AUTOIMMUNITY

Susceptibility to Multiple Sclerosis (MS) has been linked mainly to the HLA-DRB1 locus, with the HLA-DR15 haplotype (DRB1*1501-DQA1*0102-DQB1*0602- DRB5*0101) dominating MS-risk in Caucasians. Alth...

Summary Systemic sclerosis (SSc) is an idiopathic systemic autoimmune disease. It is characterized by a triad of hallmarks: immune dysfunction, fibrosis and vasculopathy. Immune dysfunction in SSc is characterized by the activation and recruitment of immune cells and the production of autoantibodies and cytokines. How immune abnormalities link the fibrosis and vasculopathy in SSc is poorly understood. A plethora of immune cell types are implicated in the immunopathogenesis of SSc, including T cells, B cells, dendritic cells, mast cells and macrophages. How these different cell types interact to contribute to SSc is complicated, and can involve cell‐to‐cell interactions and communication via cytokines, including transforming growth factor (TGF)‐β, interleukin (IL)‐6 and IL‐4. We will attempt to review significant and recent research demonstrating the importance of immune cell regulation in the immunopathogenesis of SSc with a particular focus on fibrosis. Introduction Systemic sclerosis (SSc) is an idiopathic systemic autoimmune disease. The pathogenesis of SSc is mostly unknown. Immunological abnormality, which includes autoimmunity and infiltration and activation of immune cells, is one of three key features of the disease. The other two hallmarks of SSc are fibrosis and vasculopathy. How autoimmunity is linked to fibrosis and vascular damage remains poorly understood, with no single hypothesis or mechanism to explain the links. It is therefore likely that a combination of mechanisms is involved in the pathogenesis of SSc, and picking apart these mechanisms to establish how the triad of fibrosis, vasculopathy and immune cell activation progress in SSc is a mammoth task. Many studies have linked immune abnormality as a cause of or at least a contributor to fibrosis in SSc, while fibrosis may also contribute to immune cell activation. Fibrosis in SSc occurs mainly in the skin but may progress to visceral organs, including heart and lungs. Fibrosis is the result of activation of fibroblasts and excessive extracellular matrix (ECM) deposition, both of which are hallmarks of SSc. As well as fibrosis, immune abnormalities are also linked to vasculopathy in SSc, with most research implicating vascular damage as an activator of immune cells. However, this review will focus mainly on the role of immunological abnormalities in the development of fibrosis in SSc; we recommend the paper by Asano and Sato for a comprehensive review of vasculopathy in SSc 1. Multiple different cell types of the immune system have been implicated in SSc, including T cells, B cells, dendritic cells, mast cells and macrophages. It appears that both innate and adaptive immunity is critical in the disease. We will attempt to review the old and emerging literature to examine the potential roles of these individual cell types in the immunopathogenesis of SSc. First, we will summarize major cytokines implicated in the immunopathogenesis of SSc, because these major players reappear in many studies, linking immune cell activation with other phenotypes in SSc, especially fibrosis. Cytokines in SSc Many cytokines are elevated in SSc, but interleukin (IL)‐4, transforming growth factor (TGF)‐β and IL‐6 are considered to be the major fibrogenic cytokines, or are at least the most studied. Genetic deletion of both IL‐4 and TGF‐β prevents skin fibrosis in a mouse model of SSc 2. Many other in‐vitro studies support the idea that IL‐4 promotes fibrosis through its ability to enhance the production of collagen 3, 4 and other ECM proteins 5, 6 while antibodies against IL‐4 prevent dermal fibrosis in the tight skin (Tsk) mouse model 7 targeted deletion of IL‐4 receptor in the Tsk mouse also reduces fibrosis 2. Skin and lung in SSc have high levels of IL‐4 8 and increased levels of IL‐4 in the blood are a common feature in patients with SSc 9-11 suggesting systemic release. TGF‐β is a well‐known potent inducer of fibrosis, with TGF‐β‐stimulated fibroblasts resembling those from SSc patients 12. Activation of the TGF‐β receptor following the binding of TGF‐β results in the phosphorylation and activation of SMAD proteins in the cytoplasm 13. TGF‐β also activates the three mitogen‐activated protein kinase (MAPK) signalling branches, c‐Jun N‐terminal kinase (JNK), p38 and extracellular signal‐regulated kinases 1 and 2 (ERK1 and 2) 12 all of which can promote inflammatory signalling. TGF‐β‐induced collagen production from both healthy and SSc dermal fibroblasts was found to be dependent on p38 14. JNK activation has also been implicated in fibrosis 15. However, in one study ERK activation inhibited skin fibroblast collagens I and III production while, p38 activation up‐regulated collagen I 16. IL‐6 is a classic proinflammatory cytokine and is also considered to be an important protein in the immunopathogenesis of SSc. For example, IL‐6 levels are increased in SSc patient sera 9 and skin 17. IL‐6 levels also correlate with SSc disease severity 18. A mouse model with development of autoimmune disease with SSc‐like skin thickening and lung fibrosis was found to be mediated by IL‐6 signalling 19. Bleomycin‐induced lung inflammation with collagen deposition was significantly attenuated in IL‐6‐deficient mice 20. IL‐6 signalling through trans‐signalling appears to be important, and we found that IL‐6 and the soluble form of the IL‐6 receptor are necessary for collagen production 21. We further showed in the same study that this was critical, dependent on the downstream signalling molecule signal transducer and activator of transcription (STAT)‐3. A crucial early step hypothesized to trigger the immune abnormalities and fibrosis in SSc is vasculopathy, including the damage and apoptosis of endothelial cells, resulting in the release of internal damage‐associated molecular patterns (DAMPs), which go on to activate and recruit immune cells 22. IL‐6 was found to mediate endothelial activation and apoptosis caused by the serum of patients with SSc 23, suggesting that it may play a major role in the very early stages of SSc. However, IL‐6 was found to be up‐regulated at the late stage of the disease using immunohistological analysis of skin biopsies from SSc patients 17. In both IL‐6 knock‐out (KO) mice and mice exposed to an IL‐6 blocking antibody, bleomycin‐induced dermal fibrosis was greatly induced by supressing fibroblast activation 24. The anti‐IL‐6 receptor antibody tocilizumab has had promising results with softening of the skin in two patients with SSc in one study 25 while a Phase II trial provided SSc patients with improvement in fibrosis of the skin 26 although statistically this was not significant. Thus, IL‐6 antibody therapy could be the first biological licensed for SSc. T cells T cells have been identified early in SSc progression before any evidence of fibrosis 27. SSc skin has a greater propensity to recruit/adhere T cells compared to healthy controls because of a greater expression of intercellular adhesion molecule (ICAM‐1), which is a ligand for the lymphocyte function‐associated antigen 1 (LFA1) receptor found on the surface of lymphocytes such as T cells 28. T cells from SSc skin biopsies have increased expression of the early T cell activation marker CD69 29. TGF‐β, which is elevated in SSc, was also found to be important for the recruitment of T cells to the skin in an SSc mouse model 30. A recent paper demonstrated that abatacept, which is an antibody that interferes with T cell activation, reduced fibrosis in not one, but two animal models of fibrosis 31. This was associated with reduced T cell activation and reduced levels of IL‐6, which may be mediated by blockade of cross‐talk between T cells and antigen‐presenting cells such as monocytes. Abatacept works by blocking the interaction of CD80/86 with cytotoxic T lymphocyte antigen (CTLA)‐4 on T cells, which is required for co‐stimulatory activation of T cells alongside major histocompatibility complex (MHC) and antigen presentation from antigen‐presenting cells such as dendritic cells, macrophages and B cells 32 Abatacept was initiated in three patients with morphea, and all three showed regression of the modified Rodnan skin score 33. T cells can be further classified into diverse subsets characterized by their distinct function, activators and the cytokines they subsequently release. Some of these T cell types have been found to play a role in SSc, which has been reviewed in more detail 34. However, many studies have identified the T helper type 2 (Th2) subtype as playing the largest role in SSc. Th2 cells are a major IL‐4‐producing cell type, but also secrete IL‐5, IL‐6, IL‐10 and IL‐13. Depletion of CD3+ T cells in a bleomycin mouse model reduced both fibrosis and IL‐4 secretion 35. As described above, IL‐4 can directly induce fibroblasts to promote fibrosis, but in addition IL‐4 can induce TGF‐β production in fibroblasts 36 and macrophages 37, which then leads to further signalling to fibroblasts to produce more collagen. Therefore, it is very likely that T cells have a major role to play in the onset of fibrosis in SSc through activation of macrophages and fibroblasts. An unusual population of CD4+CD8+ double‐positive T cells has been described in the skin that has very high levels of IL‐4 38. Gross et al. showed almost 30 years ago that IL‐4 is required to induce naive T cells to produce further IL‐4 39. If IL‐4 is required for T cells to produce IL‐4, this leads to the question: ‘what is the initial source of IL‐4 in SSc?’. This question remains unanswered, but potentially implicates other immune cells in the onset or progression of SSc. IL‐13 is produced predominantly by activated Th2 cells 34. IL‐13 was established as a major profibrotic agent in a model of hepatic fibrosis 40. IL‐13 inhibited IL‐1β‐induced matrix metalloproteinase (MMP)‐1 and MMP‐3 production and enhanced tissue inhibitor of metalloproteinase (TIMP)‐1 and collagen generation in fibroblasts 41. In the bleomycin model of SSc pulmonary fibrosis IL‐13 levels increased with pathogenesis, while neutralization of IL‐13 attenuated bleomycin‐induced pulmonary fibrosis 42. The potent fibrosis‐inducer, TGF‐β, may also contribute to an increase in Th2‐originating IL‐13 because TGF‐β up‐regulates GATA binding protein 3 (GATA‐3) expression in the T cells of patients with SSc, resulting in an increase in IL‐13 synthesis 43. As well as CD4+CD8+ double‐positive T cells, CD8+ single‐positive cells have been described that produce exuberant levels of IL‐13 44. These cells appear to be memory CD8+ T cells 45. We have shown that IL‐13 is directly profibrotic and is STAT‐6‐dependent 46. Thus, IL‐13 produced from T cells probably contributes to fibrosis in SSc and a monoclonal IL‐13 clinical‐grade antibody exists. Very recently, IL‐13 was shown to decrease MMP‐1 expression in both healthy and SSc fibroblasts and therefore may have an anti‐fibrotic role 47. However, this was only demonstrated in tumour necrosis factor (TNF)‐α‐induced fibroblasts, thus the effect of IL‐13 may be specific to the mechanism with which TNF‐α induces MMP‐1 expression. Another cytokine, IL‐17, prevalent in the serum of SSc patients in some studies, also implicates T cells 48. Th17 cells, which are characterized by their production of IL‐17A, IL‐17F, IL‐21 and IL‐22, are elevated in SSc skin compared to healthy controls 49, 50. Thus, Th17 cells and the IL‐17 they produce may play an important role in SSc. However, the roles of Th17 and IL‐17 in SSc are controversial, and have been recently reviewed in detail 51. Importantly, some studies have not detected differences in IL‐17 levels between SSc patients and healthy controls 52, 53 while IL‐17 has been shown to both increase 50 and decrease 54, 55 collagen production. The reasons for the differences in these studies between pro‐ or anti‐fibrotic effects of IL‐17 are not clear, and may reflect the source of the recombinant protein. A reduction in regulatory T cells has been recorded in the skin lesions of patients with SSc 56 suggesting that there is compromised capacity to regulate immune responses in SSc. A recent study identified an imbalance of regulatory T cells and Th17 cells, with a decrease in the former and an increase in the latter in the peripheral blood of SSc patients compared to healthy controls 57. In addition, a more recent study has confirmed that Th2 and Th17 cells are found in higher frequencies in SSc patients, and this was found to positively correlate with IL‐35 levels, although a causal link has not been established 58. Further evidence of disruption to T cell homeostasis comes from a recent study showing that SSc patients displaying severe peripheral vascular complications have an expansion of the recently discovered angiogenic T cells 59. This expansion in angiogenic T cells may also demonstrate a link between immune cell activation and vasculopathy in SSc. Macrophages More than 30 years ago circulating monocytes were found to be strongly activated in patients with SSc 60. Since then, many studies have found evidence of monocyte/macrophage activation in SSc. Higher numbers of macrophages have been observed in skin from SSc patients 27, 61 while cells positive for CD163, a putative marker for M2 macrophages 62, have been found to be increased in the serum of SSc patients 61, 63-65. M2 macrophages are therefore prominent in SSc, with a very recent study involving more than 200 SSc patients, not only confirming this but also demonstrating that soluble CD163 (sCD163) is significantly elevated in the serum of SSc patients and therefore may be of use as a biomarker 66. Microarray gene expression data have also supported a role for monocytes/macrophages in the immunopathogenesis of SSc, with studies demonstrating that peripheral blood mononuclear cells (PBMCs) from SSc patients have an increased expression of genes associated with monocyte/macrophages 67-69. Such large‐scale genomic studies have identified multiple innate immune regulators in SSc. Macrophages are also a major source of TGF‐β 70, which is a potent inducer of fibrosis 71. Many studies have implicated macrophages in the initiation or progression of fibrosis in SSc. Microarray data from SSc lung tissue confirm what has been observed in the blood with markers of macrophage activation 72 and emigration, which correlate with progressive lung fibrosis 73. In another recent study, a novel multi‐network approach to compare gene expression profiles has identified a gene expression signature indicative of profibrotic M2 macrophages in SSc tissues 74. Interestingly, the profibrotic macrophage gene expression profile differed between skin and lung, suggesting that although a role for macrophages in the immunopathogenesis of fibrosis in SSc seems likely in both skin and lung, there may still be subtle differences in what those roles are. Recently nintedanib, an inhibitor of the receptor tyrosine kinase platelet‐derived growth factor receptor, fibroblast growth factor receptor and vascular endothelial growth factor receptor, blocked myofibroblast differentiation and subsequent fibrosis in a mouse model of SSc. Interestingly, these nintedanib‐mediated anti‐fibrotic effects were associated with reduced numbers of M2 macrophages 75. Many chemokines which both recruit and can be primarily secreted by macrophages are up‐regulated in SSc skin 76-79 which is not surprising, given that infiltration of macrophages in SSc skin has been known since the early 1990s 80. However, the chemokine (C‐C motif) ligand 19 (CCL19) was up‐regulated in SSc skin and co‐localized with CD163‐positive macrophages, suggesting that it has a role in the recruitment of macrophages 76. The authors of the study also suggest that macrophages are the source of the CCL19, and in‐vivo experiments demonstrate that Toll‐like receptor (TLR)‐3, ‐4 or ‐9 activation is required for CCL19 expression in monocytes. TLR activation is considered a major event in the immunopathogenesis of SSc 81, 82. Cell types which highly express TLRs such as macrophages are thought to play a major role in SSc. However, many different cell types express TLRs, and thus TLR‐mediated signalling in the immunopathogenesis of SSc may not be limited to macrophages. It is possible that TLR activation in any cell type expressing TLRs is also a source of CCL19, and therefore a potential cause of macrophage recruitment in SSc. Indeed, Mathes et al. also observed that CCL19 expression was up‐regulated from TLR activation in T cells and other cell types from PBMCs, but monocytes gave the most robust increase 76. We have found that macrophages that respond to the TLR‐8 stimulus single‐stranded RNA in SSc result in up‐regulation of TIMP‐1 that is functional and leads to increased collagen deposition 83. Interestingly, these monocytes seemed to be perivascular. IL‐6 appears to play an important part in the role of macrophages in SSc because inhibition of phosphodiesterase‐4, which blocks M2 differentiation, also inhibits IL‐6 production, fibroblast activation and fibrosis in an SSc mouse model 84. IL‐6 may also be an important activator of M2 macrophages, as indicated by IL‐6 receptor blockage by tocilizumab 85 causing down‐regulation of genes associated with M2 macrophages in SSc skin. Oncostatin M is another IL‐6‐like cytokine that appears to be involved in fibrosis, as cells treated with OSM induced ECM accumulation in fibroblasts 86. SSc patients with higher expression of the M2 macrophage marker, MRC1, were also found to have elevated levels of IL‐13 in their plasma 69 suggesting that macrophage activation in SSc results in production of cytokines from macrophages themselves or indirectly through interaction with other immune cells, because macrophages can both secrete IL‐13 87 and, through being antigen‐presenting cells, can activate T cells which could lead to IL‐13 production. There have been more than 30 years of research implicating macrophages in the immunopathogenesis of SSc, with various cytokines produced by macrophages and other immune cells, internal molecules from damaged endothelial cells and chemokines all contributing to the recruitment and activation of macrophages, resulting in a profibrotic environment (Fig. 1). However, how macrophages are recruited to and activated in the tissues of SSc patients is not fully understood. What role is played by macrophages in the progression of fibrosis is also not fully known, but activation of TLRs and the subsequent profibrotic signalling remains an obvious choice. Targeting of TLRs may be a therapeutic option. B cells B cells are heavily implicated in the immunopathogenesis of SSc. B cells are known inducers of fibrosis generally and in SSc 88 and mounting evidence suggests that they are modified and activated in patients with SSc. SSc patients have abnormalities of B cell homeostasis in the blood, which includes an expansion of naive B cells and activated but diminished memory B cells 89. The cytokine B cell activating factor (BAFF), which belongs to the TNF ligand family, is a potent activator of B cells 90. Not only are BAFF levels elevated in SSc; levels also correlate with disease severity, while B cells isolated from SSc patients produce more IL‐6 when exposed to BAFF 91. A BAFF antagonist inhibited IL‐6 and IL‐10 expression in the skin of the Tsk mouse model of SSc, while stimulation of B cells with BAFF greatly increased IL‐6 92. IL‐6 can direct the differentiation of T cells into IL‐4‐producing Th2 cells 93, therefore B cell‐secreted IL‐6 could contribute to the Th2 phenotype detected in SSc. As well as being involved in IL‐6 cytokine production, B cells are also suggested to play an important role in fibrosis. Co‐cultures of B cells and fibroblasts from SSc patients induced a fibrotic response, including collagen, TGF‐β1 and IL‐6 secretion. Exposure of co‐cultures to BAFF (and anti‐IgM) further increased secretion of the profibrotic compounds 88. The majority of B cells are IL‐6‐producing, termed B effector cells (Beffs), and have a proinflammatory and autoimmunity role. However, a small subset of B cells termed regulatory B cells (Bregs) are potent negative regulators of inflammation and autoimmunity, in part through their ability to express IL‐10 94. These Bregs therefore have the potential to inhibit diseases with immune abnormalities such as SSc. Indeed, Bregs were able to inhibit the initiation of a mouse model of multiple sclerosis and depletion of Bregs increased symptom severity 95. Interestingly, the frequency of blood Bregs was found to be significantly lower in SSc patients compared to healthy controls 96 while in a very recent study by Matsushita et al., Bregs were found to have a similar role in a mouse model of SSc. Depletion of IL‐6‐producing Beffs reduced skin and lung fibrosis, while depletion of IL‐10‐producing Bregs caused more severe fibrosis 97 backing up findings from a previous study that Bregs can suppress skin fibrosis in a graft‐versus‐host disease mouse model of SSc 98. A tipping of the balance of Bregs and Beffs in favour of fewer Bregs is therefore a potential important event in the initiation of fibrosis in SSc. However, the cause of this shift in B cell homeostasis remains unknown, and future studies to identify the trigger/s may provide useful early therapeutic targets. Various response regulators on the surface of B cells exist, with some increasing and others decreasing B cell receptor (BCR) signals. Of these, the positive B cell regulator, CD19, is the most established in SSc disease research. CD19 is an important regulatory molecule expressed by B cells. CD19+ B cells increase in SSc, with expression of CD19 on the surface of B cells significantly higher in SSc patients compared to healthy controls 99. CD19‐deficient mice have decreased sera levels of several autoantibodies 100 suggesting that B cells could be responsible for the autoantibodies detected in SSc patients. In agreement, a transgenic mouse model, with an elevation in CD19 expression similar to SSc human patients, shows an increase in the levels of autoantibodies 101, 102. It is also worth noting that CD19 over‐expression has not been observed in the Tsk mouse model, suggesting that a role for CD19 in B cell activation in SSc is not straightforward or that the animal models can only replicate the human disease phenotypes to a degree. However, the apparent lack of CD19 over‐expression may be explained partly through constitutive activation of CD19 and the downstream signalling pathway via up‐regulated CD19 tyrosine phosphorylation, which could negate the need for over‐expression 102. Autoantibody production is prevalent, and can be used diagnostically in SSc. Furthermore, autoantibodies against platelet‐derived growth factor receptor (PDGFR) were found in SSc patients, and caused collagen I‐increased expression and transition of normal human primary fibroblasts to a myofibroblast phenotype through activation of an intracellular loop that involves Ha‐Ras, ERK1/2 and reactive oxygen species (ROS) 103. However, it is still possible that B cell activation is the cause of both autoantibody production and fibrosis, as the two may be linked. Interestingly, B cell activation via CD19 specifically may also be a cause of other immune cell type infiltration in SSc. CD19‐deficient mice showed less infiltration of mast cells, macrophages, T cells and B cells after bleomycin treatment compared to wild‐type cells exposed to bleomycin 104. In the same study, it was discovered that B cells, via hyaluronan‐induced TLR‐4 activation, produced various cytokines which were dramatically reduced in CD19‐deficient cells. Therefore, B cells have the potential to contribute to fibrosis directly as well as indirectly through recruitment and activation of other immune cells to promote a cycle of cytokine and fibrogenic compound release. Mast cells Mast cell infiltration was detected from lip biopsy in very early stages of SSc and preceded the onset of skin changes 105 and more recently cutaneous mastocytosis was found to precede the onset of SSc in a 36‐year‐old woman 106 suggesting that mast cells may play an early role in the onset of SSc. Mast cell infiltration has also been detected late in the development of SSc 107 and in a subset of SSc patients with localized scleroderma‐like lesions 108. An increase in mast cell numbers has also been detected in SSc skin 109. Mast cells are able to produce TGF‐β and are found localized to fibroblasts in the skin of SSc patients 110. Interestingly, mast cells are not only a source of TGF‐β; they can also directly transfer TGF‐β to fibroblasts mediated by transgranulation via cell–cell contacts 111, which has also been observed in the dermis of SSc patients 112. The triggers of mast infiltration in SSc or other fibrotic diseases are not known, but a very recent study has identified Snail‐dependent production of plasminogen activator inhibitor‐1 (PAI1) in keratinocytes as a chemoattractant of mast cells, a mediator of mast cell–fibroblast adhesion and a promoter of fibrogenesis 113. Interestingly, PAI1 up‐regulated tenascin‐C (TENC) expression, which is the main activator of TLR‐4 to promote fibrosis and found to be elevated in SSc 114. Therefore, the role of PAI1 in SSc could involve mast cell recruitment and activation of fibroblasts to promote an inflammatory environment. Dendritic cells Two major types of dendritic cell are known: conventional (cDC) and plasmacytoid (pDC). pDC are a specialized cell type that when activated produce large amounts of interferon (IFN). A very recent study observed that pDCs infiltrate the skin of SSc patients and are chronically activated, producing IFN‐α and chemokine (C‐X‐C motif) ligand 4 (CXCL4). Fibrosis was reverted in a mouse model of SSc after pDCs depletion 115. In the same study, TLR‐8 signalling was found to be important for CXCL4 production and recruitment of pDCs to fibrotic skin. TLR involvement has also been found to be important for mediating the increased cytokine production in DCs in SSc, which has been comprehensively reviewed 116. pDCs isolated from SSc patients show up‐regulation of CXCL4 115, 117, which can also be detected at elevated levels and correlates with disease severity in SSc patient plasma 118 and has been suggested as a biomarker in SSc. In another recent study, pDC depletion in a bleomycin mouse model improved the clinical score, lung histopathology score, skin thickness and collagen content. The expression of genes involved in chemotaxis, dendritic cell differentiation, inflammation and fibrosis in the lungs of pDC‐depleted mice was also significantly reduced. Alongside this, B and T cells were also found to be reduced in the lungs, suggesting an important role for ongoing immune abnormalities induced by DCs 119. In another bleomycin‐induced pulmonary fibrosis mouse model, DC accumulation in the lung was reduced through blocking of TGF‐β with the use of inhibitor SB431542 120. TGF‐β was also found to be important for the recruitment of pDCs (and T cells) to the skin in an SSc mouse model 30 suggesting an important role for TGF‐β in DC location and activation. An important role for microRNAs in the pathogenesis SSc is emerging 121. Recently, microRNA expression profiling has identified miRNA‐618 (miR‐618) as being up‐regulated in pDCs from SSc patients, while over‐expression in healthy pDCs resulted in an SSc‐like pDC 122. Thus, microRNAs are disturbed, leading to altered functionality of immune cells. Alterations of microRNAs by restoration using microRNA therapeutics may be one way to reset dendritic cells to a ‘tolerogenic’ phenotype. Anti‐nuclear antibodies, which are a subset of autoantibodies, are found to be elevated in SSc patients 123. DNA topoisomerase I (TopoI) is an intracellular target of anti‐nuclear antibody anti‐TopoI. The presence of anti‐TopoI antibodies has been clinically associated with a more severe form of SSc 124. Immunization of mice with TopoI peptide‐loaded DCs induces anti‐TopoI autoantibody response and long‐term fibrosis of skin and lung, which is preceded by inflammation with increased IL‐17A and CXCL4 expression 125. TopoI peptide‐loaded DCs also induce proliferation of SSc and healthy‐derived T cells, but activation of T cells requires a fragmented form of TopoI and activation of full‐length TopoI is IL‐2‐dependent 126. Interestingly, antigen‐presenting cells within PBMCs were able to activate T cells more efficiently than dendritic cells, and even processed TopoI peptides differently, suggesting that there is a complex regulation of T cells by dendritic cells and other antigen‐presenting cells with regard to anti‐nuclear antibodies. Overall, experimental manipulation of DCS can mimic some major events in the pathogenesis of SSc, and therefore raises the intriguing prospect that DCs may be capable of initiating SSc. A broken loop that may perpetuate inflammation with cytokine‐mediated up‐regulation of co‐stimulatory molecules may also exist in SSc. Other immune cells Dysregulation and activation of immune cells is clearly a major hallmark of SSc involving many cell types. This complex network of signalling between different immune cells is only now beginning to become unravelled, with new evidence emerging regularly to provide new players in SSc immunopathogenesis. For example, only a few studies have highlighted the potential for platelets 127-129, neutrophils 130, 131, natural killer cells 132, 133 and innate lymphoid cells 134, 135 to be dysregulated and contribute to the pathogenesis of SSc. Type 2 innate lymphoid cells are found in higher numbers in SSc and correlate strongly with the Rodnan skin score and also the presence of lung fibrosis. Although not considered to be innate immune cells, fibroblasts play a crucial role in the fibrotic phenotype of SSc, but they also express TLRs and are therefore capable of responding to DAMPs. Expression of TLR‐2 was found to be increased in fibroblasts from SSc skin 136. Recently, fibroblasts exposed to TGF‐β and IL‐17A responded with a 100‐fold increase in the production of IL‐6 137, a known profibrotic mediator. Conclusion Various cell types of the immune system appear to be involved to some degree in the immunopathogenesis of SSc through promoting other immune cell activation, fibrosis or vascular damage. Very recent studies have highlighted the importance of this plethora of immune cell types in SSc (Table 1). However, research to date provides evidence for macrophages, T cells and B cells having the most important role in SSc. Whether this accurately represents their importance in SSc or is a result of research interests combined with the practicality of investigating specific cell types remains to be seen. Nevertheless, immune cell activation is a recurring observation in SSc research and is leading to the development and testing of therapeutic interventions. Although the over‐production of cytokines is a well‐accepted symptom of SSc, TGF‐β, IL‐6 and IL‐4 specifically seem to be integral to many of the immune abnormalities recorded. Thoroughly understanding how these and other cytokines fully regulate, and are regulated by, all the different immune cells and their subcategories in SSc will be a difficult task, but every study into immune cell activation, regardless of cell type, brings us closer to achieving it and as a result developing therapeutic interventions. Current trials such as the Fassinate trial in SSc blocking IL‐6 seem promising, with the Phase II trial positive, although the primary end‐point was not reached. Phosphodiesterase type 4 (PDE4) inhibitors appear to regulate macrophages leading to reduced fibroblast activation. There is an ongoing trial concerning morphea. Immune cell Recent findings Reference T cell Angiogenic T cells were elevated in SSc patients displaying digital ulcers, which is a severe peripheral vascular complication 59 Macrophage The soluble form of the M2 macrophage marker CD163 is elevated in the serum of patients with SSc, highlighting it as a potential biomarker 66 B cell Used a mouse model of SSc to demonstrate the importance of B cell homeostasis. Depletion of IL‐6‐producing B effector cells reduced fibrosis while depleting of IL‐10‐producing B regulatory cells had the opposite effect 97 Mast cell A subset of SSc patients with localized scleroderma‐like lesions were found to have an inflammatory phenotype leading to the activation of mast cells in the dermis of mechanically stressed skin 108 Dendritic cell After depletion of pDCs fibrosis was reverted in mice with established SSc‐like disease. pDC depletion prior to induction of disease also prevented fibrosis 115 Platelets Activated platelets induced the production of the profibrotic mediator thymic stromal lymphopoietin (TSLP) in human dermal endothelial cells 128 Neutrophils Neutrophil activation was induced by SSc microparticles. Microparticles were derived from platelets and expressed the damage‐associated molecular pattern HMGB1. An inhibitor of HMGB1 attenuated neutrophil activation 131 Natural killer A peculiar natural killer cell phenotype in SSc patients was identified characterized by decreased chemokine and activation receptors expression. These SSc‐derived natural killer cells were potent inducers of endothelial microparticle release suggesting that there may be a role for natural killer cells in the activation of endothelial cells in SSc 133 Innate lymphoid Type 2 innate lymphoid (ILC2) cells are elevated in patients with SSc. ILC2 counts correlated with skin fibrosis. This study highlights that there may be a profibrotic role for ILC2 cells in SSc 134 Fibroblast Although not an immune cell type, fibroblasts exposed to TGF‐β and IL‐17a responded with a 100‐fold increase in the production of IL‐6 137 DCs = dendritic cells; IL = interleukin; TGF = transforming growth factor; HMGB1 = high mobility group box 1. Disclosures None. References

Abstract Anti‐Ro52 autoantibodies (Ro52‐autoAbs) appear in the sera of connective tissue disease (CTD) patients with interstitial lung disease (ILD). Studies using patient sera have shown a correlation between the generation of Ro52‐autoAbs and the clinical morbidity and severity of CTD with ILD. In this study, we used a single B‐cell manipulating technology and obtained 12 different monoclonal Ro52‐autoAbs (mRo52‐autoAbs) from the selected four patients suffering from severe ILD with a high titer of Ro52‐autoAbs in their sera. Western blot analysis revealed that 11 of 12 mRo52‐autoAbs bound to the coiled‐coil domain of Ro52. Competitive ELISA demonstrated that mRo52‐autoAbs competed with each other to bind to Ro52. Epitope mapping showed that two of them specifically bound to a peptide (PEP08) in the coiled‐coil domain. We then examined the titer of Ro52‐autoAbs in the sera of 192 CTD patients and assessed the relationship between the serum levels of Ro52‐autoAbs that were reactive to PEP08 peptide and the clinical morbidity and severity of ILD. Statistical analysis revealed that the production of PEP08‐reactive Ro52‐autoAbs correlated with the morbidity and severity of ILD in CTD. Assessment of the production of PEP08‐reactive Ro52‐autoAbs in autoimmune diseases is useful for predicting the clinical morbidity of ILD. Introduction Ro52/TRIM21 is known as an E3 ligase involved in the process of ubiquitination and plays a prominent regulatory role in inflammation 1-7. Ro52 is the common target of autoantibodies in connective tissue diseases (CTDs) 8, 9, such as systemic lupus erythematosus (SLE) 10-12, Sjögren's syndrome (SS) 13-17, polymyositis/dermatomyositis (PM/DM) 18, 19, and systemic sclerosis (SSc) 20. Ro52‐specific autoantibodies (Ro52‐autoAbs) have been detected in the sera of many patients with various CTDs 9, 21. Some of these patients develop severe interstitial lung disease (ILD), and several reports have indicated that the production of Ro52‐autoAbs in patient sera is related to the incidence of ILD in CTD 22, 23. In SSc patients, the generation of Ro52‐autoAbs has been correlated to the morbidity and severity of ILD 23. In mixed connective tissue disease (MCTD) patients, Ro52‐autoAbs have been detected in 50% of patients with lung fibrosis 22. The Ro52 protein consists of multi‐domains, including an N‐terminal RING finger domain with E3 ubiquitin ligase activity, a B‐box domain, a coiled‐coil (C‐C) domain involved in oligomerization, and the C‐terminal PRYSPRY domain that commonly determines functions by acting as a target 24, 25. Some epitopes of Ro52‐autoAbs in SLE and SS patients have been reported 26, most of which reside in the C‐C domain of Ro52 27-34. The ring and PRYSPRY domains are also targets of Ro52‐autoAbs 29, 35. These epitopes were determined in patient sera using polyclonal antibodies. Concerning monoclonal Ro52‐autoAbs (mRo52‐autoAbs), Mietzner et al. generated monoclonal antibodies (mAbs) from IgG memory B cells of SLE patients and showed that some of them reacted to Ro52 36. However, there have been no reports of epitopes for human mRo52‐autoAbs produced in patients with ILD. In this study, we used our single cell analysis method denoted the immunospot array assay on a chip (ISAAC) 37, 38 to obtain mRo52‐autoAbs from the PBMCs of four DM patients suffering from severe ILD. We obtained twelve human mRo52‐autoAbs and showed that eleven of them bound to the C‐C domain of Ro52. We further determined an epitope for two of these mRo52‐autoAbs. Finally, we examined the relationship between the production of Ro52‐autoAbs that were reactive to the determined epitope in patient sera as well as the clinical morbidity and severity of ILD in CTD. Results Isolation of mRo52‐autoAbs from DM patients with severe ILD using the ISAAC system First, we examined 192 CTD patients to determine the titer of Ro52‐autoAbs in their sera by EUROLINE scan. We also examined them by scanning high‐resolution computed tomography (HRCT) and found that 57% (109/192) of these patients had ILD. The results agreed with previous studies 22, 23. We then tried to analyze the correlation of Ro52‐autoAb‐production and ILD. To achieve this goal, we tried to obtain mRo52‐autoAbs from DM patients because they are frequently complicated with ILD 39. We randomly selected 4 DM patients with severe ILD who also suffered from anti‐synthetase syndrome and had anti‐SS‐A, anti‐Jo‐1, or anti‐PL‐7 autoantibodies as well as Ro52‐autoAbs in their sera (Supporting Information Table 1). One hundred six antibody‐secreting cells (ASCs) that produced Ro52‐autoAbs were detected in the PBMCs of these four DM patients using ISAAC and retrieved from the chip (Fig. 1A). The antibody cDNAs were cloned from the single cells, and mAbs were produced. Fifteen recombinant mAbs specific to recombinant Ro52 (rRo52) protein were obtained. Sequence analysis revealed that three of 15 mRo52‐autoAbs had the same amino acid sequence as another clone, which indicated that these clones were derived from the same B‐cell clone (Table 1). We eventually obtained 12 different mRo52‐autoAbs from four DM patients with ILD (Fig. 1B). Heavy‐chain Light‐chain Pt. No. Name of mRo52‐autoAb Subclass V D J CDR3‐IMGT Family V J CDR3‐IMGT 100,112 110,112‐20 IgG1 IGHV3 IGHD6 IGHJ4 AKDWAQFVLRYFFDH κ IGKV1 IGKJ1 QQYDSYPWT 100,112 110,112‐19a IgG1 IGHV3 IGHD6 IGHJ4 AKDWAQFVLRYFFDH κ IGKV1 IGKJ1 QQYDSYPWT 100,112 110,112‐26 IgG1 IGHV7 IGHD6 IGHJ4 ARQGMFSTGWYLAH κ IGKV1 IGKJ1 QNYKSYSRT 100,112 110,112‐29 IgG1 IGHV7 IGHD4 IGHJ5 ARGALTMPTRVDP κ IGKV1 IGKJ5 QQTYSTLPIT 55 55‐37 IgG1 IGHV1 IGHD3 IGHJ4 ARSFYFGSGSYPY κ IGKV3 IGKJ2 QQYGSSPYT 55 55‐34a IgG1 IGHV1 IGHD3 IGHJ4 ARSFYFGSGSYPY κ IGKV3 IGKJ2 QQYGSSPYT 150 150‐08 IgG1 IGHV3 IGHD1 IGHJ4 ARGKSWLDY κ IGKV3 IGKJ4 QQYGSSLT 150 150‐14 IgG1 IGHV1 IGHD2 IGHJ5 ARGVYRRSGGNSWGFDP κ IGKV2 IGKJ4 MQALQTPPT 150 150‐42 IgG1 IGHV3 IGHD1 IGHJ4 ARDLGGGRRIYYFDY κ IGKV3 IGKJ4 QQYGSSPL 150 150‐44 IgG1 IGHV3 IGHD3 IGHJ4 AKVIGITFGGVIFPFDY κ IGKV3 IGKJ1 QQYGTAPT 150 150‐45 IgG1 IGHV3 IGHD3 IGHJ4 AKGFGTTFGGVIFPFDY κ IGKV3 IGKJ1 QQYSTSPT 150 150‐46 IgG1 IGHV3 IGHD5 IGHJ4 ARGKYWLDY κ IGKV3 IGKJ4 QQYGSSLT 150 150‐60 IgG1 IGHV3 IGHD4 IGHJ3 ARARFRTTVTTYAFDI κ IGKV2 IGKJ1 MQALQTPPWT 150 150‐61 IgG1 IGHV3 IGHD2 IGHJ4 ARERGGFCSGGSCYSCDY κ IGKV2 IGKJ1 MQALQTPRT 150 150‐58a IgG1 IGHV3 IGHD2 IGHJ4 ARERGGFCSGGSCYSCDY κ IGKV2 IGKJ1 MQALQTPRT a same as the above‐mentioned mRo52‐autoAbs were obtained PBMCs from 4 patients (#55, #100, #112, and #150). PBMCs of 2 patients (#100, #112) were mixed since the number of PBMCs of these patients were too small for ISAAC. 15 mAbs were cloned and found to bind rRo52 protein in which 3 mAbs had the same amino acid sequence. Final number of mRo52‐autoAbs became 12. V, D, J gene segments and CDR3‐IMGT in heavy chain as well as V, J gene segments and CDR3‐IMGT in light chain are shown. Epitope mapping To determine the epitopes of mRo52‐autoAbs, we first synthesized the various truncated fragments of rRo52 and then examined the reactivity of each mRo52‐autoAb to the fragments (Fig. 2A and B). Eleven of 12 mRo52‐autoAbs reacted with the C‐C domain as well as the full‐length rRo52 protein. One mRo52‐autoAb (Ab100, 112–20) reacted with the full‐length protein but not with any of the other fragments. We then examined whether these eleven mRo52‐autoAbs competed for binding to a specific site of the C‐C domain using competitive ELISA as previously reported 40. mRo52‐autoAbs together with one of the peroxidase (POD)‐conjugated mRo52‐autoAbs were added to wells that were coated with rRo52 proteins. The bound enzyme activity was then measured after washing the samples. As shown in Fig. 3A, all eleven mRo52‐autoAbs, but not the control mAb, inhibited the binding of POD‐conjugated Ab55‐37 mAb. Similarly, all 11 mRo52‐autoAbs inhibited the binding of other POD‐conjugated mRo52‐autoAbs (Ab150‐61, Ab150‐42, and Ab150‐45). These results suggest that the mRo52‐autoAbs that were obtained from DM patients with severe ILD (DM‐ILD patients) bind to the same or juxtaposed epitopes of Ro52. We then synthesized the peptide library of the C‐C domain of Ro52 (Fig. 3B and Supporting Information Table 2) to determine the precise epitope for mRo52‐autoAbs and examined the binding of mRo52‐autoAbs to these peptides. Two of 11 mRo52‐autoAbs (Ab55‐37 and Ab150‐61) were used for the binding assay because they showed clear binding activity to the C‐C domain (Fig. 2B). Those two mRo52‐autoAbs reacted with a peptide (PEP08, RIHAEFVQQKNFLVEE) but not with the other peptides of the C‐C domain or the control HIV‐env peptide (Fig. 3C). It is notable that these two mRo52‐autoAbs were obtained from PBMCs of different patients (Table 1). These data show that PEP08 contains an epitope of the mRo52‐autoAbs. These data, together with the competitive‐binding data (Fig. 3) indicate that all the mRo52‐autoAbs obtained by the ISAAC system bind to this epitope or a juxtaposed epitope. Relationship between the production of PEP08‐reactive Ro52‐autoAb and the morbidity of ILD in CTD As described above, the production of Ro52‐autoAb was reported to be clinically associated with ILD in SSc and MCTD 22, 23. To investigate the relationship between the production of the Ro52‐autoAbs that bind to PEP08 in the sera of CTD patients and the incidence and severity of ILD, 192 CTD patients were selected from our hospital whose Ro52‐autoAb titer was examined by EUROLINE scan (Fig. 4A). Overall, 43% (82/192) of them were positive for Ro52‐autoAb, while the rest (110/192) were negative. The reactivity of Ro52‐autoAb sero‐positive sera to PEP08 was analyzed. The results showed that the Ro52‐autoAb from 45% (37/82) of the patients bound to PEP08, whereas that from 55% (45/82) of the patients did not (Fig. 4A and B). All the profiles of PEP08‐Ab‐positive, PEP08‐Ab‐negative, and Ro52‐autoAb‐negative patients with CTD are shown in Supporting Information Table 3. We then compared the morbidity and severity of ILD in 38 patients in the PEP08‐Ab‐positive group and 154 patients in the PEP08‐Ab‐negative group using an odds ratio and Fisher's exact test. In these analyses, we assumed that 110 patients who were Ro52‐autoAb‐negative were negative for PEP08‐Ab (Fig. 4A). The assumption was confirmed by analyzing 14 Ro52‐autoAb‐negative patients in Supporting Information Table 3. They were also negative for PEP08‐Ab (Fig. 4C). For the morbidity of ILD, 30/38 (79%) patients in the PEP08‐Ab‐positive group had ILD. In contrast, 80/154 (52%) patients in the PEP08‐Ab‐negative group had ILD (Fig. 4A and D, odds ratio = 3.47, P = 0.0031, CI: 1.50–8.05). We also compared the severity of ILD in the PEP08‐Ab‐positive group and the PEP08‐Ab‐negative group using the KL‐6 value that had been considered to be an indication of severity of ILD 41, 42.Overall, 26/33 (79%) patients in the PEP08‐Ab‐positive group had a high KL‐6 value (KL‐6 ≥ 500). In contrast, 82/143 (57%) patients in the PEP08‐Ab‐negative group had a high KL‐6 value (KL‐6 ≥ 500) (Fig. 4A and E, odds ratio = 2.65, P = 0.0285, CI: 1.13–6.78). There was no correlation between the titer of PEP08‐Ab and the KL‐6 value (Supporting Information Fig. 1). The results indicate that PEP08‐Ab‐positive CTD patients had a higher indication of morbidity and severity of ILD than PEP08‐Ab‐negative CTD patients. Discussion Ro52 is the common target of autoAbs in CTD, and the production of Ro52‐autoAbs in patient sera has been reported to be related to the incidence of ILD with CTD 22, 23; however, there have been no reports concerning the epitopes of Ro52 protein related to ILD. To gain an understanding of the relationship of the Ro52‐autoAb epitopes and ILD, we obtained 12 human mRo52‐autoAbs from peripheral blood B cells of DM‐ILD patients (Table 1 and Fig. 1) and determined their antigenic epitopes. We found that 11 of them reacted with the C‐C domain of the Ro52 protein (Fig. 2). Previous studies have also demonstrated that Ro52‐autoAbs reacts with the C‐C domain using serum of SS or SLE patients 27-34. The C‐C domain plays an important role in protein–protein interactions and is usually protruding from the main body of proteins. Thus, it tends to be recognized by B cells using their antigen receptors on the cell surface. We could not determine the epitopes of one mRo52‐autoAb, Ab100/112‐20, which reacted only with the full‐length protein, indicating that it may recognize the discontinuous conformational epitope formed by the two domains. In contrast, the remaining mRo52‐autoAbs may bind to linear epitopes since they recognized the antigen in immunoblotting with SDS‐denatured and linearized proteins. However, we could not exclude the possibility that they recognized refolded proteins during immunoblotting. Regarding the epitopes of Ro52‐autoAbs in various CTDs, their epitopes have been reported to be distributed in various rRo52 fragments (Supporting Information Table 4). In this study, by competitive ELISA (Fig. 3A) and epitope mapping using a peptide library (Fig. 3B and C), we found that two mRo52‐autoAbs from DM‐ILD patients bound to a Ro52 epitope (PEP08), and the remaining nine Abs bound to its juxtaposed epitopes. Interestingly, those two mRo52‐autoAbs were obtained from different patients. This outcome posed a strong possibility that PEP08 is one of the important epitopes for ILD in CTD, although we must consider that mRo52‐autoAbs obtained from DM patients may have biased the results. Wahlen et al. reported an epitope of Ro52‐autoAbs that was found to show production correlated with the onset of congenital heart block (CHB) named “P200” (aa200–239) 43. The maternal mRo52‐autoAbs of CHB patients did not react to the peptide named “P176” (aa176‐196). In contrast, our mRo52‐autoAbs from DM‐ILD patients reacted with the PEP08 (aa177‐192) involved in P176. Although all these peptides were included in the C‐C domain (aa133‐255), each epitope may play a different role in oligomerization and might be involved in the onset of different CTD. Concerning the functional role of Ro52‐autoAbs in the diseases, we investigated how the generation of PEP08‐specific Ro52‐autoAbs in the CTD patients affected the incidence and severity of ILD (Fig. 4). We showed that the PEP08‐Ab‐positive group had a higher morbidity of ILD (odds ratio = 3.47, p = 0.0031, CI: 1.50–8.05) (Fig. 4D). The group also exhibited a significant relationship to the severity of ILD (odds ratio = 2.76, p = 0.0285, CI: 1.13–6.78) (Fig. 4E). The results showed that the production of PEP08‐reactive Ro52‐autoAbs influenced the morbidity of ILD in CTD. Although the molecular mechanisms in which Ro52‐autoAbs induce a high risk of ILD are not known, further analysis of the in vivo effect of mRo52‐autoAbs using an animal model may facilitate further understanding of involvement of PEP08‐reactive Ro52‐autoAbs to the onset of ILD in CTD. This study has the following limitations: (1) The titer of KL‐6 was not assessed for some patients who did not develop ILD, as shown in Supporting Information Table 3). The epitopes of Ro52‐autoAbs produced in various CTDs were distributed on various Ro52 fragments (Supporting Information Table 4). Here, we determined the PEP08 epitope using mRo52‐autoAbs from DM‐ILD patients. Thus, the selection of DM patients may have biased the results. (3) In this study, we focused on anti‐Ro52 autoantibodies of CTD patients and investigated the clinical correlation of a mRo52‐autoAbs epitope to ILD. Although PEP08‐Ab‐positive CTD patients had a higher indication of morbidity and severity of ILD than PEP08‐Ab‐negative CTD patients, it remains unknown why many PEP08‐negative patients still had ILD and a high KL‐6 value. For the onset of ILD, multiple risk factors 41 should be considered to be involved. One of the risk factors is autoantibodies; however, it has been reported that ILD‐related autoantibodies are not only anti‐Ro52 autoantibodies but also anti‐PL‐7 and anti‐Jo‐1 autoantibodies 39, 44, among others. Thus, analysis of the involvement of anti‐PL‐7 and anti‐Jo‐1 autoantibodies in ILD may contribute to understanding the morbidity and severity of many PEP08‐negative patients. It was recently reported that the production of Ro52‐autoAbs in patient sera were related to undifferentiated connective tissue diseases (UCTD) 45, 46. Measurement of PEP08‐Ab in sera of UCTD patients may contribute to the prediction of ILD. In summary, we obtained twelve mRo52‐autoAbs from four DM‐ILD patients. Eleven of these mRo52‐autoAbs bound to the C‐C domain in immunoblotting, and they competed with each other to bind to the C‐C domain. We determined the binding site of mRo52‐autoAb using the peptide library and analyzed the relationship of the production of PEP08‐Ab and the clinical morbidity and severity of ILD. We showed that the PEP08‐Ab‐positive patients exhibited a higher incidence risk of ILD than PEP08‐Ab‐negative patients. Future studies using the mRo52‐autoAbs in vivo system may be required to understand the pathological function of Ro52‐Abs in ILD with CTD. Materials and methods Patients and their characterization Patients who visited the First Department of Internal Medicine at Toyama University Hospital and were diagnosed with CTD were included in the study (Fig. 4). Informed consent was obtained from all patients in accordance with a protocol approved by the Ethical Review Board of the University of Toyama. The characteristics of the patients at the time of enrollment are summarized in Supporting Information Table 3. The patients included 145 females (76%) and 47 males (24%). Their median age was 63 years. Sixty patients (31%) had PM/DM, 44 (23%) had SSc, 40 (21%) had rheumatoid arthritis, 17 (9%) had SS, 13 (7%) had MCTD, 10 (5%) had SLE, 6 (3%) had amyopathic dermatomyositis, and 2 (1%) had vasculitis syndrome. The Ro52‐autoAb‐titer in their sera was examined by a EUROLINE scan (Cosmic Corporation, Japan). In the EULOLINE scan, recombinant Ro‐52 (52 kDa) produced with the baculovirus system in insect cells is used to detect Ro52‐autoAbs. We defined the Ro52‐autoAb titer as “high” when the EUROLINE scan determined the titer as 2± or 3±. Diagnosis of ILD was performed by examining the interstitial markings in the lung in HRCT. The severity of ILD was examined by measuring the KL‐6 value (SRL, Tokyo, Japan). In this study, KL‐6 was used as a potential biomarker of ILD in CTD 41, 47, 48. KL‐6 was measured using electrochemiluminescence immunoassay as described previously 49. The sensitivity, specificity, and diagnostic accuracy were 93.9%, 96.3%, and 95.7% for KL‐6; 81.8%, 86.6%, and 85.2% for SP‐A; 69.7%, and 95.1%, and 87.8% for SP‐D; and 51.5%, 92.7%, and 80.9% for MCP‐1; respectively 41, 47, 48, indicating that KL‐6 is a superior serum biomarker for ILD compared with SP‐A or SP‐D. It has also been reported that KL‐6 is negatively well correlated with DLco and FVC 50 and positively correlated with the HRCT score 51 in cases of ILD with CTD. Severe ILD was defined when the KL‐6 value was greater than 500 U/mL. Detection of ASCs using the immunospot array assay on a chip (ISAAC) PBMCs were prepared from 10 mL of heparinized peripheral blood from patients with a high serum titer of Ro52‐autoAbs after centrifugation at 400 × g using a Ficoll‐Hypaque gradient. CD138+ cells were enriched with human CD138 MicroBeads (Miltenyi Biotec) using an autoMACS Pro‐separator (Miltenyi Biotec) according to the manufacturer's instructions. The ISAAC method is covered by patents that have been exclusively licensed to Valneva (Nantes, France). Details and instructions regarding the microwell array chip and the ISAAC method have been previously described 37, 38. Briefly, we coated the surface of the chip with 10 μg/mL recombinant Ro52 (rRo52) protein (ProSpec) in PBS by incubating the chip overnight at 4°C. After removing the antibody solution, we blocked the chip with 0.01% Biolipidure (NOF Corporation, Japan) for 30 min at room temperature (r.t.) and subsequently washed the chip with the culture medium (RPMI 1640 containing 10% fetal calf serum, 50 μM 2‐mercaptoethanol, 100 μg/mL streptomycin, and 100 U/mL penicillin). We then arrayed the CD138+ cells in culture medium on the chip and removed the residual cells outside the wells with gentle washing. We cultured the cells on the chip for 3 h at 37°C. After gentle washing, we added 1 μg/mL Cy3‐conjugated anti‐human IgG‐Fc antibody (Millipore) for 30 min. Finally, we stained the cells with 1 μM Oregon Green (Molecular Probes) for 5 min at r.t. After gentle washing, antigen‐specific antibody‐secreting cells were retrieved under a fluorescence microscope (BX51WI, Olympus) using a capillary with a micromanipulator. Amplification of antibody cDNA, production of antibody, and purification of antibody We amplified antibody cDNAs encoding the VH and VL fragments using a single‐cell 5’‐RACE method with primers for the γ, κ, and λ chains. We then inserted these cDNAs into expression vectors that contained cDNA encoding the whole immunoglobulin constant region for IgG1 or the light chains, as previously described 38, 52. Then, we co‐transfected Expi293FTM cells with both the heavy and light chain expression vectors using the Expi293FTM Expression System (Life Technologies). We then collected the culture supernatant and purified the antibody using a protein G column as previously described 38. The immunoglobulin gene repertoire was analyzed with the IMGT/V‐Quest tool (http://www.imgt.org/). Identification of mRo52‐autoAbs We coated the wells of 96‐well plates (Nunc) with 50 μL of 0.5 μg/mL rRo52 protein in PBS overnight at 4°C. The next day, we blocked the wells with 3% BSA in PBS containing 0.1% Tween 20 (PBST). After washing the wells, we added to the wells monoclonal antibodies (1 μg/mL) and patient sera in PBST and incubated the plate at r.t. for 1 h. After washing the wells, we added alkaline phosphatase‐conjugated anti‐human IgG Fc‐antibody (Sigma) and incubated the plate at r.t for 1 h. We washed the wells and detected the amount of bound antibodies using p−nitrophenylphosphate (Sigma). Subsequently, we measured the optical absorbance at 405 nm with an ELISA reader according to the manufacturer's instructions. Western blotting of mutant recombinant Ro52 (rRo52) proteins with mRo52‐autoAbs We conjugated peroxidase to mRo52‐autoAbs or an anti‐influenza nucleoprotein antibody 38, 53 (control antibody) using the Peroxidase Labeling Kit‐NH2 (Dojindo, Japan) according to the manufacturer's instructions. Full‐length or truncated rRo52 proteins were produced in vitro with the cell‐free protein synthesis system using purified wheat embryos as previously described 54. Briefly, we amplified cDNAs encoding full‐length or truncated rRo52 proteins by PCR and inserted them into a pEU‐E01 expression vector (CellFree Sciences, Ehime, Japan). All proteins were fused to a D1CE epitope tag at the C‐terminus 55. We then produced recombinant proteins using the WEPRO7240 Expression Kit (CellFree Sciences) according to the manufacturer's instructions. Next, we resolved the rRo52 proteins using SDS‐PAGE with 5–20% gradient acrylamide gel (BIO CRAFT, Tokyo, Japan) under reducing conditions and transferred them to an Immobilon‐P nylon membrane (Millipore). We treated the membrane with Blocking One (Nacalai Tesque, Kyoto, Japan) and probed it with peroxidase‐conjugated antibodies. Visualization was performed using the ECL detection system (GE Healthcare). To confirm the loading of recombinant proteins on the blotting membrane, we washed the membrane with PBST for a few minutes and inactivated the peroxidase of the peroxidase‐conjugated mRo52‐autoAb 0.01% sodium azide for 30 min followed by a wash with PBST for a few minutes. We then treated the membrane with Blocking One and probed it with peroxidase‐conjugated anti‐D1CE tag antibodies. Competitive ELISA assay with mRo52‐autoAbs We coated the wells of 96‐well Maxisorp plates (Nunc) with 50 μL of 5 μg/mL rRo52 in PBS and incubated them at 4°C overnight. The next day, we blocked the wells with 3% BSA in PBS at r.t. for 1 h. We prepared the solution containing 10 clones of mRo52‐autoAbs or control mAb at 10 μg/mL and one of the clones of mRo52‐autoAbs conjugated with peroxidase at 0.1 μg/mL. After washing the wells, we added 50 μL of a clone of unconjugated mRo52‐autoAb to each well with 50 μL of a clone of peroxidase‐conjugated mRo52‐autoAb and incubated them at r.t. for 1 h. After washing the samples, we added 50 μL of TMB solution (KPL) and incubated the samples for 30 min at r.t.; then, we added 50 μL of stop solution (KPL). We measured the optical absorbance at 450 nm with an ELISA reader (BMG LABTECH) according to the manufacturer's instructions. ELISA of mAbs with a peptide library We first prepared the peptide library for the C‐C domain of Ro52 (Eurofins Genomics). The amino acid sequences are shown in Supporting Information Table 2. This library had 21 peptides. Each peptide consisted of 16 amino acids and overlapped by eight amino acids with an adjacent peptide. We then coated the wells of 96‐well Maxisorp plates (Nunc) with 50 μL of 5 μg/mL peptides in PBS and incubated them at 4°C overnight. The next day, we blocked the wells with 3% BSA in PBS. After washing the samples, we added the peroxidase‐conjugated mRo52‐autoAb or control mAb to the wells and incubated them at r.t. for 1 h. After washing the cells, we detected the antibodies that were bound to antigens using a TMP detection system (KPL). Subsequently, we measured the optical absorbance at 450 nm with an ELISA reader (BMG LABTECH) according to the manufacturer's instructions. ELISA of patient serum samples with the peptides We coated the wells of 96‐well Maxisorp plates (Nunc) with 50 μL of 10 μg/mL peptide (PEP08: RIHAEFVQQKNFLVEE) or HIV env gp160 peptide (RYLRDQQLL) (MBL) in PBS and incubated the cells overnight at 4°C. The next day, we blocked the wells with 3% BSA in PBS. After washing the wells, we added the patient sera to the wells and incubated the plate at r.t. for 1 h. After washing the wells, we added alkaline phosphatase‐conjugated anti‐human IgG Fc‐antibody (Sigma) and incubated the plate at r.t for 1 h. We washed the wells and detected the amount of bound antibodies using p−nitrophenylphosphate (Sigma). Subsequently, we measured the optical absorbance at 405 nm with an ELISA reader according to the manufacturer's instructions. We performed an ELISA three times per patient. When the mean of the OD(PEP08)/OD(HIV) was higher than 1 + SD, we classified the patients in the “PEP08‐positive group.” When the mean of OD(PEP08)/OD(HIV) was less than 1 + SD, the patients were classified in the “PEP08‐negative group.” Acknowledgments We thank Sanae Hirota for technical assistance and Kaoru Hata for secretarial work. This research was supported by research grants from Chugai Pharmaceutical Co., LTD. (H.T.) and from the Platform Project for Supporting in Drug Discovery and Life Science Research (Platform for Drug Discovery, Informatics, and Structural Life Science) from the Ministry of Education, Culture, Sports, Science (MEXT), Japan Agency for Medical Research and development (AMED) (A.M.); Platform Project for Supporting Drug Discovery and Life Science Research (Basis for Supporting Innovative Drug Discovery and Life Science Research (BINDS)) from AMED under Grant Number JP18am0101077 (T.O.), and JSPS KAKENHI Grant Number 16H06499 (H.K.). All authors designed the experiments. M.O., T.O., Y.N., and H.K. performed the experiments. M.O., R.T., K.S., H.T., and K.T. acquired clinical data. M.O., T.O., H.K., and A.M. analyzed the experiments and wrote the manuscript. Conflicts of interest Okumura, Ozawa, Kishi, Taki, Tobe, and Muraguchi have a patent application pending for the Ro52 autoantibody (JP2016‐220793). The rest of the authors declare no commercial or financial conflict of interest. Supporting Information References Abbreviations ASC antibody‐secreting cell CTD connective tissue disease HRCT high‐resolution computed tomography ILD interstitial lung disease ISAAC immunospot array assay on a chip MCTD mixed connective tissue disease POD peroxidase SLE systemic lupus erythematosus UCTD undifferentiated connective tissue disease