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Differential regulation of CD36 expression in antigen‐presenting cells: Oct‐2 dependence in B lymphocytes but not dendritic cells or macrophages

Differential regulation of CD36 expression in antigen‐presenting cells: Oct‐2 dependence in B... Abstract In mice, three antigen‐presenting cell types [B lymphocytes, macrophages and dendritic cells (DC)] express the scavenger receptor CD36. This molecule has been implicated in many important functions, including DC maturation and antigen presentation. In murine B cells, the CD36 gene requires the Oct‐2 transcription factor for its expression. We previously found that B cells from Oct‐2‐null mice display defects in maturation, survival and proliferation. Here we have looked for a possible role for CD36 in B cells, but found that CD36 is dispensable for all responses tested. Although loss of CD36 did not directly affect B cell function, it did modulate slightly the isotype and level of IgG produced in vivo in naive mice, and IgM in Leishmania‐infected mice. We also show that in DC and macrophages, CD36 expression is independent of Oct‐2. We conclude that CD36 does not play a major role in B cell function, but that CD36 may contribute indirectly to humoral immunity through cells of the innate immune system. B lymphocyte, dendritic cell, Leishmania major, monocyte/macrophage, transgenic/knockout Introduction The B cell‐restricted transcriptional activator, Oct‐2, binds to a conserved site found in most Ig gene promoters. However, through targeted mutation of the gene for Oct‐2, we have shown that Ig gene promoters do not require this factor for expression, at least in newly formed peripheral B cells. Nevertheless, Oct‐2‐null B cells display several abnormalities (1–3). Without Oct‐2, immature B cells emigrating from the bone marrow fail to differentiate efficiently into long‐lived cells, resulting in a deficit of mature peripheral B cells. Serum Ig levels are markedly reduced in naive and immunized oct‐2–/– mice and the peritoneal B‐1 lymphocyte compartment is absent. In vitro, oct‐2–/– B cells maintain properties of immature ‘transitional’ B cells (3–5) which do not proliferate, but undergo apoptosis in response to a BCR signal (6,7). Presumably, these effects are due to the dysregulation of Oct‐2‐dependent target genes in B cells. One of these, the first Oct‐2‐dependent gene identified, is that encoding the cell surface protein CD36 (8). CD36 is expressed on many cell types, including some hematopoietic cells (9). It acts as a multi‐ligand scavenger receptor that can mediate adhesion via thrombospondin and scavenging of apoptotic cells. A role for CD36 in dendritic cell (DC) maturation and antigen cross‐presentation has been proposed (10,11), and its critical role in fatty acid metabolism has recently been proven (12). However, no function has yet been ascribed to CD36 in B lymphocytes. Indeed, while mouse B cells express CD36, most normal human B cells do not. Instead, B cell leukemias are often CD36+, with expression correlating with more advanced disease (13). These observations prompted three questions. First, is the CD36 gene dependent upon Oct‐2 in B cells, macrophages and DC, the three antigen‐presenting cells where it is expressed? Second, does the inability to express CD36 contribute to the abnormal phenotype of Oct‐2‐null B cells? Third, does the loss of CD36 impact on the immune response to a pathogen in vivo? This examination of hematopoietic cells from Oct‐2‐null mice shows that CD36 relies on Oct‐2 for its expression only in B lymphocytes. To investigate the consequences of CD36 loss to the B cell system, we assessed the B cell compartment of CD36‐null mice, examining cell subpopulations and phenotypes, B cell responses to stimulation in vitro, and responses in a T cell‐dependent model of infection with the parasite Leishmania major. Murine cutaneous leishmaniasis is caused by L. major, an intracellular parasite of mononuclear phagocytes. Intradermal infection with L. major produces a local skin lesion and self‐limiting granuloma that heals in 10–12 weeks in genetically resistant mice (e.g. C57BL/6 mice) or it may disseminate systemically and kill the host (14,15). Recovery is dependent upon the induction of Th1‐type pro‐inflammatory cytokines such as IL‐12 and IFN‐γ, and the generation of CD4+ Th1‐type cells. Th1 cytokines induce macrophage activation and killing of the intracellular organisms by NO (16,17). B cells also play a role in T cell‐mediated healing (18), but depletion of CD5+ B cells has no effect on the course of infection (19). Given the central roles of the T cell‐activated macrophage and B cells in host resistance against leishmaniasis, we included this model infection in our characterization of CD36 activities in vivo. Methods Mice and tissues Oct‐2‐null mice die at birth, so adoptive transfer of embryonic day 13 (E13) fetal liver into RAG‐1‐deficient recipients was used to generate mature lymphoid cells, as described (2). To access oct‐2–/– macrophages and DC, oct‐2+/– mice were crossed, E18 fetal liver taken and genotyped. Cells were stained directly ex vivo for lineage markers and for CD36 expression as described below. DC and macrophages were isolated as previously described (20,21), with slight modifications to maximize yields, using collagenase digestion and immunomagnetic bead depletion of either non‐DC or non‐macrophages. For DC, cells were depleted with mAb anti‐CD3 (KT3‐1.1), anti‐Thy1 (T24/31.7), anti‐B220 (RA36B2), anti‐Gr1 (RB68C5) and anti‐erythrocyte (TER119). For macrophages, depletion used mAb anti‐CD3 (KT3‐1.1), anti‐CD4 (GK1.5), anti‐CD8α (53‐6.7), anti‐B220 (RA36B2) and anti‐erythroid (TER119). Macrophages were also expanded from unfractionated E13 fetal liver in the presence of macrophage colony stimulating factor (M‐CSF; 1000 U/ml in DME + 10% FBS). Duplicate cultures were maintained for 6 days and one was supplemented with lipopolysaccharide (LPS, 1 µg/ml; Difco, Detroit, MI) for the final 24 h. The adherent cells were dislodged by pipetting in EDTA/FBS. The vast majority of the cells in all cultures were macrophages (CD11b+; data not shown). The CD36 knockout mice were generated on the 129/Sv background (12). We had shown that this strain can display anomalous B cell behavior (22), so we backcrossed the mice for six generations to C57BL/6 mice before performing the analyses described here. ELISAs Serum was diluted through a mid‐log10 series from an original 1/100 dilution. ELISAs were performed as described (2,24). For the anti‐Leishmania antibody responses, the dilution was 1:10,000 and the Ig isotype determined using an isotyping kit according to the manufacturer’s recommendations (Bio‐Rad, Hercules, CA). For the Ig2c isotype, a specific antibody was used as in (24). Flow cytometry Peripheral blood lymphocytes from heparinized, red cell‐depleted blood were stained with anti‐B220 (RA36B2). Splenocytes and peritoneal cells were prepared and stained as described (3). Details of the mAb and labeling procedure for DC and macrophages have been described elsewhere (20,21). All samples were pre‐incubated with a mix of whole mouse Ig and anti‐FcRII/III (2.4G2) to minimize non‐specific staining. The mAb used to identify DC and macrophages were anti‐CD11c (N418) and anti‐CD11b (M1/70) respectively. Propidium iodide (PI) was included in the final wash at 1 µg/ml to label dead cells. Analyses were carried out on either a FACStar Plus or FACScan (Becton Dickinson, San Jose, CA). Cell surface CD36 expression was then determined by gating on live CD11c+ DC, CD11b+ macrophages and B220+ B cells. CD36 was detected by anti‐CD36 (clone63; Cascade BioScience, Winchester, MA) and an anti‐mouse IgA second stage (Caltag, Burlingame, CA) after blocking with whole rat Ig. Lymphocyte proliferation assays Spleen cell suspensions were cultured at 1 × 106 cells/ml for 72 h with or without B cell mitogens and pulsed with 1 µCi [3H]thymidine during the final 5 h of culture, as described (2). Parasites Parasites were of the virulent cloned line of L. major LRC‐L137 (MHOM/IL/67/JerichoII). Promastigotes were maintained in vitro at 26°C in Schneider’s Drosophila medium with 10% FBS and used in stationary phase. Cutaneous infection of wild‐type and CD36‐null mice Mice were injected intradermally with 105 promastigotes. Lesion development was assessed weekly for 10–12 weeks, according to the lesion‐scoring system described (23,25). Mice were bled periodically, and their Leishmania‐specific antibody titers and isotypes measured as described above. A soluble parasite lysate (SLA) obtained by freezing and thawing promastigotes was used as antigen (26,27). In vitro infection of macrophages with L. major promastigotes Bone marrow‐derived macrophages from wild‐type and CD36‐null mice were cultured for 5–7 days in RPMI medium with 10% FBS and 10% supernatant from cultures of the 929 cell line as a source of M‐CSF (28). Macrophages were transferred onto coverslips (2 × 105 cells/well in 0.5 ml of medium) and allowed to adhere for 18 h at 37°C. Non‐adherent cells were washed away and monolayers were infected with L. major promastigotes at a ratio of 5:1. After 30 min, free parasites were removed and the cells were re‐cultured for 2, 24, 48 or 72 h. The cells were then fixed in methanol and stained with Giemsa. For each time point 500 cells were counted, and the percent infected cells and number of parasites present were calculated. Experiments were performed at least 3 times with duplicate samples. Results and discussion Expression of CD36 in myeloid cells is Oct‐2 independent Earlier experiments (8,29) proved that transcription of the CD36 gene was directly dependent upon Oct‐2 in a pre‐B lymphoma cell line. Konig et al. (8) also showed a strong correlation between CD36 and Oct‐2 expression in a number of B and monocyte/macrophage cell lines and in primary hematopoietic tissues. However, a direct effect of Oct‐2 on CD36 expression was not proven for the myeloid cells. Here we have examined the surface expression of CD36 in three primary hematopoietic tissues from oct‐2–/– mice to determine the contribution of Oct‐2 to CD36 expression in each cell type. As expected, B cells require Oct‐2 for CD36 expression (Fig. 1A). However, when DC (CD11c+) and macrophages (CD11b+) from fetal liver were examined, there was no correlation between CD36 expression and oct‐2 genotype. All DC were CD36+ at the time of isolation and expressed equivalent levels of the protein (Fig. 1B, top). Surprisingly, none of the CD11b+ macrophages directly ex vivo expressed CD36 (Fig. 1B, middle). We therefore expanded macrophages from E13 fetal liver in the presence of M‐CSF and added LPS during the final 24 h to half the cultures, to activate the cells. We found that expanding the cells in M‐CSF was sufficient to induce a high level of CD36 expression in the cultured macrophages, which was not altered by LPS addition (Fig. 1B, bottom). We also learned that Oct‐2 is not required for CD36 expression in macrophages, as the oct‐2‐null cells expressed normal CD36 levels. These data indicate that the CD36 gene is regulated differentially in antigen‐presenting cells, with B cells expressing intermediate levels, and DC and macrophages expressing at least 10‐fold higher levels of surface CD36. Despite the presence of Oct‐2 in cells of both the myeloid and B lymphoid lineages (8,30), only B cells require Oct‐2 as a critical CD36 regulator, with the Oct‐2 binding site in the promoter acting as the central regulatory element (29). These studies do not exclude the possible involvement of other octamer‐binding factors in the regulations of the CD36 gene in myeloid cells, in particular the ubiquitously expressed Oct‐1. Studies on CD36 expression in Oct‐1‐deficient animals will clarify this issue. Interestingly, however, immediately adjacent to the Oct‐2 site in the CD36 promoter is a binding site for the transcriptional regulator AML‐1. AML‐1 activates the transcription of several tissue‐specific genes in myeloid cells (31) and may contribute to the high level of CD36 gene expression in macrophages and DC. Features of the B cell compartment in CD36‐null mice To explore whether Oct‐2‐mediated CD36 expression in B cells had evolved to enable some humoral immune function, we examined B cells and their products in CD36‐null mice. We compared serum Ig levels in naive C57BL/6 and CD36–/– mice, and found that CD36‐deficient mice had normal or even slightly higher than normal titers of all Ig isotypes tested (Fig. 2A). In C57BL/6 mice, the IgG2a gene has been deleted and the alternative IgG2c gene is expressed (24). 129/Sv mice have the opposite profile. The CD36–/– mice, generated in the 129/Sv strain and backcrossed onto the C57BL/6 background, express IgG2c, but not IgG2a, and express the same level of IgG2c as the control C57BL/6 mice (Fig. 2A). This overall picture contrasts with Oct‐2‐deficient mice, which have ∼10% of normal levels of most Ig isotypes (2). These observations suggest that loss of CD36 expression in B cells does not compromise Ig gene expression or secretion and so is not a primary limiting factor in Oct‐2‐deficient B cells. However, loss of CD36 does influence the profile of Ig isotypes expressed in vivo, with IgG1 and IgG2b being somewhat favored in this set of mice. In other experiments, the absolute differences between IgG1 and IgG2b levels for C56BL/6 control and CD36–/– mice varied, but levels in CD36‐null mice were always higher than controls (data not shown). A characteristic feature of peripheral oct‐2–/– B cells is their immaturity. Mature cells down‐regulate expression of the heat‐stable antigen (HSA; CD24) from the high level expressed on recent bone marrow emigrants (4,5). In Oct‐2‐null mice, mature (HSAlo) cells are virtually absent (3). In contrast, mature and immature B cells in the spleen of CD36‐null mice were present in normal ratios (Fig. 2B). Clearly, CD36 is not essential for the signal that mediates this maturation step. Peritoneal B‐1 cells were also present in normal numbers in CD36‐null mice (Fig. 2C), while they are absent in Oct‐2‐null mice (3). Like the peripheral maturation of immature B cells, generation and maintenance of B‐1 lymphocytes require signals through the BCR (32,33). Oct‐2 is required for the successful receipt of such signals in both the B‐1 and the B‐2 compartments, perhaps by regulating expression of a component(s) of the BCR signaling pathway. Our conclusion is that CD36 is not a critical Oct‐2 target gene in the context of these important cell survival and maturation signals. Responses of CD36‐null B cells to mitogenic stimulation are normal Proliferative responses of CD36‐deficient splenocytes to a variety of B cell mitogens were examined and found to be normal (Fig. 2D). This is in stark contrast to Oct‐2‐deficient B cells, which are severely hyporesponsive to LPS, to anti‐Rp105 signaling and to IL‐5 stimulation, and significantly hyporesponsive to anti‐µ stimulation [(2) and our unpublished observations]. The data in Figs 1 and 2 strongly suggest that, despite the clear dependence of the CD36 gene on Oct‐2 for its expression in B lymphocytes (29), CD36 insufficiency is not a major factor in the defective phenotype of Oct‐2‐deficient B cells. Indeed, we found no evidence that CD36 plays a significant role in any of the B cell‐autonomous attributes we examined here. However, the combined loss of CD36 and other Oct‐2 target genes may be responsible for some of the deficiencies exhibited by oct‐2‐null B cells. Humoral immune response in L. major‐infected CD36‐null mice L. major‐infected CD36‐null mice had elevated Leishmania‐specific total IgG compared to controls (Fig. 3A). However, there was no significant difference in the levels of specific IgG isotypes examined (Fig. 3B). This mirrored the pattern seen in naive CD36–/– mice (Fig. 2A). In contrast, the level of Leishmania‐specific IgM detected in the wild‐type mice was significantly higher than in the CD36‐null mice. L. major‐infected CD36‐null mice developed smaller lesions and appeared to cure somewhat faster than control mice, with all mice cured by week 7. The smaller lesion size was significant in the early weeks post‐infection as determined by the Mann–Whitney statistical test (P < 0.05 at weeks 3, 5 and 6; Fig. 3C). The disease pattern suggests that loss of CD36 expression engenders an even more pronounced Th1 immune response to the parasite than is exhibited by the already genetically resistant C57BL/6 mouse. Indeed, Urban et al. (10) have shown that CD36 ligation can strongly influence cytokine production by T cells. The hypothesis that CD36 promotes resistance to infection by biasing the Th cell phenotype can be tested in genetically susceptible BALB/c mice lacking the CD36 gene. To what extent these responses are affected by the antigen‐processing and ‐presentation ability of the CD36‐null macrophages remains to be established. However, we could find no significant difference in the ability of cells from the CD36‐null mice to support parasite growth compared to wild‐type mice (Fig. 3D). L. major promastigotes were able to establish infection in CD36‐null bone marrow‐derived macrophages as efficiently as in wild‐type cells (Fig. 3D) and the number of parasites per infected macrophage was similar during a 72 h period of infection (data not shown). Therefore, CD36 expression is not a limiting factor for parasite invasion and survival in macrophages. In conclusion, we have shown that the Oct‐2 transcription factor is necessary for the expression of the CD36 protein on murine B cells, but that specific loss of CD36 from these cells is relatively inconsequential. We found no evidence of B cell dysfunction in CD36–/– mice in the assays performed, which focused on those capacities that are compromised in Oct‐2‐deficient B cells. In contrast, CD36 expression is Oct‐2‐independent in myeloid cells (DC and macrophages, in particular), arguing that the gene is differentially regulated in these antigen‐presenting cells. We observed a potential influence of CD36 on the humoral immune response, in the altered spectrum of Ig isotypes and the response to L. major infection. CD36 may therefore play a wider role in immune regulation, indirectly influencing the quality of a humoral immune response via influences on the balance of Th1/Th2 cells, as well as through cells of the innate immune system. These observations are consistent with the proposal that CD36 is an ancient functional component of innate immunity (9). Acknowledgements We thank Wendy Deitrich and Eren Loza for technical assistance and animal care respectively, and Dr D. Tarlinton for critical reading of the manuscript. Professor D. Metcalf generously provided murine M‐CSF, Dr A. Lew provided anti‐IgG2 and Dr A. Strasser provided a number of mAb for cell surface staining. This work was supported by the Australian National Health and Medical Research Council, and the UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases (TDR). Abbreviations DC—dendritic cell E—embryonic day HSA—heat‐stable antigen LPS—lipopolysaccharide M‐CSF—macrophage stimulating factor PI—propidium iodide View largeDownload slide Fig. 1. CD36 expression on B lymphoid and myeloid cells. (A) Surface CD36 expression on B220+ peripheral blood cells. (B) CD36 staining on dendritic cells (CD11c+) and macrophages (CD11b+) from E18 fetal liver or on macrophages expanded from E13 fetal liver in the presence of M‐CSF. LPS was added for the final 24 h of some cultures (+LPS). In all panels, the dotted line represents a background control (no primary antibody) and the solid line corresponds to staining with a CD36‐specific antibody. View largeDownload slide Fig. 1. CD36 expression on B lymphoid and myeloid cells. (A) Surface CD36 expression on B220+ peripheral blood cells. (B) CD36 staining on dendritic cells (CD11c+) and macrophages (CD11b+) from E18 fetal liver or on macrophages expanded from E13 fetal liver in the presence of M‐CSF. LPS was added for the final 24 h of some cultures (+LPS). In all panels, the dotted line represents a background control (no primary antibody) and the solid line corresponds to staining with a CD36‐specific antibody. View largeDownload slide Fig. 2. Assessment of CD36‐null B cells, their products and behavior. (A) Serum Ig titers in naive mice. Filled circles, C57BL/6 mice; open circles, CD36‐null mice; open squares with dotted lines, 129/Sv mice. Each line corresponds to the mean (±SD) of three or more animals. (B) HSA levels on splenic B220+ cells. The dotted line delineates immature (HSAhi) from mature (HSAlo) cells, as determined by Allman et al. (4,5). The data are representative of an analysis of four mice of each genotype, with the number indicating the mean percentage of cells in the indicated region for all mice. (C) Lymphocyte populations in peritoneal cells harvested from C57BL/6 and CD36‐null mice, stained to highlight B‐1 lymphocytes (boxed). This panel shows representative data from four mice of each type (the number indicates the mean of cells in the B‐1 region for all four mice). Staining with B220 and Mac‐1 or with B220 and CD5 gave the same result, i.e. no difference between the control and mutants. (D) Proliferative responses of splenocytes to a number of B cell mitogens. Values are the mean (±SD) for triplicate wells and each bar represents a single mouse. Filled bars, C57BL/6 mice; hatched bars, CD36‐null mice. Stimulation index is the ratio of c.p.m in stimulated cultures divided by the c.p.m. in matched, unstimulated cultures. View largeDownload slide Fig. 2. Assessment of CD36‐null B cells, their products and behavior. (A) Serum Ig titers in naive mice. Filled circles, C57BL/6 mice; open circles, CD36‐null mice; open squares with dotted lines, 129/Sv mice. Each line corresponds to the mean (±SD) of three or more animals. (B) HSA levels on splenic B220+ cells. The dotted line delineates immature (HSAhi) from mature (HSAlo) cells, as determined by Allman et al. (4,5). The data are representative of an analysis of four mice of each genotype, with the number indicating the mean percentage of cells in the indicated region for all mice. (C) Lymphocyte populations in peritoneal cells harvested from C57BL/6 and CD36‐null mice, stained to highlight B‐1 lymphocytes (boxed). This panel shows representative data from four mice of each type (the number indicates the mean of cells in the B‐1 region for all four mice). Staining with B220 and Mac‐1 or with B220 and CD5 gave the same result, i.e. no difference between the control and mutants. (D) Proliferative responses of splenocytes to a number of B cell mitogens. Values are the mean (±SD) for triplicate wells and each bar represents a single mouse. Filled bars, C57BL/6 mice; hatched bars, CD36‐null mice. Stimulation index is the ratio of c.p.m in stimulated cultures divided by the c.p.m. in matched, unstimulated cultures. View largeDownload slide Fig. 3. Anti‐Leishmania antibody responses. (A) Anti‐Leishmania serum IgG at a dilution of 1:10,000 as determined by absorption at 405 nm. (B) Isotypes detected in these sera at a dilution of 1:100 in wild‐type C57BL/6 and CD36‐null mice. Each point represents an individual mouse. (C) Lesion development in C57BL/6 and CD36‐null mice infected intradermally with 105L. major promastigotes and monitored weekly. Each dot represents the jittered lesion score of an individual mouse, such that otherwise superimposed points can be separated. This allows the simultaneous display of all data points (23). (D) Data from one representative experiment in which macrophages were infected with L. major promastigotes and 500 cells were counted to determine the percent infected cells over a 72‐h period as described in Methods. Filled symbols, C57BL; open symbols, CD36–/– mice. View largeDownload slide Fig. 3. Anti‐Leishmania antibody responses. (A) Anti‐Leishmania serum IgG at a dilution of 1:10,000 as determined by absorption at 405 nm. (B) Isotypes detected in these sera at a dilution of 1:100 in wild‐type C57BL/6 and CD36‐null mice. Each point represents an individual mouse. (C) Lesion development in C57BL/6 and CD36‐null mice infected intradermally with 105L. major promastigotes and monitored weekly. Each dot represents the jittered lesion score of an individual mouse, such that otherwise superimposed points can be separated. This allows the simultaneous display of all data points (23). (D) Data from one representative experiment in which macrophages were infected with L. major promastigotes and 500 cells were counted to determine the percent infected cells over a 72‐h period as described in Methods. Filled symbols, C57BL; open symbols, CD36–/– mice. References 1 Corcoran, L. M., Karvelas, M., Nossal, G. J., Ye, Z. S., Jacks, T. and Baltimore, D. 1993. Oct‐2, although not required for early B‐cell development, is critical for later B‐cell maturation and for postnatal survival. Genes Dev.  7: 570. Google Scholar 2 Corcoran, L. M. and Karvelas, M. 1994. Oct‐2 is required early in T cell‐independent B cell activation for G1 progression and for proliferation. Immunity  1: 635. Google Scholar 3 Humbert, P. O. and Corcoran, L. M. 1997. Oct‐2 gene disruption eliminates the peritoneal B‐1 lymphocyte lineage and attenuates B‐2 cell maturation and function. J. Immunol.  159: 5273. Google Scholar 4 Allman, D. M., Ferguson, S. E. and Cancro, M. P. 1992. Peripheral B cell maturation. I. Immature peripheral B cells in adults are heat‐stable antigenhi and exhibit unique signaling characteristics. J. Immunol.  149: 2533. Google Scholar 5 Allman, D. M., Ferguson, S. E., Lentz, V. M. and Cancro, M. P. 1993. Peripheral B cell maturation. II. Heat‐stable antigenhi splenic B cells are an immature developmental intermediate in the production of long‐lived marrow‐derived B cells. J. Immunol.  151: 4431. Google Scholar 6 Carsetti, R., Kohler, G. and Lamers, M. C. 1995. Transitional B cells are the target of negative selection in the B cell compartment. J. Exp. Med.  181: 2129. Google Scholar 7 Sater, R. A., Sandel, P. C. and Monroe, J. G. 1998. BCR‐induced apoptosis in primary transitional murine B cells: signaling requirements and modulation by T cell help. Int. Immunol.  10: 1673. Google Scholar 8 Konig, H., Pfisterer, P., Corcoran, L. M. and Wirth, T. 1995. Identification of CD36 as the first gene dependent on the B‐cell differentiation factor Oct‐2. Genes Dev.  9: 1598. Google Scholar 9 Febbraio, M., Hajjar, D. P. and Silverstein, R. L. 2001. CD 36: a class B scavenger receptor involved in angiogenesis, atherosclerosis, inflammation, and lipid metabolism. J. Clin. Invest.  108: 785. Google Scholar 10 Urban, B. C., Willcox, N. and Roberts, D. J. 2001. A role for CD36 in the regulation of dendritic cell function. Proc. Natl Acad. Sci. USA  98: 8750. Google Scholar 11 Albert, M. L., Pearce, S. F., Francisco, L. M., Sauter, B., Roy, P., Silverstein, R. L. and Bhardwaj, N. 1998. Immature dendritic cells phagocytose apoptotic cells via alphavbeta5 and CD36, and cross‐present antigens to cytotoxic T lymphocytes. J. Exp. Med.  188: 1359. Google Scholar 12 Febbraio, M., Abumrad, N. A., Hajjar, D. P., Sharma, K., Cheng, W., Pearce, S. F. and Silverstein, R. L. 1999. A null mutation in murine CD36 reveals an important role in fatty acid and lipoprotein metabolism. J. Biol. Chem.  274: 19055. Google Scholar 13 Rutella, S., Rumi, C., Puggioni, P., Barberi, T., Di Mario, A., Larocca, L. M. and Leone, G. 1999. Expression of thrombospondin receptor (CD36) in B‐cell chronic lymphocytic leukemia as an indicator of tumor cell dissemination. Haematologica  84: 419. Google Scholar 14 Reiner, S. L. and Locksley, R. M. 1995. The regulation of immunity to Leishmania major. Annu. Rev. Immunol.  13: 151. Google Scholar 15 Roberts, L. J., Baldwin, T. M., Curtis, J. M., Handman, E. and Foote, S. J. 1997. Resistance to Leishmania major is linked to the H2 region on chromosome 17 and to chromosome 9. J. Exp. Med.  185: 1705. Google Scholar 16 Liew, F. Y., Li, Y., Moss, D., Parkinson, C., Rogers, M. V. and Moncada, S. 1991. Resistance to Leishmania major infection correlates with the induction of nitric oxide synthase in murine macrophages. Eur. J. Immunol.  21: 3009. Google Scholar 17 Liew, F. Y. and O’Donnell, C. A. 1993. Immunology of leishmaniasis. Adv. Parasitol.  32: 161. Google Scholar 18 Scott, P., Natovitz, P. and Sher, A. 1986. B lymphocytes are required for the generation of T cells that mediate healing of cutaneous leishmaniasis. J. Immunol.  137: 1017. Google Scholar 19 Babai, B., Louzir, H., Cazenave, P. A. and Dellagi, K. 1999. Depletion of peritoneal CD5+ B cells has no effect on the course of Leishmania major infection in susceptible and resistant mice. Clin. Exp. Immunol.  117: 123. Google Scholar 20 Vremec, D., Zorbas, M., Scollay, R., Saunders, D. J., Ardavin, C. F., Wu, L. and Shortman, K. 1992. The surface phenotype of dendritic cells purified from mouse thymus and spleen: investigation of the CD8 expression by a subpopulation of dendritic cells. J. Exp. Med.  176: 47. Google Scholar 21 Vremec, D., Pooley, J., Hochrein, H., Wu, L. and Shortman, K. 2000. CD4 and CD8 expression by dendritic cell subtypes in mouse thymus and spleen. J. Immunol.  164: 2978. Google Scholar 22 Corcoran, L. M. and Metcalf, D. 1999. IL‐5 and Rp105 signaling defects in B cells from commonly used 129 mouse substrains. J. Immunol.  163: 5836. Google Scholar 23 Mitchell, G. F., Curtis, J. M., Handman, E. and McKenzie, I. F. 1980. Cutaneous leishmaniasis in mice: disease patterns in reconstituted nude mice of several genotypes infected with Leishmania tropica. Aust. J. Exp. Biol. Med. Sci.  58: 521. Google Scholar 24 Martin, R. M., Brady, J. L. and Lew, A. M. 1998. The need for IgG2c specific antiserum when isotyping antibodies from C57BL/6 and NOD mice. J. Immunol. Methods  15: 187. Google Scholar 25 Roberts, L. J., Foote, S. J. and Handman, E. 2000. A new standard for the assessment of disease progression in murine cutaneous leishmaniasis. Parasite Immunol.  22: 231. Google Scholar 26 Scott, P., Pearce, E. Natovitz, P. and Sher, A. 1987. Vaccination against cutaneous leishmaniasis in a murine model. II. Immunologic properties of protective and nonprotective subfractions of soluble promastigote extract. J. Immunol.  139: 3118. Google Scholar 27 Morris, L., Troutt, A. B., Handman, E. and Kelso, A. 1992. Changes in the precursor frequencies of IL‐4 and IFN‐gamma secreting CD4+ cells correlate with resolution of lesions in murine cutaneous leishmaniasis. J. Immunol.  149: 2715. Google Scholar 28 Alexander, J., Satoskar, A. R. and Russell, D. G. 1999. Leishmania species: models of intracellular parasitism. J. Cell Sci.  112: 2993. Google Scholar 29 Shore, P., Dietrich, W. and Corcoran, L. M. 2002. Oct‐2 regulates CD36 gene expression via a consenus octamer which excludes the co‐activator OBF‐1. Nucleic Acids Res.  30: 1767. Google Scholar 30 Cockerill, P. N. and Klinken, S. P. 1990. Octamer‐binding proteins in diverse hemopoietic cells. Mol. Cell. Biol.  10: 1293. Google Scholar 31 Lutterbach, B. and Hiebert, S. W. 2000. Role of the transcription factor AML‐1 in acute leukemia and hematopoietic differentiation. Gene  245: 223. Google Scholar 32 Yankee, T. M. and Clark, E. A. 2000. Signaling through the B cell antigen receptor in developing B cells. Rev. Immunogenet.  2: 185. Google Scholar 33 Hardy, R. R., Li, Y. S., Allman, D., Asano, M., Gui, M. and Hayakawa, K. 2000. B‐cell commitment, development and selection. Immunol. Rev.  175: 23. Google Scholar Author notes 1The Walter and Eliza Hall Institute of Medical Research, PO Royal Melbourne Hospital, Victoria 3050, Australia 2Department of Medicine, Division of Hematology and Medical Oncology, Cornell University, New York, NY 10021, USA http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png International Immunology Oxford University Press

Differential regulation of CD36 expression in antigen‐presenting cells: Oct‐2 dependence in B lymphocytes but not dendritic cells or macrophages

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Oxford University Press
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0953-8178
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1460-2377
DOI
10.1093/intimm/dxf075
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Abstract

Abstract In mice, three antigen‐presenting cell types [B lymphocytes, macrophages and dendritic cells (DC)] express the scavenger receptor CD36. This molecule has been implicated in many important functions, including DC maturation and antigen presentation. In murine B cells, the CD36 gene requires the Oct‐2 transcription factor for its expression. We previously found that B cells from Oct‐2‐null mice display defects in maturation, survival and proliferation. Here we have looked for a possible role for CD36 in B cells, but found that CD36 is dispensable for all responses tested. Although loss of CD36 did not directly affect B cell function, it did modulate slightly the isotype and level of IgG produced in vivo in naive mice, and IgM in Leishmania‐infected mice. We also show that in DC and macrophages, CD36 expression is independent of Oct‐2. We conclude that CD36 does not play a major role in B cell function, but that CD36 may contribute indirectly to humoral immunity through cells of the innate immune system. B lymphocyte, dendritic cell, Leishmania major, monocyte/macrophage, transgenic/knockout Introduction The B cell‐restricted transcriptional activator, Oct‐2, binds to a conserved site found in most Ig gene promoters. However, through targeted mutation of the gene for Oct‐2, we have shown that Ig gene promoters do not require this factor for expression, at least in newly formed peripheral B cells. Nevertheless, Oct‐2‐null B cells display several abnormalities (1–3). Without Oct‐2, immature B cells emigrating from the bone marrow fail to differentiate efficiently into long‐lived cells, resulting in a deficit of mature peripheral B cells. Serum Ig levels are markedly reduced in naive and immunized oct‐2–/– mice and the peritoneal B‐1 lymphocyte compartment is absent. In vitro, oct‐2–/– B cells maintain properties of immature ‘transitional’ B cells (3–5) which do not proliferate, but undergo apoptosis in response to a BCR signal (6,7). Presumably, these effects are due to the dysregulation of Oct‐2‐dependent target genes in B cells. One of these, the first Oct‐2‐dependent gene identified, is that encoding the cell surface protein CD36 (8). CD36 is expressed on many cell types, including some hematopoietic cells (9). It acts as a multi‐ligand scavenger receptor that can mediate adhesion via thrombospondin and scavenging of apoptotic cells. A role for CD36 in dendritic cell (DC) maturation and antigen cross‐presentation has been proposed (10,11), and its critical role in fatty acid metabolism has recently been proven (12). However, no function has yet been ascribed to CD36 in B lymphocytes. Indeed, while mouse B cells express CD36, most normal human B cells do not. Instead, B cell leukemias are often CD36+, with expression correlating with more advanced disease (13). These observations prompted three questions. First, is the CD36 gene dependent upon Oct‐2 in B cells, macrophages and DC, the three antigen‐presenting cells where it is expressed? Second, does the inability to express CD36 contribute to the abnormal phenotype of Oct‐2‐null B cells? Third, does the loss of CD36 impact on the immune response to a pathogen in vivo? This examination of hematopoietic cells from Oct‐2‐null mice shows that CD36 relies on Oct‐2 for its expression only in B lymphocytes. To investigate the consequences of CD36 loss to the B cell system, we assessed the B cell compartment of CD36‐null mice, examining cell subpopulations and phenotypes, B cell responses to stimulation in vitro, and responses in a T cell‐dependent model of infection with the parasite Leishmania major. Murine cutaneous leishmaniasis is caused by L. major, an intracellular parasite of mononuclear phagocytes. Intradermal infection with L. major produces a local skin lesion and self‐limiting granuloma that heals in 10–12 weeks in genetically resistant mice (e.g. C57BL/6 mice) or it may disseminate systemically and kill the host (14,15). Recovery is dependent upon the induction of Th1‐type pro‐inflammatory cytokines such as IL‐12 and IFN‐γ, and the generation of CD4+ Th1‐type cells. Th1 cytokines induce macrophage activation and killing of the intracellular organisms by NO (16,17). B cells also play a role in T cell‐mediated healing (18), but depletion of CD5+ B cells has no effect on the course of infection (19). Given the central roles of the T cell‐activated macrophage and B cells in host resistance against leishmaniasis, we included this model infection in our characterization of CD36 activities in vivo. Methods Mice and tissues Oct‐2‐null mice die at birth, so adoptive transfer of embryonic day 13 (E13) fetal liver into RAG‐1‐deficient recipients was used to generate mature lymphoid cells, as described (2). To access oct‐2–/– macrophages and DC, oct‐2+/– mice were crossed, E18 fetal liver taken and genotyped. Cells were stained directly ex vivo for lineage markers and for CD36 expression as described below. DC and macrophages were isolated as previously described (20,21), with slight modifications to maximize yields, using collagenase digestion and immunomagnetic bead depletion of either non‐DC or non‐macrophages. For DC, cells were depleted with mAb anti‐CD3 (KT3‐1.1), anti‐Thy1 (T24/31.7), anti‐B220 (RA36B2), anti‐Gr1 (RB68C5) and anti‐erythrocyte (TER119). For macrophages, depletion used mAb anti‐CD3 (KT3‐1.1), anti‐CD4 (GK1.5), anti‐CD8α (53‐6.7), anti‐B220 (RA36B2) and anti‐erythroid (TER119). Macrophages were also expanded from unfractionated E13 fetal liver in the presence of macrophage colony stimulating factor (M‐CSF; 1000 U/ml in DME + 10% FBS). Duplicate cultures were maintained for 6 days and one was supplemented with lipopolysaccharide (LPS, 1 µg/ml; Difco, Detroit, MI) for the final 24 h. The adherent cells were dislodged by pipetting in EDTA/FBS. The vast majority of the cells in all cultures were macrophages (CD11b+; data not shown). The CD36 knockout mice were generated on the 129/Sv background (12). We had shown that this strain can display anomalous B cell behavior (22), so we backcrossed the mice for six generations to C57BL/6 mice before performing the analyses described here. ELISAs Serum was diluted through a mid‐log10 series from an original 1/100 dilution. ELISAs were performed as described (2,24). For the anti‐Leishmania antibody responses, the dilution was 1:10,000 and the Ig isotype determined using an isotyping kit according to the manufacturer’s recommendations (Bio‐Rad, Hercules, CA). For the Ig2c isotype, a specific antibody was used as in (24). Flow cytometry Peripheral blood lymphocytes from heparinized, red cell‐depleted blood were stained with anti‐B220 (RA36B2). Splenocytes and peritoneal cells were prepared and stained as described (3). Details of the mAb and labeling procedure for DC and macrophages have been described elsewhere (20,21). All samples were pre‐incubated with a mix of whole mouse Ig and anti‐FcRII/III (2.4G2) to minimize non‐specific staining. The mAb used to identify DC and macrophages were anti‐CD11c (N418) and anti‐CD11b (M1/70) respectively. Propidium iodide (PI) was included in the final wash at 1 µg/ml to label dead cells. Analyses were carried out on either a FACStar Plus or FACScan (Becton Dickinson, San Jose, CA). Cell surface CD36 expression was then determined by gating on live CD11c+ DC, CD11b+ macrophages and B220+ B cells. CD36 was detected by anti‐CD36 (clone63; Cascade BioScience, Winchester, MA) and an anti‐mouse IgA second stage (Caltag, Burlingame, CA) after blocking with whole rat Ig. Lymphocyte proliferation assays Spleen cell suspensions were cultured at 1 × 106 cells/ml for 72 h with or without B cell mitogens and pulsed with 1 µCi [3H]thymidine during the final 5 h of culture, as described (2). Parasites Parasites were of the virulent cloned line of L. major LRC‐L137 (MHOM/IL/67/JerichoII). Promastigotes were maintained in vitro at 26°C in Schneider’s Drosophila medium with 10% FBS and used in stationary phase. Cutaneous infection of wild‐type and CD36‐null mice Mice were injected intradermally with 105 promastigotes. Lesion development was assessed weekly for 10–12 weeks, according to the lesion‐scoring system described (23,25). Mice were bled periodically, and their Leishmania‐specific antibody titers and isotypes measured as described above. A soluble parasite lysate (SLA) obtained by freezing and thawing promastigotes was used as antigen (26,27). In vitro infection of macrophages with L. major promastigotes Bone marrow‐derived macrophages from wild‐type and CD36‐null mice were cultured for 5–7 days in RPMI medium with 10% FBS and 10% supernatant from cultures of the 929 cell line as a source of M‐CSF (28). Macrophages were transferred onto coverslips (2 × 105 cells/well in 0.5 ml of medium) and allowed to adhere for 18 h at 37°C. Non‐adherent cells were washed away and monolayers were infected with L. major promastigotes at a ratio of 5:1. After 30 min, free parasites were removed and the cells were re‐cultured for 2, 24, 48 or 72 h. The cells were then fixed in methanol and stained with Giemsa. For each time point 500 cells were counted, and the percent infected cells and number of parasites present were calculated. Experiments were performed at least 3 times with duplicate samples. Results and discussion Expression of CD36 in myeloid cells is Oct‐2 independent Earlier experiments (8,29) proved that transcription of the CD36 gene was directly dependent upon Oct‐2 in a pre‐B lymphoma cell line. Konig et al. (8) also showed a strong correlation between CD36 and Oct‐2 expression in a number of B and monocyte/macrophage cell lines and in primary hematopoietic tissues. However, a direct effect of Oct‐2 on CD36 expression was not proven for the myeloid cells. Here we have examined the surface expression of CD36 in three primary hematopoietic tissues from oct‐2–/– mice to determine the contribution of Oct‐2 to CD36 expression in each cell type. As expected, B cells require Oct‐2 for CD36 expression (Fig. 1A). However, when DC (CD11c+) and macrophages (CD11b+) from fetal liver were examined, there was no correlation between CD36 expression and oct‐2 genotype. All DC were CD36+ at the time of isolation and expressed equivalent levels of the protein (Fig. 1B, top). Surprisingly, none of the CD11b+ macrophages directly ex vivo expressed CD36 (Fig. 1B, middle). We therefore expanded macrophages from E13 fetal liver in the presence of M‐CSF and added LPS during the final 24 h to half the cultures, to activate the cells. We found that expanding the cells in M‐CSF was sufficient to induce a high level of CD36 expression in the cultured macrophages, which was not altered by LPS addition (Fig. 1B, bottom). We also learned that Oct‐2 is not required for CD36 expression in macrophages, as the oct‐2‐null cells expressed normal CD36 levels. These data indicate that the CD36 gene is regulated differentially in antigen‐presenting cells, with B cells expressing intermediate levels, and DC and macrophages expressing at least 10‐fold higher levels of surface CD36. Despite the presence of Oct‐2 in cells of both the myeloid and B lymphoid lineages (8,30), only B cells require Oct‐2 as a critical CD36 regulator, with the Oct‐2 binding site in the promoter acting as the central regulatory element (29). These studies do not exclude the possible involvement of other octamer‐binding factors in the regulations of the CD36 gene in myeloid cells, in particular the ubiquitously expressed Oct‐1. Studies on CD36 expression in Oct‐1‐deficient animals will clarify this issue. Interestingly, however, immediately adjacent to the Oct‐2 site in the CD36 promoter is a binding site for the transcriptional regulator AML‐1. AML‐1 activates the transcription of several tissue‐specific genes in myeloid cells (31) and may contribute to the high level of CD36 gene expression in macrophages and DC. Features of the B cell compartment in CD36‐null mice To explore whether Oct‐2‐mediated CD36 expression in B cells had evolved to enable some humoral immune function, we examined B cells and their products in CD36‐null mice. We compared serum Ig levels in naive C57BL/6 and CD36–/– mice, and found that CD36‐deficient mice had normal or even slightly higher than normal titers of all Ig isotypes tested (Fig. 2A). In C57BL/6 mice, the IgG2a gene has been deleted and the alternative IgG2c gene is expressed (24). 129/Sv mice have the opposite profile. The CD36–/– mice, generated in the 129/Sv strain and backcrossed onto the C57BL/6 background, express IgG2c, but not IgG2a, and express the same level of IgG2c as the control C57BL/6 mice (Fig. 2A). This overall picture contrasts with Oct‐2‐deficient mice, which have ∼10% of normal levels of most Ig isotypes (2). These observations suggest that loss of CD36 expression in B cells does not compromise Ig gene expression or secretion and so is not a primary limiting factor in Oct‐2‐deficient B cells. However, loss of CD36 does influence the profile of Ig isotypes expressed in vivo, with IgG1 and IgG2b being somewhat favored in this set of mice. In other experiments, the absolute differences between IgG1 and IgG2b levels for C56BL/6 control and CD36–/– mice varied, but levels in CD36‐null mice were always higher than controls (data not shown). A characteristic feature of peripheral oct‐2–/– B cells is their immaturity. Mature cells down‐regulate expression of the heat‐stable antigen (HSA; CD24) from the high level expressed on recent bone marrow emigrants (4,5). In Oct‐2‐null mice, mature (HSAlo) cells are virtually absent (3). In contrast, mature and immature B cells in the spleen of CD36‐null mice were present in normal ratios (Fig. 2B). Clearly, CD36 is not essential for the signal that mediates this maturation step. Peritoneal B‐1 cells were also present in normal numbers in CD36‐null mice (Fig. 2C), while they are absent in Oct‐2‐null mice (3). Like the peripheral maturation of immature B cells, generation and maintenance of B‐1 lymphocytes require signals through the BCR (32,33). Oct‐2 is required for the successful receipt of such signals in both the B‐1 and the B‐2 compartments, perhaps by regulating expression of a component(s) of the BCR signaling pathway. Our conclusion is that CD36 is not a critical Oct‐2 target gene in the context of these important cell survival and maturation signals. Responses of CD36‐null B cells to mitogenic stimulation are normal Proliferative responses of CD36‐deficient splenocytes to a variety of B cell mitogens were examined and found to be normal (Fig. 2D). This is in stark contrast to Oct‐2‐deficient B cells, which are severely hyporesponsive to LPS, to anti‐Rp105 signaling and to IL‐5 stimulation, and significantly hyporesponsive to anti‐µ stimulation [(2) and our unpublished observations]. The data in Figs 1 and 2 strongly suggest that, despite the clear dependence of the CD36 gene on Oct‐2 for its expression in B lymphocytes (29), CD36 insufficiency is not a major factor in the defective phenotype of Oct‐2‐deficient B cells. Indeed, we found no evidence that CD36 plays a significant role in any of the B cell‐autonomous attributes we examined here. However, the combined loss of CD36 and other Oct‐2 target genes may be responsible for some of the deficiencies exhibited by oct‐2‐null B cells. Humoral immune response in L. major‐infected CD36‐null mice L. major‐infected CD36‐null mice had elevated Leishmania‐specific total IgG compared to controls (Fig. 3A). However, there was no significant difference in the levels of specific IgG isotypes examined (Fig. 3B). This mirrored the pattern seen in naive CD36–/– mice (Fig. 2A). In contrast, the level of Leishmania‐specific IgM detected in the wild‐type mice was significantly higher than in the CD36‐null mice. L. major‐infected CD36‐null mice developed smaller lesions and appeared to cure somewhat faster than control mice, with all mice cured by week 7. The smaller lesion size was significant in the early weeks post‐infection as determined by the Mann–Whitney statistical test (P < 0.05 at weeks 3, 5 and 6; Fig. 3C). The disease pattern suggests that loss of CD36 expression engenders an even more pronounced Th1 immune response to the parasite than is exhibited by the already genetically resistant C57BL/6 mouse. Indeed, Urban et al. (10) have shown that CD36 ligation can strongly influence cytokine production by T cells. The hypothesis that CD36 promotes resistance to infection by biasing the Th cell phenotype can be tested in genetically susceptible BALB/c mice lacking the CD36 gene. To what extent these responses are affected by the antigen‐processing and ‐presentation ability of the CD36‐null macrophages remains to be established. However, we could find no significant difference in the ability of cells from the CD36‐null mice to support parasite growth compared to wild‐type mice (Fig. 3D). L. major promastigotes were able to establish infection in CD36‐null bone marrow‐derived macrophages as efficiently as in wild‐type cells (Fig. 3D) and the number of parasites per infected macrophage was similar during a 72 h period of infection (data not shown). Therefore, CD36 expression is not a limiting factor for parasite invasion and survival in macrophages. In conclusion, we have shown that the Oct‐2 transcription factor is necessary for the expression of the CD36 protein on murine B cells, but that specific loss of CD36 from these cells is relatively inconsequential. We found no evidence of B cell dysfunction in CD36–/– mice in the assays performed, which focused on those capacities that are compromised in Oct‐2‐deficient B cells. In contrast, CD36 expression is Oct‐2‐independent in myeloid cells (DC and macrophages, in particular), arguing that the gene is differentially regulated in these antigen‐presenting cells. We observed a potential influence of CD36 on the humoral immune response, in the altered spectrum of Ig isotypes and the response to L. major infection. CD36 may therefore play a wider role in immune regulation, indirectly influencing the quality of a humoral immune response via influences on the balance of Th1/Th2 cells, as well as through cells of the innate immune system. These observations are consistent with the proposal that CD36 is an ancient functional component of innate immunity (9). Acknowledgements We thank Wendy Deitrich and Eren Loza for technical assistance and animal care respectively, and Dr D. Tarlinton for critical reading of the manuscript. Professor D. Metcalf generously provided murine M‐CSF, Dr A. Lew provided anti‐IgG2 and Dr A. Strasser provided a number of mAb for cell surface staining. This work was supported by the Australian National Health and Medical Research Council, and the UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases (TDR). Abbreviations DC—dendritic cell E—embryonic day HSA—heat‐stable antigen LPS—lipopolysaccharide M‐CSF—macrophage stimulating factor PI—propidium iodide View largeDownload slide Fig. 1. CD36 expression on B lymphoid and myeloid cells. (A) Surface CD36 expression on B220+ peripheral blood cells. (B) CD36 staining on dendritic cells (CD11c+) and macrophages (CD11b+) from E18 fetal liver or on macrophages expanded from E13 fetal liver in the presence of M‐CSF. LPS was added for the final 24 h of some cultures (+LPS). In all panels, the dotted line represents a background control (no primary antibody) and the solid line corresponds to staining with a CD36‐specific antibody. View largeDownload slide Fig. 1. CD36 expression on B lymphoid and myeloid cells. (A) Surface CD36 expression on B220+ peripheral blood cells. (B) CD36 staining on dendritic cells (CD11c+) and macrophages (CD11b+) from E18 fetal liver or on macrophages expanded from E13 fetal liver in the presence of M‐CSF. LPS was added for the final 24 h of some cultures (+LPS). In all panels, the dotted line represents a background control (no primary antibody) and the solid line corresponds to staining with a CD36‐specific antibody. View largeDownload slide Fig. 2. Assessment of CD36‐null B cells, their products and behavior. (A) Serum Ig titers in naive mice. Filled circles, C57BL/6 mice; open circles, CD36‐null mice; open squares with dotted lines, 129/Sv mice. Each line corresponds to the mean (±SD) of three or more animals. (B) HSA levels on splenic B220+ cells. The dotted line delineates immature (HSAhi) from mature (HSAlo) cells, as determined by Allman et al. (4,5). The data are representative of an analysis of four mice of each genotype, with the number indicating the mean percentage of cells in the indicated region for all mice. (C) Lymphocyte populations in peritoneal cells harvested from C57BL/6 and CD36‐null mice, stained to highlight B‐1 lymphocytes (boxed). This panel shows representative data from four mice of each type (the number indicates the mean of cells in the B‐1 region for all four mice). Staining with B220 and Mac‐1 or with B220 and CD5 gave the same result, i.e. no difference between the control and mutants. (D) Proliferative responses of splenocytes to a number of B cell mitogens. Values are the mean (±SD) for triplicate wells and each bar represents a single mouse. Filled bars, C57BL/6 mice; hatched bars, CD36‐null mice. Stimulation index is the ratio of c.p.m in stimulated cultures divided by the c.p.m. in matched, unstimulated cultures. View largeDownload slide Fig. 2. Assessment of CD36‐null B cells, their products and behavior. (A) Serum Ig titers in naive mice. Filled circles, C57BL/6 mice; open circles, CD36‐null mice; open squares with dotted lines, 129/Sv mice. Each line corresponds to the mean (±SD) of three or more animals. (B) HSA levels on splenic B220+ cells. The dotted line delineates immature (HSAhi) from mature (HSAlo) cells, as determined by Allman et al. (4,5). The data are representative of an analysis of four mice of each genotype, with the number indicating the mean percentage of cells in the indicated region for all mice. (C) Lymphocyte populations in peritoneal cells harvested from C57BL/6 and CD36‐null mice, stained to highlight B‐1 lymphocytes (boxed). This panel shows representative data from four mice of each type (the number indicates the mean of cells in the B‐1 region for all four mice). Staining with B220 and Mac‐1 or with B220 and CD5 gave the same result, i.e. no difference between the control and mutants. (D) Proliferative responses of splenocytes to a number of B cell mitogens. Values are the mean (±SD) for triplicate wells and each bar represents a single mouse. Filled bars, C57BL/6 mice; hatched bars, CD36‐null mice. Stimulation index is the ratio of c.p.m in stimulated cultures divided by the c.p.m. in matched, unstimulated cultures. View largeDownload slide Fig. 3. Anti‐Leishmania antibody responses. (A) Anti‐Leishmania serum IgG at a dilution of 1:10,000 as determined by absorption at 405 nm. (B) Isotypes detected in these sera at a dilution of 1:100 in wild‐type C57BL/6 and CD36‐null mice. Each point represents an individual mouse. (C) Lesion development in C57BL/6 and CD36‐null mice infected intradermally with 105L. major promastigotes and monitored weekly. Each dot represents the jittered lesion score of an individual mouse, such that otherwise superimposed points can be separated. This allows the simultaneous display of all data points (23). (D) Data from one representative experiment in which macrophages were infected with L. major promastigotes and 500 cells were counted to determine the percent infected cells over a 72‐h period as described in Methods. Filled symbols, C57BL; open symbols, CD36–/– mice. View largeDownload slide Fig. 3. Anti‐Leishmania antibody responses. (A) Anti‐Leishmania serum IgG at a dilution of 1:10,000 as determined by absorption at 405 nm. (B) Isotypes detected in these sera at a dilution of 1:100 in wild‐type C57BL/6 and CD36‐null mice. Each point represents an individual mouse. (C) Lesion development in C57BL/6 and CD36‐null mice infected intradermally with 105L. major promastigotes and monitored weekly. Each dot represents the jittered lesion score of an individual mouse, such that otherwise superimposed points can be separated. This allows the simultaneous display of all data points (23). (D) Data from one representative experiment in which macrophages were infected with L. major promastigotes and 500 cells were counted to determine the percent infected cells over a 72‐h period as described in Methods. Filled symbols, C57BL; open symbols, CD36–/– mice. References 1 Corcoran, L. M., Karvelas, M., Nossal, G. J., Ye, Z. S., Jacks, T. and Baltimore, D. 1993. Oct‐2, although not required for early B‐cell development, is critical for later B‐cell maturation and for postnatal survival. Genes Dev.  7: 570. Google Scholar 2 Corcoran, L. M. and Karvelas, M. 1994. Oct‐2 is required early in T cell‐independent B cell activation for G1 progression and for proliferation. Immunity  1: 635. Google Scholar 3 Humbert, P. O. and Corcoran, L. M. 1997. Oct‐2 gene disruption eliminates the peritoneal B‐1 lymphocyte lineage and attenuates B‐2 cell maturation and function. J. Immunol.  159: 5273. Google Scholar 4 Allman, D. M., Ferguson, S. E. and Cancro, M. P. 1992. Peripheral B cell maturation. I. Immature peripheral B cells in adults are heat‐stable antigenhi and exhibit unique signaling characteristics. J. Immunol.  149: 2533. Google Scholar 5 Allman, D. M., Ferguson, S. E., Lentz, V. M. and Cancro, M. P. 1993. Peripheral B cell maturation. II. Heat‐stable antigenhi splenic B cells are an immature developmental intermediate in the production of long‐lived marrow‐derived B cells. J. Immunol.  151: 4431. Google Scholar 6 Carsetti, R., Kohler, G. and Lamers, M. C. 1995. Transitional B cells are the target of negative selection in the B cell compartment. J. Exp. Med.  181: 2129. Google Scholar 7 Sater, R. A., Sandel, P. C. and Monroe, J. G. 1998. BCR‐induced apoptosis in primary transitional murine B cells: signaling requirements and modulation by T cell help. Int. Immunol.  10: 1673. Google Scholar 8 Konig, H., Pfisterer, P., Corcoran, L. M. and Wirth, T. 1995. Identification of CD36 as the first gene dependent on the B‐cell differentiation factor Oct‐2. Genes Dev.  9: 1598. Google Scholar 9 Febbraio, M., Hajjar, D. P. and Silverstein, R. L. 2001. CD 36: a class B scavenger receptor involved in angiogenesis, atherosclerosis, inflammation, and lipid metabolism. J. Clin. Invest.  108: 785. Google Scholar 10 Urban, B. C., Willcox, N. and Roberts, D. J. 2001. A role for CD36 in the regulation of dendritic cell function. Proc. Natl Acad. Sci. USA  98: 8750. Google Scholar 11 Albert, M. L., Pearce, S. F., Francisco, L. M., Sauter, B., Roy, P., Silverstein, R. L. and Bhardwaj, N. 1998. Immature dendritic cells phagocytose apoptotic cells via alphavbeta5 and CD36, and cross‐present antigens to cytotoxic T lymphocytes. J. Exp. Med.  188: 1359. Google Scholar 12 Febbraio, M., Abumrad, N. A., Hajjar, D. P., Sharma, K., Cheng, W., Pearce, S. F. and Silverstein, R. L. 1999. A null mutation in murine CD36 reveals an important role in fatty acid and lipoprotein metabolism. J. Biol. Chem.  274: 19055. Google Scholar 13 Rutella, S., Rumi, C., Puggioni, P., Barberi, T., Di Mario, A., Larocca, L. M. and Leone, G. 1999. Expression of thrombospondin receptor (CD36) in B‐cell chronic lymphocytic leukemia as an indicator of tumor cell dissemination. Haematologica  84: 419. Google Scholar 14 Reiner, S. L. and Locksley, R. M. 1995. The regulation of immunity to Leishmania major. Annu. Rev. Immunol.  13: 151. Google Scholar 15 Roberts, L. J., Baldwin, T. M., Curtis, J. M., Handman, E. and Foote, S. J. 1997. Resistance to Leishmania major is linked to the H2 region on chromosome 17 and to chromosome 9. J. Exp. Med.  185: 1705. Google Scholar 16 Liew, F. Y., Li, Y., Moss, D., Parkinson, C., Rogers, M. V. and Moncada, S. 1991. Resistance to Leishmania major infection correlates with the induction of nitric oxide synthase in murine macrophages. Eur. J. Immunol.  21: 3009. Google Scholar 17 Liew, F. Y. and O’Donnell, C. A. 1993. Immunology of leishmaniasis. Adv. Parasitol.  32: 161. Google Scholar 18 Scott, P., Natovitz, P. and Sher, A. 1986. B lymphocytes are required for the generation of T cells that mediate healing of cutaneous leishmaniasis. J. Immunol.  137: 1017. Google Scholar 19 Babai, B., Louzir, H., Cazenave, P. A. and Dellagi, K. 1999. Depletion of peritoneal CD5+ B cells has no effect on the course of Leishmania major infection in susceptible and resistant mice. Clin. Exp. Immunol.  117: 123. Google Scholar 20 Vremec, D., Zorbas, M., Scollay, R., Saunders, D. J., Ardavin, C. F., Wu, L. and Shortman, K. 1992. The surface phenotype of dendritic cells purified from mouse thymus and spleen: investigation of the CD8 expression by a subpopulation of dendritic cells. J. Exp. Med.  176: 47. Google Scholar 21 Vremec, D., Pooley, J., Hochrein, H., Wu, L. and Shortman, K. 2000. CD4 and CD8 expression by dendritic cell subtypes in mouse thymus and spleen. J. Immunol.  164: 2978. Google Scholar 22 Corcoran, L. M. and Metcalf, D. 1999. IL‐5 and Rp105 signaling defects in B cells from commonly used 129 mouse substrains. J. Immunol.  163: 5836. Google Scholar 23 Mitchell, G. F., Curtis, J. M., Handman, E. and McKenzie, I. F. 1980. Cutaneous leishmaniasis in mice: disease patterns in reconstituted nude mice of several genotypes infected with Leishmania tropica. Aust. J. Exp. Biol. Med. Sci.  58: 521. Google Scholar 24 Martin, R. M., Brady, J. L. and Lew, A. M. 1998. The need for IgG2c specific antiserum when isotyping antibodies from C57BL/6 and NOD mice. J. Immunol. Methods  15: 187. Google Scholar 25 Roberts, L. J., Foote, S. J. and Handman, E. 2000. A new standard for the assessment of disease progression in murine cutaneous leishmaniasis. Parasite Immunol.  22: 231. Google Scholar 26 Scott, P., Pearce, E. Natovitz, P. and Sher, A. 1987. Vaccination against cutaneous leishmaniasis in a murine model. II. Immunologic properties of protective and nonprotective subfractions of soluble promastigote extract. J. Immunol.  139: 3118. Google Scholar 27 Morris, L., Troutt, A. B., Handman, E. and Kelso, A. 1992. Changes in the precursor frequencies of IL‐4 and IFN‐gamma secreting CD4+ cells correlate with resolution of lesions in murine cutaneous leishmaniasis. J. Immunol.  149: 2715. Google Scholar 28 Alexander, J., Satoskar, A. R. and Russell, D. G. 1999. Leishmania species: models of intracellular parasitism. J. Cell Sci.  112: 2993. Google Scholar 29 Shore, P., Dietrich, W. and Corcoran, L. M. 2002. Oct‐2 regulates CD36 gene expression via a consenus octamer which excludes the co‐activator OBF‐1. Nucleic Acids Res.  30: 1767. Google Scholar 30 Cockerill, P. N. and Klinken, S. P. 1990. Octamer‐binding proteins in diverse hemopoietic cells. Mol. Cell. Biol.  10: 1293. Google Scholar 31 Lutterbach, B. and Hiebert, S. W. 2000. Role of the transcription factor AML‐1 in acute leukemia and hematopoietic differentiation. Gene  245: 223. Google Scholar 32 Yankee, T. M. and Clark, E. A. 2000. Signaling through the B cell antigen receptor in developing B cells. Rev. Immunogenet.  2: 185. Google Scholar 33 Hardy, R. R., Li, Y. S., Allman, D., Asano, M., Gui, M. and Hayakawa, K. 2000. B‐cell commitment, development and selection. Immunol. Rev.  175: 23. Google Scholar Author notes 1The Walter and Eliza Hall Institute of Medical Research, PO Royal Melbourne Hospital, Victoria 3050, Australia 2Department of Medicine, Division of Hematology and Medical Oncology, Cornell University, New York, NY 10021, USA

Journal

International Immunology – Oxford University Press

Published: Oct 1, 2002

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