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Lymphocyte Development and Function in the Absence of Retinoic Acid-Related Orphan Receptor ¹

Ivan Dzhagalov^*, Vincent Giguère and You-Wen He^2,*

^* Department of Immunology, Duke University Medical Center, Durham, NC 27710; and Molecular Oncology Group, McGill University Health Center, Montreal, Quebec, Canada

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
The orphan nuclear receptor, retinoid acid-related orphan receptor (ROR), is essential for the development of cerebellar Purkinje cells and bone tissue. ROR may also play a critical role in lymphocyte development and function because staggerer mice, a natural mutant strain with a disrupted expression of ROR, have reduced thymic and splenic cellularity. In this report, we analyzed the role of ROR in lymphocyte development by examining lymphoid compartments in ROR^–/– mice and Rag-2^–/– mice reconstituted with ROR^–/– bone marrow. We found that T and B cell development was severely defective in ROR^–/– mice, but not in Rag-2^–/–/ROR^–/– chimeric mice. We also analyzed cellular and humoral immune responses in Rag-2^–/–/ROR^–/– chimeric mice. Our results show that serum IgG levels were elevated in Rag-2^–/–/ROR^–/– chimeric mice after immunization with a T-dependent Ag compared with control chimeras. IFN- production by ROR^–/– CD8⁺ T cells after TCR stimulation was also increased. Furthermore, ROR^–/– mast cells and macrophages produced an increased amount of TNF- and IL-6 upon activation. These results indicate that ROR indirectly regulates lymphocyte development by providing an appropriate microenvironment and controls immune responses by negatively regulating cytokine production in innate immune cells and lymphocytes.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Orphan nuclear receptors regulate multiple cellular processes including cell growth, differentiation, and apoptosis in a variety of tissues and organs (1, 2). The retinoid acid-related orphan receptor (ROR)³ subfamily of orphan nuclear receptors consists of three members, , , and , which share a similar structure (3, 4, 5). A highly conserved DNA-binding domain containing 66 aa, and a less conserved putative ligand-binding domain are located at the amino and carboxyl terminus of the proteins, respectively. Multiple isoforms that differ from each other only at the amino terminus have been identified for each member of RORs. The different isoforms of RORs are generated either by alternative mRNA splicing or differential promoter usage. ROR is expressed in at least four different isoforms, 1, 2, 3, and 4 in humans or mice (6, 7). Two isoforms have been identified for ROR (1 and 2) and ROR (1 and 2/t; Refs. 3, 4, 5). Recent studies demonstrate that ROR plays important roles in thymocyte development and lymphoid organogenesis (8, 9, 10, 11). ROR-deficient mice have massive apoptosis of thymocytes and lack lymph nodes and Peyer’s patches. Furthermore, the TCR repertoire was impaired in ROR mutant mice. The expression of ROR is largely restricted to the neuronal system (12).

ROR is expressed in several tissues including the spleen, skeletal muscle, testis, lens, retina, and the Purkinje cells of the cerebellum (6, 7). Consistent with its wide expression, ROR regulates neuronal cell development, bone metabolism, and arteriosclerosis (13, 14, 15). Staggerer mice, a natural mutant strain with a disrupted expression of ROR, show tremor, body imbalance, hypotonia, and small size, and die within 3–4 wk after birth. Mice with a targeted disruption of ROR expression recapitulated the neurological defects in staggerer mice (16, 17). These defects are caused by impaired generation and maintenance of the Purkinje cells. Staggerer mice also have thinner bones and are more susceptible to arteriosclerosis when maintained on an atherogenic diet, suggesting that ROR is critically involved in regulating bone metabolism and susceptibility to atherosclerosis.

Several studies have implicated ROR as a key regulator during lymphocyte development (14, 18). These studies reported smaller size of the thymus and spleen as well as reduced cellularity of the lymphoid organs in staggerer mice. However, the nature of the defects in lymphocyte development in staggerer mice was not defined. ROR may regulate immune responses because macrophages from staggerer mice produced more IL-1 and TNF- than controls after LPS stimulation (19, 20). Moreover, over-expression of ROR in smooth muscle cells inhibited TNF--induced expression of IL-6, IL-8, and cyclooxygenase-2, and up-regulated the expression of IB (21). Although the mutated gene in staggerer mice has been defined as ROR, it remains possible that mutations in other loci may contribute to the thymic and splenic defects in staggerer mice.

To clarify the role of ROR in lymphocyte development and function, we first determined the expression of ROR in T and B lymphocytes. We then analyzed T and B cell development in both ROR^–/– mice and Rag-2^–/–/ROR^–/– bone marrow (BM) chimeric mice. Moreover, we examined cellular and humoral immune responses in the Rag-2^–/–/ROR^–/– BM chimeras. Our results show that ROR indirectly regulates the development of T and B cells by providing an appropriate microenvironment and controls immune responses by negatively regulating cytokine production in innate immune cells and lymphocytes.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Mice

Homozygous ROR-deficient mice (–/–) were generated by the breeding of heterozygous mice (+/–) backcrossed to C57BL/6 for six generations (17). Because heterozygous mice did not have obvious defects and were indistinguishable from wild-type mice, both heterozygous and wild-type (+/+) littermates were used as controls throughout the study. Rag-2^–/–CD45.1⁺ congenic mice were kindly provided by Dr. M. Kondo (Duke University, Durham, NC). C57BL/6 mice were purchased from The Jackson Laboratory (Bar Harbor, ME). All mice were maintained under specific pathogen-free conditions at the Duke University Vivarium and used at the indicated age. All experiments were performed according to protocols approved by Duke University Animal Care and Use Committee.

Generation of BM chimeric mice

BM transfer was performed as described (22). Briefly, single cell suspensions from the BM of 2- to 3-wk-old ROR^–/– mice or littermate controls were lysed of RBC and injected into 6-wk or older Rag-2^–/– mice that received sublethal (500 rad) irradiation 24 h before the transfer. Rag-2^–/–/ROR^–/– and Rag-2^–/–/ROR^+/+ BM chimeric mice were used 6 wk after the BM transfer.

Flow cytometric analyses

Single-cell suspensions of the spleen, BM, thymus, and lymph nodes were lysed of erythrocytes, incubated with an FcR blocker (2.4G2), and stained with fluorochrome-labeled mAbs in PBS containing 2% FCS and 0.02% sodium azide. The following mAbs directly conjugated to FITC, PE, Cy5-PE, or APC were used for immunofluorescence analyses: CD4, CD8, CD43, CD45R (B220), BP-1 (Ly51), CD24 (heat stable Ag (HSA)), IgD, and IgM from BD Pharmingen (San Diego, CA), eBioscience (San Diego, CA), or Biolegend (San Diego, CA). Analyses were performed on a FACSCalibur flow cytometer using CellQuest software (BD Biosciences, San Jose, CA). Apoptotic cells were determined by annexin V and 7-aminoactinomycin D staining using an Annexin V-PE kit (BD Pharmingen).

Cell proliferation assays

For CFSE-labeled cell proliferation, single cell suspensions from the spleen of Rag-2^–/–/ROR^–/– and Rag-2^–/–/ROR^+/– chimeric mice were mixed with CFSE (Molecular Probes, Eugene, OR), incubated for 5 min at room temperature, and washed three times in 5% FCS in PBS. CFSE-labeled cells were resuspended in PBS and 300 µl were injected into sublethally irradiated C57BL/6 mice i.v. (40 x 10⁶ cells per mouse). At different time points, the spleen and LN were harvested and analyzed by FACS for in vivo homeostatic proliferation. Alternatively, the cells were resuspended at 2 x 10⁶ /ml in complete medium, stimulated with anti-CD3 (2C11) or LPS (Sigma-Aldrich, St. Louis, MO) in 6-well plates (Corning, Corning, NY) for indicated times, and analyzed by FACS. The [³H]thymidine incorporation assay was performed as described (23).

Immunization and Ab titration

Rag-2^–/–/ROR^–/– and Rag-2^–/–/ROR^+/+ BM chimeric mice were immunized i.p. with 100 µg (0.2 ml per mouse) DNP-keyhole limpet hemocyanin (KLH; Calbiochem, La Jolla, CA) mixed 1/1 with complete Freund’s adjuvant. Sera were collected at days 0, 7, 14, 21, 28, and 35. Secondary challenge with the same dose of DNP-KLH in incomplete Freund’s adjuvant was performed at day 28. Isotype-specific anti-DNP Abs were determined by ELISA as described (24). ELISA plates (Corning) were coated with 500 µg/ml DNP-albumin (Calbiochem) overnight, washed, and blocked with 3% BSA (Sigma-Aldrich) in PBS for 1 h at 37°C. One hundred microliters of serial dilution of sera were incubated on plates and washed. Secondary alkaline phosphatase-conjugated Abs to mouse IgG1, IgM, IgG2a, IgG2b, IgG3, and IgA (Southern Biotechnology Associates, Birmingham, AL) diluted in 1% BSA in PBS were added for 1 h at room temperature. The reactions were detected with 100 µl per well of p-nitrophenylphosphate substrate (Southern Biotechnology Associates) in diethanolamine buffer. The titer was expressed as relative units as compared with a "master" sample.

Quantitative RT-PCR analysis

Lymphocyte populations from the thymus and spleen of C57BL/6 mice were purified by fluorescence-activated cell sorting (>99% pure), and total RNA from 10⁶ cells was extracted with RNeasy Mini kit (Qiagen, Valencia, CA). BM-derived macrophages were produced as described (25) and stimulated with 100 ng/ml LPS (Sigma-Aldrich) for 4 h. First-strand cDNA was reverse transcribed with iScript Reverse Transcriptase kit (Bio-Rad, Hercules, CA). Quantitative RT-PCR was performed in triplicates on LightCycler (Roche, Indianapolis, IN) with the following primers: ROR forward: 5'-GTC AGC AGC TTC TAC CTG GAC ATC and ROR reverse: 5'-GTG TTG TTC TGA GAG TCA AAG GCA CG for all isoforms of ROR, and -actin forward: 5'-CAG CTT CTT TGC AGC TCC TT and -actin reverse: 5'-TCA CCC ACA TAG GAG TCC TT. The expression level of ROR was calculated and normalized to -actin using Relative Expression Software Tool software kindly provided by Dr. M. Pfaffl (Technical University of Munich, Munich, Germany).

Cytokine assays

Splenocytes from Rag-2^–/–/ROR^–/– and Rag-2^–/–/ROR^+/+ BM chimeric mice were stimulated with 5 µg/ml anti-CD3 (2C11) in RPMI 1640 complete medium for 48 h. The supernatant was assayed for IL-6 using an ELISA kit (eBioscience). To determine IL-2, TNF-, and IFN- production, the anti-CD3-activated cells were cultured with 50 U/ml human IL-2 (BioLegend) for 72 more hours. The cells were washed and incubated for 6 h with plate-bound CD3 Ab (5 µg/ml) to mimic Ag-induced cell death. Aliquots of the supernatants were collected and stored at –80°C until assayed. After that, the cells were first stained for surface CD8 or CD4 and then fixed with 2% paraformaldehyde in PBS for 30 min at 4°C. The cells were permeabilized with 0.5% saponin (Sigma-Aldrich) and stained with anti-IFN- FITC and PE anti-IL-4 (BD Pharmingen) for 30 min at 4°C, followed by flow cytometry analysis. TNF- production was assayed with an ELISA kit (eBioscience). IL-2 production was determined following a standard ELISA protocol using 4 µg/ml anti-IL-2 capture Ab (BioLegend), 1 µg/ml biotin anti-IL-2 detection Ab (BioLegend), and 1/250 dilution of streptavidin-HRP (eBioscience). The reaction was developed with TMB Peroxidase EIA Substrate kit (Bio-Rad). Cytokine production from ROR^–/– BM-derived mast cells and macrophages was performed as described (25). For intracellular IFN- staining in OVA-specific CD8⁺ T cells, 4 x 10⁶ splenocytes in 24-well plates were first stimulated with media alone or 10^–8 M of OVA peptide (American Peptide, Sunnyvale, CA) for 5 h in complete RPMI 1640 medium in the presence of GolgiStop (BD Pharmingen).

Bacteria and infections

The recombinant Listeria monocytogenes strain, which secrets chicken OVA (rLmOVA) and contains an erythromycin-resistance marker, was provided by M. Bevan (University of Washington, Seattle, WA; Refs. 26 and 27). Frozen stocks of the rLmOVA was grown in brain-heart infusion broth supplemented with 5 µg/ml erythromycin. Bacterial culture samples were grown to mid-log phase, measured by OD₆₀₀, aliquoted, and frozen at –80°C. Bacteria were diluted in PBS for injection. All mice were infected i.v. with priming doses equivalent to 10,000 rLmOVA as determined by spreading bacterial samples on brain-heart infusion agar plates.

⁵¹Cr-release assays

To determine the ability of splenocytes to lyse OVA peptide-loaded target cells, 1/2 serial dilutions of splenocytes were prepared in triplicate in 96-well round-bottom plates. EL-4 target cells were labeled with ⁵¹Cr and cultured with or without 10^–7 M SIINFEKL peptide for 1 h at 37°C. Target cells were then washed and added to effector cells at 10,000 cells per well. After a 6-h incubation at 37°C, the plates were centrifuged at 1500 rpm for 5 min and 100 µl of the supernatants were collected. The samples were counted on 1272 CLINIGAMMA (PerkinElmer Wallac, Turku, Finland) to determine the amount of ⁵¹Cr release. The percentage of specific lysis was calculated as 100 x (experimental cpm – spontaneous cpm)/(maximum cpm – spontaneous cpm).

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
ROR expression in lymphocytes

Although ROR was detected in a variety of tissues in Northern blot analysis (6, 7, 28), its expression pattern in lymphocytes is not defined. To examine ROR expression in lymphoid cells, we designed primers that can amplify all four isoforms of ROR. We performed real-time RT-PCR analysis on FACS-sorted thymocyte subsets and mature CD4⁺, CD8⁺ T, and B220⁺ B lymphocytes from the spleen of adult C57BL/6 mice. ROR is expressed at a low level in CD4⁺CD8⁺ double-positive (DP) thymocytes and high levels in CD4⁺ single-positive (SP) and CD8⁺ SP thymocytes (Fig. 1A). Interestingly, ROR mRNA is up-regulated in CD4⁺, but down-regulated in CD8⁺ mature T cells (Fig. 1A). TCR-mediated activation of CD4⁺ and CD8⁺ T lymphocytes did not obviously modulate ROR expression (Fig. 1A). In addition, ROR expression in B220⁺ B cells was detected at a very low level (Fig. 1A). We also examined ROR expression in resting and LPS-activated macrophages. ROR is expressed at a 5- to 6-fold higher level in resting macrophages than activated macrophages, suggesting that LPS-mediated TLR signaling down-regulates ROR expression (Fig. 1B). These results demonstrate that ROR is expressed in lymphocytes and macrophages in a developmentally regulated fashion.

View larger version (10K): [in this window] [in a new window]   FIGURE 1. Quantitative RT-PCR analysis of ROR mRNA expression in lymphocytes and macrophages. T cell subsets and B cells from C57BL/6 mice were purified by FACS sorting and extracted for total RNA. Real-time RT-PCR was performed as described in Materials and Methods. ROR expression was detected with primers that amplify all four ROR isoforms and expressed as a relative mRNA level normalized for -actin expression. A, ROR expression in lymphocytes. *, ROR expression value is 0.01 in B cells. B, ROR expression in BM-derived macrophages with or without LPS stimulation (100 ng/ml).

Lymphocyte development in ROR^–/– mice

Next, we examined T and B lymphocyte development in ROR^–/– mice. ROR^–/– mice exhibit tremor and imbalance and are visibly smaller than littermate controls as previously reported (16, 17). The mutant mice die within 3–4 wk after birth. As shown in Fig. 2A, the spleen and thymus from ROR^–/– mice were dramatically smaller than littermate controls. The total cell numbers of thymocytes and splenocytes in ROR^–/– mice was on average 30–40% of those in littermate controls (Fig. 2B). These results are similar to those reported on staggerer mice (18), suggesting that a similar defect in these two types of mice caused the reduced size of the thymus and spleen. We then performed FACS analysis to determine the phenotypes of lymphocytes in the thymus, spleen, and BM of ROR^–/– mice. ROR^–/– mice had a nearly complete loss of the DP thymocyte compartment and a corresponding increase of the percentages of DN, CD4⁺, and CD8⁺ SP thymocytes (Fig. 2C). Furthermore, the percentage of immature pre-B cells (CD43⁺B220⁺BP-1^–HSA⁺) in the BM of ROR^–/– mice was reduced from 72 to 47% (Fig. 2C). In contrast, the fraction representing more mature B cells (CD43^–B220⁺IgD⁺IgM⁺) was dramatically increased (5–63%; Fig. 2C). The percentages of CD4⁺ and CD8⁺ T cells in the spleen of ROR^–/– mice appear to be higher than those in control spleens (Fig. 2C); however, the total number of mature T cells in the ROR^–/– spleen was lower than that in controls (8.4 x 10⁶ in knockout mice, 24.7 x 10⁶ in controls). The percentage of IgM⁺B220⁺ mature B cells in the spleen of ROR^–/– mice was reduced from 67 to 43% when compared with controls (Fig. 2C). Taken together, these results demonstrate that T and B lymphocyte development was severely disrupted in ROR^–/– mice and suggest that ROR may play a critical role in lymphocyte development.

View larger version (47K): [in this window] [in a new window]   FIGURE 2. Analysis of the lymphocyte compartment in ROR–/– mice. A, Thymus and spleen from 2-wk-old ROR–/– (–/–) and littermate control mouse (+/+). B, Total cell number of thymus and spleen of 2- to 3-wk-old ROR–/– (–/–) and littermate controls (+/+ or +/–). C, FACS profiles of thymus, spleen, and BM from 2- to 3-wk-old ROR–/– (–/–) and littermate controls (+/+). CD43+B220+ BM cells were gated for BP-1 and HSA expression, and CD43–B220+ BM cells were gated for IgM and IgD expression.

Lymphocyte development in Rag-2^–/– mice reconstituted with ROR^–/– BM

To determine whether T and B cells in ROR^–/– mice have intrinsic defects, we generated BM chimeras using Rag-2^–/–CD45.1⁺ congenic mice as hosts and neonatal ROR^–/– or littermate control mice as BM donors. Four to 6 wk after the BM transfer, we analyzed T and B lymphocyte development in the chimeric mice. Surprisingly, the total cell numbers in the thymus and spleen of Rag-2^–/– mice that received ROR^–/– BM were comparable to those recovered from Rag-2^–/– hosts transferred with littermate control BM (Fig. 3A). Furthermore, FACS analysis of the T and B cell compartment in the thymus, BM, and spleen of Rag-2^–/–/ROR^–/– chimeric mice revealed a completely normal development of these cells when compared with Rag-2^–/–/ROR^+/+ control chimeric mice (Fig, 3B). These results demonstrate that T and B cells develop normally in the absence of ROR and suggest that the altered pattern of the T and B compartment in ROR^–/– mice is not due to intrinsic defects in the cells.

View larger version (44K): [in this window] [in a new window]   FIGURE 3. Lymphocyte development in Rag-2–/–/ROR–/– BM chimeric mice. A, Total cell number of thymus and spleen of Rag-2–/–/ROR–/– and Rag-2–/–/ROR+/+ BM chimeras. Mice were analyzed 6 wk after BM transfer. B, FACS profiles of thymus, spleen, and BM of the above chimeric mice. CD43+B220+ BM cells were gated for BP-1 and HSA expression and CD43–B220+ BM cells were gated for IgM and IgD expression.

Lymphocyte proliferation in the absence of ROR

Given that ROR is expressed in resting T and B cells (Fig. 1), we examined the proliferation of ROR^–/– T and B cells after Ag-receptor stimulation. Total splenocytes from Rag-2^–/–/ROR^–/– or Rag-2^–/–/ROR^+/+ chimeric mice were stimulated either with anti-CD3 or LPS for 2 days and the proliferation of T and B cells was measured with [³H]thymidine incorporation. Anti-CD3- and LPS-induced proliferation of ROR^–/– splenocytes was comparable to that of ROR^+/+ splenocytes (Fig. 4A). To further determine the proliferative capability of CD4⁺ and CD8⁺ T cells and B220⁺ B cells, we labeled the splenocytes from Rag-2^–/–/ROR^–/– or Rag-2^–/–/ROR^+/+ chimeras with CFSE, stimulated with anti-CD3 and LPS in vitro, and determined their proliferation by FACS analysis. As shown in Fig. 4B, no significant difference was observed in the proliferation of CD4⁺ and CD8⁺ T cells or B220⁺ B cells between ROR^–/– and ROR^+/+ groups. These results indicate that ROR is not essential for the proliferation of T and B lymphocytes in vitro.

View larger version (21K): [in this window] [in a new window]   FIGURE 4. Proliferation of ROR–/– lymphocytes. A, In vitro proliferation of T and B lymphocytes. Splenocytes from Rag-2–/–/ROR–/– and Rag-2–/–/ROR+/+ BM chimeras were stimulated with anti-CD3 or LPS and measured for proliferation by their [3H]thymidine incorporation. B, In vitro proliferation of CD4+, CD8+ T cells, and B220+ B cells. Splenocytes from the above BM chimeric mice were labeled with CFSE and stimulated with anti-CD3 (5 µg/ml) and LPS (30 µg/ml). The proliferation of CD8+ T cells was determined by FACS after 3 day in culture and the proliferation of CD4+ T cells and B cells was determined after 4 day in culture. C, Homeostatic-driven proliferation of ROR–/– T cells. Splenocytes from the BM chimeras were labeled with CFSE and injected into sublethally irradiated C57BL/6 hosts. CD4+ and CD8+ homeostatic proliferation was measured at day 17 and day 6, respectively.

Increased IgG Ab production in Rag-2^–/–/ROR^–/– chimeric mice

The observation that ROR negatively regulates inflammatory cytokine production in macrophages and in a muscle cell line (19, 20, 21) suggests that it may regulate immune responses. The early death and abnormal lymphocyte compartment in ROR^–/– mice prevented a direct test of Ab production in these animals. To circumvent these problems, we used Rag-2^–/–/ROR^–/– and Rag-2^–/–/ROR^+/+ chimeras to examine the role of ROR in Ab production. Six weeks after BM transfer, we immunized the chimeras with a T cell-dependent Ag, DNP-KLH, and boosted the immune response 4 wk later. We collected mouse sera at different time points and measured Ag-specific Abs in the mice by ELISA. Rag-2^–/–/ROR^–/– chimeric mice produced similar levels of anti-DNP IgM and IgA to those detected in Rag-2^–/–/ROR^+/+ control mice. However, Rag-2^–/–/ROR^–/– mice consistently produced significantly higher levels of anti-DNP IgG Abs than Rag-2^–/–/ROR^+/+ control mice (Fig. 5), demonstrating that the lack of ROR resulted in an increased Ab response in vivo.

View larger version (27K): [in this window] [in a new window]   FIGURE 5. Ab production in Rag-2–/–/ROR–/– BM chimeric mice immunized with DNP-KLH. Rag-2–/–/ROR–/– and Rag-2–/–/ROR+/+ mice (five per group) were immunized with DNP-KLH in CFA by i.p. injection. Serum Ag-specific Abs were determined by ELISA. The relative units were determined by comparing the titer from each mouse to a master sample. Shown are mean plus SD.

Cytokine production by ROR^–/– T cells

It was reported that macrophages from staggerer mice exhibit a hyperactive state and produce more IL-1 upon LPS stimulation (19, 20). Given the expression of ROR in T lymphocytes, we examined cytokine production by ROR^–/– T cells. Splenic T cells from Rag-2^–/–/ROR^+/+ and Rag-2^–/–/ROR^–/– mice were stimulated with anti-CD3 and tested for cytokine production. T lymphocytes from both types of mice produced a similar amount of IL-2, TNF-, and IL-6 as determined by ELISA (Fig. 6A) and intracellular cytokine staining (not shown). However, more ROR^–/– CD8⁺ T cells produced IFN- than control CD8⁺ T cells (6 vs 14%; Fig. 6B). Furthermore, the ROR^–/–IFN-⁺CD8⁺ T cells had a higher mean fluorescent intensity than the control CD8⁺ T cells (mean fluorescent intensity: 61 vs 46), suggesting an increase in both the frequency of cells producing IFN- and the amount of IFN- produced per cell. The increased production of IFN- was not seen in ROR^–/– CD4⁺ T cells (data not shown).

View larger version (31K): [in this window] [in a new window]   FIGURE 6. Cytokine production and apoptosis in ROR–/– T cells. The production of cytokines was determined following 2-day stimulation of splenocytes from BM chimeras with anti-CD3 Ab (5 µg/ml) followed by a 3-day culture with human IL-2 (50 U/ml). The cells were incubated with plate-bound anti-CD3 for 6 more hours to induce activation-induced cell death. A, The amount of IL-2, TNF-, and IL-6 production after anti-CD3 stimulation as determined by ELISA. B, Intracellular cytokine staining of IFN- and IL-4 in anti-CD3-activated CD8+ cells. Anti-CD3-activated CD8+ T cells were analyzed by three-color staining. Cells gated on CD8+ population are shown. C, TNF- and IL-6 production by ROR–/– mast cells after LPS stimulation. Mast cells generated by culturing BM cells from ROR–/– and ROR+/+ mice were stimulated with LPS for 6 h. IL-6 and TNF- production was measured with ELISA. D, T lymphocytes with or without anti-CD3 stimulation were analyzed for their apoptosis rate by annexin V staining.

Given the role of ROR in regulating IB production and the role of NF-B in T cell apoptosis (30, 31), we examined ROR^–/– T cell apoptosis under different conditions. As shown in Fig. 6D, resting and anti-CD3-activated T cells from Rag-2^–/–/ROR^–/– hosts exhibited similar apoptotic rates to those from control chimeras. Furthermore, activation-induced T cell death in both groups was comparable (Fig. 6D), indicating that ROR is not critically involved in T cell apoptosis.

ROR^–/–CD8⁺ T cell response to L. monocytogenes infection

Given the expression of ROR in mature CD8⁺ T cells, we tested the capability of ROR^–/–CD8⁺ T cell responding to L. monocytogenes infection. We infected Rag-2^–/– /ROR^–/– and Rag-2^–/–/ROR^+/+ chimeras with 1 x 10⁴ L. monocytogenes expressing OVA. Seven days later, the primary CD8⁺ T cells response was examined by intracellular staining of IFN-. This dose of L. monocytogenes induced an average of 3.0% IFN-⁺CD8⁺ T cells in C57BL/6 hosts (not shown), but a lower CD8⁺ response (on average 0.7%) in Rag-2^–/–/ROR^+/+ hosts for unknown reasons (Fig. 7A). However, L. monocytogenes infection induced a 3-fold increase of IFN-⁺CD8⁺ T cells in Rag-2^–/–/ROR^–/– hosts (Fig. 7A). We further tested the cytotoxicity of CD8⁺ T cells in L. monocytogenes infected chimeric mice. Splenocytes from Rag-2^–/–/ROR^–/– chimeras exhibited a dramatically elevated cytotoxic activity against OVA peptide-pulsed target cells (Fig. 7B). These results demonstrate that ROR regulates CD8⁺ effector formation in vivo presumably by controlling the IFN- production in Ag-specific CD8⁺ T cells.

View larger version (29K): [in this window] [in a new window]   FIGURE 7. ROR–/–CD8+ T cell response to L. monocytogenes infection. Rag-2–/–/ROR–/– and Rag-2–/–/ROR+/+ BM chimeric mice (five mice per group) were infected i.v. with 1 x 104 L. monocytogenes expressing OVA. The CD8+ T cell response was evaluated 7 days after the infection. A, IFN- producing OVA257–264-specific CD8+ T cells determined by intracellular staining following a 5-h incubation with or without the peptide. B, Ex vivo cytotoxicity as measured by 51Cr release assay. The target cells were EL-4 cells pulsed with or without OVA257–264 peptide. Data are representative of three independent infections.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

Our findings that ROR^–/– mice also have a dramatically reduced size of the thymus and spleen are consistent with those seen in staggerer mice, suggesting that ROR mutation has caused the reduced lymphoid cellularity in staggerer mice. However, the defective T and B cell development in ROR^–/– mice is not intrinsic to lymphocytes because ROR^–/– BM develops into these lymphoid lineages normally in Rag-2^–/– hosts. We consider that a likely explanation for the reduced thymus and spleen in ROR^–/– mice is due to chronic stress. The selective loss of DP thymocytes and pro-B cells, both of which are extremely sensitive to glucocorticoid-induced cell death, in ROR^–/– mice suggests that these mice may produce more glucocorticoids due to imbalance of their bodies. However, we cannot rule out that defective neuronal development in ROR^–/– mice may result in an increased production of glucocorticoid or other factors that may induce the selective loss of lymphoid cells. Nevertheless, we conclude that ROR indirectly regulates T and B lymphocyte development by providing an appropriate microenvironment.

The elevated levels of Ag-specific IgG in Rag-2^–/–/ROR^–/– BM chimeras demonstrate that ROR negatively regulates the humoral immune response. Two explanations could account for the increased Ab production to a T-dependent Ag in the mutant chimeras, which are increased activation and expansion of CD4⁺ T and B cells and/or increased cytokine production by immune cells. Three lines of evidence suggest that the increased Ab production in the Rag2^–/–/ROR^–/– BM chimeras is due to an increased cytokine production but not increased activation and expansion of CD4⁺ T and B220⁺ B cells. First, both mast cells and macrophages from ROR^–/– mice produced dramatically increased TNF- and IL-6 upon LPS stimulation. Second, the activation and proliferation of ROR^–/– CD4⁺ T and B220⁺ B cells are comparable to controls. Third, the fact that Ag-specific IgM and IgA were not increased in the immunized mutant chimeras suggests that the Ab production in the mutant chimeras is specifically influenced by cytokines toward the IgG isotypes. Our analysis revealed that although 100% of lymphocytes were derived from donor BM in the chimeras, only 40–60% of myeloid cells were from the donor origin. The low percentages of myeloid cells from the donor origin are likely due to the low irradiation dose we used in the experiments. As such, one would predict an even greater increase of Ag-specific Ab production in the mutant chimeras with a 100% myeloid cell reconstitution by ROR^–/– BM.

Our results suggest that ROR acts as a negative regulator of IFN-, but not IL-6 and TNF- expression in CD8⁺ T lymphocytes. A higher fraction of ROR^–/– CD8⁺ T cells produced IFN- than control CD8⁺ T cells after both anti-CD3 stimulation in vitro and antigenic stimulation in vivo. The increased expression of TNF- and IL-6 in ROR^–/– mast cells and macrophages, but not in T cells, suggest that ROR differentially affects these pathways in different types of cells. Transcriptional regulation of IFN- expression in CD4⁺ T lymphocytes has been extensively investigated (32). Multiple transcription factors including NF-B, Stat4, and T-bet have been shown to be involved in positive regulation of IFN- expression. Recent reports have identified additional transcription factors in regulating IFN- expression in CD4⁺ and CD8⁺ cells. These include the NF-B repressor, Foxj1, and the T-box factor, Eomesodermin (33, 34). The molecular mechanisms by which ROR regulates IFN- production is currently under investigation. However, based on previous results, we speculate that ROR negatively regulates IFN- production by activating the transcription of IB, which in turn inhibits the activation of NF-B. Alternatively, ROR may interact with other transcription factors that are involved in IFN- production.

What is the physiological role of ROR in a normal immune system? The fact that ROR is highly expressed in resting macrophages but down-regulated upon LPS stimulation suggests that ROR may function to prevent resting macrophages from producing inflammatory cytokines. In contrast, ROR expression is not down-regulated in both activated CD4⁺ and CD8⁺ T lymphocytes, indicating a continued requirement for the presence of this orphan receptor to regulate their function. The absence of ROR results in increased CD8⁺ T response to a model pathogen. Thus, one function of ROR in T lymphocytes may be to down-size activated Ag-specific CD8⁺ T cell response.

	Acknowledgments

 
We thank Dr. Mike Cook in the Flow Cytometry Facility of Duke University Medical Center for FACS analysis, and Dr. Dave Draper for critical review of this manuscript.

	Footnotes

 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ This work was supported by an American Cancer Society Grant RSG-0125201 and National Institutes of Health Grant CA92123.

² Address correspondence and reprint requests to Dr. You-Wen He, Department of Immunology, Duke University Medical Center, Durham, NC 27710. E-mail address: he000004{at}mc.duke.edu

³ Abbreviations used in this paper: ROR, retinoid acid-related orphan receptor; BM, bone marrow; HSA, heat stable Ag; rLmOVA, recombinant Listeria monocytogenes strain which secrets chicken OVA; KLH, keyhole limpet hemocyanin; DP, double-positive; SP, single-positive.

Received for publication March 9, 2004. Accepted for publication June 17, 2004.

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