RORs and development Critical role of RORα in cerebellar development Although RORα is expressed in a variety of tissues, including testis, kidney, adipose and liver, it is most highly expressed in the brain, particularly in the cerebellum and thalamus [ Becker-Andre et al., 1993; Carlberg et al., 1994; Dussault et al., 1998; Hamilton et al., 1996; Matysiak-Scholze and Nehls, 1997; Nakagawa et al., 1997; Nakagawa et al., 1998; Nakagawa and O'Leary, 2003; Vogel et al., 2000]. The function of RORα in brain development has not been extensively studied beyond the cerebellum. A high level of RORα expression was observed in the cerebellar Purkinje cells, while RORα is not detected in the granule cell layer [ Ino, 2004; Matsui et al., 1995; Nakagawa et al., 1997; Sashihara et al., 1996; Sotelo and Wassef, 1991]. During Purkinje cell development, RORα expression is first observed at E12.5, just after Purkinje cell precursors exit the mitotic cycle and leave the ventricular zone to migrate along radial glia to form a temporary cerebellar plate-like structure [ Gold et al., 2003; Gold et al., 2007; Goldowitz and Hamre, 1998; Jetten and Joo, 2006]. RORα null mice, generated by targeted disruption of the RORα gene, display essentially an identical phenotype as homozygous staggerer mice (RORα sg/sg) [ Dussault et al., 1998; Hamilton et al., 1996; Herrup and Mullen, 1981; Landis and Sidman, 1978; Sidman et al., 1962; Steinmayr et al., 1998]. This natural mutant strain carries a 6.4 kb intragenic insertion causing deletion of the fifth exon, encoding the beginning of the LBD, and a frameshift that results in a premature stop codon [ Gold et al., 2007; Hamilton et al., 1996]. RORα-deficient mice exhibit several abnormalities, including thin long bones, suggesting a role for RORα in bone formation [ Meyer et al., 2000]. In addition, these mice display ataxia and severe cerebellar atrophy, characterized by significantly fewer Purkinje cells and a loss of cerebellar granule cells [ Doulazmi et al., 2001; Doulazmi et al., 1999; Dussault et al., 1998; Gold et al., 2007; Hamilton et al., 1996; Herrup and Mullen, 1979; Herrup and Mullen, 1981; Landis and Reese, 1977; Landis and Sidman, 1978; Sidman et al., 1962; Steinmayr et al., 1998]. Although the morphology of the cerebellar cortex in young heterozygous (RORα sg/+) mice appears normal, upon aging, an accelerated loss of Purkinje and granule cells was observed [ Doulazmi et al., 1999; Hadj-Sahraoui et al., 2001; Hadj-Sahraoui et al., 1997; Zanjani et al., 1992]. At E17.5 of development, the number of Purkinje cells generated appears to be similar between WT and RORα sg/sg mice [ Doulazmi et al., 2001; Vogel et al., 2000]. However, by PND5, the number of Purkinje cells is dramatically reduced and cells appear disorganized. Surviving RORα sg/sg Purkinje cells contain stunted dendritic arbors lacking distal spiny branchlets and are deficient in their assembly of mature synapses with granule cells, suggesting that dendritogenesis is impaired. This is accompanied by a lack of parallel fiber (PF) input and a persistence of multiple climbing fiber (CF) innervations [ Boukhtouche et al., 2006a; Janmaat et al., 2009; Mariani, 1982; Mariani and Changeux, 1980]. Dysfunctional synapse formation between PF and Purkinje cells is associated with a delayed shift from the glutamate transporter Slc17a6 (VGluT2) into Slc17a7 (VGluT1) in PF terminals and to an altered distribution of the glutamate receptors, Grm1 and Grm2, in Purkinje cells [ Janmaat et al., 2009]. Expression of RORα in RORα sg/sg Purkinje cells was shown to restore normal dendritogenesis in organotypic cultures [ Boukhtouche et al., 2006a]. These observations indicated that RORα does not regulate the genesis of Purkinje cells, but rather their maturation, particularly dendritic differentiation ( Figure 3). This concept was further supported by observations showing that several genes down-regulated in Purkinje cells of RORα sg/sg mice encode proteins involved in the maturation of Purkinje cells. These include genes involved in signal-dependent calcium release, such as the calmodulin inhibitor Pcp4, the IP3 receptor ( Itpr1) and its interacting partner Cals1, the calcium-transporting ATPase Atp2a2, and major intracellular calcium buffer 1 ( Calb1) [ Gold et al., 2003; Gold et al., 2007; Hamilton et al., 1996; Messer et al., 1990]. In addition, several genes linked to the glutamatergic pathway were expressed at reduced levels in RORα sg/sg mice, including the glutamate receptor Grm1, the glutamate transporter Slc1a6, and its anchor Spnb3, encoding a brain-specific β-spectrin. ![Figure 3 Figure 3](picrender.fcgi?artid=2670432&blobname=nrs07003.f3.gif) | Figure 3 Regulation of Purkinje cell maturation and cerebellar development by RORα. |
The promoter regulatory regions of several genes down-regulated in RORα sg/sg Purkinje cells, including those of Pcp2, Pcp4, Itpr1, Shh, and Slc1a6, contain ROREs that bind RORα [ Gold et al., 2003; Gold et al., 2007; Serinagaoglu et al., 2007]. Chromatin immunoprecipitation (ChIP) analysis demonstrated that RORα is associated with these ROREs in vivo, suggesting that these genes are directly regulated by RORα. Moreover, these studies revealed the identity of several transcriptional mediators that are part of RORα transcription activation complexes at these ROREs. Interestingly, the composition of these RORα:coactivator complexes associated with different promoters exhibited a certain promoter specificity. For example, the RORα transactivation complex associated with the Pcp2-RORE contained TIP60 (Tat-interactive protein, 60 kD), and SRC-1; the Slc1a6-RORE bound complex contained CBP, TIP60, SRC-1, and GRIP-1; while the Pcp4-RORE associated complex contained β-catenin, SRC-1, p300, and TIP60 [ Gold et al., 2003]. These results suggest that the sequence and the promoter context of the RORE play a critical role in determining the composition of the RORα:coactivator complex recruited to each promoter. The development of Purkinje cells and granule cells is mutually dependent. Positioning and morphological differentiation of Purkinje cells depends on granule cells, while proliferation of granule precursors requires factors released by Purkinje cells [ Goldowitz and Hamre, 1998; Vogel et al., 2000]. At birth, the number of granule cells is somewhat reduced, and this becomes more prominent during the first weeks after birth. In adult RORα-deficient mice, the granular layer is very thin and depleted of granule cells [ Doulazmi et al., 2001; Doulazmi et al., 1999; Dussault et al., 1998; Gold et al., 2007; Hamilton et al., 1996; Herrup and Mullen, 1979; Herrup and Mullen, 1981; Landis and Reese, 1977; Landis and Sidman, 1978; Sidman et al., 1962; Steinmayr et al., 1998; Yoon, 1972]. Studies using WT:staggerer chimeras indicated that the depletion of cerebellar granule cells in RORα sg/sg mice is related to abnormalities inherent to RORα sg/sg Purkinje cells [ Herrup and Mullen, 1979; Herrup and Mullen, 1981]. This is consistent with the high level of expression of RORα in Purkinje cells and the lack of RORα in cerebellar granule cells. Sonic hedgehog (Shh) released by Purkinje cells plays a critical role in the regulation of the proliferation of cerebellar granule cells [ Dahmane and Ruiz i Altaba, 1999; Gold et al., 2003; Kenney et al., 2003; Wallace, 1999] ( Figure 3). Interaction of Shh with Patched (Ptch) receptors on cerebellar granule precursors results in the activation of GLI Krüppel-like zinc finger proteins and the subsequent transcriptional activation of N-MYC and other cell cycle regulatory genes. The expression of Shh in RORα sg/sg Purkinje cells was found to be reduced several fold, suggesting that the repression of Shh may be a critical factor in the diminished proliferation of granule precursors [ Gold et al., 2003; Gold et al., 2007]. This is consistent with the observed repression of several Shh target genes, including N-MYC, in the cerebellum of RORα sg/sg mice. This hypothesis is further supported by experiments showing that Shh was able to enhance proliferation of granule precursor cells in sections of cerebellum from PND4 RORα sg/sg mice and partially prevented the reduction in granule cells [ Boukhtouche et al., 2006a]. These observations indicate that the down-regulation of Shh in RORα sg/sg mice is an important contributing factor in the development of cerebellar hypoplasia in these mice and that RORα functions as a positive regulator of Shh expression. ChIP analysis indicated that RORα regulates Shh expression directly through ROREs in the promoter regulatory region of the Shh gene. Moreover, these studies identified β-catenin and p300 as constituents of the RORα transactivation complex associated with the RORE in Shh. The association of β-catenin with RORα transactivation complexes suggests a link between Wnt and ROR signaling, and is consistent with a reported role for Wnt signaling in cerebellar development [ Salinas et al., 1994; Wang et al., 2001]. Link between RORα and spinocerebellar ataxia type 1 (SCA1) Comparison of gene expression profiles indicated a possible link between RORα and spinocerebellar ataxia type 1 (SCA1), an autosomal dominant inherited neurodegenerative disorder that is characterized by progressive loss of motor coordination, speech impairment, and problems swallowing [ Serra et al., 2006; Zoghbi and Orr, 2000]. SCA1 is caused by aberrant effects triggered by an expanded polyglutamine (polyQ) tract within the protein ataxin 1 (ATXN1), resulting in variable degrees of neurodegeneration, predominantly in the cerebellum (due to loss of Purkinje cells), brainstem, and spinal cord. Sca1[82Q] mutant mice, which express an Atxn1 protein containing an 82 glutamine repeat, develop severe ataxia and degeneration of Purkinje cells [ Burright et al., 1995]. A considerable overlap was observed between genes repressed in the cerebellum of Sca1[82Q] mice and genes found to be regulated by RORα in Purkinje cells, suggesting a possible connection between the two [ Gold et al., 2003; Serra et al., 2004]. Immunohistochemical analysis indicated that the level of RORα was significantly reduced in Purkinje cells from Sca1[82Q] mice [ Serra et al., 2006]. This reduction was found to be, at least in part, due to a decrease in RORα protein stability. In addition, it was demonstrated that RORα and ATXN1 are part of the same protein complex. This association was indirect and mediated by a direct interaction of ATXN1 with the coactivator TIP60, which has been show to be part of several RORα-coactivator complexes in Purkinje cells [ Gold et al., 2003; Serra et al., 2004]. It was concluded that decreased levels of RORα protein in SCA1 Purkinje cells causes reduced expression of RORα-target genes that are critical for Purkinje cell development and function, and consequently is an important contributory factor in the neurodegeneration observed in SCA1 [ Serra et al., 2006]. Physiological Function of RORβ in brain and retina RORβ exhibits a restricted pattern of expression that is limited to certain regions of the brain and retina [ Andre et al., 1998a; Andre et al., 1998b; Azadi et al., 2002; Nakagawa and O'Leary, 2003; Schaeren-Wiemers et al., 1997]. During the period between E12.5 and PND5 of mouse development, RORβ is expressed in the neocortex and dorsal thalamus in a highly dynamic, spatiotemporal manner [ Jetten and Joo, 2006; Nakagawa and O'Leary, 2003]. In adult brain, RORβ mRNA is most highly expressed in the cerebral cortex, thalamus, hypothalamus, suprachiasmatic nucleus (SCN), and the pineal gland, while little expression was detected in the cerebellum or hippocampus [ Andre et al., 1998a; Andre et al., 1998b; Schaeren-Wiemers et al., 1997; Sumi et al., 2002]. RORβ is also expressed in the anterior olfactory nucleus posterior and sensory cortices. These observations suggested that RORβ is highly restricted to several regions involved in the processing of sensory information. Study of RORβ null mice provided several clues about the physiological function of RORβ. RORβ null mice exhibit several motor defects, including a ‘duck-like’ gait and an impairment in several neurological reflexes, e.g., hind paw clasping reflex [ Andre et al., 1998a; Masana et al., 2007]. RORβ null mice display several behavioral changes, including increased exploratory activity and reduced anxiety behavior, and olfactory deficits [ Masana et al., 2007]. In addition, RORβ -/- mice show a delayed onset of male fertility [ Andre et al., 1998a]. The relatively high expression in the retina and the development of retinal degeneration in RORβ -/- mice suggested an important regulatory function for RORβ in this tissue [ Andre et al., 1998a]. At E17.5 of mouse and rat development, RORβ expression is detected in the formative ganglion cell layer and the neuroblastic layer [ Andre et al., 1998a; Schaeren-Wiemers et al., 1997; Srinivas et al., 2006]. By PND6, the neuroblastic layer has split into the inner nuclear layer of interneurons and the outer nuclear layer, where the photoreceptor cells, rods and cones, reside. At PND16 and older, RORβ is still expressed in all retinal layers, but most highly in the outermost part of the outer nuclear layer. RORβ -/- mice exhibit several retinal developmental defects. At PND6, cell loss was observed in the ganglion layer, while the interneurons in the inner layer appeared in disarray. Although an outer nuclear layer was formed, it was significantly thinner than in WT retina, but maintained for several weeks. Rods and cones photoreceptors in the outer layer appeared disorganized and lacked outer and inner segments. These observations suggested that RORβ plays a key role in the maturation of photoreceptors, particularly the formation of the outer segments, the light sensing structures of cones and rods that contain opsin and rhodopsin, respectively [ Srinivas et al., 2006]. In most mammals, color vision depends on the photopigments, S and M opsin, which are synthesized by distinct cone populations [ Ebrey and Koutalos, 2001]. In WT mice, S opsin expression is observed at PND6, and by PND13 is concentrated to the outer segments of maturing cones; however, in RORβ -/- mice, induction of S opsin ( Opn1sw) is significantly delayed. After PND23, S opsin is detected at greatly reduced levels. Little difference in the induction of M opsin ( Opn1mw) and thyroid hormone receptor TRβ2, which is essential for Opn1mw expression, was observed between WT and RORβ -/- mice. The transcription factor CRX (cone-rod homeobox factor), which is critical for both S and M opsin gene expression, is normally expressed in rods and cones [ Srinivas et al., 2006]. These data and histological studies suggested that RORβ is not a requirement for the initiation of differentiation of progenitors into photoreceptors, but specifically regulates their maturation, as illustrated by the reduced level of S opsin. Study of the Opn1sw promoter provided further evidence that RORβ is an important regulator of Opn1sw expression. The proximal promoter region of Opn1sw contains two putative ROREs that are able to bind RORβ. Optimal activation of this promoter region was found to require a synergy between RORβ and CRX [ Srinivas et al., 2006]. The latter is consistent with the co-expression of both transcription factors in the photoreceptor layer during the period when Opn1sw expression is induced. The activation of the Opn1sw promoter by RORβ was mediated principally through interaction with the most proximal RORE and, as expected, required both the DBD and AF2 domains of RORβ. Analysis of a transgene consisting of LacZ controlled by the Opn1sw proximal promoter region in mice showed that its expression was restricted to the photoreceptors with features of cones in the outer nuclear layer, while mutations in the ROREs totally abolished this expression. These observations indicate that RORβ is required for the induction of S opsin and support the conclusion that RORβ regulates Opn1sw transcription in a direct manner through ROREs within its proximal promoter region. In addition, they explain the greatly diminished expression of Opn1sw observed in the retina of RORβ -/- mice. Role for RORα in cone development Recent studies have indicated that RORα also plays a role in mouse retinal development [ Fujieda et al., 2009; Ino, 2004]. In the retina, RORα expression is highly dynamic. It is first observed at E17 in the developing ganglion layer, while it is not detectable in the neuroblastic layer. At PND3, RORα was also found in the outer and inner border of the neuroblastic layer. In retina of adult mice, RORα is intensely expressed in the ganglion cell layer, the inner nuclear layer, and in cone photoreceptors in the outer layer, as indicated by its co-localization with both S and M opsin. Analysis of Opn1sw and Opn1mw expression showed that their level of expression was significantly reduced in RORα sg/sg retina. Expression of cone arrestin ( Arr3) was also diminished, whereas the expression of several other cone genes, including phosphodiesterases 6c and 6h ( Pde6c and Pde6h) and cyclic nucleotide gated channel b3 ( Gngb3), and the expression of the rod-specific gene rhodopsin ( Rho) were unchanged. Expression of RORβ was also unaffected in RORα sg/sg retina. As demonstrated for RORβ, RORα was shown to act in synergy with CRX to induce Opn1sw expression [ Fujieda et al., 2009]. In vivo ChIP analysis supported the conclusion that Opn1sw, Opn1mw, and Arr3 are direct targets of RORα regulation. Thus, both RORα and RORβ function as regulators of cone development and Opn1sw; however, in contrast to RORα, RORβ does not affect Opn1mw expression. These data suggest that although RORα and RORβ exhibit some functional redundancy, their functions in cone development appear largely distinct from each other. Role of RORγ in the development of secondary lymphoid tissues RORγ1 and RORγt (RORγ2) exhibit distinct patterns of tissue-specific expression. RORγ1 is expressed in many tissues, including liver, adipose, skeletal muscle, and kidney, while the expression of RORγt is exclusively expressed in a few distinct cell types of the immune system [ Eberl and Littman, 2003; Eberl and Littman, 2004; Jetten, 2004; Jetten and Joo, 2006; Lipp and Muller, 2004]. Mice deficient in RORγ expression lack lymph nodes and Peyer's patches (PPs), suggesting that RORγ is indispensable for lymph node organogenesis and development of PPs [ Eberl and Littman, 2003; Eberl et al., 2004; He, 2002; Jetten and Joo, 2006; Jetten et al., 2001; Jetten and Ueda, 2002; Kurebayashi et al., 2000; Lipp and Muller, 2004; Littman et al., 1999; Sun et al., 2000]. Lymphoid tissue inducer (LTi) cells play a critical role in the development of lymph nodes and PPs. LTi cells originate from hematopoietic precursor cells in the fetal liver and have been recently characterized as CD45 intCD4 +CD3 -CD127(IL-7Rα) +Lin - cells in mice and in humans as lineage-negative CD45 intCD4 -CD3 -CD127 hiLin - cells [ Cupedo et al., 2009; Eberl and Littman, 2003; Mebius et al., 2001; Mebius et al., 1997]. LTi cells are recruited to the lymph node anlagen through their interaction with mesenchymal organizer cells, specialized cells derived from differentiating mesenchymal cells [ Cupedo et al., 2002; Cupedo and Mebius, 2005; Eberl, 2005; Eberl and Littman, 2003; Lipp and Muller, 2004] ( Figure 4). This cell:cell interaction involves several positive feedback loops and amplifying mechanisms. Lymph node organogenesis is initiated by interaction of lymphotoxin α1β2 at the surface of LTi cells with their respective receptors on mesenchymal organizer cells. This leads to the induction of several adhesion molecules, including vascular cell adhesion molecule 1 (VCAM-1) and mucosal addressin cellular adhesion molecule 1 (MadCAM-1) and the subsequent production of homeostatic chemokines by mesenchymal organizer cells [ Cupedo et al., 2002; Cupedo and Mebius, 2005; Eberl, 2005; Lipp and Muller, 2004; Muller et al., 2003; Yoshida et al., 2002] ( Figure 4). In addition to reinforcing the interaction between LTi and mesenchymal organizer cells, these interactions amplify the recruitment of additional LTi cells and promote the recruitment of other cells, such as monocytes and T and B lymphocytes, from the circulation [ Cupedo and Mebius, 2005; Muller et al., 2003]. Subsequent studies showed that LTi cells are absent from spleen, mesentery, and intestine of RORγ -/- E18.5 embryos, indicating that RORγt plays a critical role in the generation and/or survival of LTi cells [ Eberl, 2005; Eberl and Littman, 2003]. As a consequence, the lack of lymph nodes and PPs in RORγt- or RORγ-deficient mice is due to the absence of LTi cells. ![Figure 4 Figure 4](picrender.fcgi?artid=2670432&blobname=nrs07003.f4.gif) | Figure 4 RORγt is essential for the development of secondary lymphoid tissues. |
Besides PPs and mesenteric lymph nodes, the intestinal immune system contains several other lymphoid cell compartments, including cryptopatches and isolated lymphoid follicles (ILFs) [ Eberl, 2005; Taylor and Williams, 2005] ( Figure 4). ILFs develop from cryptopatches in response to inflammatory innate immune signals generated by the colonization of the intestine by bacteria. Lin -cKit +CD127 +CD44 + cells in cryptopatches and in ILFs express high levels of RORγt and appear to constitute the adult counterpart of LTi cells [ Eberl, 2005; Eberl and Littman, 2004]. The deficiency in cryptopatches and ILFs observed in RORγ -/- mice indicate that RORγt is essential for the development of these cell compartments and is due to the absence of these LTi-like cells in these mice. Recent studies identified a novel lymphocyte population in the intestinal lamina propria that produces IL-22 and co-expresses RORγt and CD127, and natural killer (NK) cell markers NKp46 and NK1.1 (CD3 -RORγt hiCD127 +NKp46 +NK1.1 int) [ Cupedo et al., 2009; Luci et al., 2009; Sanos et al., 2009] ( Figure 4). These cells are absent in RORγt-deficient mice. Because of its dependence on RORγt and its CD127 +c-Kit + phenotype, these NK cells resemble LTi cells, suggesting a possible link between the two cell types. LTi cells have been reported to be able to differentiate into NK and antigen presenting cells [ Mebius et al., 2001; Mebius et al., 1997]. The basic helix-loop-helix transcription factor Id2, which as RORγt is expressed in LTi cells, is essential for both LTi and NK cell development [ Yokota et al., 1999]. In contrast, peripheral NK cells do not express RORγt and their number is unaffected in RORγ-deficient mice [ Sun et al., 2000]. It is has been suggested that RORγt hiNKp46 +NK1.1 int cells do not belong to the NK cell lineage, but may be derived from the Lin -Kit +CD127 +RORγt hi LTi-like cells in cryptopatches [ Cupedo et al., 2009; Luci et al., 2009; Sanos et al., 2009]. The precise role of these cells in normal intestinal mucosa immunity and pathology, including colitis, needs further study. In contrast to lymph nodes and Peyer’s patches, the development of several other lymphoid tissues, including nasal- and bronchial-associated lymphoid tissue (BALT and NALT), was undisturbed in RORγ -/- mice, suggesting that the organogenesis of these lymphoid tissues involves a different mechanism [ Harmsen et al., 2002]. Critical functions of RORγ in thymopoiesis Several studies have demonstrated that RORγt plays a critical role in the regulation of thymopoiesis [ Guo et al., 2002; He, 2000; He et al., 1998; Hirose et al., 1994; Jetten, 2004; Jetten and Joo, 2006; Kurebayashi et al., 2000; Medvedev et al., 1996; Ortiz et al., 1995; Sun et al., 2000]. During thymopoiesis, T cell precursor CD25 -CD44 +CD4 -CD8 - cells (DN1) differentiate successively via two intermediate stages, CD25 +CD44 + (DN2) and CD25 +CD44 - (DN3), into CD44 -CD25 - (DN4) thymocytes ( Figure 5). These cells then differentiate via immature single positive (ISP) cells (CD3 -CD4 -CD8 low) into double positive CD4 +CD8 + (DP) thymocytes [ Miyazaki, 1997]. After successful T cell receptor α (TCRα) gene rearrangement, DP cells expressing TCRαβ receptor undergo a careful selection process to eliminate thymocytes expressing nonfunctional or autoreactive TCR [ Starr et al., 2003; Zhang et al., 2005]. The positive selected DP thymocytes mature into single positive (SP) CD4 +CD8 - helper and CD4 -CD8 + cytotoxic T cells that then colonize the secondary lymphoid organs, including the spleen, lymph nodes, and PPs. RORγt is transiently expressed during thymopoiesis [ Guo et al., 2002; He, 2002; He et al., 2000; He et al., 1998; Jetten, 2004; Jetten and Joo, 2006; Jetten et al., 2001; Sun et al., 2000] ( Figure 5). It is undetectable in DN thymocytes and highly induced when ISP cells differentiate into DP thymocytes and again down-regulated when DP thymocytes differentiate into mature T lymphocytes. At birth and during early stages of life, RORγ null mice have a significantly smaller thymus compared to wild type mice as a result of a drastic reduction in the number of double positive DP and SP thymocytes, while the percentage of ISP thymocytes is greatly increased [ Eberl and Littman, 2003; Jetten et al., 2001; Jetten and Ueda, 2002; Kurebayashi et al., 2000; Sun et al., 2000]. The accumulation of ISP cells in RORγ -/- mice appears to be due to a delay in the differentiation of ISP into DP cells, suggesting a role for RORγt in the regulation of the ISP-DP transition [ Guo et al., 2002]. In addition, DP thymocytes undergo massive apoptosis in vivo and in vitro [ Jetten, 2004; Jetten and Joo, 2006; Jetten et al., 2001; Kurebayashi et al., 2000; Sun et al., 2000]. The accelerated apoptosis of RORγ -/- DP thymocytes is related to reduced expression of the anti-apoptotic gene Bcl-XL. This repression is an early and key event in accelerated apoptosis in RORγ -/- thymocytes [ Sun et al., 2000; Xie et al., 2005]. Thus, RORγ functions as a positive regulator of Bcl-XL expression and, as a result, promotes the survival of DP thymocytes, thereby enabling gene rearrangement [ Eberl et al., 2004; He et al., 1998; Jetten, 2004; Jetten and Joo, 2006; Jetten et al., 2001; Jetten and Ueda, 2002; Kurebayashi et al., 2000; Ma et al., 1995]. Little is still understood about the mechanism by which Bcl-XL and RORγt expression are regulated during thymopoiesis. The reduced expression of Bcl-XL in RORγ -/- mice and the observation that the induction and down-regulation of Bcl-XL and RORγt expression occur in parallel, are consistent with the concept that RORγt regulates Bcl-XL expression. Whether this regulation involves a direct or indirect mechanism has yet to be established. A recent study provided evidence for a link between early growth response gene 3 ( EGR3) expression and the regulation of RORγt and Bcl-XL [ Xi and Kersh, 2004; Xi et al., 2006]. Pre-TCR signals transiently induce EGR3, which promotes proliferation, whereas the subsequent down-regulation of EGR3 leads to growth cessation and induction of RORγt and Bcl-XL needed for cell survival and proper gene rearrangement. Analysis of gene expression profiles of thymocytes from WT and RORγ -/- mice identified additional changes in gene expression, including repression of the transcription factor EGR1, Ngfi-A binding protein 2 ( NAB2), and the glucocorticoid-induced transcript 1 ( GLCCI1) [ Kang et al., 2006]. One of the genes most dramatically repressed in thymocytes from RORγ -/- mice is oligonucleotide/oligosaccharide-binding fold-containing 2a ( Obfc2a, also named NABP1). The pattern of Obfc2a expression during thymopoiesis correlates with that of RORγt. Therefore, Obfc2a might be a potential RORγ target gene; alternatively, its down-regulation is just a consequence of the reduction in DP cells observed in RORγ -/- mice. In summary, these studies indicate that RORγ plays a critical role in thymocyte homeostasis by regulating differentiation, proliferation, and apoptosis of thymocytes [ He, 2002; Jetten, 2004; Jetten and Joo, 2006; Jetten et al., 2001; Winoto and Littman, 2002]. Role of RORα in lymphocyte development Both the spleen and the thymus are significantly smaller in RORα-deficient mice, suggesting that RORα may have a role in the regulation of thymopoiesis and lymphocyte development [ Dzhagalov et al., 2004; Trenkner and Hoffmann, 1986]. RORα mRNA is expressed at low levels in DP thymocytes and at high levels in SP thymocytes, while it is decreased in mature CD8 + and increased in mature CD4 + T lymphocytes. B220 + B lymphocytes express low levels of RORα. The total number of thymocytes and splenocytes is greatly diminished in RORα -/- mice and is associated with a loss of DP thymocytes and a corresponding increase in the percentage of DN and SP thymocytes [ Dzhagalov et al., 2004]. In RORα -/- spleen, both mature T and B lymphocytes were significantly reduced, suggesting that RORα plays a critical role in lymphocyte development. No differences were observed between RORα -/- and WT mice in the induction of CD4 + and CD8 + T lymphocyte proliferation in response to anti-CD3 or lipopolysaccharide (LPS), nor in the induction of cytokines by TNFα [ Dzhagalov et al., 2004]. In contrast, the induction of IL-6 and tumor necrosis factor α was greatly increased in RORα -/- mast cells and macrophages after LPS treatment [ Dzhagalov et al., 2004; Kopmels et al., 1992]. These results indicate that at least in certain immune cells, RORα functions as a negative regulator of cytokine expression. Although the mechanism of this repression is not fully understood, RORα has been reported to positively regulate the expression of IKBα, a negative regulator of the NF-κB signaling pathway [ Delerive et al., 2001]. On this basis, it was concluded that RORα might function as a negative regulator of inflammation. |
References |