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Peroxisome proliferator-activated receptors (PPARs) are members of the nuclear hormone receptor superfamily. PPARγ, a ligand activated transcription factor, has important anti-inflammatory and anti-proliferative functions and it has been associated with diseases including diabetes, scarring and atherosclerosis among others. PPARγ is expressed in most bone marrow derived cells and influences their function. PPARγ ligands can stimulate human B cell differentiation and promote antibody production. A knowledge gap is that the role of PPARγ in B cells under physiological conditions is not known. We developed a new B cell-specific PPARγ (B-PPARγ) knockout mouse and explored the role of PPARγ during both the primary and secondary immune response. Here, we show that PPARγ deficiency in B cells decreases germinal center B cells and plasma cell development as well as the levels of circulating antigen-specific antibodies during a primary challenge. Inability to generate germinal center B cells and plasma cells is correlated to decreased MHC class II expression and decreased Bcl-6 and Blimp-1 levels. Furthermore, B-PPARγ-deficient mice have an impaired memory response, characterized by low titers of antigen-specific antibodies and low numbers of antigen-experienced antibody-secreting cells. However, B-PPARγ-deficient mice have no differences in B cell population distribution within neither primary nor secondary lymphoid organs during development. This is the first report to show under physiological conditions that PPARγ expression in B cells is required for an efficient B cell-mediated immune response as it regulates B cell differentiation and antibody production.
Peroxisome proliferator-activated receptors (PPARs) are members of the nuclear hormone receptor superfamily (1). These ligand activated transcription factors are divided into three subtypes, PPARα, PPARβ/δ and PPARγ (2). PPAR signaling is activated by natural and synthetic ligands. Of interest, PPARγ can be activated by the endogenous prostaglandin 15-deoxy-Δ12,14 prostaglandin J2, or by the synthetic anti-diabetic thiazolidinediones (3–5). PPARγ is generally considered an anti-inflammatory and anti-proliferative transcription factor (2). PPARγ and its ligands have been implicated in diseases such as diabetes, scarring and atherosclerosis, among others (6–8).
PPARγ plays an important role in driving adipogenesis and lipid metabolism, and in dampening inflammation (2, 9, 10). PPARγ is also involved in regulating aspects of the immune system. T cells, B cells, macrophages and dendritic cells express PPARγ (11–14). PPARγ is involved in monocyte and dendritic cell differentiation (12, 13, 15). Furthermore, in dendritic cells PPARγ signaling downregulates IL-12 production (12, 13). Similarly in T cells, PPARγ ligands decrease IL-12 and IFNγ production, as well as cell survival (16–18).
B cells play an important role during both the innate and adaptive immune response. After initial antigen encounter, experienced B cells differentiate into antibody-secreting cells or to memory cells, ensuring an efficient response upon antigen re-encounter (19). Thus, mounting a strong but controlled humoral response is crucial for establishing long-term immune protection (20). We previously demonstrated that normal and malignant B cells express PPARγ (14). Using human B cells, we determined that PPARγ plays a role in regulating B cell function, particularly in promoting antibody production and B cell differentiation towards a plasma cell in vitro (21). In addition, malignant B-lineage cancer cells lines, with PPARγ deliberately over-expressed, have decreased proliferation and enhanced apoptosis (22–24).
Despite its role in B cell function in vitro, the role of PPARγ in B cell biology under physiological conditions is not known. Much of the animal work has been limited by technical challenges including the inability to generate complete PPARγ-deficient animals, as they are embryonically lethal (9). An alternative approach, which utilized global PPARγ haploinsufficient mice, has shown that a systemic reduction in PPARγ levels increases B cell proliferation and the antigen-specific immune response (25). However, even though that study showed the involvement of PPARγ in the humoral response, the experimental model used suffers from the fact that all cells were PPARγ haploinsufficient, not just B cells. Therefore, to further explore the role of PPARγ in B cell function under physiological conditions, we generated a new B cell-specific PPARγ knockout mouse and analyzed the role of PPARγ during both the primary and secondary immune response.
Mice carrying a recombinant PPARγ gene with loxP sites flanking exon 2 were a gift from Dr. Frank Gonzalez (26). Strain C.129P2-Cd19tm1(cre)Cgn/J, which contains the Cre recombinase under control of the Cd19 promoter on the Balb/C genetic background, was purchased from The Jackson Laboratory (Bar Harbor, Maine). These strains were crossed to generate Cd19-Cre+/− PPARγfl/- heterozygous mice, which were then backcrossed to C57BL/6J for 5 generations. Sibling crosses were then used to generate Cd19-Cre+/− PPARγfl/fl mice, and this genotype was maintained by sibling and cousin mating using male Cd19-Cre+/− PPARγfl/fl and female Cd19-Cre−/− PPARγfl/fl mice. The progeny are either Cd19-Cre+/− PPARγfl/fl (B cell PPARγ knockout) or Cd19-Cre−/− PPARγfl/fl (normal B cell litter mate controls). Progeny were genotyped by a commercial service (Transnetyx, Cordova, TN) using PCR oligos that span the junction of the Cd19 promoter and the Cre coding sequence and the junction of PPARγ intron 1 and the loxP site.
Because Cre is knocked into the CD19 locus, Cd19-Cre+/− mice have only one functional copy of CD19. To control for Cd19 copy number effects, we bred Cd19-Cre+/− males to C57BL/6J females, and the resulting Cd19-Cre+/− PPARγwt/wt offspring were used as controls for some experiments. B cells and antibody titers in naive mice were analyzed as in Figures 1, ,33 and Supplemental figure 2, and the OVA immune response was analyzed as in Figures 4, ,55 and Supplemental figure 3. In both cases, the results were similar to experiments performed using Cd19-Cre−/− PPARγfl/fl controls (data not shown).
To confirm B cell deletion of PPARγ, DNA was extracted from highly purified B cells obtained by fluorescence activated cell sorting (FACS) and subjected to quantitative PCR analysis. Excision of exon 2 in Cre-positive mice was confirmed using the sense 5’-GTAGAACCTGCATCTCCACC-3’ and antisense 5’-CTTGCATCCTTCACAAGCATG-3’ primers, exon 1 was used as a control and amplified with sense 5’-CATGGTTGACACAGAGATGC-3’ and antisense 5’-GTGTGGAGCAGAAATGCTGG -3’ primers. Lack of PPARγ protein expression was further confirmed by western blot analysis. Purified B cells were lysed with Nuclear Extract Kit (Active Motif, Carlsbad, CA). Ten µg of nuclear extract were loaded onto gradient SDS-PAGE gels (Pierce/Thermo Fisher Scientific, Rockford, IL). Western blots were probed using anti-PPARγ antibody (D69) (Cell Signaling Technology, Beverly, MA), and anti-actin antibody (Calbiochem/EMD Chemicals, Gibbstown, NJ). HRP-conjugated goat anti-rabbit or goat anti-mouse antibodies (Jackson ImmunoResearch) were used. Western blots were visualized with ECL (PerkinElmer Life Sciences).
To assess the efficiency of Cre recombination, we used the reporter strain B6.129X1-Gt(ROSA)26Sortm1(EYFP)Cos/J (R26-Stop-EYFP, The Jackson Laboratory). This strain contains a gene expressing enhanced yellow fluorescent protein (EYFP), preceded by a floxed STOP sequence, inserted in the Gt(ROSA)26Sor locus (27). When crossed with a strain expressing Cre recombinase, the STOP sequence is deleted, allowing expression of EYFP in the Cre-expressing cells or tissues. Cd19-Cre+ males were bred to R26-Stop-EYFP+/+ females, and Cre+/EYFP+/− offspring were identified by tail snip DNA PCR as above. Peripheral blood mononucleated cells were harvested and analyzed by flow cytometry.
Spleens were harvested, processed into a single cell suspension and B cells were isolated using CD19 magnetic beads (Miltenyi; Auburn, CA). The purity was >98% as determined by CD19 surface staining. B cells (1×106 cells/ml, unless otherwise specified) were cultured in RPMI1640 media (Invitrogen Life Technologies) supplemented with 5% FBS, 50 µM β-mercaptoethanol (Eastman Kodak, Rochester, NY), 10 mM HEPES (U.S. Biochemical, Cleveland, OH), 2 mM L-glutamine (Invitrogen Life Technologies, Carlsbad, CA), 50 µg/ml gentamicin (Invitrogen Life Technologies, Carlsbad, CA). B cells were stimulated with LPS (E. coli 055:B5, Sigma) (1 µg/ml), or anti-mouse CD40 (HM40-3, BD Pharmingen) (5 µg/ml) plus IL-4 (eBioscience) (25 ng/ml).
Activated B cells (105 cells/ ml) were cultured in triplicate using 96-well round bottom plates. Cell cultures were pulsed with [3H]-Thymidine (1 µCi/well) for 24 hours prior to harvest. [3H]-Thymidine incorporation was measured by scintillation spectroscopy using a Topcount Luminometer (PerkinElmer, Boston, MA)
Sera from mice or supernatants from activated B cells were collected for total IgM, IgG or IgA quantification by ELISA kits as specified by the manufacturer (Bethyl Laboratories, Montgomery, TX). Ovalbumin (OVA)-specific antibodies were measured using pre-coated OVA (10 µg/ml) plates and mouse-specific antibody ELISA kit (Bethyl Laboratories, Montgomery, TX).
Cells were incubated in OVA-coated ELISpot plates (Millipore, Billerica, MA) for 5 hours at 37°C. Alkaline phosphatase-conjugated goat anti-mouse IgM, IgG or IgA antibodies (Southern Biotech, Birmingham, AL) were used as recommended by the manufacturer. ELISpot plates were developed with Vector AP substrate kit III (Vector Laboratories, Burlingame, CA) and quantified using the CTL plate reader and ImmunoSpot software (Cellular Technologies, Shaker Heights, OH).
Primary mouse immunization was done using 10 µg OVA absorbed on complete Freund’s adjuvant (CFA) (1:1 ratio) by intraperitoneal (i.p.) injection and samples collected 2 weeks after injection. Secondary stimulation was done i.p. with 10 µg OVA (in PBS) 10 weeks later and samples collected 2 weeks after immunization. Cd19-Cre−/− PPARγfl/fl or Cd19-Cre+/− PPARγwt/wt animals were used as controls. Primary footpad immunizations were done using 25 µg OVA adsorbed on CFA (1:1 ratio), 20 µl final volume. Two weeks after immunization, popliteal lymph nodes were collected for analysis.
Single cell suspensions were prepared from mouse bone marrow, blood, spleen and lymph nodes and stained with a mixture of fluorochrome-conjugated anti-mouse monoclonal antibodies: CD93 (clone AA4.1, eBioscience), CD1d (clone 1B1, BD Pharmingen), CD24 (clone M1/69, eBioscience), CD5 (clone 53–7.3, BioLegend), IgM (cloneII/41, eBioscience), CD23 (clone B3B4, BioLegend), CD21(clone 7E9, BioLegend), IgD (clone 11.26c.2a, BioLegend), CD19 (clone 6D5, BioLegend), B220 (clone RA3-6B2, eBioscience), CD43 (clone 1B11, BioLegend), CD25 (clone PC61.5, eBioscience), CD3 (clone 17A2, BioLegend), FasL 9 (clone MFL3, BioLegend), CD138 (clone 281-2, BD Pharmingen), CD62L (clone MEL-14, eBioscience), CD95 (clone Jo2, BD Pharmingen), CD86 (clone GL-1, BioLegend), GL7 (clone GL7, eBioscience), CD44 (clone IM7, eBioscience), I-A/I-E (clone M5/114.15.2, BD Pharmingen), CD80 (clone 16-10A1, eBioscience), CXCR5 (clone 2G8, BD Pharmingen), IgG (eBioscience), IgM (clone II/41, eBioscience), CD3 (clone 145-2C11, BioLegend), CD4 (clone GK1.5, BioLegend), CD69 (clone H1.2F3, BioLegend), CD138 (clone 281-2, BD Pharmingen) followed by staining with fluorochrome-conjugated streptavidin (Invitrogen) if biotin-conjugated antibody was included in the staining panel. All the samples were stained for dead cell exclusion using Live/Dead fixable violet dead cell staining kit (Invitrogen). Samples were run on a 12-color LSRII cytometer (BD Pharmingen) and analyzed by FlowJo software (Tree Star Inc., Ashland, OR).
Total RNA was isolated using miRNAeasy Mini Kit (Qiagen, Valencia, CA). iScript cDNA Synthesis Kit (Bio-Rad, Hercules, CA) was used to reverse transcribe RNA to cDNA. Steady-state levels of Bcl-6, Blimp-1 and GAPDH RNA were assessed by real-time PCR using iQ SYBR Green Supermix (Bio-Rad). Primers used for Bcl-6 sense, 5’-AGACGCACAGTGACAAACCATACAA-3’, and antisense 5’-GCTCCACAAATGTTACAGCGATAGG-3’; for Blimp-1 were sense, 5’- TTCTTGTGTGGTATTGTCGGGACTT-3’ and antisense, 5’- TTGGGGACACTCTTTGGGTAGAGTT-3’; and for GAPDH sense 5’-AGCCTCGTCCCGTAGACAAA-3’ and antisense, 5’-CCTTGACTGTGCCGTTGAAT-3’ (28). Results were analyzed using Bio-Rad iCycler software. Bcl-6 and Blimp-1 mRNA steady-state levels were normalized to GAPDH.
Freshly dissected spleen were embedded in optimum cutting temperature compound (Sakura) and stored at −80°C. Cryostat sections (10 µm) were prepared from frozen tissues. Sections were fixed in cold acetone and re-hydrated in PBS, blocked with 5% whole rat serum and stained with APC-antiCD3ε (clone 145-2C11, BioLegend) and PE-anti B220 (clone RA3-6B2, eBioscience). Images were acquired using a fluorescence microscope connected to a monochrome CCD digital camera and were analyzed using ImagePro Plus software (MediaCybernetics, Bethesda,MD)
Data are expressed as mean ± SEM. Significance was determined by one-way ANOVA with a Tukey’s post test, or a two-tailed unpaired Student t-test where applicable. Statistical analyses were done using GraphPad Prism 5.0 (GraphPad Software, La Jolla, CA.). Probability values of p ≤ 0.05 were considered statistically significant.
Global PPARγ knockout animals are embryonically lethal, limiting in vivo studies on PPARγ function. Therefore, we developed a new B cell-specific PPARγ knockout mouse utilizing transgenic mice in which Cre recombinase expression is under control of the CD19 promoter, which is only expressed in the B cell lineage (29, 30). The efficiency of Cre-mediated recombination in B cells using an EYFP reporter was >90% (Supplemental Figure 1A). The CD19-Cre mice were bred with a targeted strain in which LoxP sites are positioned flanking exon 2 of the PPARγ gene (26) to generate conditional knock-outs in which only B cells harbor a Cre-dependent deletion of PPARγ (B-PPARγ-deficient mice). In these Cre-positive B cells, exon 2 is excised thus rendering the PPARγ receptor non-functional (Supplemental Figure 1B). Efficiency of exon 2 deletion in highly purified B cells was near 100%, determined by PCR analysis of genomic DNA (Supplemental Figure 1C). Knock-out of PPARγ expression in B cells from B-PPARγ-deficient mice was further confirmed by Western blot analysis using purified B cells (Supplemental Figure 1D).
Considering the potential importance of PPARγ in B cell function, we first evaluated any possible deficiencies in B cell subpopulations. The total number of splenocytes, as well as the distribution of splenic CD19+ B cells in the B-PPARγ-deficient mice were measured and compared to control mice (Figure 1A–B). Naïve B-PPARγ-deficient mice have a normal count of total splenocytes and CD19+ cells.
Early B cell development takes place in the bone marrow while late stages of differentiation can take place in secondary lymphoid organs. We performed a comprehensive evaluation of B cell subpopulations (as defined in Table 1 and Supplemental Figure 2A–B) in B-PPARγ-deficient mice (29, 31, 32). Pro-B cell, pre-B cell, immature B cell and transitional B cell population frequencies do not change in the bone marrow of B-PPARγ-deficient mice compared to control animals (Figure 1C–E). In addition, follicular, marginal zone B and B1 cell subpopulations in spleen and lymph node were also found unchanged (Supplemental Figure 2C–H).
We have previously shown that PPARγ signaling is important for human B cell antibody production in vitro (21). We therefore evaluated the primary B cell response under in vitro conditions. Antibody production was measured in highly-purified B cells isolated from B-PPARγ-deficient and control mouse spleens and treated with LPS (Figure 2). Interestingly, LPS-treated B cells from B-PPARγ-deficient mice produced similar levels of IgM, but significantly lower levels of IgG (Figure 2A–B).
PPARγ affects normal and malignant B cell differentiation and proliferation (21, 33). Considering that B cells from PPARγ-deficient mice produced less IgG upon activation, we investigated if there were any differences in cell proliferation. Accordingly, LPS-activated purified PPARγ-deficient B cells had decreased proliferation compared to control cells (Figure 2C).
Next, we measured antibody concentration in sera of naïve B-PPARγ-deficient animals. Both B-PPARγ-deficient and control groups, showed similar total IgM, IgG and IgA serum antibody titers (Figure 3A–C).
Because the observed decrease in antibody levels and proliferation were seen under stimulating culture conditions (Figure 2), we next asked whether B-PPARγ-deficient animals would also have an impaired antibody-mediated immune response upon antigen exposure. To test our hypothesis, B-PPARγ-deficient and control mice were challenged using the OVA antigen model. The primary immune response was measured 2 weeks after immunization. Interestingly, B-PPARγ-deficient mice had an impaired antibody-mediated immune response, seen by a significantly lower amount of both antigen-specific IgM and IgG (Figure 4A–B). This defect cannot be ascribed to either structural disruption in lymphoid organ architecture in B-PPARγ-deficient mice (Figure 4C–H) or deficiencies in development of peripheral B cell populations (Figure 1 and supplemental Figure 3).
Adaptive immunity is mediated in part by plasma and memory cells. These fully differentiated cells are responsible for the quick antibody-mediated immune response upon antigen re-encounter. To further study the impact of B-PPARγ-deficiency on the adaptive immune response, mice were rechallenged with OVA 10 weeks after initial immunization. Two weeks after antigen re-exposure, the antibody memory response was analyzed and measured (Figure 5A–B). Control mice had an enhanced memory response and higher antigen-specific antibody titers. Again, B-PPARγ-deficient mice had considerably lower antigen-specific serum IgM and IgG levels. IgA levels were also measured during the primary and secondary response however, IgA levels were below detection.
The B cell memory response was further analyzed by measuring the number of spleen antigen-specific antibody-producing cells present in re-immunized B-PPARγ-deficient mice (Figure 5C–F). Correlating with the decreased serum antibody titers, B cells from immunized B-PPARγ-deficient animals did not respond to antigen re-exposure. In fact, the number of antigen-specific antibody-secreting cells was comparable to that of non-immunized animals. This significant defect was true for IgM, IgG and IgA isotypes (Figure 5D–F).
It is possible that the decreased antibody response observed is a consequence of impaired B cell differentiation following antigen presentation. To test this hypothesis, B-PPARγ-deficient mice were challenged by delivery of OVA antigen into the footpad. B and T cells were analyzed in the immunized and non-immunized contralateral popliteal lymph nodes (PLNs) 2 weeks after the OVA challenge. Distribution of germinal center (GC) B cell, plasma cell and follicular helper T cell populations were analyzed (Figure 6). Immunized B-PPARγ-deficient mice had a significant decrease in the percentage of GC B cells, similar to those of the non-immunized control group (Figure 6A–B). Furthermore, the plasma cell population was also absent in the PLNs of immunized B-PPARγ-deficient mice (Figure 6C–D). Follicular helper T cells (Tfh), which are crucial in the germinal center reaction (34), had a normal distribution in B-PPARγ-deficient mice compared to control (Figure 6E–F).
Given the differences in GC B cell and plasma cell differentiation, we further analyzed B cell activation in vitro. Purified splenic B cells from control and B-PPARγ-deficient mice were stimulated with anti-CD40 and IL-4, a stimulation resembling T cell-dependent activation in vivo. Similar to LPS treatment (Figure 2), PPARγ-deficient B cells stimulated with anti-CD40 plus IL-4 produced significantly lower levels of IgM and IgG compared to B cells from control mice (Figure 7A–B).
B cell-T cell interactions within secondary lymphoid organs are necessary for B cell activation and initiation of the antigen-specific immune response (35). Upon stimulation, B cells increase expression of activation markers including MHC class II, CD80, CD86 and CD69. However, stimulated PPARγ-deficient B cells did not upregulate MHC class II (Figure 7C–D). CD80, CD86 and CD69 expression levels in PPARγ-deficient B cells were similar to those of the control group (Supplemental Figure 4).
To further confirm the mechanisms responsible for the decreased B cell activation and differentiation seen in vivo and in vitro, we measured Bcl-6 and Blimp-1 expression. Bcl-6 and Blimp-1 are key transcription factors involved in B cell differentiation. Bcl-6 is necessary for GC B cell development (36, 37), while Blimp-1 regulates plasma cell differentiation (38, 39). Bcl-6 and Blimp-1 steady state mRNA levels were significantly lower in PPARγ-deficient B cells compared to control (Figure 7E–F).
PPARγ is a widely expressed transcription factor that influences many areas of biology. Here, we show evidence demonstrating a physiological role of PPARγ in B cells. Using a newly developed B cell-specific PPARγ knockout mouse model, we have shown that PPARγ expression in B cells is necessary for optimal humoral immune responses. A B cell-specific PPARγ-deficient mouse model provides a novel and direct system to study the role of PPARγ as a regulator of antibody production. Here, mice lacking PPARγ expression in B cells are shown to have decreased proliferation and IgG production in vitro, as well as low levels of circulating antigen-specific antibodies during a primary response. Furthermore, PPARγ-deficient mice have an impaired immune memory response, characterized by low titers of antigen-specific antibodies and low numbers of antigen-experienced antibody-secreting cells. Our results denote the importance of PPARγ expression in B cells during both the primary and secondary immune response.
PPARγ is involved in key B cell processes, including differentiation and antibody production (21). Interestingly, our results show that B cell-specific PPARγ-deficient mice have normal basal antibody levels as well as a normal percentage of B cells present in the spleen. Detailed analysis of B cell development in primary and secondary lymphoid organs showed neither differences in the population distribution of the PPARγ knockout animals, nor changes in their spleen follicle architecture. We conclude that the decreased primary humoral response is not due to changes in B cell development.
Upon antigen encounter, B cells activate, proliferate and differentiate. Antigen experienced B cells will differentiate into memory or plasma cells, which produce antigen-specific antibodies during the adaptive immune response (19). Our results show that the number of antigen-specific antibody-secreting cells in immunized B-PPARγ-deficient mice is equivalent to background levels seen in the non-immunized group. The lack of antigen-specific antibody-secreting cells in immunized B-PPARγ-deficient mice is true for all antibody isotypes measured. These findings correlate to the decreased serum antibody titers during, the primary and secondary immune response. Interestingly, immunized animals have a normal distribution of follicular, transitional, marginal zone and B1 cell populations, confirming that PPARγ is not required for early B cell development.
It should be noted that the Cre gene was introduced into the CD19 locus by homologous recombination, so that Cd19-Cre−/− PPARγfl/fl mice, used in this study as normal B cell litter mate controls, have two functional copies of the CD19 gene, in contrast to the Cd19-Cre+/− PPARγfl/fl mice (B-PPARγ-deficient) which have one functional CD19 locus. To control for CD19 copy number effects, some experiments were repeated using Cd19-Cre+/− PPARγwt/wt mice as controls. B cell number and antibody titer in naive mice were analyzed as in Figures 1, ,33 and Supplemental figure 2, and the OVA immune response was analyzed as in Figures 4, ,55 and Supplemental figure 3. In both cases, the results were similar to experiments performed using Cd19-Cre−/− PPARγfl/fl controls (data not shown). We conclude that deletion of one copy of the CD19 gene has no effect on B cell development or function in these mice.
Initial B cell priming takes place within lymphoid organs, leading to the generation of germinal centers (35). The germinal center reaction is necessary for B cell somatic hypermutation, affinity maturation and class switch recombination (35). Even though early B cell development in B-PPARγ-deficient mice is normal, our results show that PPARγ is required for B cell activation during antigen presentation. Our analysis has revealed that B-PPARγ-deficient mice generate Tfh cells upon antigen stimulation. However, B-PPARγ-deficient mice are unable to develop GC B cells, as well as differentiated plasma cells. We have not directly surveyed the memory cell populations. However, absence of GC B cell and plasma cell populations during the primary immune response and lack of antigen-specific antibody producing cells during both primary and secondary immunizations makes it very likely that B-PPARγ-deficient mice also have decreased number of memory B cells.
Furthermore, PPARγ-deficient B cells don’t upregulate MHC class II upon activation. Decreased MHC class II expression could 1) disrupt the immunological synapse between antigen-primed T cells and naïve B cells, thus preventing GC B cell development, 2) decrease B cell antigen presentation, consequently decreasing the antigen-specific immune response. Further in vivo investigation of B cell activation is required.
At a molecular level, we demonstrate that PPARγ deficiency in B cells decreases Bcl-6 and Blimp-1 expression, which are crucial during GC B cell and plasma cell development (36–39). There are both direct and indirect mechanisms which can explain the effects of PPARγ on Bcl-6 and Blimp-1. Signaling through CD40 and IL-4 receptor activates NF-κB signaling. Bcl-6 expression can be downregulated via NF-κB (40, 41). Furthermore, PPARγ is known to inhibit NF-κB translocation to the nucleus (42). Thus, it is possible that PPARγ regulates Bcl-6 expression through NF-κB signaling, which initiates B cell activation and Blimp-1-mediated plasma cell differentiation. Alternatively, Bcl-6 and Blimp-1 contain multiple PPAR response elements (PPRE) within their 3’-untranslated region with high PPARγ binding efficiency. Therefore, it is also possible for PPARγ to directly regulate Bcl-6 and Blimp-1.
Here, we provide conclusive evidence that PPARγ has a physiological role during the primary and secondary immune response, using a novel B cell specific PPARγ-deficient mouse model. PPARγ has been associated with multiple diseases including autoimmune disorders, which involve B cell function (25, 43, 44). The in vivo study of PPARγ and PPARγ-ligands has been limited due to challenges in the development of animal models. Our novel B cell-specific PPARγ-deficient mouse model will serve as a powerful tool for future studies on the role of PPARγ as well as PPARγ-ligand-dependent and -independent signaling mechanisms in multiple disease models.
The authors have no additional acknowledgments.
1This work was supported in part by DE011390 (to R.P.P.), ES01247 (to R.P.P.), NIH HL75432 (to P.J.S.), NIH T32 DE007202 and T32 HL66988
The authors have no financial conflicts of interest