B1 cells are the actual TLR-responsive cells, and that TLR signaling activates these cells and results in downregulation of CD5. Potentially, this leads to disruption of CD5-IgM-BCR association and allows for subsequent BCR-medicated signaling, also the latter is not demonstrated.

This manuscript proposes a model in which CD5- B1 cells (B1b cells) are derived from CD5+ B1 cells (B1a cells), and reflect a post-activation state. This opposes the theory of 'division of labor' of B1a and B1b cells.

The manuscript is well written and the data are well presented. The overall message is interesting and adds to our knowledge of these innate-like B cells.

Questions/remarks:

1) As a general remark, the authors mostly focus on secondary lymphoid tissues, in which the differentiation into plasma cells probably occurs. Why do the authors not investigate the body cavities, where initial activation of B1 cells occurs?Downregulation of CD5 likely already occurs at this site.

We thank the reviewer for this comment. As we outlined in the Introduction of the manuscript, our previous work has shown that the cells entering the MedLN after influenza infection are nearly exclusively CD5+ B-1 cells (Choi and Baumgarth, 2008; Waffarn et al., 2015). The activation signal that traps these cells in the draining lymph node was shown by us to be the activation of the integrin CD11b, which was flipped into an active state via Type I IFNR signaling of pleural cavity B-1 cells by 2-days following influenza infection. At that timepoint the cells express CD5 at normal levels. While total numbers of CD5+ B-1 cells decline in the pleural cavity, the CD5- cells stay unaltered, suggesting that the CD5+ cells leave the pleural cavity before downregulating CD5 (Waffarn et al., 2015). We also generated CD5- only chimeras to demonstrate that CD5- B-1 cells cannot enter the lymph nodes (Choi et al., 2008). Based on these data, and the fact that IgM responses occur in the lymph nodes and not the body cavities, our focus has been on the secondary lymphoid tissues.

2) What is the rationale of choosing the neonatal chimeric model as described in the Materials and methods for pleural cavity and mediastinal lymph node B1 cells, as the model is peritoneum-oriented (anti-IgM treatment and transfer of cells is i.p.)? This should be better explained.

We apologize for this lack of clarity. The only way to inject a newborn mouse with B-1 cells is via the i.p. route. As was established in the original protocols we cited in the Materials and methods section and our own subsequent work (Lalor et al., 1989, Baumgarth et al., 1999, Baumgarth et al., 2000, Choi and Baumgarth, 2008, Choi et al., 2012, Waffarn et al., 2015 and Savage et al., 2017), subsequent to their injection i.p. B-1 cells in the neonate will expand into all compartments of the mouse, including both pleural and peritoneal cavities, spleen, bone marrow, all lymph nodes tested (mesenteric, mediastinal, inguinal, axillaris, cervical) as well as the gastrointestinal tract (Kroese et al., 1989) and lung (Baumgarth, unpublished). Thus, rather than seeing this chimera as a mouse with peritoneal cavity B-1 cells of a different allotype, the chimera has nearly all of its B-1 cell compartment exchanged for that of the donor. We have expanded the Materials and methods subsection “Chimera generation” to provide more information on these chimeras. We have also clarified the rationale for the use of the chimeras in the manuscript text in the first and fifth paragraphs of the revised Results section.

3) The authors should include the transfer of CD5- cells as a control (now only 100% CD5+, 99% or 95% CD5+). Do CD5-B1 cells also upregulate CD138 upon TLR signaling?

We thank the reviewer for this comment. We have previously conducted such a study and demonstrated that mice reconstituted with only CD5- B-1 cells have only very few MedLN B-1 cells and of those few that made it (about 200 B-1 cells in the entire lymph node), 65% of which expressed CD5, potential contaminants of our FACS-sorts, although we cannot exclude “reversion” to CD5+ status (Choi et al., 2008). For that reason we felt it was better in this manuscript to provide a range of mixes of CD5+ and CD5-, as this would expand on the data, by enabling us to correlate the secretion of IgM and CD5 expression with reconstitution frequencies. Importantly, our hypothesis to be tested was that CD5+ cells would lose CD5 expression. We now provide a more detailed description of that previous experiment and the rationale for the study, as conducted in subsection “CD5+ B-1 cells become CD5- IgM ASC in the MedLN after Influenza infection”.

4) Is there plasticity of CD5+ and CD5- B1 cells or is their fate set (in the pre-PC differentiated state)? mRNA (Figure 2) suggests transcriptional regulation of CD5 expression by TLR signaling (is this significant?). Is CD5 mRNA and surface expression restored upon removal of TLR agonists in vitro? This could be easily tested, and would add to the message of this manuscript.

We thank the reviewer for these comments. We have analyzed CD5 expression in cells that were pulsed with CpGs followed by 2-day exposure to anti-IgM stimulation or medium only. During that timeframe we did not see what appeared to be reexpression CD5 (not shown). The difficulty with interpretation of the data from those experiments is, however, that not all cells are stimulated to lose CD5 and it is not clear whether those expressing CD5 never lost its expression or potentially re-expressed it. Thus, although a “simple question” in principle, experimentally this is not quite as easy to address. Additional experiments that would test the effects of BCR-stimulation and/or removal of CpG stimulation followed by conditions that allowed B-1 cell survival without further stimulation would need to be developed, which we believe to be outside the scope of this manuscript.

5) Figure 6: the PLA data are difficult to interpret and may be redundant, as CD5 surface expression was already been shown to be lost upon TLR signals. This obviously affects the assay and it is thus questionable as of whether these results are useful. In addition, how can CD5 be detected in the PLA in CD5- B1 cells? (Figure 6A, panel 3).

We believe the PLA data to be of utmost important for interpretation of our studies. They demonstrate effects of CpG stimulation and BCR-engagement that are much more fine-grained than we can measure by surface staining only. What they demonstrate is that CD5 association with IgM is actually rapidly enhanced after BCR-signaling, while it is lost upon stimulation with CpGs – even at a time where the CD5 protein is still expressed on the cell surface. Thus, we see changes in the BCR complex configuration well before we see a loss of surface expression of CD5. Furthemore, we demonstrate effects of CpG stimulation on CD79a:Syk interaction, clearly linking TLR and BCR-complex signals.

To more clearly make this point we have rewritten the corresponding text in paragraph three of subsection “Changes in BCR signaling following innate activation of B-1 cells”. The very low frequencies expression of CD5 on the CD5- cell are presumably small impurities in the FACS-sort. We chose a figure that include one of those impurities for fullest disclosure, but as can be seen in the summary figure there frequencies are very low.

6) Many plots are gated from IgM+cells (i.e. Figure 1A and Figure 8A): is it possible that the authors miss populations, as the B1 cells may have already undergone CSR, for example to IgA? Did they check for IgA+ or IgG+ B1-like cells? (dump-CD23-CD43+CD19+)

We thank the reviewer for this comment. Previous work by us (Baumgarth et al., 1999) showed that for those antibodies for which we can discern allotypic differences (IgM, IgA, IgG1, IgG2a/c), the major contribution of B-1 cells was the generation of IgM. Hence we have focused on this response in our work and the goal of this manuscript is the study of IgM responses by B-1 cells. We agree with the reviewer, however, that B-1 cells may generate other isotypes. Of particular interest is IgG3. We showed data supporting strong contributions of IgG3 by B-1 cells in the steady state (Savage et al., 2017) and our colleagues in the Barton lab found previously that much microbial antibodies are of the IgG3 isotype that are generated by B-1 cells (Koch et al., 2016). The companion paper outlines their follow-up work (Kreuk et al., 2019). Unfortunately, IgG3 has no allotypic variation among mouse strains, and thus our chimera approach cannot distinguish B-1 from B-2-derived antibody secretion. In unpublished studies we looked at IgA production in the lung tissue before and after influenza infection and found the levels of IgA-production by B-1 cells to not be affected by the infection (Baumgarth, not published). However, a more detailed analysis on the MedLN might be warranted.

https://doi.org/10.7554/eLife.46997.015

Article and author information

Author details

  1. Hannah P Savage

    1. Center for Comparative Medicine, University of California, Davis, Davis, United States
    2. Graduate Group in Immunology, University of California, Davis, Davis, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1057-7239
  2. Kathrin Kläsener

    1. BIOSS Centre for Biological Signalling Studies, University of Freiburg, Freiburg, Germany
    2. Department of Molecular Immunology, Institute of Biology III at the Faculty of Biology of the University of Freiburg, Freiburg, Germany
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5969-2553
  3. Fauna L Smith

    1. Graduate Group in Immunology, University of California, Davis, Davis, United States
    2. Integrated Pathobiology Graduate Group, University of California, Davis, Davis, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  4. Zheng Luo

    Center for Comparative Medicine, University of California, Davis, Davis, United States
    Contribution
    Resources, Formal analysis, Validation
    Competing interests
    No competing interests declared
  5. Michael Reth

    1. BIOSS Centre for Biological Signalling Studies, University of Freiburg, Freiburg, Germany
    2. Department of Molecular Immunology, Institute of Biology III at the Faculty of Biology of the University of Freiburg, Freiburg, Germany
    Contribution
    Conceptualization, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  6. Nicole Baumgarth

    1. Center for Comparative Medicine, University of California, Davis, Davis, United States
    2. Graduate Group in Immunology, University of California, Davis, Davis, United States
    3. Integrated Pathobiology Graduate Group, University of California, Davis, Davis, United States
    4. Department of Pathology, Microbiology and Immunology, School of Veterinary Medicine, University of California, Davis, Davis, United States
    Contribution
    Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing—original draft, Writing—review and editing
    For correspondence
    nbaumgarth@ucdavis.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2891-4483

Funding

National Institute of Allergy and Infectious Diseases (U19-AI109962)

  • Nicole Baumgarth

National Institute of Allergy and Infectious Diseases (R01-AI117890)

  • Nicole Baumgarth

National Institutes of Health (T32 OD011147)

  • Fauna L Smith

National Institutes of Health (T32 AI060555)

  • Hannah P Savage

National Institutes of Health (2T32OD010931-09)

  • Hannah P Savage

National Institutes of Health (5T35OD010956-14)

  • Hannah P Savage

National Center for Advancing Translational Sciences (TL1 TR000133)

  • Hannah P Savage

The German Federal and State Governments Excellence Initiative (EXC 294)

  • Kathrin Kläsener
  • Michael Reth

Deutsche Forschungsgemeinschaft (TRR130-P02)

  • Kathrin Kläsener
  • Michael Reth

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

Acknowledgements

We thank Drs. Joseph Benoun and Stephen McSorley (UC Davis) for expert help in conducting infections with S typhimurium, Dr. Aaron Kantor (Stanford University) for PtC liposomes, and Dr. Greg Barton (UC Berkeley) for TLR-deficient mice and discussions. This work was supported through NIH/NIAID grants U19-AI109962 (NB) and R01-AI117890 (NB), the National Center for Advancing Translational Sciences, NIH, through grant number UL1 TR000002 and linked award TL1 TR000133 (HPS), the NIH −2T32OD010931-09 (HPS), NIH – 5T35OD010956 (HPS) and the T-32 AI060555 (HPS) and NIH- T32 OD011147 (FLS), and through the Excellence Initiative of the German Federal and State Governments (EXC 294) and TRR130-P02 of the Deutsche Forschungsgemeinschaft (KK and MR).

Ethics

Animal experimentation: This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All procedures and experiments involving animals were approved by the Animal Use and Care Committee of University of California, Davis (protocol #20556).

Senior Editor

  1. Wendy S Garrett, Harvard T.H. Chan School of Public Health, United States

Reviewing Editor

  1. Andrew J MacPherson, University of Bern, Switzerland

Publication history

  1. Received: March 19, 2019
  2. Accepted: July 16, 2019
  3. Version of Record published: August 21, 2019 (version 1)

Copyright

© 2019, Savage 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.

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