|Mutation Type||critical splice donor site (2 bp from exon)|
|Coordinate||125,313,388 bp (GRCm38)|
|Base Change||A ⇒ C (forward strand)|
|Gene Name||lymphotoxin B receptor|
|Synonym(s)||TNF-R-III, TNFRrp, TNFCR, TNFR2-RP, LTbetaR, TNF receptor-related protein, LT-beta receptor, LT beta-R, Tnfbr, Tnfrsf3, Ltar|
|Chromosomal Location||125,306,571-125,313,885 bp (-)|
FUNCTION: [Summary is not available for the mouse gene. This summary is for the human ortholog.] This gene encodes a member of the tumor necrosis factor receptor superfamily. The major ligands of this receptor include lymphotoxin alpha/beta and tumor necrosis factor ligand superfamily member 14. The encoded protein plays a role in signalling during the development of lymphoid and other organs, lipid metabolism, immune response, and programmed cell death. Activity of this receptor has also been linked to carcinogenesis. Alternatively spliced transcript variants encoding multiple isoforms have been observed. [provided by RefSeq, Aug 2012]
PHENOTYPE: Homozygotes for a targeted null mutation lack Peyer's patches, colon-associated lymphoid tissues, and lymph nodes. Mutants also exhibit severely reduced numbers of NK cells and increased susceptibility to Theiler's murine encephalomyelitis virus. [provided by MGI curators]
|Amino Acid Change|
|Institutional Source||Beutler Lab|
|Gene Model||predicted gene model for protein(s): [ENSMUSP00000032489]|
|Predicted Effect||probably null|
|Alleles Listed at MGI|
|Mode of Inheritance||Autosomal Recessive|
|Local Stock||Live Mice|
|Last Updated||2018-04-02 9:58 PM by Diantha La Vine|
|Record Created||2013-09-26 6:07 PM by Kuan-Wen Wang|
The kama phenotype was initially identified among G3 mice of the pedigree R0553, some of which exhibited diminished T-dependent antibody responses to recombinant Semliki Forest virus (rSFV)-encoded β-galactosidase (rSFV-β-gal; Figure 1).
|Nature of Mutation|
Whole exome HiSeq sequencing of the G1 grandsire identified 43 mutations. The reduced T-dependent antibody response to rSFV-β-gal phenotype was linked by continuous variable mapping to a mutation in Ltbr: a T to G transversion at base pair 125,313,388 on chromosome 6, corresponding to base pair 483 in GenBank genomic region NC_000072. Linkage was fond with a recessive model of inheritance (P = 4.479 x 10-13), wherein 10 variant homozygotes departed phenotypically from 34 unaffected mice that were either heterozygous (n = 25) or homozygous (n = 9) for the reference allele (Figure 2). A substantial semidominant effect was observed in the T-dependent antibody response to rSFV-β-gal assay, but the mutation is preponderantly recessive. The mutation is located in the donor splice site of intron 2, two nucleotides from the previous exon; the transcript contains 10 total exons. The effect of the mutation at the cDNA and protein level is unknown. One possibility, shown below, is that aberrant splicing may result in the skipping of the 97 base pair exon 2 and splicing from exon 1 to exon 3. The aberrant splicing would lead to a deletion of 32 amino acids and a frameshift; the mis-spliced protein would end prematurely in exon 7.
The donor splice site of intron 2 of Ltbr, which is destroyed by the mutation, is indicated in blue; the mutated nucleotide is indicated in red.
Ltbr encodes the lymphotoxin β receptor (LTβR), a member of the tumor necrosis factor receptor (TNFR) superfamily (1). Similar to other members of the TNFR family, LTβR has an extracellular domain (ECD; amino acids 28-221; SMART), a transmembrane domain (TMD; amino acids 222-244), and an intracellular domain (ICD; amino acis 245-415) [Figure 3; (2); reviewed in (3)]. Amino acids 1-27 of LTβR comprise a signal peptide (2). LTβR has two conserved putative Asn-linked glycosylation sites at Asn40 and Asn179 (2).
Within the ECD, LTβR has four cysteine-rich domains (CRDs; amino acids 43-80, 83-124, 126-169, and 172-212) [(4); reviewed in (3)]. The CRDs mediate ligand (i.e., LTα1β2 and LIGHT) specificity and affinity [(5;6); reviewed in (3)]. The second and third CRDs (i.e., amino acids 83-124 and 126-169) mediate most receptor-ligand interactions (6). The interaction between human LTβR and the lymphotoxin (LT)α1β2 ligand was determined by crystallization of a LTα1β2-LTβR-anti-LTα Fab complex [Figure 4; PDB:4MXW; (6)]. The architecture of LTβR is similar to TNFR1 (PDB:1TNR); the CRD1 and CRD2 domains of LTβR are superimposable with the corresponding domains in TNFR1 (6). LTα1β2 has two LTβR binding sites: a lower affinity site at the LTβ–LTβ’ interface and a higher affinity site at the LTα–LTβ interface (6). Mutation of either site disrupted receptor binding at the site and prevented subsequent signal transduction (6). LTβR is bound in the groove formed by the two LTβ molecules (6). The orientation of CRD3 with respect to CRD2 is variable (6). CRD2 mainly forms the predominantly polar interface between LTβR and LTβ–LTβ’; there are minimal additional contacts from CRD1 (6). Gln85, Arg102, and Asp105 from CRD2 form electrostatic interactions with Lys108, Gln109, and Arg142 in LTβ’ (6).
The self-association domain (SAD; amino acids 324-377) within the ICD is required for LTβR-associated apoptotic signaling as well as for the association of LTβR with the serine/threonine kinases p50 and p80 (1;7). The TNFR superfamily of proteins diverge within the ICD domain whereby they can be subdivided as death and non-death receptors [reviewed in (3)]. LTβR, along with TNFR2, HVEM, and TROY, are non-death receptors in that they do not contain death domains, but have one or two TNFR-associated factor (TRAF) binding sites (amino acids 385-408 in LTβR) [(8;9); reviewed in (3)]. Amino acids 388-393 form several contacts with TRAF3 (see the record for hulk) (9). Mutation of amino acids 388-393 to alanines revealed that amino acids 404-406 are also required for TRAF3/TRAF2 recruitment and signaling (9). LTβR and the C-terminal domain of TRAF3 form a 3:3 trimeric complex (10;11). LTβR can also bind TRAF2 and TRAF5, but not TRAF6 (12;13).
Northern blot analysis determined that Ltbr was expressed in the lung, kidney, stomach, small intestine, large intestine, liver, and adrenal gland (2;14;15). Nakamura et al. determined that Ltbr was moderately expressed in the heart and testis and weakly expressed in the brain, thymus, spleen, and lymph nodes (2). In the mouse, Ltbr is expressed as early as embryonic day 7 (E7) and is expressed throughout embryonic development (2). In situ hybridization of whole E14.5 embryos, detected Ltbr in the acinar glandular cells of the submaxillary gland, cortical and medullary thymic regions, muscle, respiratory epithelial cells of the lung and sinus, epithelial lining of the stomach, and epithelial cells of the gut and villi of the E16.5 embryo; highest levels of Ltbr were detected in the epithelial cells of the gut, while the weakest levels of Ltbr were detected in the epithelium of the gut and lung (14). The distribution of Ltbr was not changed by E18.5 (14). By birth, the level of Ltbr in the large and small intestinal epithelium are low that do not change throughout adulthood (14). CD11b+ splenic monocytes as well as CD11b+CD11c+CD11b−CD11c+ and CD11c+CD8a+ splenic dendritic cells expressed Ltbr (14). LTBR is expressed in several solid tumors including colorectal, larynx/pharynx, and lung tumors as well as melanomas (16). LTBR is highly expressed in melanoma cell lines D5, Hs294T, SKMel-5, WM115, and SKMel-28 (17;18).
LTβR is expressed on stromal and mesenchymal cells of the lymphoid tissues, dendritic cells and monocytes [(19;20); reviewed in (21)]. During development LTβR is expressed by fetal stromal cells within the lymph node anlagen as well as by myeloid cells, hepatocytes, intestinal epithelial cells, fibroblasts, and endothelial cells [(14;19;22); reviewed in (23)]. LTβR is not expressed on T or B lymphocytes (15;24). Quiescent and activated hematopoietic stem cells express the LTβR on the cell surface as well as in the perinuclear region (25).
Activated LTβR is internalized through an AP2/clathrin-dependent as well as a clathrin-independent/dynamin-2-dependent pathway [(26); reviewed in (3)]. Internalization via the clathrin-independent pathway is required for the activation of the alternative NF-κB pathway (26).
LTα1β2 and LIGHT (alternatively, TNF superfamily (TNFSF)14) are the known ligands of LTβR; both LTα1β2 and LIGHT are members of the TNF superfamily [(4;27-31); see the record PanR1 (Tnf) and walla (Cd40lg)]. LTα1β2 is a membrane-bound heterotrimeric complex composed of LTα (alternatively, TNFSF1) and a LTβ (alternatively, TNFSF3) homodimer. LTα1β2 is expressed on activated T and B lymphocytes as well as natural killer (NK) cells, a subset of follicular B cells, and lymphoid tissue inducer (LTi) cells (CD4+IL-7R+CD3− CD45+RORγt+; see the record for chestnut) (24;28;32). LIGHT is a homotrimer expressed on T lymphocytes, monocytes, granulocytes, and immature dendritic cells (24). The interaction of LTα1β2 or LIGHT with LTβR forms a connection between lymphocytes and the surrounding parenchymal and stromal cells [reviewed in (33)].
LTβR signals through both the canonical (classical; see the record for finlay) and non-canonical (alternative; see the record for xander) nuclear factor κB (NF-κB) signaling pathways [Figure 5; (34-36); reviewed in (23)]. In canonical NF-κB signaling pathway, TRAF2/5 is quickly recruited to LTβR upon ligation (12;37). The activation of the inhibitor of κB kinase (IKK) complex (NEMO/IKKγ (see the record for panr2), IKKα and IKKβ) promotes the proteasome-mediated degradation of IκBα and the release of the p50:p65 heterodimer to the nucleus (27;35). In the noncanonical NF-κB signaling pathway, LTβR ligation activates IKKα through NF-κB inducing kinase (NIK). TRAF3 is subsequently degraded, releasing NF-κB-inducing kinase (NIK; see the record for lucky) to phosphorylate IKKα, which subsequently phosphorylates p100 (NF-κB2; see the record for xander), leading to the processing of p100 to p52. The p52 fragment forms a heterodimer with RelB and translocates to the nucleus to control the transcription of target genes (9). Activation of the non-canonical pathway is necessary for sustained NF-κB activity in response to LTβR stimulation (34). LTβR-mediated activation of the NF-κB signaling pathways results in the expression of genes that encode adhesion molecules, chemokines (e.g., CCL21 and CXCL13), and lymphokines involved in inflammation and secondary lymphoid organogenesis and homeostasis [Table 1; (35); reviewed in (3)]. For a detailed review on the NF-κB signaling pathways see the records for finlay and xander.
Table 1. Factors regulated by LTβR-associated signaling
LTβR-associated signaling mediates several functions including lymphoid tissue development and maintenance (46-49), formation of germinal centers (50), dendritic cell-mediated immune function (51;52), apoptosis, chemokine secretion, maintenance of splenic architecture, maintenance of T and B cell segregation into discrete compartments, protection against autoimmune diseases, homeostasis of the intestinal immune system (35) including protection against Citrobacter rodentium-induced colitis (29;53), and protection against Mycbacterium bovis bacillus Calmette-Guérin (BCG), Mycobacterium tuberculosis, Listeria monocytogenes, and cytomegalovirus infections (54-56).
Lymphoid tissue development and maintenance
LTβR-associated signaling is essential for the formation of lymph node stromal cell microenvironments during lymph node development. Embryonic LTi cells produce TNF superfamily member RANK ligand (RANKL; alternatively, TRANCE or TNFSF11), IL-7Rα, and LTα1β2 that stimulate LTβR-expressing mesenchymal lymphoid tissue organizer (LTo) cells to produce factors (e.g., chemokines, adhesion molecules, IL-7, and VEGF-C) that organize lymphoid cells into lymph nodes and Peyer’s patches (47;48;57;58). TRANCE and IL-7 subsequently induce LTα1β2 expression on immature LTi cells, enhancing LTβR-associated signaling (57). LTβR ligation results in the activation of the canonical NF-κB pathway and the subsequent expression of proinflammatory molecules including chemokines MIP1β, MIP-2, and the adhesion molecule VCAM-1 [Table 1; (57)]. Prolonged triggering of LTβR leads to the activation of the noncanonical NF-κB pathway and the subsequent transcription of the lymphoid chemokines including CCL21, CCL19, and CXCL13 that recruit immune cells to the developing anlage as well as promote stromal cell proliferation and survival [Table 1; (34;35;49;59;60); reviewed in (61)]. In the adult, LTβR-associated signaling maintains the integrity of secondary lymphoid tissues [reviewed in (21)].
LTβR activation on lymph node endothelial cells is essential for lymphoid tissue organization and lymphocyte homeostasis (20). LTα1β2/LTβR-associated signaling is essential for the differentiation of vascular endothelial cells into high endothelial venules (HEV), the formation of the HEV network, and maintenance of functional HEVs (20;62). Blockade of LTβR-associated signaling with a soluble decoy receptor (LTβR-Ig) resulted in reduced lymph node cellularity in adult mice compared to wild-type mice (62). The reduced cellularity was due to impaired lymphocyte entry into lymph nodes due to reduced levels of peripheral lymph node addressin (PNAd) and MAdCAM1 on HEVs (62). Endothelial cell-specific deletion of Ltbr in mice (VE-cadherin-Cre x Ltβrfl/fl) resulted in a failure to develop 20-45% of their peripheral lymph nodes; the formation of mesenteric lymph nodes was not affected (20). Remaining lymph nodes exhibited impaired formation (i.e., reduced segment length and branching points) of HEVs (20). Adhesion molecule and chemokine expression (i.e., Ccl21 and Ccl19) were also reduced in the HEVs (20). Changes in endothelial cell-lymphocyte interactions resulted in impaired homing of lymphocytes to peripheral lymph nodes (20).
Lymphatic vessels transport fluids and nutrients into the circulation to maintain homeostasis. In addition, lymphatic vessels transport antigen-presenting cells, memory and effector T cells, and inflammatory mediators from peripheral tissues to the lymph nodes [reviewed in (63)]. LTα1β2/LTβR-associated signaling functions in lymphangiogenesis during the formation of tertiary lymphoid structures (22). The role of LTα1β2/LTβR-associated signaling in de novo lymphangiogenesis in lymphatic vessels was examined in Ltbr-/- crossed with transgenic mice (TGCCL21) that overexpressed CCL21 in the thyroid (22). The TGCCL21/Ltbr-/- mice had reduced numbers of lymphatic vessels compared to TGCCL21/Ltbr+/- littermates (22). Loss of Ltbr expression negatively affected de novo lymphoangiogenesis in areas with lymphocytic infiltrates, but lymphatic vessels were identified in the adjacent normal tissue; LTβR is not required for embryonic development of lymphatic vessels (22).
Medullary thymic epithelial cell differentiation and Aire regulation
LTβR-associated signaling upregulates the expression of RANK in the thymic stroma, promoting accessibility to RANKL; RANKL/RANK signaling is necessary for medullary thymic epithelial cell (mTEC) differentiation during embryogenesis (64). Postnatal mTECs contain both CCL21- and Aire (autoimmune regulator transcription factor)-expressing subsets (65). Aire is an autoimmune regulator that mediates central tolerance for peripherally restricted antigens (66). The role of LTβR-associated signaling on Aire expression is conflicting. Chin et al. found that loss of Ltbr expression resulted in reduced steady-state levels of Aire as well as its downstream target genes (66). As a result, Aire-associated protection from autoimmunity was lost and the Ltbr-/- mice produced autoantibodies, including rheumatoid factor and anti-DNA antibodies (66). In contrast, Boehm et al. found that the expression level of Aire was normal in thymic MECs from Ltbr-/- mice (67). In addition, Lkhagvasuren et al. determined that the numbers of CCL21-expressing, but not Aire-expressing, mTECs of the stroma are reduced in Ltbr-/- mice (65).
Organization of splenic microenvironments
LTβR-associated signaling maintains the structure of the spleen, including the marginal zone that separates the red and white pulp regions of the spleen (68;69). LT, primarily from B cells, controls the microstructure of the spleen to promote antibody responses (70;71). LTβR is required to maintain the levels of metallophilic macrophages in the corticomedullary zones (CMZ) of both the thymus and spleen (72).
LTβR and retinoic acid receptor-related orphan receptor-γt (RORγt) transcription factor (see the record for chestnut) regulate the differentiation of γδ T cells (58;73). An anti-LTβR agonist antibody did not alter the development and numbers of TCRγδ+ cells, but the expression of γδ-associated genes (e.g., Dap12) in the cells were rescued (73). In Ltbr-/- mice, there was abnormal activation-induced upregulation of GzmA, Dap12, and cyclin D3 (73). Further studies have proposed that the LTα1β2/LTβR signaling pathway functions in the development of both γδ T cells and invariant natural killer T (iNKT) cells (58). Ltbr-/- mice are deficient in IgE production, indicating an imbalance in CD4+ T helper (TH) cell subsets (74). In addition, upon C. rodentium or L. major infection, Ltbr-/- mice favor the development of IL-4-producing CD4+ T cells, rather than interferon (IFN)γ producing T cells [reviewed in (23)]. In response to mouse cytomegalovirus (MCMV) challenge, LTβR and LTα prevent CD8+ T cell apoptosis (55). Blockade of LT-associated signaling resulted in decreased expansion of the OVA-specific CD8+ T (OT-I) cell population in response to MCMV, indicating that LTβR functions in CD8+ T cell homeostasis (75). In addition, agonist antibodies to LTβR induced IFNβ production from bone marrow-derived dendritic cells (BMDCs) in response to lipopolysaccharide (75).
Dendritic cell homeostasis
LTβR is required for the homeostasis of lymphoid tissue DCs (51;52). Splenic dendritic cell homeostasis is maintained upon their migration to the spleen via the chemokine microenvironment induced by LTβR signaling (52). LIGHT/LTβR-associated survival and proliferation signals regulate the number of dendritic cells, while chemokines regulate the localization of the dendritic cells in the spleen (52). Ltbr-/- mice lack specific subsets of splenic dendritic cells (e.g., Notch-dependent endothelial cell-selective adhesion molecule (ESAM)hi dendritic cells) [(51;52;76;77); reviewed in (23)]. OVA-specific CD4+ T cells (OT-II) exhibit reduced division rates in mice that lack the LTβR-dependent ESAMhi dendritic cell population compared to wild-type mice, indicating that LTβR-associated signaling may promote CD4+ T cell priming by regulating the maturation status of antigen presenting cells (76). DCs from Ltbr-/- mice are also less efficient at inducing CD8+ T cell expansion; exposure to type I IFNs alleviates the defect (75).
LTβR-associated signaling can activate both caspase-dependent and caspase-independent apoptotic pathways (78). LTβR-associated cytotoxic signaling can be initiated by both LTα1β2 and LIGHT in conjuction with IFNγ (79-81). LTβR induces a slow apoptotic death, similar to TNFR2 and CD30 (80;81). TRAF3 (see the record for hulk) mediates LTβR-induced apoptosis within 36-72 hours (compared to a few hours in the case of Fas ligand (see the record for riogrande) or 24 hours in the case of TNF) in HT29 adenocarcinoma cells (12;82). The time delay between TRAF3-LTβR association and cell death indicates that the effects of TRAF3 association with LTβR may be indirectly linked to the process of cell death (12). LTβR does not contain a death domain, therefore, it does not associate with death domain-containing TRADD and FADD as observed in TNFR-associated NF-κB signaling (44). Overexpression of LTβR, without ligand stimulation or an agonist antibody, is sufficient to trigger apoptosis (1). LTβR-induced apoptosis can be inhibited by several caspase inhibitors (i.e., Z-VAD-FMK (a broad spectrum caspase inhibitor), DEVD-FMK (a CPP32 inhibitor), and the CrmA (a serine proteinase inhibitor), indicating that CPP32 or interleukin-1β-converting enzyme (ICE)-related caspases may be involved in LTβR-induced apoptotic signaling (1).
LTα1β2-induced activation of LTβR leads to downregulation of proinflammatory cytokine expression upon TLR restimulation, indicating that LTβR-associated signaling induces TLR cross-tolerance and downregulates the inflammatory response (28). LTβR-induced tripartite motif (TRIM)-containing 30α expression on bone marrow-derived macrophages is mediated by IκBα-dependent signaling downstream of TRAF3 (28). TRIM30α induction is a negative regulator of NF-κB activation upon induction by TLR signaling (28;83). siRNA-mediated knockdown of TRIM30α resulted in loss of LTβR-dependent induction of TRIM30α and LTβR-mediated TLR cross-tolerance (28). Loss of LTβR-associated signaling resulted in the loss of TRIM30α induction with a concomitant increase in the expression of pro-inflammatory cytokines (28;83). Macrophage/neutrophil-specific knockout of Ltbr (LTβRflox/flox × LysM-Cre) resulted in impaired LTβR-induced cross-tolerance to TLR4 and TLR9 ligands, indicating that LTβR activation on macrophages and neutrophils induces TRIM30α expression (28).
LTβR in the intestine
LTβR-associated signaling is required for B cell recruitment to the intestinal lamina propria and for serum and intestinal IgA expression [(84); reviewed in (21)]. Inducible nitric oxid synthase (iNOS)-producing dendritic cells contribute to intestinal IgA production and are absent in the Ltbr-/- mice resulting in loss of both serum and intestinal IgA expression [(84;85); reviewed in (23)].
LTα1β2/LTβR-associated signaling is essential in controlling the spread of enteric pathogens (29;86;87). For example, Ltbr-/- mice exhibit a greater systemic spread of Salmonella typhimurium; B cell localization in response to S. typhimurium is dependent on LTβR-associated signaling (86). Ltbr-/- mice exhibited reduced humoral responses in response to C. rodentium and were highly susceptible to C. rodentium by 10 days post-infection (29;53). In contrast to wild-type splenocytes that produce IFNγ in response to C. rodentium, Ltbr-/- splenocytes produced IL-4 and IL-10 (53). Mice with gut epithelial cell-specific knockout of Ltbr (Vil-Ltbr−/−) exhibited deficiency in clearing C. rodentium infection and had 15-20 times the amount of bacterial titers in the spleen and feces compared to wild-type mice 10 days post-infection (29). In contrast to Ltbr-/- mice, the Vil-Ltbr−/− mice survived the C. rodentium infection, indicating that LTβR signaling from other cell types may contribute to the severity of the disease (29). Macrophage and neutrophil- specific LTβR deficient (LysM-Ltbr−/−) mice also exhibited increased bacterial titers in the blood, but survived the infection, indicating that LTβR-associated signaling on macrophages and/or neutrophils is required for bacterial clearance (29). Taken together, LTα1β2-expressing cells instruct, via LTβR-associated signaling, intestinal epithelial cells to mobilize the innate immune response against microbial infection (29).
Ltbr−/− mice exhibited susceptibility to dextran sodium sulfate (DSS)-induced colitis (88). Jungbeck et al. determined that LTβR activation by LTα1β2 was necessary for downregulation of the inflammatory response observed in the DSS-colitis model (89). Inhibition of the LTβR-associated signaling pathway reduced inflammation in chronic DSS-induced colitis through the downregulation of MAdCAM-1 expression (90). Macrophage and neutrophil-specific Ltbr−/− (Ltbrflox/flox × LysM-Cre) mice exhibited exacerbated intestinal inflammation in response to DSS, indicating that LTβR expression on macrophages and/or neutrophils is necessary for the control and down-regulation of the inflammatory reaction (83). Wimmer et al. determined that LTα1β2 expressed on CD4+ T cells was responsible for LTβR activation on macrophages, resulting in a TRIM30α-dependent signaling pathway activation and downregulation of the inflammatory response (83;88).
Hyperlipidemia is often associated with inflammation and is a risk factor for coronary heart disease (i.e., atherosclerosis). LTα1β2 and LIGHT are regulators of enzymes that control lipid metabolism (91). Deregulation of LIGHT on T cells results in hypertriglyceridemia and hypercholesterolemia (91). Transgenic mice that expressed increased levels of LIGHT in the T cell lineage (Lck LIGHT Tg) on a Ltbr−/− background (LIGHT Tg/Ltbr−/−) exhibited corrected LIGHT-induced dyslipidemia (91). Increased plasma levels of LIGHT have been observed in patients with stable or unstable angina (92) and acute atherothrombotic stroke (93). In addition, mutations in the LT-α gene (LTA) have been linked to increased susceptibility to myocardial infarction in Japanese patients (94). Higher levels of soluble LTβR in humans is significantly associated with cardiovascular risk factors, renal function, biomarkers associated with atherosclerosis, and measures of atherosclerosis burden (i.e., coronary calcium, aortic plaque, and aortic wall thickness); membrane levels of LTβR were not examined (95).
In a choline-deficient, ethionine-supplemented (CDE) dietary model of chronic liver injury, LTβR was identified as necessary to mediate wound healing (25;96). Ltbr-/- mice fed the CDE diet exhibited reduced fibrosis and dysregulated immune responses (25). In the CDE-induced chronic liver injury model, While RANTES was significantly upregulated upon CDE treatment in the Ltbr-/- mice, the expression of TNF and IFN-γ were not induced (25). Ruddell et al. proposed that activated LTβR on hepatic stellate cells mediated the recruitment of progenitor cells, hepatic stellate cells, and leukocytes needed for wound healing and regeneration during chronic liver injury (25).
In melanoma cells and tumors, LTβR induces non-canonical NF-κB activation to regulate gene expression that leads to increased cell growth (17). shRNA-induced reduction in Ltbr expression decreased NF-κB promoter activity as well as decreased growth and invasiveness compared to the control (17). LTα1β2- and LIGHT-mediated activation of LTβR on mouse fibrosarcoma cells (BFS-1) resulted in increased solid tumor growth with a concomitant increase in CXCL2-induced angiogenesis (36;42). Inhibition of LTα1β2/LTβR signaling prevented tumor angiogenesis and neovascularization, leading to tumor growth arrest (42). T cell- or B cell-specific knockout of LTβ resulted in reduced solid fibrosarcosomas compared to wild-type mice, indicating that both T and B lymphocytes function in the activation of LTβR in these tumors (36). Daller et al. determined that both IκBα and NIK are involved in pro-angiogenesis signaling after LTβR activation (36). In contrast to fibrosarcosomas, activation of LTβR on some adenocarcinoma cell lines (HT-29 and WiDr) results in the induction of cell death (80). In addition, an agonistic anti-LTβR antibody blocked tumor growth in models of colon and cervical carcinoma (16). In melanoma cell lines, LTβR-mediated activation of NFκB resulted in tumor cell proliferation (17). In addition, LTβR-associated signaling promotes the development of hepatocellular carcinoma (97).
Additional Ltbr knockout mouse phenotypes
Ltbr knockout (Ltbr-/-; Ltbrtm1Kpf, MGI:2384140) mice appear healthy, are born at the expected Menelian frequency, and are fertile (48). Ltbr-/- mice exhibited defects in secondary lymphoid organogenesis including the absence of cervical, axillary, inguinal, paraaortic/sacral, popliteal, and mesenteric lymph nodes, Peyer’s patches, and germinal centers, disorganization of the splenic architecture (i.e., disturbed microarchitecture of the white pulp, no separate B- and T-cell areas, no follicles, and disruptions to the marginal zone), and disruption of the thymic stroma architecture (40;47;48;58). All of the Ltbr-/- mice had a spleen and a thymus; the spleens of the Ltbr-/- mice were 1.5 to 2 times bigger than those in wild-type mice (48). Thymocyte maturation was normal in the Ltbr-/- mice (48). Within the spleen, the marginal zone B cell population (B220+, IgMhigh, IgDdull), MOMA-1+ metallophilic macrophages, sialoadhesin+ MZ macrophages, and MAdCAM-1+ sinus lining cells were completely undetectable (48). Flow cytometric analysis of lymphocytes from lungs and spleens of the Ltbr-/- mice determined that αEβ7high integrin+ T cells were absent (48). Infiltrations of lymphocytes (mainly CD4+ T cells and B220+ B cells) were observed in the Ltbr-/- mice around the perivascular areas in the lungs, liver, pancreas, submandibular glands, the fatty tissue of the mediastinum, mesenterium, cortex of the suprarenal glands, and kidney (48). Ltbr-/- mice exhibited lethality three weeks after exposure to Plasmodium berghei ANKA (PbA)-induced experimental cerebral malaria (ECM) with a concomitant high parasitaemia and severe anemia; wild-type mice succumb after approximately one week (98). Ltbr-/- mice did not develop ECM-associated neurological signs such as postural disorders, ataxia, impaired reflexes, loss of grip strength, progressive paralysis, and coma (98).
LTβR-associated signaling has been linked to several human conditions including autoimmunity, atherosclerosis, and cancer (see descriptions, above). Mutations in LTBR have been linked to increased risk for IgA nephropathy (OMIM: %161950), a form of glomerulonephritis that leads to end-stage renal disease, in Korean children (99).
LTβR-associated signaling is known to maintain the marginal zone to promote antibody responses and is required for the formation of germinal centers during antigen-dependent responses (35;48;68;69;70;71). Ltbr-/- mice exhibited higher levels of IgM in response to the T-dependent antigen, NP19-CG (4-hydroxy-3-nitrophenyl-acetyl-chicken gamma globulin absorbed to alum) compared to wild-type mice, however the amount of anti-NP IgG1 levels produced were lower than wild-type mice (48). The kama mice exhibit a negligible T-dependent IgG response to rSFV-β-gal; the IgM levels were not measure in kama. The morphology of the spleen in the kama mice has not been examined, but the diminished antibody response indicates defects in germinal center formation in the kama mice.
kama(F):5'- GAGTCCCCACTTACCAATGTCACAG -3'
kama(R):5'- ATGGACCAGGTTTCTCTCTAGCCC -3'
kama_seq(F):5'- GCAAGTCTTGCAAACCGTG -3'
kama_seq(R):5'- CTTCTGCCCTCCAGAGCTG -3'
Kama genotyping is performed by amplifying the region containing the mutation using PCR, followed by sequencing of the amplified region to detect the single nucleotide transversion.
Kama(F): 5’- GAGTCCCCACTTACCAATGTCACAG-3’
Kama(R): 5’- ATGGACCAGGTTTCTCTCTAGCCC-3’
Kama_seq(F): 5’- GCAAGTCTTGCAAACCGTG-3’
Kama_seq(R): 5’- CTTCTGCCCTCCAGAGCTG-3’
1) 94°C 2:00
2) 94°C 0:30
3) 55°C 0:30
4) 72°C 1:00
5) repeat steps (2-4) 40X
6) 72°C 10:00
7) 4°C ∞
The following sequence of 589 nucleotides are amplified (Chr.6: 125312980-125313568, GRCm38):
gagtccccac ttaccaatgt cacaggggcg gcacagctgg caggtggaga gatggttcca gtgttcatta taggaattat gggggcaagt cttgcaaacc gtgtcttggc tgcggctgca taccgcaaag acaaactcgc ctatgaggcg aatggggaaa gagggttatc aggagacaag aagtccatct ctgcgtccgc tgcctctgct tgctgcatgc ctcggagcct ctccttggtc tctttgtccc catcctgtct tcatggctac atgccgctac ctcttcaaat ctcagtgtcc gtgggacaat gctaatgctt cgtagactct ccttcctggc tagttttagc agtgctggct cattctggct cacctctacc acccaagcca cttggagcca ctgttctcac ctgggggaca gcgggagcag cagacgtcgt gcatgggctc gtagtattcc ttgtcctggt cccagcaagt ctggttttct atgcgataag ggggcacctt ccagaggagg cggagaaaga gaagaatgag gcagggtcag ctctggaggg cagaagggct agagagaaac ctggtccat
Primer binding sites are underlined and the sequencing primers are highlighted; the mutated nucleotide is shown in red text (A>C, Chr.+ strand; T>G, sense strand).
1. Wu, M. Y., Wang, P. Y., Han, S. H., and Hsieh, S. L. (1999) The Cytoplasmic Domain of the Lymphotoxin-Beta Receptor Mediates Cell Death in HeLa Cells. J Biol Chem. 274, 11868-11873.
2. Nakamura, T., Tashiro, K., Nazarea, M., Nakano, T., Sasayama, S., and Honjo, T. (1995) The Murine Lymphotoxin-Beta Receptor cDNA: Isolation by the Signal Sequence Trap and Chromosomal Mapping. Genomics. 30, 312-319.
3. Remouchamps, C., Boutaffala, L., Ganeff, C., and Dejardin, E. (2011) Biology and Signal Transduction Pathways of the Lymphotoxin-alphabeta/LTbetaR System. Cytokine Growth Factor Rev. 22, 301-310.
4. Crowe, P. D., VanArsdale, T. L., Walter, B. N., Ware, C. F., Hession, C., Ehrenfels, B., Browning, J. L., Din, W. S., Goodwin, R. G., and Smith, C. A. (1994) A Lymphotoxin-Beta-Specific Receptor. Science. 264, 707-710.
5. Eldredge, J., Berkowitz, S., Corin, A. F., Day, E. S., Hayes, D., Meier, W., Strauch, K., Zafari, M., Tadi, M., and Farrington, G. K. (2006) Stoichiometry of LTbetaR Binding to LIGHT. Biochemistry. 45, 10117-10128.
6. Sudhamsu, J., Yin, J., Chiang, E. Y., Starovasnik, M. A., Grogan, J. L., and Hymowitz, S. G. (2013) Dimerization of LTbetaR by LTalpha1beta2 is Necessary and Sufficient for Signal Transduction. Proc Natl Acad Sci U S A. 110, 19896-19901.
7. Wu, M. Y., Hsu, T. L., Lin, W. W., Campbell, R. D., and Hsieh, S. L. (1997) Serine/threonine Kinase Activity Associated with the Cytoplasmic Domain of the Lymphotoxin-Beta Receptor in HepG2 Cells. J Biol Chem. 272, 17154-17159.
8. Force, W. R., Glass, A. A., Benedict, C. A., Cheung, T. C., Lama, J., and Ware, C. F. (2000) Discrete Signaling Regions in the Lymphotoxin-Beta Receptor for Tumor Necrosis Factor Receptor-Associated Factor Binding, Subcellular Localization, and Activation of Cell Death and NF-kappaB Pathways. J Biol Chem. 275, 11121-11129.
9. Sanjo, H., Zajonc, D. M., Braden, R., Norris, P. S., and Ware, C. F. (2010) Allosteric Regulation of the Ubiquitin:NIK and Ubiquitin:TRAF3 E3 Ligases by the Lymphotoxin-Beta Receptor. J Biol Chem. 285, 17148-17155.
10. Li, C., Norris, P. S., Ni, C. Z., Havert, M. L., Chiong, E. M., Tran, B. R., Cabezas, E., Reed, J. C., Satterthwait, A. C., Ware, C. F., and Ely, K. R. (2003) Structurally Distinct Recognition Motifs in Lymphotoxin-Beta Receptor and CD40 for Tumor Necrosis Factor Receptor-Associated Factor (TRAF)-Mediated Signaling. J Biol Chem. 278, 50523-50529.
11. Wu, H. (2013) Higher-Order Assemblies in a New Paradigm of Signal Transduction. Cell. 153, 287-292.
12. VanArsdale, T. L., VanArsdale, S. L., Force, W. R., Walter, B. N., Mosialos, G., Kieff, E., Reed, J. C., and Ware, C. F. (1997) Lymphotoxin-Beta Receptor Signaling Complex: Role of Tumor Necrosis Factor Receptor-Associated Factor 3 Recruitment in Cell Death and Activation of Nuclear Factor kappaB. Proc Natl Acad Sci U S A. 94, 2460-2465.
13. Inoue, J., Ishida, T., Tsukamoto, N., Kobayashi, N., Naito, A., Azuma, S., and Yamamoto, T. (2000) Tumor Necrosis Factor Receptor-Associated Factor (TRAF) Family: Adapter Proteins that Mediate Cytokine Signaling. Exp Cell Res. 254, 14-24.
14. Browning, J. L., and French, L. E. (2002) Visualization of Lymphotoxin-Beta and Lymphotoxin-Beta Receptor Expression in Mouse Embryos. J Immunol. 168, 5079-5087.
15. Force, W. R., Walter, B. N., Hession, C., Tizard, R., Kozak, C. A., Browning, J. L., and Ware, C. F. (1995) Mouse Lymphotoxin-Beta Receptor. Molecular Genetics, Ligand Binding, and Expression. J Immunol. 155, 5280-5288.
16. Lukashev, M., LePage, D., Wilson, C., Bailly, V., Garber, E., Lukashin, A., Ngam-ek, A., Zeng, W., Allaire, N., Perrin, S., Xu, X., Szeliga, K., Wortham, K., Kelly, R., Bottiglio, C., Ding, J., Griffith, L., Heaney, G., Silverio, E., Yang, W., Jarpe, M., Fawell, S., Reff, M., Carmillo, A., Miatkowski, K., Amatucci, J., Crowell, T., Prentice, H., Meier, W., Violette, S. M., Mackay, F., Yang, D., Hoffman, R., and Browning, J. L. (2006) Targeting the Lymphotoxin-Beta Receptor with Agonist Antibodies as a Potential Cancer Therapy. Cancer Res. 66, 9617-9624.
17. Dhawan, P., Su, Y., Thu, Y. M., Yu, Y., Baugher, P., Ellis, D. L., Sobolik-Delmaire, T., Kelley, M., Cheung, T. C., Ware, C. F., and Richmond, A. (2008) The Lymphotoxin-Beta Receptor is an Upstream Activator of NF-kappaB-Mediated Transcription in Melanoma Cells. J Biol Chem. 283, 15399-15408.
18. Winter, H., van den Engel, N. K., Poehlein, C. H., Hatz, R. A., Fox, B. A., and Hu, H. M. (2007) Tumor-Specific T Cells Signal Tumor Destruction Via the Lymphotoxin Beta Receptor. J Transl Med. 5, 14.
19. Murphy, M., Walter, B. N., Pike-Nobile, L., Fanger, N. A., Guyre, P. M., Browning, J. L., Ware, C. F., and Epstein, L. B. (1998) Expression of the Lymphotoxin Beta Receptor on Follicular Stromal Cells in Human Lymphoid Tissues. Cell Death Differ. 5, 497-505.
20. Onder, L., Danuser, R., Scandella, E., Firner, S., Chai, Q., Hehlgans, T., Stein, J. V., and Ludewig, B. (2013) Endothelial Cell-Specific Lymphotoxin-Beta Receptor Signaling is Critical for Lymph Node and High Endothelial Venule Formation. J Exp Med. 210, 465-473.
21. McCarthy, D. D., Summers-Deluca, L., Vu, F., Chiu, S., Gao, Y., and Gommerman, J. L. (2006) The Lymphotoxin Pathway: Beyond Lymph Node Development. Immunol Res. 35, 41-54.
22. Furtado, G. C., Marinkovic, T., Martin, A. P., Garin, A., Hoch, B., Hubner, W., Chen, B. K., Genden, E., Skobe, M., and Lira, S. A. (2007) Lymphotoxin Beta Receptor Signaling is Required for Inflammatory Lymphangiogenesis in the Thyroid. Proc Natl Acad Sci U S A. 104, 5026-5031.
23. Upadhyay, V., and Fu, Y. X. (2013) Lymphotoxin Signalling in Immune Homeostasis and the Control of Microorganisms. Nat Rev Immunol. 13, 270-279.
24. Ware, C. F., VanArsdale, T. L., Crowe, P. D., and Browning, J. L. (1995) The Ligands and Receptors of the Lymphotoxin System. Curr Top Microbiol Immunol. 198, 175-218.
25. Ruddell, R. G., Knight, B., Tirnitz-Parker, J. E., Akhurst, B., Summerville, L., Subramaniam, V. N., Olynyk, J. K., and Ramm, G. A. (2009) Lymphotoxin-Beta Receptor Signaling Regulates Hepatic Stellate Cell Function and Wound Healing in a Murine Model of Chronic Liver Injury. Hepatology. 49, 227-239.
26. Ganeff, C., Remouchamps, C., Boutaffala, L., Benezech, C., Galopin, G., Vandepaer, S., Bouillenne, F., Ormenese, S., Chariot, A., Schneider, P., Caamano, J., Piette, J., and Dejardin, E. (2011) Induction of the Alternative NF-kappaB Pathway by Lymphotoxin Alphabeta (LTalphabeta) Relies on Internalization of LTbeta Receptor. Mol Cell Biol. 31, 4319-4334.
27. Schneider, K., Potter, K. G., and Ware, C. F. (2004) Lymphotoxin and LIGHT Signaling Pathways and Target Genes. Immunol Rev. 202, 49-66.
28. Wimmer, N., Huber, B., Barabas, N., Rohrl, J., Pfeffer, K., and Hehlgans, T. (2012) Lymphotoxin Beta Receptor Activation on Macrophages Induces Cross-Tolerance to TLR4 and TLR9 Ligands. J Immunol. 188, 3426-3433.
29. Wang, Y., Koroleva, E. P., Kruglov, A. A., Kuprash, D. V., Nedospasov, S. A., Fu, Y. X., and Tumanov, A. V. (2010) Lymphotoxin Beta Receptor Signaling in Intestinal Epithelial Cells Orchestrates Innate Immune Responses Against Mucosal Bacterial Infection. Immunity. 32, 403-413.
30. Browning, J. L., Dougas, I., Ngam-ek, A., Bourdon, P. R., Ehrenfels, B. N., Miatkowski, K., Zafari, M., Yampaglia, A. M., Lawton, P., and Meier, W. (1995) Characterization of Surface Lymphotoxin Forms. use of Specific Monoclonal Antibodies and Soluble Receptors. J Immunol. 154, 33-46.
31. Mauri, D. N., Ebner, R., Montgomery, R. I., Kochel, K. D., Cheung, T. C., Yu, G. L., Ruben, S., Murphy, M., Eisenberg, R. J., Cohen, G. H., Spear, P. G., and Ware, C. F. (1998) LIGHT, a New Member of the TNF Superfamily, and Lymphotoxin Alpha are Ligands for Herpesvirus Entry Mediator. Immunity. 8, 21-30.
32. Lane, P. J., Gaspal, F. M., and Kim, M. Y. (2005) Two Sides of a Cellular Coin: CD4(+)CD3- Cells Regulate Memory Responses and Lymph-Node Organization. Nat Rev Immunol. 5, 655-660.
33. Ware, C. F. (2005) Network Communications: Lymphotoxins, LIGHT, and TNF. Annu Rev Immunol. 23, 787-819.
34. Muller, J. R., and Siebenlist, U. (2003) Lymphotoxin Beta Receptor Induces Sequential Activation of Distinct NF-Kappa B Factors Via Separate Signaling Pathways. J Biol Chem. 278, 12006-12012.
35. Dejardin, E., Droin, N. M., Delhase, M., Haas, E., Cao, Y., Makris, C., Li, Z. W., Karin, M., Ware, C. F., and Green, D. R. (2002) The Lymphotoxin-Beta Receptor Induces Different Patterns of Gene Expression Via Two NF-kappaB Pathways. Immunity. 17, 525-535.
36. Daller, B., Musch, W., Rohrl, J., Tumanov, A. V., Nedospasov, S. A., Mannel, D. N., Schneider-Brachert, W., and Hehlgans, T. (2011) Lymphotoxin-Beta Receptor Activation by Lymphotoxin-Alpha(1)Beta(2) and LIGHT Promotes Tumor Growth in an NFkappaB-Dependent Manner. Int J Cancer. 128, 1363-1370.
37. Nakano, H., Oshima, H., Chung, W., Williams-Abbott, L., Ware, C. F., Yagita, H., and Okumura, K. (1996) TRAF5, an Activator of NF-kappaB and Putative Signal Transducer for the Lymphotoxin-Beta Receptor. J Biol Chem. 271, 14661-14664.
38. Zhu, M., Chin, R. K., Tumanov, A. V., Liu, X., and Fu, Y. X. (2007) Lymphotoxin Beta Receptor is Required for the Migration and Selection of Autoreactive T Cells in Thymic Medulla. J Immunol. 179, 8069-8075.
39. Ngo, V. N., Korner, H., Gunn, M. D., Schmidt, K. N., Riminton, D. S., Cooper, M. D., Browning, J. L., Sedgwick, J. D., and Cyster, J. G. (1999) Lymphotoxin alpha/beta and Tumor Necrosis Factor are Required for Stromal Cell Expression of Homing Chemokines in B and T Cell Areas of the Spleen. J Exp Med. 189, 403-412.
40. Seymour, R., Sundberg, J. P., and Hogenesch, H. (2006) Abnormal Lymphoid Organ Development in Immunodeficient Mutant Mice. Vet Pathol. 43, 401-423.
41. Cyster, J. G. (1999) Chemokines and Cell Migration in Secondary Lymphoid Organs. Science. 286, 2098-2102.
42. Hehlgans, T., Stoelcker, B., Stopfer, P., Muller, P., Cernaianu, G., Guba, M., Steinbauer, M., Nedospasov, S. A., Pfeffer, K., and Mannel, D. N. (2002) Lymphotoxin-Beta Receptor Immune Interaction Promotes Tumor Growth by Inducing Angiogenesis. Cancer Res. 62, 4034-4040.
43. Degli-Esposti, M. A., Davis-Smith, T., Din, W. S., Smolak, P. J., Goodwin, R. G., and Smith, C. A. (1997) Activation of the Lymphotoxin Beta Receptor by Cross-Linking Induces Chemokine Production and Growth Arrest in A375 Melanoma Cells. J Immunol. 158, 1756-1762.
44. Chang, Y. H., Hsieh, S. L., Chen, M. C., and Lin, W. W. (2002) Lymphotoxin Beta Receptor Induces Interleukin 8 Gene Expression Via NF-kappaB and AP-1 Activation. Exp Cell Res. 278, 166-174.
45. Lu, T. T., and Cyster, J. G. (2002) Integrin-Mediated Long-Term B Cell Retention in the Splenic Marginal Zone. Science. 297, 409-412.
46. Banks, T. A., Rouse, B. T., Kerley, M. K., Blair, P. J., Godfrey, V. L., Kuklin, N. A., Bouley, D. M., Thomas, J., Kanangat, S., and Mucenski, M. L. (1995) Lymphotoxin-Alpha-Deficient Mice. Effects on Secondary Lymphoid Organ Development and Humoral Immune Responsiveness. J Immunol. 155, 1685-1693.
47. De Togni, P., Goellner, J., Ruddle, N. H., Streeter, P. R., Fick, A., Mariathasan, S., Smith, S. C., Carlson, R., Shornick, L. P., and Strauss-Schoenberger, J. (1994) Abnormal Development of Peripheral Lymphoid Organs in Mice Deficient in Lymphotoxin. Science. 264, 703-707.
48. Futterer, A., Mink, K., Luz, A., Kosco-Vilbois, M. H., and Pfeffer, K. (1998) The Lymphotoxin Beta Receptor Controls Organogenesis and Affinity Maturation in Peripheral Lymphoid Tissues. Immunity. 9, 59-70.
49. Rennert, P. D., James, D., Mackay, F., Browning, J. L., and Hochman, P. S. (1998) Lymph Node Genesis is Induced by Signaling through the Lymphotoxin Beta Receptor. Immunity. 9, 71-79.
50. Matsumoto, M., Mariathasan, S., Nahm, M. H., Baranyay, F., Peschon, J. J., and Chaplin, D. D. (1996) Role of Lymphotoxin and the Type I TNF Receptor in the Formation of Germinal Centers. Science. 271, 1289-1291.
51. Kabashima, K., Banks, T. A., Ansel, K. M., Lu, T. T., Ware, C. F., and Cyster, J. G. (2005) Intrinsic Lymphotoxin-Beta Receptor Requirement for Homeostasis of Lymphoid Tissue Dendritic Cells. Immunity. 22, 439-450.
52. Wang, Y. G., Kim, K. D., Wang, J., Yu, P., and Fu, Y. X. (2005) Stimulating Lymphotoxin Beta Receptor on the Dendritic Cells is Critical for their Homeostasis and Expansion. J Immunol. 175, 6997-7002.
53. Spahn, T. W., Maaser, C., Eckmann, L., Heidemann, J., Lugering, A., Newberry, R., Domschke, W., Herbst, H., and Kucharzik, T. (2004) The Lymphotoxin-Beta Receptor is Critical for Control of Murine Citrobacter Rodentium-Induced Colitis. Gastroenterology. 127, 1463-1473.
54. Ehlers, S., Holscher, C., Scheu, S., Tertilt, C., Hehlgans, T., Suwinski, J., Endres, R., and Pfeffer, K. (2003) The Lymphotoxin Beta Receptor is Critically Involved in Controlling Infections with the Intracellular Pathogens Mycobacterium Tuberculosis and Listeria Monocytogenes. J Immunol. 170, 5210-5218.
55. Banks, T. A., Rickert, S., and Ware, C. F. (2006) Restoring Immune Defenses Via Lymphotoxin Signaling: Lessons from Cytomegalovirus. Immunol Res. 34, 243-254.
56. Lucas, R., Tacchini-Cottier, F., Guler, R., Vesin, D., Jemelin, S., Olleros, M. L., Marchal, G., Browning, J. L., Vassalli, P., and Garcia, I. (1999) A Role for Lymphotoxin Beta Receptor in Host Defense Against Mycobacterium Bovis BCG Infection. Eur J Immunol. 29, 4002-4010.
57. Vondenhoff, M. F., Greuter, M., Goverse, G., Elewaut, D., Dewint, P., Ware, C. F., Hoorweg, K., Kraal, G., and Mebius, R. E. (2009) LTbetaR Signaling Induces Cytokine Expression and Up-Regulates Lymphangiogenic Factors in Lymph Node Anlagen. J Immunol. 182, 5439-5445.
58. Elewaut, D., and Ware, C. F. (2007) The Unconventional Role of LT Alpha Beta in T Cell Differentiation. Trends Immunol. 28, 169-175.
59. Muller, G., and Lipp, M. (2003) Concerted Action of the Chemokine and Lymphotoxin System in Secondary Lymphoid-Organ Development. Curr Opin Immunol. 15, 217-224.
60. White, A., Carragher, D., Parnell, S., Msaki, A., Perkins, N., Lane, P., Jenkinson, E., Anderson, G., and Caamano, J. H. (2007) Lymphotoxin a-Dependent and -Independent Signals Regulate Stromal Organizer Cell Homeostasis during Lymph Node Organogenesis. Blood. 110, 1950-1959.
62. Browning, J. L., Allaire, N., Ngam-Ek, A., Notidis, E., Hunt, J., Perrin, S., and Fava, R. A. (2005) Lymphotoxin-Beta Receptor Signaling is Required for the Homeostatic Control of HEV Differentiation and Function. Immunity. 23, 539-550.
63. Alitalo, K., Tammela, T., and Petrova, T. V. (2005) Lymphangiogenesis in Development and Human Disease. Nature. 438, 946-953.
64. Mouri, Y., Yano, M., Shinzawa, M., Shimo, Y., Hirota, F., Nishikawa, Y., Nii, T., Kiyonari, H., Abe, T., Uehara, H., Izumi, K., Tamada, K., Chen, L., Penninger, J. M., Inoue, J., Akiyama, T., and Matsumoto, M. (2011) Lymphotoxin Signal Promotes Thymic Organogenesis by Eliciting RANK Expression in the Embryonic Thymic Stroma. J Immunol. 186, 5047-5057.
65. Lkhagvasuren, E., Sakata, M., Ohigashi, I., and Takahama, Y. (2013) Lymphotoxin Beta Receptor Regulates the Development of CCL21-Expressing Subset of Postnatal Medullary Thymic Epithelial Cells. J Immunol. 190, 5110-5117.
66. Chin, R. K., Lo, J. C., Kim, O., Blink, S. E., Christiansen, P. A., Peterson, P., Wang, Y., Ware, C., and Fu, Y. X. (2003) Lymphotoxin Pathway Directs Thymic Aire Expression. Nat Immunol. 4, 1121-1127.
67. Boehm, T., Scheu, S., Pfeffer, K., and Bleul, C. C. (2003) Thymic Medullary Epithelial Cell Differentiation, Thymocyte Emigration, and the Control of Autoimmunity Require Lympho-Epithelial Cross Talk Via LTbetaR. J Exp Med. 198, 757-769.
68. Gommerman, J. L., and Browning, J. L. (2003) Lymphotoxin/light, Lymphoid Microenvironments and Autoimmune Disease. Nat Rev Immunol. 3, 642-655.
69. Zindl, C. L., Kim, T. H., Zeng, M., Archambault, A. S., Grayson, M. H., Choi, K., Schreiber, R. D., and Chaplin, D. D. (2009) The Lymphotoxin LTalpha(1)Beta(2) Controls Postnatal and Adult Spleen Marginal Sinus Vascular Structure and Function. Immunity. 30, 408-420.
70. Fu, Y. X., Huang, G., Wang, Y., and Chaplin, D. D. (1998) B Lymphocytes Induce the Formation of Follicular Dendritic Cell Clusters in a Lymphotoxin Alpha-Dependent Fashion. J Exp Med. 187, 1009-1018.
71. Gonzalez, M., Mackay, F., Browning, J. L., Kosco-Vilbois, M. H., and Noelle, R. J. (1998) The Sequential Role of Lymphotoxin and B Cells in the Development of Splenic Follicles. J Exp Med. 187, 997-1007.
72. Milicevic, N. M., Nohroudi, K., Labudovic-Borovic, M., Milicevic, Z., Pfeffer, K., and Westermann, J. (2006) Metallophilic Macrophages are Lacking in the Thymus of Lymphotoxin-Beta Receptor-Deficient Mice. Histochem Cell Biol. 126, 687-693.
73. Silva-Santos, B., Pennington, D. J., and Hayday, A. C. (2005) Lymphotoxin-Mediated Regulation of Gammadelta Cell Differentiation by Alphabeta T Cell Progenitors. Science. 307, 925-928.
74. Kang, H. S., Blink, S. E., Chin, R. K., Lee, Y., Kim, O., Weinstock, J., Waldschmidt, T., Conrad, D., Chen, B., Solway, J., Sperling, A. I., and Fu, Y. X. (2003) Lymphotoxin is Required for Maintaining Physiological Levels of Serum IgE that Minimizes Th1-Mediated Airway Inflammation. J Exp Med. 198, 1643-1652.
75. Summers deLuca, L., Ng, D., Gao, Y., Wortzman, M. E., Watts, T. H., and Gommerman, J. L. (2011) LTbetaR Signaling in Dendritic Cells Induces a Type I IFN Response that is Required for Optimal Clonal Expansion of CD8+ T Cells. Proc Natl Acad Sci U S A. 108, 2046-2051.
76. Lewis, K. L., Caton, M. L., Bogunovic, M., Greter, M., Grajkowska, L. T., Ng, D., Klinakis, A., Charo, I. F., Jung, S., Gommerman, J. L., Ivanov, I. I., Liu, K., Merad, M., and Reizis, B. (2011) Notch2 Receptor Signaling Controls Functional Differentiation of Dendritic Cells in the Spleen and Intestine. Immunity. 35, 780-791.
77. Wu, Q., Wang, Y., Wang, J., Hedgeman, E. O., Browning, J. L., and Fu, Y. X. (1999) The Requirement of Membrane Lymphotoxin for the Presence of Dendritic Cells in Lymphoid Tissues. J Exp Med. 190, 629-638.
78. Chen, M. C., Hsu, T. L., Luh, T. Y., and Hsieh, S. L. (2000) Overexpression of Bcl-2 Enhances LIGHT- and Interferon-Gamma -Mediated Apoptosis in Hep3BT2 Cells. J Biol Chem. 275, 38794-38801.
79. Chang, Y. H., Chao, Y., Hsieh, S. L., and Lin, W. W. (2004) Mechanism of LIGHT/interferon-Gamma-Induced Cell Death in HT-29 Cells. J Cell Biochem. 93, 1188-1202.
80. Browning, J. L., Miatkowski, K., Sizing, I., Griffiths, D., Zafari, M., Benjamin, C. D., Meier, W., and Mackay, F. (1996) Signaling through the Lymphotoxin Beta Receptor Induces the Death of some Adenocarcinoma Tumor Lines. J Exp Med. 183, 867-878.
81. Rooney, I. A., Butrovich, K. D., Glass, A. A., Borboroglu, S., Benedict, C. A., Whitbeck, J. C., Cohen, G. H., Eisenberg, R. J., and Ware, C. F. (2000) The Lymphotoxin-Beta Receptor is Necessary and Sufficient for LIGHT-Mediated Apoptosis of Tumor Cells. J Biol Chem. 275, 14307-14315.
82. Force, W. R., Cheung, T. C., and Ware, C. F. (1997) Dominant Negative Mutants of TRAF3 Reveal an Important Role for the Coiled Coil Domains in Cell Death Signaling by the Lymphotoxin-Beta Receptor. J Biol Chem. 272, 30835-30840.
83. Wimmer, N., Huber, B., Wege, A. K., Barabas, N., Rohrl, J., Pfeffer, K., and Hehlgans, T. (2012) Lymphotoxin-Beta Receptor Activation on Macrophages Ameliorates Acute DSS-Induced Intestinal Inflammation in a TRIM30alpha-Dependent Manner. Mol Immunol. 51, 128-135.
84. Kang, H. S., Chin, R. K., Wang, Y., Yu, P., Wang, J., Newell, K. A., and Fu, Y. X. (2002) Signaling Via LTbetaR on the Lamina Propria Stromal Cells of the Gut is Required for IgA Production. Nat Immunol. 3, 576-582.
85. Fritz, J. H., Rojas, O. L., Simard, N., McCarthy, D. D., Hapfelmeier, S., Rubino, S., Robertson, S. J., Larijani, M., Gosselin, J., Ivanov, I. I., Martin, A., Casellas, R., Philpott, D. J., Girardin, S. E., McCoy, K. D., Macpherson, A. J., Paige, C. J., and Gommerman, J. L. (2011) Acquisition of a Multifunctional IgA+ Plasma Cell Phenotype in the Gut. Nature. 481, 199-203.
86. Barthel, M., Hapfelmeier, S., Quintanilla-Martinez, L., Kremer, M., Rohde, M., Hogardt, M., Pfeffer, K., Russmann, H., and Hardt, W. D. (2003) Pretreatment of Mice with Streptomycin Provides a Salmonella Enterica Serovar Typhimurium Colitis Model that Allows Analysis of both Pathogen and Host. Infect Immun. 71, 2839-2858.
87. Tumanov, A. V., Koroleva, E. P., Guo, X., Wang, Y., Kruglov, A., Nedospasov, S., and Fu, Y. X. (2011) Lymphotoxin Controls the IL-22 Protection Pathway in Gut Innate Lymphoid Cells during Mucosal Pathogen Challenge. Cell Host Microbe. 10, 44-53.
88. Jungbeck, M., Stopfer, P., Bataille, F., Nedospasov, S. A., Mannel, D. N., and Hehlgans, T. (2008) Blocking Lymphotoxin Beta Receptor Signalling Exacerbates Acute DSS-Induced Intestinal Inflammation--Opposite Functions for Surface Lymphotoxin Expressed by T and B Lymphocytes. Mol Immunol. 45, 34-41.
89. Jungbeck, M., Daller, B., Federhofer, J., Wege, A. K., Wimmer, N., Mannel, D. N., and Hehlgans, T. (2009) Neutralization of LIGHT Ameliorates Acute Dextran Sodium Sulphate-Induced Intestinal Inflammation. Immunology. 128, 451-458.
90. Stopfer, P., Obermeier, F., Dunger, N., Falk, W., Farkas, S., Janotta, M., Moller, A., Mannel, D. N., and Hehlgans, T. (2004) Blocking Lymphotoxin-Beta Receptor Activation Diminishes Inflammation Via Reduced Mucosal Addressin Cell Adhesion Molecule-1 (MAdCAM-1) Expression and Leucocyte Margination in Chronic DSS-Induced Colitis. Clin Exp Immunol. 136, 21-29.
91. Lo, J. C., Wang, Y., Tumanov, A. V., Bamji, M., Yao, Z., Reardon, C. A., Getz, G. S., and Fu, Y. X. (2007) Lymphotoxin Beta Receptor-Dependent Control of Lipid Homeostasis. Science. 316, 285-288.
92. Scholz, H., Sandberg, W., Damas, J. K., Smith, C., Andreassen, A. K., Gullestad, L., Froland, S. S., Yndestad, A., Aukrust, P., and Halvorsen, B. (2005) Enhanced Plasma Levels of LIGHT in Unstable Angina: Possible Pathogenic Role in Foam Cell Formation and Thrombosis. Circulation. 112, 2121-2129.
93. Liu, G. Z., Fang, L. B., Hjelmstrom, P., and Gao, X. G. (2008) Enhanced Plasma Levels of LIGHT in Patients with Acute Atherothrombotic Stroke. Acta Neurol Scand. 118, 256-259.
94. Ozaki, K., Ohnishi, Y., Iida, A., Sekine, A., Yamada, R., Tsunoda, T., Sato, H., Sato, H., Hori, M., Nakamura, Y., and Tanaka, T. (2002) Functional SNPs in the Lymphotoxin-Alpha Gene that are Associated with Susceptibility to Myocardial Infarction. Nat Genet. 32, 650-654.
95. Owens, A. W., Matulevicius, S., Rohatgi, A., Ayers, C. R., Das, S. R., Khera, A., McGuire, D. K., and de Lemos, J. A. (2010) Circulating Lymphotoxin Beta Receptor and Atherosclerosis: Observations from the Dallas Heart Study. Atherosclerosis. 212, 601-606.
96. Akhurst, B., Matthews, V., Husk, K., Smyth, M. J., Abraham, L. J., and Yeoh, G. C. (2005) Differential Lymphotoxin-Beta and Interferon Gamma Signaling during Mouse Liver Regeneration Induced by Chronic and Acute Injury. Hepatology. 41, 327-335.
97. Haybaeck, J., Zeller, N., Wolf, M. J., Weber, A., Wagner, U., Kurrer, M. O., Bremer, J., Iezzi, G., Graf, R., Clavien, P. A., Thimme, R., Blum, H., Nedospasov, S. A., Zatloukal, K., Ramzan, M., Ciesek, S., Pietschmann, T., Marche, P. N., Karin, M., Kopf, M., Browning, J. L., Aguzzi, A., and Heikenwalder, M. (2009) A Lymphotoxin-Driven Pathway to Hepatocellular Carcinoma. Cancer Cell. 16, 295-308.
98. Togbe, D., de Sousa, P. L., Fauconnier, M., Boissay, V., Fick, L., Scheu, S., Pfeffer, K., Menard, R., Grau, G. E., Doan, B. T., Beloeil, J. C., Renia, L., Hansen, A. M., Ball, H. J., Hunt, N. H., Ryffel, B., and Quesniaux, V. F. (2008) Both Functional LTbeta Receptor and TNF Receptor 2 are Required for the Development of Experimental Cerebral Malaria. PLoS One. 3, e2608.
|Science Writers||Anne Murray|
|Authors||Bruce Beutler, Jin Huk Choi, Kuan-Wen Wang, Ming Zeng|