ABSTRACT

   Systemic lupus erythematosus (SLE) is an autoimmune disease of unknown
   origin. Recent evidence has linked butyrophilin 3A1 (BTN3A1) to immune
   dysregulation. This study was to elucidate the relationship of BTN3A1
   in SLE. Expression of BTN3A1 in plasma and peripheral blood mononuclear
   cells from SLE patients and healthy controls explored the association
   between BTN3A1 and SLE. We found that BTN3A1 mRNA, plasma levels, and
   expression in CD4^+ T cells were significantly elevated in SLE
   patients. In BTN3A1 gene knock‐in (BTN3A1^KI) mice, inflammation and
   lupus‐like manifestations occurred, including increased proportions of
   Th1, Th2, and Th17 cells, decreased Treg cells, elevated levels of
   inflammatory cytokines and anti‐dsDNA antibodies, renal injury, and
   suppressed IL‐38 serum levels. Intraperitoneal injection of IL‐38 in
   pristane‐treated BTN3A1^KI mice notably alleviated these pathological
   changes. Mechanistic investigations revealed that CD4^+ T cells and the
   ferroptosis pathway were closely associated with the effects mediated
   by the BTN3A1‐IL‐38 axis. In vitro experiments showed that IL‐38
   stimulation reduced proliferation, apoptosis, and decreased the
   expression of ferroptosis‐related proteins, Fe^2⁺, glutathione, and
   malondialdehyde in CD4^+BTN3A1^+/+ T and BTN3A1^+/+ Jurkat T cells.
   Overall, BTN3A1 plays a crucial role in SLE pathogenesis by regulating
   CD4^+ T cell function.

   Keywords: butyrophilin 3A1, IL‐38, inflammation, lupus
     __________________________________________________________________

   We found that BTN3A1 was abnormally expressed in peripheral blood of
   SLE patients. BTN3A1 promotes the release of inflammatory factors and
   antibodies in mice and the pathological damage of the kidney, and
   multi‐omics reveal the abnormalities of CD4 + T cells and the
   occurrence of ferroptosis. Finally, we demonstrated that BTN3A1
   promotes inflammation and induces lupus‐like diseases by inhibiting the
   IL‐38‐iron death axis through BTN3A1.

   graphic file with name MCO2-6-e70356-g001.jpg

1. Introduction

   Systemic lupus erythematosus (SLE) is a complex autoimmune disease, and
   its main causes may include immune dysregulation, genetics, and
   environmental irritation. SLE is highly prevalent in women often
   attacks systemic organs, including the kidneys, blood, joints, and
   nervous system. During the development of SLE, CD4^+ T cells, the
   master population of a class of αβ T cells, are hyperactivated, leading
   to loss of immune tolerance. In addition, autoantibodies are produced
   by memory B cells or plasma cells whose differentiation is induced by
   CD4^+ T helper (Th) cells [[42]1]. The imbalance of CD4^+ T cells,
   especially Th17/Treg, is a key feature of disease activity in SLE, and
   there is activated CD4^+ T cell infiltration in the affected kidney,
   skin, and other organs [[43]2, [44]3]. Studies have confirmed that the
   occurrence of ferroptosis in CD4^+ T cells has an important impact on
   the immune imbalance of lupus nephritis [[45]4]. Thus, the balance of
   CD4^+ T cells is critical to the pathogenesis of SLE. Moreover,
   searching for biomarkers regulating CD4^+ T cell function and
   discussing how the potential biomarkers are involved in CD4^+ T cell
   signaling events could provide new therapeutic options for SLE
   treatment.

   The Butyrophilin 3A (BTN3A, also called CD277) subfamily, consisting of
   BTN3A1, BTN3A2, and BTN3A3, plays a key role in regulating immunity
   [[46]5]. BTN3A receptors expressed on antigen‐presenting cells (APCs)
   can promote negative co‐stimulatory functions upon binding to ligands
   expressed on T cells. It has been demonstrated that BTN3A1 is
   significantly associated with T cell function in the tumor
   microenvironment [[47]6, [48]7]. BTN3A1, a major histocompatibility
   complex‐associated gene, is located on the short arm of human
   chromosome 6 (6p22.1). It encodes a membrane protein with two
   extracellular immunoglobulin structural domains and an intracellular
   B30.2 structural domain [[49]8]. The positively charged pocket formed
   by the B30.2 structural domain binds directly to a range of negatively
   charged small molecules, such as the associated phosphoantigen (pAg)
   [[50]9]. BTN3A1 is expressed on the surface of some immune cell
   subpopulations, such as basophils, γδ T cells, memory CD8^+ T cells,
   CD4^+ T cells, and B cells [[51]10]. In addition, expression of BTN3A1
   was significantly positively correlated with infiltration of CD4^+ T
   cells in gliomas [[52]11]. Vγ9Vδ^2+ T cells, as the main subset of γδ T
   cells in human peripheral blood, are involved in immune responses in
   infections, tumors, and inflammation, and their function is associated
   with BTN3A1 [[53]12, [54]13]. The activation of BTN3A1 significantly
   enhances TCR‐induced T cell proliferation and the secretion of
   cytokines, and BTN3A1 plays a key role in the type I IFN response via
   the enhancement of interferon regulatory factor 3 (IRF3)
   phosphorylation and IFN‐γ secretion [[55]14, [56]15].

   BTN3A1 is closely associated with cancers, and its expression is
   strongly correlated with the clinical prognosis of cancers. For
   example, BTN3A1 is associated with tumor‐infiltrating immune cells and
   is co‐expressed with a variety of immune checkpoints in patients with
   breast cancer (BRCA) and non‐small cell lung cancer (NSCLC). Currently,
   there are limited studies on BTN3A1 and autoimmune diseases. Psoriasis
   patients had higher BTN3A1 expression in monocytes relative to the
   healthy subjects, with high expression of this factor in CD14^+ cells
   of skin lesions [[57]16]. A study discussed the rs3802604 polymorphism
   of BTN3A1 gene, indicating that this locus is a risk polymorphism for
   type 1 diabetes [[58]17]. In addition, our previous study has shown
   that the gene polymorphism of BTN3A1 is associated with the risk of SLE
   in the Chinese Han population [[59]18]. Interleukin‐38 (IL‐38) has been
   shown to inhibit inflammation. For instance, IL‐38 inhibits Th17 cells
   and myeloid cells and then binds to IL‐1R6 to repress inflammation.
   THP‐1 monocyte cell line expresses low expression of IL‐6, TNF‐α,
   IL‐1β, IL‐17, and IL‐23 after IL‐38 stimulation [[60]19]. In addition,
   IL‐38 has been shown to diminish the accumulation of eosinophils and
   the number of Th2 and Th17 cells, and to enhance the proportion of Treg
   cells [[61]20]. Our previous studies showed that IL‐38 can alleviate
   liver, spleen, and kidney enlargement in lupus mice, reverse lupus‐like
   pathological changes, improve immune homeostasis imbalance, and inhibit
   the occurrence and development of lupus through regulating ERK1/2,
   JNK1/2, p38, NF‐κB p65, and STAT5 signaling pathways [[62]21]. However,
   it remains to be elucidated whether BTN3A1 is involved in the
   pathogenesis of autoimmune diseases such as SLE by regulating IL‐38.

   While BTN3A1 has been extensively investigated in cancers and γδ T
   cells, studies in CD4^+ T cells and SLE are absent. The association
   between BTN3A1 and SLE pathogenesis remains to be discussed. In our
   study, we initially explored the expression of BTN3A1 in the peripheral
   blood of SLE patients, as well as the expression of BTN3A1 in various
   immune cells. Since the BTN3A1 gene is not expressed in mouse, next, we
   knock in the BTN3A1 gene into wild‐type (WT) C57BL/6 mice (BTN3A1^KI)
   to clarify the role of BTN3A1 in inflammation and SLE occurrence.
   Moreover, our previous findings showed abnormal expression of IL‐38 in
   SLE patients, and IL‐38 inhibited lupus development. We then
   investigated the association of BTN3A1 and IL‐38, and discussed whether
   IL‐38 is involved in the BTN3A1‐mediated effects. These results
   highlight the implication of BTN3A1 for inflammation and SLE
   development, and suggest a novel mechanism by which BTN3A1 is involved
   in ferroptosis of CD4^+ T cells through inhibition of IL‐38.

2. Results

2.1. Highly Expressed BTN3A1 in SLE Patients

   To investigate whether expression of BTN3A1 was abnormal in SLE
   patients and whether expression of BTN3A1 was related to SLE
   pathogenesis, we examined plasma levels of BTN3A1 and BTN3A1 mRNA
   expression from PBMCs in SLE patients. Plasma levels of BTN3A1 were
   significantly increased in SLE patients compared with healthy controls
   (Figure [63]1A), and BTN3A1 mRNA expression was significantly increased
   in SLE patients compared with healthy controls (Figure [64]1B). There
   were higher plasma levels of BTN3A1 in SLE patients with lupus
   nephritis compared to patients without lupus nephritis (Figure [65]1C).
   The plasma levels of BTN3A1 were positively correlated with SLE disease
   activity index (SLEDAI) score (r ^2 = 0.3728, p < 0.0001,
   Figure [66]1D). Subgroup analysis showed that plasma levels of BTN3A1
   were significantly related to several clinical and laboratory features,
   such as vasculitis, arthritis, cylindruria (Figure [67]S1A–H), and were
   significantly correlated with C3, RF expression (Figure [68]S1I,J).
   Interestingly, the proportion of CD3^+, CD4^+, CD8^+ T cells was higher
   in SLE patients compared with healthy controls (Figures [69]1E–G). The
   proportion of CD3^+BTN3A1^+, CD4^+BTN3A1^+, CD8^+BTN3A1^+ T cells was
   also higher in SLE patients (Figures [70]1E–G). These were similar to
   CD19^+, CD14^+, CD11b^+, CD11c^+ cells, where the proportion of CD19^+,
   CD14^+, CD11b^+, CD11c^+ cells was higher in SLE patients (Figure
   [71]S2A), and proportion of CD19^+BTN3A1^+, CD14^+BTN3A1^+,
   CD11b^+BTN3A1^+, CD11c^+BTN3A1^+ cells was significantly higher in SLE
   patients compared with healthy controls, respectively (Figure [72]S2B).
   The findings suggest that expression of BTN3A1 was highly expressed in
   SLE patients, and may correlate with SLE risk.

FIGURE 1.

   FIGURE 1
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   BTN3A1 expression in systemic lupus erythematosus patients (SLE). (A)
   Plasma levels of BTN3A1 from healthy controls (N = 100) and SLE
   patients (N = 280) were detected by ELISA. (B) Detection of mRNA
   expression of BTN3A1 in healthy controls (N = 30) and SLE patients (N =
   30) by qRT‐PCR. (C) Plasma levels of BTN3A1 in SLE patients with lupus
   nephritis (N = 123) and without lupus nephritis (N = 157). (D)
   Relationship between BTN3A1 plasma levels and disease activity index
   score in SLE patients. (E, F) Representative flowchart of different T
   cells and BTN3A1 expression in different T cells from healthy controls
   (N = 18) and SLE patients (N = 18). (G) Statistical analysis of T cell
   subpopulations and BTN3A1 expression in different T cell
   subpopulations. *p < 0.05, **p < 0.01, and ****p < 0.0001.

2.2. Knock‐in BTN3A1 Promotes Inflammation and Induces a Lupus‐Like Disease

   Since the BTN3A1 gene does not exist in WT mice, we constructed the
   human‐derived BTN3A1 gene knock‐in (BTN3A1^KI) mice (Figure [74]S3A,B).
   First, we discussed whether the knock‐in BTN3A1 gene may contribute to
   inflammation. Second, we evaluated whether the knock‐in BTN3A1 gene may
   induce a lupus‐like disease (Figure [75]S3C). As shown in
   Figure [76]2A–C and Figure [77]S4A,B, the BTN3A1^KI mice injected with
   PBS had an upregulated proportion of CD3^+, CD8^+, Th1, Th2, Th17
   cells, CD19^+, CD14^+, CD11b^+, CD11c^+ cells, and a downregulated
   proportion of Treg cells compared with the WT mice injected with PBS.
   Expression of inflammatory cytokines IL‐4, IL‐6, IL‐10, IL‐17A, IFN‐γ,
   and TNF‐α was increased in the BTN3A1^KI‐PBS group as well
   (Figure [78]2D). After pristane injection, mice in the
   BTN3A1^KI‐Pristane group showed a higher proportion of CD3^+, CD8^+,
   Th1, Th2, Th17 cells, CD19^+, CD14^+, CD11b^+, CD11c^+ cells, and a
   lower proportion of Treg cells compared with the WT‐Pristane group
   (Figure [79]2A–C and Figure [80]S4A,B). Expression of the inflammatory
   cytokines (IL‐4, IL‐6, IL‐10, IL‐17A, IFN‐γ, and TNF‐α) was increased
   in the BTN3A1^KI‐Pristane group (Figure [81]2D). Thus, the knock‐in
   BTN3A1 gene contributed to inflammation.

FIGURE 2.

   FIGURE 2
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   BTN3A1 promotes inflammation and induces lupus‐like features. (A–C)
   Representative flowchart and statistical analysis of CD3^+, CD8^+ T
   cells, Th1, Th2, Th17, Treg cells in mice from the wild‐type (WT)‐PBS
   (N = 8), WT‐Pristane (N = 8), BTN3A1^KI‐PBS (N = 8) and
   BTN3A1^KI‐Pristane groups (N = 8). (D) Inflammatory cytokines IL‐4,
   IL‐6, TNF‐α, IFN‐γ, IL‐10 and IL‐17A were detected by CBA kits in the
   serum of different groups of mice. (E–G) Representative figure and
   statistical analysis of mice spleen, liver and kidneys. (H) The dsDNA
   levels were detected by ELISA. (I, J) HE staining, Masson staining and
   statistical analysis of mice kidneys showed the extent of kidney
   damages and fibrosis. (K) Ultrastructure of glomerular and renal
   tubular damages in mice as revealed by transmission electron microscope
   (TEM). (L) Representative figure of IgG deposition in mice kidneys
   assessed by immunofluorescence intensity and statistical analysis.
   *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001.

   We further discussed the role of the knock‐in BTN3A1 gene in lupus
   development. Morphologically, the BTN3A1^KI‐PBS group displayed
   significant splenomegaly, hepatomegaly, and bilateral renal enlargement
   (Figures [83]2E–G). Organ weight analysis revealed that the liver and
   kidneys of BTN3A1^KI‐PBS mice were notably heavier than those of the
   WT‐PBS group. In addition, white spherical calcified particles were
   observed adhering to the hepatic lobules, further highlighting the
   pathological changes. These signs of splenomegaly, hepatomegaly, and
   renal enlargement were more pronounced in the BTN3A1^KI‐Pristane group
   (Figures [84]2E–G). In terms of autoantibody expression, the
   BTN3A1^KI‐PBS group exhibited a marked increase in dsDNA autoantibody
   levels compared to the WT‐PBS group, with BTN3A1^KI lupus mice showing
   the highest dsDNA expression (Figure [85]2H). Histopathological
   examination of the kidneys demonstrated that the BTN3A1^KI‐PBS group
   had a significantly higher HE score and fibrosis percentage than the
   WT‐PBS group (Figures [86]2I,J). Microscopic analysis revealed
   glomerular segmental or global sclerosis, thickening of the renal
   capsule basement membrane, epithelial nuclear swelling and hyperplasia,
   and nephrogenic fibrosis. Transmission electron microscopy (TEM)
   further confirmed severe damage to the glomeruli and tubular structures
   in BTN3A1^KI‐PBS mice (Figure [87]2K). Compared with the WT‐Pristane
   group, the BTN3A1^KI‐Pristane group presented more severe renal injury
   (Figures [88]2I–K), accompanied by increased IgG deposits
   (Figure [89]2L). Collectively, these findings indicate that the BTN3A1
   gene knock‐in exacerbates inflammation and elicits lupus‐like
   manifestations.

2.3. BTN3A1 Induces the Lupus Development by Inhibiting IL‐38 Expression

   In our previous studies, we found higher plasma levels of IL‐38 and
   higher IL‐38 mRNA expression in PBMCs from SLE patients compared with
   healthy controls [[90]21], and IL‐38 inhibited lupus development
   [[91]22]. To discuss whether BTN3A1 may regulate IL‐38 expression, and
   then promote inflammation and induce the lupus‐like disease, mice in
   the BTN3A1^KI‐Pristane group were injected with recombinant mouse
   IL‐38. First, we validated expression of IL‐38 in plasma and PBMCs from
   SLE patients, showing consistent results with previous findings (Figure
   [92]S5A,B). Serum levels of IL‐38 were examined in the WT‐PBS,
   WT‐Pristane, BTN3A1^KI‐PBS, and BTN3A1^KI‐Pristane groups, and serum
   IL‐38 levels were lower in the BTN3A1^KI‐Pristane group and
   BTN3A1^KI‐PBS group than in the WT‐Pristane group and WT‐PBS group,
   respectively (Figure [93]S6). This suggests that BTN3A1 inhibited IL‐38
   expression in the mice with lupus‐like disease. Second, mice in the
   BTN3A1^KI‐Pristane group were injected with different doses of
   recombinant mouse IL‐38. Proportion of inflammation‐related immune
   cells and expression of inflammatory cytokines were reversed by
   addition of IL‐38, evidenced by less proportion of CD3^+, CD8^+, Th1,
   Th2, Th17 cells, CD19^+, CD14^+, CD11b^+, CD11c^+ cells, more
   proportion of Treg cells, lower expression of the inflammatory
   cytokines (IL‐4, IL‐6, IL‐10, IL‐17A, IFN‐γ, and TNF‐α) in the
   BTN3A1^KI‐Pristane‐IL‐38 group (Figures [94]3A–D, Figure [95]S7A,B).
   Splenomegaly, hepatomegaly, and swelling of both kidneys was reversed
   in the BTN3A1^KI‐Pristane‐IL‐38 group, along with less weight of the
   above organs (Figures [96]3E–G). Serum levels of autoantibody dsDNA
   were reduced in the BTN3A1^KI‐Pristane‐IL‐38 group (Figure [97]3H).
   Interestingly, histopathology of the kidneys was improved in the
   BTN3A1^KI‐Pristane‐IL‐38 group, evidenced by lower HE score, fibrosis
   percentage, and less structural damage in the kidneys
   (Figures [98]3I–K). IgG deposition in the kidneys was inhibited in the
   BTN3A1^KI‐Pristane‐IL‐38 group (Figure [99]3L). Together, BTN3A1
   inhibited IL‐38 expression and then contributed to inflammation and
   induced the lupus‐like disease.

FIGURE 3.

   FIGURE 3
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   IL‐38 alleviates BTN3A1‐induced inflammation and the lupus‐like
   disease. (A–C) Representative flowchart and statistical analysis of
   CD3^+, CD8^+ T cells, Th1, Th2, Th17, Treg cells in mice from the
   BTN3A1^KI‐Pristane‐PBS group (N = 8) and BTN3A1^KI‐Pristane‐IL‐38
   groups (with injection of different concentrations of IL‐38, N =
   8/group). (D) The inflammatory cytokines IL‐4, IL‐6, TNF‐α, IFN‐γ,
   IL‐10 and IL‐17A were detected by the CBA kits in the serum from
   different groups of mice. (E–G) Representative figure and statistical
   analysis of mice spleen, liver, and kidneys. (H) The dsDNA levels in
   different groups of mice were detected by ELISA. (I, J) HE staining,
   Masson staining and statistical analysis of mice kidneys showed the
   extent of kidney damages and fibrosis. (K) Ultrastructure of glomerular
   and renal tubular damages in mice as revealed by TEM. (L)
   Representative figure of IgG deposition in mice kidneys assessed by
   immunofluorescence intensity and statistical analysis. *p < 0.05,
   **p < 0.01, ***p < 0.001 and ****p < 0.0001.

2.4. Ferroptosis Is a Key Signaling Pathway in the BTN3A1‐IL‐38 Axis‐Mediated
Effects

   As discussed above, BTN3A1 contributed to inflammation and induced the
   lupus‐like disease by inhibiting IL‐38. We further discussed the
   mechanisms that the BTN3A1‐IL‐38 axis involves. Spleens of the
   different groups of mice were evaluated by transcriptomics and
   proteomics analysis. The number of differentially expressed proteins
   and genes was summarized in Table [101]S1 when different group was
   compared with each other. Transcriptomics and proteomics correlation
   analysis showed that among the 200 differentially expressed proteins
   and 2001 differentially expressed genes, 39 key genes/proteins were
   shown by Venn diagrams (Figure [102]4A). The upregulation and
   downregulation of differentially expressed genes and proteins were
   shown in the Volcano plot (Figure [103]4B,C). Based on the
   transcriptomics data, the genes and proteins were categorized as
   RNA‐proteomics and RNA+proteomics, and the distribution of gene
   expression and gene abundance in the two subgroups was shown
   separately. There was higher expression and abundance of RNA+proteomics
   genes after association compared with RNA‐proteomics genes
   (Figures [104]4D,E). We subjected 39 key genes/proteins to KEGG pathway
   analysis, of which four key genes were enriched in ferroptosis,
   including downregulated Slc11a2, Steap3, and Tfrc in both
   transcriptomics and proteomics analysis, as well as downregulated
   Slc7a11 in transcriptomics and upregulated Slc7a11 in proteomics
   analysis (Figure [105]4F, Table [106]S2). Ferroptosis was significantly
   different in KEGG pathway enrichment in both transcriptomics and
   proteomics analysis (Figure [107]4G, Table [108]S2). Results of the
   other comparisons were shown in Figures [109]S8–S10, Table [110]S3.

FIGURE 4.

   FIGURE 4
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   Transcriptomics and proteomics cross‐analysis explores signaling
   pathways regulated by the BTN3A1‐IL‐38 axis. (A) Overlap of
   differentially expressed proteins and differentially expressed genes in
   mice from the BTN3A1^KI‐Pristane‐PBS group (N = 3) and the
   BTN3A1^KI‐Pristane‐IL‐38 group (with injection of recombinant IL‐38
   (50ug/kg), N = 3) (featured by Venn diagrams). (B, C) All
   differentially expressed genes and differentially expressed proteins
   visualized by Volcano plots. (D, E) Box plots and scatter plots
   visualized the abundance of genes associated with proteins in the
   BTN3A1^KI‐Pristane‐PBS group and the BTN3A1^KI‐Pristane‐IL‐38 groups.
   Red represents the distribution of gene abundance in transcriptomics
   associated to proteomics. Green represents the distribution of gene
   abundance in transcriptomics not associated to proteomics. (F, G) KEGG
   pathway enrichment showed key pathways, and KEGG enrichment heat map
   showed distribution of key genes and key proteins.

2.5. T Cells Are Significantly Related to the BTN3A1 Inhibition of IL‐38

   To better clarify which immune cells were regulated by the BTN3A1‐IL‐38
   axis, single‐cell transcriptomics explored the differentially
   distributed immune cells. After removing damaged and duplicated cells
   from the data, we retained 47067 high‐quality cellular transcriptomes
   for further analysis. For different immune cells, we used the marker
   genes identified for the corresponding cells. The relevant marker genes
   and their sources are shown in Table [112]S4. We used principal
   component analysis (PCA) and cluster analysis for the determination of
   cellular lineages. Comparing the immune cells in the
   BTN3A1^KI‐Pristane‐PBS group and the BTN3A1^KI‐Pristane‐IL‐38 group
   (injected with 50ug/kg recombinant mouse IL‐38), five cell populations
   were identified, namely T cells, B cells, monocytes, natural killer
   (NK) cells, and plasmacytoid dendritic (pDC) cells (Figure [113]5A).
   The distribution of these five cell populations in each sample was
   shown by composite bar graphs, and there is a significant difference in
   the distribution of T cells between the two groups (Figure [114]5B,
   Table [115]S5). Therefore, we performed cluster analysis again for the
   T cells, and 10 T cell subpopulations were identified. It is notable
   that CD4 was the most expressed marker in these cells
   (Figures [116]5C,D; Table [117]S6). Then, we explored the CD4^+ T cells
   in further experiments. In addition, Slc11a2, Steap3, Tfrc, and Sle7a11
   are presently expressed in T cells (Figure [118]S11). For the other
   immune cells (B cells, NK cells, monocytes, pDC), we all performed
   clustering again, and the corresponding cell subpopulations and cell
   numbers were shown in the appendix (Figures [119]S12–S15, Tables
   [120]S7–S10).

FIGURE 5.

   FIGURE 5
   [121]Open in a new tab

   Single‐cell sequencing in mice from the BTN3A1^KI‐Pristane‐PBS group
   and the BTN3A1^KI‐Pristane‐IL‐38 group (50ug/kg IL‐38) showed
   significantly different T cells distribution. (A) UMAP plot showed the
   5 major cell types identified based on marker genes in mice spleen
   tissues (N = 3/group). (B) Composite bar graphs showed percentages of T
   cells, B cells, natural killer cells, monocytes and pDCs in each
   sample. (C, D) T cells were clustered again, and the UMAP plot showed
   the results of distribution of all T cell subpopulations, as well as
   the expression of CD4 in these cluster species.

2.6. BTN3A1 Inhibits the IL‐38‐Ferroptosis Pathway and Then Promotes T Cell
Dysfunction

   We transfected WT C57BL/6 mice with BTN3A1^+/+ adenovirus and isolated
   naïve CD4^+ T cells for the following analysis. Similarly, Jurkat T
   cells were infected with BTN3A1^+/+ adenovirus. As shown in Figure
   [122]S16A,B, infection of Jurkat T cells with the BTN3A1^+/+ adenovirus
   (MOI = 200, Day 6) and injection of BTN3A1^+/+ adenovirus (10^9 PFU,
   Day 5) in WT mice showed the optimal expression of BTN3A1,
   respectively. Moreover, 100 ng/mL of IL‐38 stimulation showed the
   optimal effects on proliferation and apoptosis of CD4^+ T cells (Figure
   [123]S16C–E).

   In CD4^+BTN3A1^+/+ T cells, the mRNA expression of ferroptosis‐related
   proteins (Scl7a11, Steap3, Slc11a2, Tfrc; Figure [124]6A) was
   significantly decreased after the addition of 100 ng/mL IL‐38.
   Similarly, 100 ng/mL IL‐38 stimulation was able to significantly
   inhibit the ferroptosis pathway, by which the cellular levels of Fe^2+,
   glutathione, and malondialdehyde were significantly decreased
   (Figure [125]6B), suggesting that BTN3A1 inhibits IL‐38 and then
   regulates activation of the ferroptosis pathway. Moreover,
   CD4^+BTN3A1^+/+ T cells were treated with IL‐38 or IL‐38+Ferrostatin‐1
   (an inhibitor of the ferroptosis pathway). Compared with
   CD4^+BTN3A1^+/+ T cells (Control group/Blank), addition of 100 ng/mL
   IL‐38 stimulation inhibited apoptosis and proliferation of the cells,
   which were further significantly reduced in the presence of
   Ferrostatin‐1 (Figure [126]6C,D).

FIGURE 6.

   FIGURE 6
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   BTN3A1 inhibits the IL‐38‐ferroptosis axis and induces CD4^+ T cells
   dysfunction. Naïve CD4^+ T cells were isolated from spleens of the
   BTN3A1^+/+ mice (N = 5/group), and then were activated. Part of the
   cells were further stimulated with 100 ng/mL of IL‐38. (A) The mRNA
   expression of ferroptosis‐related proteins in the cells was detected by
   qRT‐PCR. (B) Detection of Fe^2+, glutathione and malondialdehyde levels
   in above cells reflects cellular ferroptosis. (C, D) The IL‐38‐treated
   CD4^+BTN3A1^+/+ T cells we intervened with/without Ferrostatin‐1
   treatment. Cell proliferation was measured by CCK8 kits and cell
   apoptosis was measured. Representative flowchart and statistical
   analysis of apoptosis of different groups of cells are shown.
   *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001.

   In addition, the Jurkat T cells were used to validate the above
   mechanisms (Figure [128]S17A–F). The results showed that the
   proliferation and apoptosis of BTN3A1^+/+ Jurkat T cells were inhibited
   by 100 ng/mL IL‐38 stimulation (Figure [129]S17A). Jurkat T cells
   transfected with BTN3A1^+/+ adenovirus in the presence of 100 ng/mL
   IL‐38 had significantly reduced mRNA expression of ferroptosis‐related
   proteins Slc11a2, Steap3, Slc7a11, and Tfrc (Figure [130]S17B), and
   cellular levels of Fe^2+, glutathione, and malondialdehyde (Figure
   [131]S17C). Moreover, the addition of Ferrostatin‐1 to IL‐38‐treated
   BTN3A1^+/+ Jurkat T cells had less cell proliferation and apoptosis
   compared with IL‐38‐treated BTN3A1^+/+ Jurkat T cells without
   Ferrostatin‐1 treatment (Figure [132]S17D,E). Furthermore, BTN3A1 can
   promote ferroptosis, evidenced by increased expression of
   ferroptosis‐related proteins, which in turn lead to proliferation and
   apoptosis of Jurkat cells (Figure [133]S17B–F). Thus, BTN3A1 not only
   directly promotes ferroptosis but also indirectly promotes ferroptosis
   by inhibiting the IL‐38 signaling. Together, BTN3A1 suppresses the
   IL‐38‐ferroptosis pathway and then contributes to T cell proliferation
   and apoptosis.

3. Discussion

   In this study, we found that highly expressed BTN3A1 in SLE patients
   was significantly associated with severe clinical manifestations, such
   as lupus nephritis, and correlated with SLEDAI score. BTN3A1^KI mice
   showed inflammation and the lupus‐like inflammatory manifestations,
   including renal injuries, imbalance of T cell subpopulations, a burst
   of inflammatory cytokines, and decreased levels of serum IL‐38. In
   addition, BTN3A1 contributes to CD4^+ T cells' proliferation and
   apoptosis by inhibiting the IL‐38‐ferroptosis signaling pathway.
   Collectively, BTN3A1 contributes to inflammation and induces the
   lupus‐like disease.

   BTN3A1 has been widely studied in tumors and is abundantly expressed in
   the microenvironment of human epithelial ovarian cancer and BRCA
   [[134]23]. Hypoxia‐stimulated dendritic cells revealed high expression
   of BTN3A1 [[135]7]. Our study found that plasma levels of BTN3A1 and
   BTN3A1 mRNA expression in PBMCs were elevated in SLE patients.
   Similarly, BTN3A1 expression was elevated in different immune cells
   from SLE patients, including CD3^+, CD4^+, CD8^+ T cells. CD14^+ cells
   in psoriatic skin lesions highly expressed BTN3A1 and were regulated by
   IFN‐γ [[136]16]. We also found highly expressed BTN3A1 in CD14^+ cells
   from SLE patients. These data provide supportive evidence that BTN3A1
   may be involved in the pathogenesis of SLE. To validate this
   hypothesis, BTN3A1^KI mice were constructed, and the lupus‐like disease
   was induced by pristane injection. We examined inflammatory immune
   cells and cytokines in the physiological status of BTN3A1^KI mice,
   suggesting an increased percentage of CD3^+, CD8^+ T cells, Th1, Th2,
   Th17 cells, and decreased Treg cells, as well as higher expression of
   IL‐4, IL‐6, IL‐10, IL‐17A, IFN‐γ, and TNF‐α. These inflammatory
   phenotypes were further exacerbated in the lupus‐like mice. It is
   accepted that an imbalance of Th1, Th2, Th17, Treg cells is the basis
   of the pathogenesis of SLE [[137]24]. Therefore, BTN3A1 may contribute
   to T cell dysfunction, leading to inflammation. The kidneys are the
   most vulnerable organ in SLE [[138]25]. Renal pathological analysis and
   TEM findings revealed renal structural damage in pristane‐induced lupus
   mice. Indeed, BTN3A1^KI mice were able to express the human‐derived
   BTN3A1 gene [[139]26], and the mouse model of lupus induced by pristane
   was able to develop dysregulated immune milieu and systemic organs
   damage similar to human SLE [[140]27]. Thus, BTN3A1 can induce the
   lupus‐like disease.

   IL‐38 demonstrates potent anti‐inflammatory effects in SLE [[141]21].
   IL‐38 competitively binds to IL‐36R, IL‐1R1, IL‐1 receptor accessory
   protein‐like‐1 (IL‐1RAPL1) to inhibit the activation and function of
   Th1 and Th17 cells, and promotes the proliferation of Treg cells, thus
   exerting anti‐inflammatory functions. This study also discussed whether
   BTN3A1 may regulate IL‐38 and then affect inflammation and SLE
   development. First, we examined IL‐38 expression in the plasma, PBMCs
   of SLE patients, and healthy volunteers, and the results were
   consistent with the previous findings [[142]21]. We found lower serum
   levels of IL‐38 in pristane‐treated BTN3A1^KI mice. To test our
   hypothesis, we injected pristane‐treated BTN3A1^KI mice with
   recombinant IL‐38. Our data revealed amelioration of BTN3A1‐induced
   lupus‐like changes, such as improved degree of renal injuries and
   reduced percentage of CD3^+, CD8^+ T cells, Th1, Th2, Th17 cells,
   upregulated percentage of Treg cells, and lower expression of IL‐4,
   IL‐6, IL‐10, IL‐17A, IFN‐γ, TNF‐α, and dsDNA. However, our study showed
   that a 50 µg/kg IL‐38 dose was more effective than 100 µg/kg in
   reducing BTN3A1‐induced inflammation and lupus‐like phenotype, although
   both reversed the inflammatory responses and renal damages (higher
   doses were only significantly effective in some indicators). This may
   be due to high doses of IL‐38‐mediated inactive immunomodulatory
   effects or changes in pharmacokinetics, which will lead to uneven
   tissue distribution and decreased target site of IL‐38 concentration
   [[143]28, [144]29]. We demonstrated that BTN3A1 inhibited IL‐38
   secretion, and then promoted inflammation and induced lupus‐like
   alterations.

   We conducted KEGG pathway enrichment analysis to discuss the mechanisms
   involved in the BTN3A1‐IL‐38 axis. The ferroptosis signaling pathway
   was significantly differentially expressed between the pristane‐treated
   BTN3A1^KI mice injected with/without IL‐38, indicating that the
   ferroptosis signaling may be a key to BTN3A1 inhibition of IL‐38.
   Indeed, expression of ferroptosis signaling is elevated in the
   glomeruli of patients with lupus nephritis [[145]30]. Enhanced
   neutrophil ferroptosis leads to a breakdown of immune tolerance in SLE
   patients [[146]31]. Moreover, our data revealed that CD4^+ T cells were
   significantly differently distributed in the pristane‐treated BTN3A1^KI
   mice injected with/without IL‐38 by single‐cell sequencing. CD4^+ T
   cells are involved in the pathogenesis of SLE by auto‐secreting
   cytokines and inducing B cells to secrete IL‐4, IL‐6, IL‐10 [[147]32].
   Therefore, it is hypothesized that CD4^+ T cells may play an essential
   role in SLE through inhibition of the IL‐38‐ferroptosis axis by BTN3A1.
   Ferroptosis signaling‐associated proteins (Steap3, Slc7a11, Slc11a2,
   Tfrc), glutathione, and malondialdehyde are considered to be key
   features of ferroptosis [[148]33]. Our results identified that these
   features were highly expressed in CD4^+BTN3A1^+/+ T cells, whereas
   addition of IL‐38 reduced expression of these proteins, and glutathione
   and malondialdehyde. Promotion of CD4^+ T cells activation and
   imbalanced differentiation of effector T cells was related to
   aggravation of the lupus‐like features [[149]34]. Iron overload was
   able to promote CD4^+ T cell subpopulations proliferation, inflammatory
   cytokines generation, and autoantibodies generation in lupus mice, and
   the lupus mice treated with a high‐iron diet showed a higher proportion
   of CD4^+ T cell subpopulations [[150]35]. Thus, inhibition of CD4^+ T
   cells' function such as proliferation, is essential for the
   pathogenesis of SLE. By isolating CD4^+ T cells from BTN3A1^+/+ mice,
   the addition of IL‐38 inhibited cell proliferation and apoptosis. IL‐38
   can inhibit the conversion of CD4^+ T cells into Th17, further enhance
   the immunosuppressive activity of Treg cells, and inhibit the
   inflammatory response [[151]36]. The occurrence of ferroptosis in
   different subsets of CD4^+ T cells is significantly associated with
   immune imbalance, which indicates that there is a potential
   relationship between IL‐38 and ferroptosis. These features were further
   confirmed by the addition of the ferroptosis inhibitor (Ferrostatin‐1),
   where proliferation and apoptosis of CD4^+BTN3A1^+/+ T cells were
   inhibited by treatment with IL‐38 in the presence of Ferrostatin‐1. To
   validate our findings, Jurkat T cells were used, and consistent
   findings were observed in Jurkat T cells. Overall, we confirmed that
   BTN3A1 inhibition of the IL‐38‐ferroptosis axis affects CD4^+ T cell
   biological activity and is involved in lupus pathogenesis.
   Interestingly, a study showed that BTN3A1 overexpression in APCs
   inhibited the proliferation and IFN‐γ release from CD4^+ T cells
   [[152]26]. BTN3A1 inhibits αβ T cells by binding to N‐mannosylated
   residues in the human CD45 gene, thereby eliminating effective TCR
   activation. However, we endogenously transfected with an adenovirus
   overexpressing BTN3A1, and this direct effect of overexpression of
   BTN3A1 for CD4^+ T cells may have contributed to the accelerated CD4^+
   T cell proliferation. This was confirmed in our in vivo experiments in
   which BTN3A1 increased the proportion of CD4^+ T cells (including a
   higher percentage of Th1, Th2, Th17 cells and a lower percentage of
   Treg cells) in the lupus‐like mice. Similarly, another study showed
   that upregulation of BTN3A1 in monocytes leads to activation and
   accelerated proliferation of Vγ9Vδ2^+ T cells in patients with
   psoriasis vulgaris [[153]16]. Thus, our findings strongly indicated
   that BTN3A1 is functionally indispensable in promoting CD4^+ T cells
   proliferation and apoptosis in the lupus‐like mice.

   Nevertheless, this study has several limitations. First, although it
   has been demonstrated that BTN3A1 can regulate the expression of IL‐38,
   the precise molecular mechanisms underlying this regulation remain to
   be fully elucidated. Future research should delve deeper into the
   signaling pathways and molecular interactions involved to clarify how
   BTN3A1 modulates IL‐38 expression. Second, the systemic manifestations
   of BTN3A1^KI mice require further investigation. Organs commonly
   affected in lupus, such as lymph nodes and joints, were not
   comprehensively evaluated in this study. In future studies, a more
   detailed assessment of these organs needs to be conducted to provide a
   more complete understanding of the disease phenotypes in BTN3A1^KI
   mice.

   In summary, this study found that BTN3A1 expression was increased in
   SLE patients, and BTN3A1 inhibited the IL‐38‐ferroptosis axis, and
   then, promoted inflammation and induced lupus‐like disease. Our study
   also provided insights into the involvement of CD4^+ T cells in the
   mechanisms of SLE disease pathogenesis and highlighted the potential
   role of BTN3A1 for SLE therapy. It is expected that in the future we
   will purify BTN3A1 for use in clinical trials for the treatment of SLE
   patients.

4. Materials and Methods

4.1. Patients

   A total of 328 SLE patients from the Affiliated Hospital of Southwest
   Medical University were included to detect the plasma (N = 280), mRNA
   (N = 30) expression of BTN3A1, and BTN3A1 expression in different
   immune cells (N = 18). One hundred and forty‐eight healthy controls
   from the Affiliated Hospital of Traditional Chinese Medicine of
   Southwest Medical University were used as controls. Plasma (N = 100),
   mRNA (N = 30) expression of BTN3A1, and expression of BTN3A1 in
   different immune cells (N = 18) were detected in healthy controls.
   Detailed information on patients and controls is shown in Table
   [154]S11 and Table [155]S12. The diagnosis of SLE meets the 1997
   American College of Rheumatology criteria. SLE patients with pregnancy,
   infections, tumors, and other rheumatic diseases were excluded. Blood
   samples were collected, and each participant signed an informed consent
   form. The study was approved by the Ethics Committee of the Affiliated
   Hospital of Southwest Medical University (KY2020180) and the Affiliated
   Hospital of Traditional Chinese Medicine of Southwest Medical
   University (BY2022005).

4.2. Animal Model

   We engineered a knock‐in mouse expressing the human BTN3A1 gene under
   the control of the mouse Itgax/Cd11c promoter (BTN3A1^KI). In brief, we
   created a β‐globin‐BTN3A1‐β‐globin knock‐in at the locus of ROSA26 in
   C57BL/6 mice by CRISPR/Cas‐mediated genome engineering (Figure
   [156]S3A,B). WT female C57BL/6 (7 weeks) mice were purchased from
   GemPharmatech (Chengdu, China). Experiments were conducted in two
   stages, and a lupus‐like disease was induced by intraperitoneal
   injection of 500ul pristane (Sigma Aldrich, St Louis, USA) for three
   months. In the first stage, BTN3A1^KI and WT mice were categorized into
   four groups (WT‐PBS group, WT‐Pristane group, BTN3A1^KI‐PBS group,
   BTN3A1^KI‐Pristane group). Mice in the WT group and BTN3A1^KI group
   were intraperitoneally injected with 500ul PBS at Week 8, and mice in
   the WT‐Pristane group and BTN3A1^KI‐Pristane group were
   intraperitoneally injected with pristane at Week 8. Samples of venous
   blood from the inner canthus, liver, spleen, and kidneys were collected
   for subsequent analysis after 3 months’ observation. In the second
   stage, BTN3A1^KI mice were divided into four groups
   (BTN3A1^KI‐Pristane‐PBS group, BTN3A1^KI‐Pristane‐25ug/kg IL‐38 group,
   BTN3A1^KI‐Pristane‐50ug/kg IL‐38 group, and BTN3A1^KI‐Pristane‐100ug/kg
   IL‐38 group). Mice in the BTN3A1^KI‐Pristane groups were injected with
   different doses of recombinant mouse IL‐38 (25, 50, and 100ug/kg, Sino
   Biological, Beijing, China) or PBS for a duration of two weeks (with
   intraperitoneal injection every other day). The mice were then
   monitored for one week and used for subsequent testing. A flowchart of
   the animal experiments was shown in Figure [157]S3C. All animal
   experiments were approved by the Ethics Committee of Southwest Medical
   University (swmu20230093).

4.3. Enzyme‐Linked Immunosorbent Assay (ELISA)

   BTN3A1 and IL‐38 levels in the plasma of SLE patients and healthy
   controls were detected by ELISA kits (Cusabio, Wuhan, China). Serum
   samples from mice were collected from venous blood of the inner
   canthus, and the levels of IL‐38 and dsDNA were detected by ELISA kits
   (Cusabio, Wuhan, China).

4.4. Flow Cytometry

   Peripheral blood mononuclear cells (PBMCs) from SLE patients and
   healthy controls were obtained by lymphocyte separation medium with
   density gradient centrifugation (TBDscience, Tianjing, China). CD3^+
   (CD3‐APC‐Cy7), CD4^+ (CD4‐BB515), CD8^+ (CD8‐APC) T cells, CD19^+
   (CD19‐APC), CD14^+ (CD14‐BV421), CD11b^+ (CD11b‐FITC), CD11c^+
   (CD11c‐BV421) cells in PBMCs were recognized by flow cytometry. In
   addition, expression of BTN3A1 (BTN3A1‐PE) was detected in all of the
   immune cells as described above.

   Mouse spleen was ground on a 70‐micrometer cell strainer (Novbio,
   China) and then lysed with erythrocyte lysate (Beyotime, Shanghai,
   China) on ice. CD3^+ (CD3‐FITC), CD4^+ (CD4‐FITC), CD8^+ (CD8‐APC) T
   cells, CD19^+ (CD19‐APC), CD14^+ (CD14‐FITC), CD11b^+ (CD11b‐APC),
   CD11c^+ (CD11c‐FITC) cells were detected directly by flow cytometry.
   After labeling CD4^+ T cells, the cells were fixed (RD system,
   Minnesota, USA) and membrane‐broken (RD system, Minnesota, USA), and
   finally, fluorescently labeled with Th1 (IFN‐γ‐PE‐CF594), Th2
   (IL‐4‐APC), Th17 (IL‐17A‐APC‐CyTM7), and Treg (Foxp3‐PE) cells.

   All flow cytometry antibodies were purchased from BD Biosciences
   (California, USA), and data were assayed under a BD FACSCalibur flow
   cytometer (Becton, USA).

4.5. Histopathology

   Kidneys of mice were fixed using 4% formaldehyde and then embedded in
   paraffin, cut with a thickness of 5 um sections. The kidneys were
   stained with hematoxylin‐eosin staining (HE) and Ponceau, Fuchsin, and
   Aniline blue staining (Masson), respectively. HE score (Table [158]S13)
   and Masson assay were performed to assess the extent of renal injury
   and renal fibrosis, respectively. Immune complexes deposited in the
   kidneys were evaluated by FITC‐coupled anti‐mouse IgG with
   immunofluorescence. Moreover, kidneys were pre‐fixed with 3%
   glutaraldehyde, re‐fixed with 1% osmium tetroxide, dehydrated and
   embedded, and sectioned. Ultrastructural alterations of renal tubular
   epithelial cells and glomeruli were visualized under TEM after staining
   with uranyl acetate and lead citrate.

4.6. Cytometric Bead Array System (CBA)

   Serum concentrations of cytokines IL‐4, IL‐6, IL‐10, IL‐17A, IFN‐γ, and
   TNF‐α in different groups of mice were measured by the CBA Mouse
   Th1/Th2/Th17 Cytokine Kit (BD Biosciences, San Diego, CA) on a cellular
   Counting Bead Array (CBA).

4.7. Single‐Cell RNA Sequencing

   The library was established after the mice spleen tissue was taken and
   a single cell suspension was prepared. During library construction, the
   number of genes identified by the cells was filtered out to be less
   than 200 or greater than the maximum number of genes ×90%. The
   proportion of mitochondrial reads of the cells was filtered out to be
   sorted in the top 15%, and to correct for the effect of the cell cycle.
   Final sequencing was performed using Combinatorial Probe‐Anchor
   Synthesis (cPAS).

4.8. Transcriptomics and Proteomics

   For transcriptomics, spleen tissue from mice was taken for RNA
   purification and collection, and the products were amplified and
   sequenced. The raw data obtained from sequencing were filtered to
   remove reads containing junctions (junction contamination); reads with
   unknown base N content greater than 5%; and low‐quality reads (reads in
   which the proportion of bases with a mass value of less than 15 to the
   total number of bases in the read is greater than 20% are considered to
   be low‐quality reads), and then the clean data were obtained.

   Regarding proteomics, spleen tissue was milled on the ice and tissue
   was lysed using RIPA lysis solution. Lysis was further assisted by the
   treatment of ultrasound. Samples of protein solution were obtained by
   centrifugation. Then, the samples were proteolytically digested and
   evaluated by DIA mass spectrometry.

4.9. CD4^+ T Cells Isolation and Culture

   Wild‐type C57BL/6 mice were subjected to human BTN3A1 gene
   overexpression (BTN3A1^+/+) adenovirus injection (Genechem, Shanghai,
   China). Five days later, naïve CD4^+ T cells were obtained from spleens
   by magnetic bead sorting (Miltenyi Biotec, Germany). Naïve CD4^+ T
   cells (10^6/500uL) were cultured in 10% fetal bovine serum (FBS)
   (Gibco, Massachusetts, USA), 1% penicillin‐streptomycin antibody
   (Beyotime, Chengdu, China) and activated with anti‐CD3 (2ug/ml)
   (Invitrogen, Massachusetts, USA), anti‐CD28 (2ug/mL) (Invitrogen,
   Massachusetts, USA), IL‐2 (2 ng/mL) (Peprotech, New Jersey, USA) for
   24 h. Then, the cells were stimulated with recombinant mouse IL‐38
   stimulation (24 h), and used for the following studies.

   In addition, the Jurkat T cells were cultured in 10% FBS, 1%
   penicillin‐streptomycin antibody. After BTN3A1 overexpressed adenovirus
   transfection (5 days) and then recombinant human IL‐38 stimulation
   (24 h), the cells were collected for the following studies.

4.10. Detection of Fe^2+, Glutathione, Malondialdehyde Levels

   CD4^+BTN3A1^+/+ T cells and BTN3A1^+/+ Jurkat T cells were collected
   and lysed by cell sonication. Then, the cells were assayed for levels
   of Fe^2+ (Abcam, Cambridge, UK), glutathione (Beyotime, Chengdu,
   China), and malondialdehyde (Beyotime, Chengdu, China), respectively.

4.11. Cell Proliferation and Apoptosis

   Cell proliferation of CD4^+BTN3A1^+/+ T cells and BTN3A1^+/+ Jurkat T
   cells was determined by CCK8 kits (Abmole, Houston, USA) in the
   presence or absence of inhibitors of ferroptosis (Ferrostatin‐1, 2uM)
   (Monmouth Junction, NJ, USA) for 24 h.

   Similarly, CD4^+BTN3A1^+/+ T cells and BTN3A1^+/+ Jurkat T cells were
   treated with/without Ferrostatin‐1 for 24 h, and cell apoptosis was
   detected using FITC Annexin V Apoptosis Detection Kit I (BD
   Biosciences, California, USA) and APC Annexin V Apoptosis Detection Kit
   (Vazyme, Nanjing, China).

4.12. Reverse Transcription‐Quantitative PCR

   RNA was extracted from the PBMCs of SLE patients and healthy controls,
   CD4^+ T cells, and Jurkat T cells using the EZ‐press RNA Purification
   Kit (HiFoBio, Shanghai, China). cDNA was constructed by PrimeScript RT
   Master Mix (Vazyme, Nanjing, China). qPCR was performed under the
   following thermal cycling conditions: 95°C for 30s, followed by 40
   cycles of 95°C for 10s and 60°C for 30s. GAPDH was used as an internal
   reference. Primer sequences are shown in Table [159]S14.

4.13. Statistical Analysis

   Normally distributed quantitative data are presented as mean±standard
   deviation, and were compared by Student's t‐test or one‐way analysis of
   variance. The Mann‐Whitney U test was used for quantitative data that
   were not normally distributed. Correlation analysis was used for both
   metrics. All analyses were performed using GraphPad Prism 5.0.

Author Contributions

   Study conception and design: WDX, DCW, YYT, QH, LCS, AFH. Acquisition
   of data, analysis and interpretation of data: WDX, DCW, YYT, QH, SYF,
   LF, YYC, LQY, LCS, AFH. Drafting the article: WDX, DCW, YYT, QH, LCS,
   AFH. Final approval of the version of the article to be published: all
   authors, and that all authors agree to be accountable for all aspects
   of the work.

Ethics Statement

   All animal experiments were approved by the Ethics Committee of
   Southwest Medical University (swmu20230093). The study was approved by
   the Ethics Committee of the Affiliated Hospital of Southwest Medical
   University (KY2020180) and the Affiliated Hospital of Traditional
   Chinese Medicine of Southwest Medical University (BY2022005).

Conflicts of Interest

   All authors declare no conflicts of interest.

Supporting information

   Supporting Information
   [160]MCO2-6-e70356-s001.docx^ (3.7MB, docx)

Acknowledgments