KMT5a‐Mediated IRF3 Stabilization Enhances Macrophage Chemotaxis and Renal Fibrosis Progression

ABSTRACT

While the histone lysine methyltransferase (KMT) family responsible for posttranslational modification has been reported to regulate multiple diseases progression like tumor and cardiovascular disease, its role in renal fibrosis progression is not completely understood. Here, via TGF‐β‐treated HK‐2 cells and an in vivo unilateral ureteral obstruction mice model, the histone lysine methyltransferase family was screened and the upregulated expression of KMT5a was identified in these renal fibrosis models. Interference assays using lentivirus transfection in HK‐2 cells and transgenic mice displayed that KMT5a silencing in HK‐2 cells impeded macrophage chemotaxis. Moreover, in vivo, transgenic KMT5a knockout inhibited macrophage infiltration, which contributed to the suppression of renal fibrosis progression. Mechanistically, by immunoprecipitation assays and mass spectrometry, we discovered that KMT5a could interact with and stabilize the transcription factor IRF3 to regulate macrophage chemotaxis and infiltration in renal tissues, which accelerated renal fibrosis progression. Notably, we confirmed that a small molecular inhibitor of KMT5a, UNC0379, reduced IRF3 protein levels and macrophage chemotaxis, thus suppressing renal fibrosis, in vitro and in vivo. Therefore, this study presented KMT5a as a mediator of renal fibrosis progression, an effect that was reversed by the KMT5a inhibitor UNC0379, which provided evidence of a novel target and potential drug for inhibiting renal fibrosis progression.

In renal tubular epithelial cells, TGF‐β‐induced upregulation of KMT5A enhances IRF3 methylation and protein stability. This augments chemokine transcription, leading to elevated production and secretion of CCL2, CCL3, and CCL7. These chemokines collectively promote macrophage chemotaxis and infiltration, thereby accelerating the progression of renal fibrosis.

1. Introduction

The global health burden of chronic kidney disease (CKD) is substantial, with approximately 8%–15% of the global population affected, leading to high morbidity, mortality, and health care costs [1]. CKD is characterized by an irreversible decline in renal function. In this process, renal fibrosis is a critical hallmark that facilitates disease progression to end‐stage renal failure in nearly all forms of CKD [2]. The accumulation of multiple extracellular matrix components, including α‐SMA, collagen, and fibronectin, in the interstitium is a unique feature of renal fibrosis [3]. This process leads to the progressive replacement of functional nephrons with scar tissue and the disruption of normal tissue architecture and ultimately culminates in end‐stage renal disease requiring dialysis or transplantation. The pathogenesis of renal fibrosis is complex and involves a cascade of cellular and molecular events. To date, numerous studies have reported various key drivers of pathogenesis, such as sustained inflammation, myofibroblast transition, and pervasive tubular injury [4]. In addition, critical signaling pathways, including the TGF‐β and Wnt signaling pathways, have been confirmed to accelerate the formation of fibrosis [5].

The KMT family is responsible for catalyzing the methylation of specific lysine residues of histone proteins, thus regulating chromatin structural stability and gene expression [6]. Notably, in addition to canonical histone methylation, KMTs can also regulate the methylation of many nonhistone substrates, such as transcription factors and signaling transducers [7, 8]. KMT‐mediated noncanonical methylation has been widely investigated in cancer, neurodevelopment disorders, and cardiovascular and metabolic diseases [9]. This noncanonical methylation directly alters the stability, activity, or interactions of key cellular proteins, offering a direct mechanism to regulate relevant pathways. Despite growing recognition, the comprehensive landscape and pathological significance of the KMT family, particularly in the context of renal fibrosis, remain largely unexplored. Elucidating the related mechanisms is crucial for contributing to the development of potential novel therapies.

In this study, we first screened KMTs in TGF‐β‐stimulated HK2 cells and determined that KMT5a expression was upregulated. In vivo results revealed that KMT5a knockout alleviated renal fibrosis in a unilateral ureteral obstruction model and impeded M2 macrophage infiltration into renal tissue. Mechanistically, KMT5a stabilized IRF3, which mediated macrophage chemotaxis. Importantly, we found that the KMT5a inhibitor UNC0379 inhibited IRF3 expression, leading to reduced M2 macrophage infiltration and impeding renal fibrosis progression.

2. Materials and Methods

2.1. Cell Culture and Reagents

The human tubular epithelial cell line HK‐2 (SCSP‐511) and THP‐1 cells (SCSP‐567) were obtained from the cell bank of the Chinese Academy of Sciences. Cells were cultured in DMEM (Gibco, USA) or RPMI‐1640 (Gibco, USA) supplemented with 1% penicillin–streptomycin (Yeasen, Shanghai) and 10% fetal bovine serum (Gibco, USA) at 37°C in 5% CO₂. Cultures were regularly tested for mycoplasma to ensure that the in vitro experiments were performed using mycoplasma‐negative cells.

2.2. Lentivirus Transfection

Lentiviruses targeting KMT5a were purchased from Genomeditech (Shanghai, China). HK‐2 cells were transfected in accordance with the manufacturer's protocol. Transfected cells were selected with puromycin (10 μg/mL) for approximately 7 days. The knockdown efficiency was assessed by western blotting and quantitative reverse transcription polymerase chain reaction (qRT–PCR). The sequence targeting KMT5a was as follows: 5′‐CCGAGGAACAGAAGATCAAAG‐3′.

2.3. qRT–PCR

The qRT‐PCR was performed according to our previous study [10]. Total RNA from HK‐2 cells was isolated using an RNA extraction kit (Beyotime, China; R0077S). Then, total RNA was reverse transcribed to cDNA using a Hifair V reverse transcriptase kit (Yeasen, China, 11300ES92). qRT–PCR was performed on an Applied Biosystems (Thermo Fisher, USA) using a Hieff qPCR SYBR Green Master Mix Kit (Yeasen, China, 11199ES03). The parameters in QuantiStudio software were set in accordance with the manufacturer's instructions. The housekeeping gene GAPDH was selected as an internal control. The primers used were as follows: GAPDH, F: 5′‐TCGGAGTCAACGGATTTGGT‐3′, R: 5′‐TTCCCGTTCTCAGCCTTGAC‐3′; KMT5a, F: 5′‐TGAGCCCGAACAAATGCTCT‐3′, R: 5′‐TTCCGGCTAATGGTTTCCCC‐3′; KMT1a, F: 5′‐GTGATGAGGGGCGGATTGAA‐3′, R: 5′‐AACCACGTACAGCCATCGAG‐3′; KMT1b, F: 5′‐AAGCTCTACAAGATGGCGGC‐3′, R: 5′‐AGTGAAACTAGGCAAGGCACA‐3′; KMT2a, F: 5′‐GGATCCACAACTCCAGGCAA‐3′, R: 5′‐ATTTGGAATGGACCCAGCGA‐3′; KMT2b, F: 5′‐GTCGCAAGCATAAGACGACC‐3′, R: 5′‐ACCATCCGTTCTGTGCCTTC‐3′; KMT2c, F: 5′‐CTGCCTAGACCCTCCATTGC‐3′, R: 5′‐TGCAGATGTTGCTCCACAGT‐3′; KMT2d, F: 5′‐CCCACAGGAACCGCTGTAAT‐3′, R: 5′‐CCAATCTCCCCCTCCCTGTA‐3′; KMT3a, F: 5′‐TGAAAATGTGCAGAAAACAGGT‐3′, R: 5′‐GCCAAGTGCAGTGAGAAACC‐3′; KMT3b, F: 5′‐TGTCTGCTGCCCTTTTCCAA‐3′, R: 5′‐ATCATCCGAAAGGGCTGTCC‐3′; KMT3c, F: 5′‐ACTGCAATGTGGAGTGTCAGA‐3′, R: 5′‐TGGCCAGAATCCTTGCTGTT‐3′; KMT4, F: 5′‐CGCTGCCGGTCTACGATAAA‐3′, R: 5′‐TCGATGGCACGGTTGTACTT‐3′; KMT5b, F: 5′‐GGAATGTCCGCCAAGGAACT‐3′, R: 5′‐TCTCCCCACTCACAGGGTTG‐3′; KMT5c, F: 5′‐TCCTTCTGGACCCTCTAGGT‐3′, R: 5′‐TCCAGGAAAGGGAGACCCTA‐3′.

2.4. Western Blotting

The procedures for western blotting were performed according to our previous study [11]. The culture medium was removed, and HK‐2 cells were washed with PBS three times on ice. The cells were lysed with western and IP lysis buffer (Beyotime, China; P0013) supplemented with a protease inhibitor cocktail (Yeasen, China, 20124ES); the lysates were collected in 1.5 mL tubes. After 30 min on ice, the samples were centrifuged at 12000 × g for 10 min, after which the supernatants were collected. The protein concentration was measured using a BCA quantification kit (Beyotime, China; P0010). Loading buffer was added to the samples, followed by boiling at 100°C for 5 min, and total protein was separated by SDS–PAGE at 150 V for 60 min. Then, the separated proteins were transferred to a PVDF membrane (Millipore, USA; 03010040001) at 0.28 A for 2 h on ice. Then, the membrane was immersed in rapid blocking buffer (Epizyme, China; PS108P) and incubated with primary antibodies for 12 h at 4°C. Then, the membrane was washed in TBST buffer for 30 min and incubated with an HRP‐conjugated secondary antibody for 1 h at room temperature. The bands of target proteins were visualized using an ECL reagent and a Tanon imaging system (Shanghai, China). The primary antibodies used were as follows: anti‐KMT5a (1:1000, Abcam, ab300517), anti‐α‐SMA (1:1000, Proteintech, 67 735–1‐Ig), anti‐fibronectin (1:1000, Abcam, ab2413), anti‐IRF3 (1:1000, Abcam, ab68481), and anti‐methyl lysine (1:1000, CST, 14 679).

2.5. Histology and IHC

Dissected kidney tissues were fixed with 4% paraformaldehyde for 48 h, dehydrated with gradient alcohol and embedded in paraffin. Sirius red (Solarbio, China; G1472) and Masson's trichrome (Solarbio, China; G1340) staining of kidney tissues was performed in accordance with the manufacturer's protocol. Staining images were evaluated using ImageJ software.

After deparaffinization in xylene and rehydration in a gradient alcohol series (95%, 85%, and 75%), tissue sections were treated with 3% hydrogen peroxide for quenching. Antigen retrieval was performed by heating the tissue section in a 5% citric acid solution at 100°C for 20 min. To minimize background staining, the tissue sections were incubated with bovine serum albumin to block nonspecific binding. Immunostaining was carried out by incubating the sections with specific primary antibodies overnight (≥ 8 h) at 4°C. For signal development, a commercial streptavidin–peroxidase immunohistochemistry detection system (ZSGB‐Bio, China) was then used in accordance with the manufacturer's instructions. Protein expression was quantified using Image‐Pro Plus 6.0 software. All histological quantifications were performed on the renal cortex, identified by the presence of proximal tubules with brush borders. Fields containing large vessels were excluded. The primary antibodies used in this experiment were as follows: anti‐F4/80 (1:200; Abcam, ab300421), anti‐CD206 (1:400; CST, 24595), anti‐IRF3 (1:200; Abcam, ab68481), anti‐α‐SMA (1:400; Proteintech, 67 735–1‐Ig) and anti‐CD86 (1:400; CST, 19589).

2.6. Enzyme‐Linked ImmunoSorbent Assay (ELISA)

The supernatant from HK‐2 cell lysates was collected for cytokine quantification. human CCL2 (R&D Systems, USA; DCP00), human CCL3 (R&D Systems, USA; DMA00), human CCL7 (R&D Systems, USA; DCC700), mouse CCL2 (R&D Systems, USA; MJE00B), mouse CCL3 (R&D Systems, USA; MMA00), mouse CCL7 (R&D Systems, USA; KA2199), and mouse TGF‐β (R&D Systems, USA; MMA00) quantikine ELISA kits were used to measure the levels of CCL2, CCL3, and CCL3 in the supernatant in accordance with the standard protocol. The concentration was quantified using TMB, and the absorbance at 450 nm was measured using a Varioskan ALF instrument (Thermo Fisher, USA).

2.7. Coimmunoprecipitation (Co‐IP) Assay

HK‐2 cells were harvested and lysed on ice for 30 min in immunoprecipitation (IP) lysis buffer (Beyotime, China; P0013) supplemented with a protease inhibitor cocktail (Yeasen, China, 20124ES). After lysis, the cell extracts were centrifuged at 12000 × g for 15 min at 4°C. For immunoprecipitation, the supernatants were incubated with 2.5 μg of either control IgG or an anti‐KMT5a antibody overnight under constant agitation at 4°C. Protein A/G magnetic beads (MedChemExpress, USA; HY‐K0202, 20 μL) were then added to each sample, followed by incubation for an additional 6 h at 4°C with gentle rotation. After incubation, bead‐bound immune complexes were isolated using a magnetic separation rack and washed thoroughly three times with ice‐cold PBS. Finally, the beads were resuspended in 60 μL of lysis buffer, and bound proteins were eluted by heating the bead complexes at 100°C for 10 min. The eluate, containing the purified immunoprecipitated protein complexes, was collected and subsequently analyzed by mass spectrometry and western blotting.

2.8. Macrophage Chemotaxis Assay

THP‐1 cells were induced to differentiate into macrophages using phorbol 12‐myristate 13‐acetate (PMA, 100 ng/mL) for 24 h. The chemotaxis assay was performed in a Transwell chamber system with a 5 μm pore size. A total of 1 × 10⁵ macrophages were seeded in the upper chamber. Simultaneously, 1 × 10⁵ HK‐2 cells were seeded in the lower chamber. After incubation at 37°C with 5% CO₂ for 12 h, nonmigrated cells on the upper membrane surface were removed by gentle wiping. Cells that migrated to the lower surface were fixed with 4% paraformaldehyde for 15 min and stained with 0.1% crystal violet for 20 min. The stained cells were counted under a light microscope in five randomly selected fields (200 × magnification) per well. The HK‐2 cells alone in the upper chamber were the negative control. The chemotaxis ability was calculated by experimental group migration number minus negative control.

2.9. Mice

Kmt5a‐KO mice (strain no. T027634) were purchased from GemPharmatech Company (Nanjing, China) and fed in a pathogen‐free environment. Kmt5a‐KO (Kmt5a^−/−) mice and their littermates (Kmt5a^+/+) were used for experiments. Kmt5a‐KO mice were genotyped at 2–3 weeks of age using PCR amplification of DNA isolated from the tail. The primers used were as follows: forward, 5′‐GTTTGACTGTGGCCTTGTGGC‐3′; reverse, 5′‐CAACTGACTGATGTGTCACCCAGG‐3′. The WT band was 2262 bp, and the KO band was 266 bp.

2.10. Renal Fibrosis Mouse Model

Unilateral ureteral obstruction (UUO) was used as an in vivo renal fibrosis model. Following anesthesia via 4% isoflurane inhalation, the dorsal fur of each mouse was clipped. The animals were positioned in ventral recumbency on a sterile surgical platform, and the exposed skin was disinfected. A small incision was made obliquely along the costal margin to reveal the lower renal pole and proximal ureter. The left ureter was then ligated using a No. 6 silk suture. Sham‐operated wild‐type (WT) littermates subjected to identical surgical conditions without ureteral ligation served as experimental controls.

To determine the efficacy of UNC0379 in alleviating renal fibrosis, 6–8‐week‐old mice were intraperitoneally administered UNC0379 (1 mg/kg/d) for 4 weeks after UUO surgery. Then, the mice were sacrificed, and renal tissues were dissected for subsequent histological analysis.

For IRF3 overexpression in Kmt5a‐KO mice, In UUO model, the renal was injected with AAV‐IRF3 and AAV‐vector (1 × 10¹¹vgc/mice). After 4 weeks, the mice were sacrificed, and renal tissues were dissected for subsequent histological analysis.

This study was approved by the Ethics Committee of The First Affiliated Hospital of Soochow University. All animal experiments should comply with the ARRIVE guidelines. All methods were carried out in accordance with relevant guidelines and regulations.

2.11. Chromatin Immunoprecipitation PCR Assay

A total of 2 × 10⁷ cells were digested, washed twice with PBS and fixed with 1 mL of paraformaldehyde (1%) for 10 min. Then, 125 mM glycine was used to stop the reaction. The cells were subsequently washed with PBS by centrifugation at 2000 rpm for 5 min. ChIP assay was performed with a SimpleChIP Plus sonication chromatin IP kit (Cell Signaling Technology, USA) according to the manufacturer's protocol. The mixture was digested, and DNA was sonicated into 200‐ to 800‐base pair (bp) fragments. After incubation with IRF3 antibody for 12 h, the DNA–protein complexes were purified with magnetic beads, and eluted DNA was analyzed by qRT‐PCR.

2.12. Statistical Analyses

Statistical analyses were conducted using GraphPad Prism 8 software (GraphPad Software, USA). The data are expressed as the mean ± standard deviation (SD) from a minimum of three independent experiments. For comparisons involving two groups, Student's t‐test was used. When three or more groups were compared, one‐way analysis of variance (ANOVA) with post hoc tests. was performed, followed by Dunn's post hoc test for multiple comparisons. A two‐tailed p value of less than 0.05 was considered to indicate statistical significance.

3. Results

3.1. KMT5a Expression Increased During Renal Fibrosis

To identify potential KMTs contributing to renal fibrosis formation, we first screened, using qRT–PCR, KMT family members in HK‐2 cells stimulated with TGF‐β (100 ng/mL). The data revealed that among KMT family members, KMT5a was the most significantly increased (Figure 1A). Consistently, western blotting indicated that the expression of fibronectin, α‐SMA, and KMT5a was upregulated in HK‐2 cells after TGF‐β stimulation, confirming that KMT5a expression was increased in an in vitro renal fibrosis model (Figure 1B,C). Moreover, the qRT–PCR and western blot data suggested that the upregulation of KMT5a expression in the in vitro renal fibrosis model was time‐dependent (Figure 1D,E). We also constructed an in vivo renal fibrosis model by performing UUO in C57BL/6J mice. No significant difference was observed in kidney function indicators like glomerular filtration, serum creatinine (Scr), and blood urea nitrogen (BUN) between the two groups attributing to the well‐established physiological compensatory response in the contralateral kidney of the UUO model (Figure S1A). Masson and Sirius red staining confirmed the successful establishment of renal fibrosis in UUO mice. IHC staining revealed that KMT5a expression was increased in the renal tissues of UUO mice (Figure 1F). For further confirmation in patients with chronic renal diseases (CKD), we collected 5 CKD tissues and 5 normal tissues. Consistently, the western blotting and qRT‐PCR data presented a higher KMT5a level in CKD tissues (Figure S1B,C). Therefore, these data suggest that KMT5a expression is increased in the renal fibrosis process.

FIGURE 1.

KMT5a expression was increased during renal fibrosis. (A) The expression of KMT family members in HK‐2 cells treated with PBS or TGF‐β for 24 h was measured by qRT–PCR. (B) Western blot results showing the changes in fibronectin, α‐SMA, and KMT5a expression in HK‐2 cells treated with TGF‐β (0 ng/mL vs. 100 ng/mL). (C) Statistical analyses of fibronectin, α‐SMA, and KMT5a protein expression in HK‐2 cells treated with TGF‐β (0 ng/mL vs. 100 ng/mL). (D) After TGF‐β (100 ng/mL) treatment, the expression of fibronectin, α‐SMA, and KMT5a at 0 h, 24 h, and 48 h was assessed by western blotting. (E) Statistical analyses of western blotting data for fibronectin, α‐SMA, and KMT5a expression at 0 h, 24 h, and 48 h after treatment with TGF‐β (100 ng/mL). (F) Masson's trichome, Sirius red, and KMT5a IHC staining of renal tissues from mice that underwent sham surgery or UUO for 28 days. The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, and ***p < 0.001; unpaired Student's t‐test (A, C, F) and one‐way ANOVA (E).

3.2. KMT5a Knockout Impeded Renal Fibrosis Progression in Vivo

We also screened the KMT family in renal tissues of UUO model and found the upregulation of KMT5a (Figure 2A). To confirm that KMT5a plays a role in renal fibrosis in vivo, we performed UUO in KMT5a knockout (KMT5a^KO) mice. After 28 days of UUO, renal tissues were collected to assess renal fibrosis. Western blotting revealed increased fibronectin and α‐SMA levels in WT mice with UUO, whereas in KMT5a^KO mice with UUO, the fibronectin and α‐SMA levels greatly decreased (Figure 2B,C). Consistent with these findings, Masson and Sirius red staining confirmed increased collagen deposition in WT mice with UUO; in KMT5a^KO mice with UUO, collagen deposition was clearly decreased (Figure 2D,E). Moreover, qRT–PCR data indicated that the expression of other collagen markers, namely, Col1a1 and Col1a3, was increased in WT mice with UUO but inhibited in the renal tissues of KMT5a^KO mice (Figure 2F). IHC staining revealed that α‐SMA expression was increased in WT mice but inhibited in KMT5a^KO mice with UUO (Figure 2G,H). Thus, these results suggest that KMT5a knockout impedes renal fibrosis progression in vivo.

FIGURE 2.

KMT5a knockout impeded renal fibrosis progression in vivo. (A) The expression of KMT family members in renal tissues of the Sham and UUO group was measured by qRT–PCR. (B) In WT and KMT5a^KO mice, fibronectin, α‐SMA and KMT5a levels in renal tissues from the Sham and UUO groups were assessed via western blotting. (C) Statistical analyses of fibronectin and α‐SMA expression in renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups. (D) Renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups were subjected to Masson and Sirius red staining. (E) Statistical analyses of the Sirius red‐positive area in renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups. (F) Col1a1 and Col1a3 expression in renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups was assessed using qRT–PCR. (G) Renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups were subjected to α‐SMA IHC staining. (H) Statistical analyses of α‐SMA IHC‐positive areas in renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups. The results are representative of at least three independent experiments *p < 0.05, **p < 0.01, ***p < 0.001, and ns, not significant, according to unpaired Student's t‐tests (C, E, F and H).

3.3. KMT5a Silencing Inhibited Macrophage Infiltration in Vivo

Previous studies have confirmed the critical role of inflammation in the development of renal fibrosis [12], and KMT5a has been reported to be responsible for immune editing [13, 14]. Thus, we investigated whether KMT5a contributes to renal fibrosis by eliciting inflammation. Macrophages are critical drivers of renal fibrosis progression [12]. Therefore, we performed IHC staining for F4/80, CD206, and CD86 in renal tissues from WT and KMT5a^KO mice with UUO and WT and KMT5a^KO sham mice. Compared with that in the control group, macrophage (F4/80‐positive), M2‐type macrophage (CD206‐positive), or M1‐type macrophage (CD86‐positive) infiltration was obviously enriched in the renal tissues of WT mice with UUO. However, in KMT5a^KO mice with UUO, macrophage infiltration was significantly inhibited (Figure 3A,B). To further investigate the relationship between KMT5a expression and macrophage infiltration, we silenced KMT5a in HK‐2 cells using lentivirus and verified the knockdown efficiency using western blotting and qRT–PCR (Figure 3C,D). Macrophage chemotaxis assays were subsequently performed by coculturing HK‐2‐NC or HK‐2‐shKMT5a cells with macrophages. Macrophage chemotaxis was strongly inhibited in cells in the shKMT5a group (Figure 3E). The levels of the cytokines CCL2, CCL3, and CCL7, which are important for macrophage chemotaxis, in the supernatant were assessed using ELISA. Compared with the control treatment, the knockdown of KMT5a reduced the CCL2, CCL3, and CCL7 concentrations (Figure 3F). Consistently, the qRT–PCR results suggested that CCL2, CCL3, and CCL7 expression was inhibited in HK‐2‐shKMT5a cells (Figure 3G). In the homogenate of renal tissues from KMT5a^KO mice with UUO, CCL2, CCL3, and CCL7 levels were significantly inhibited while a similar TGF‐β concentration was presented in those from WT mice (Figure 3H). Collectively, these results indicate that KMT5a silencing inhibits the chemotaxis and suppresses the infiltration of macrophages in renal tissues.

FIGURE 3.

KMT5a silencing exhausted macrophage infiltration in vivo. (A) IHC staining for F4/80, CD206 and CD86 in renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups. (B) Statistical analyses of F4/80+ macrophage, CD206+ macrophage and CD86+ macrophage infiltration in renal tissues from WT and KMT5a^KO mice in the Sham and UUO groups. (C) The knockdown efficiency of a lentivirus targeting KMT5a in HK‐2 cells was assessed by western blotting. (D) The knockdown efficiency of a lentivirus targeting KMT5a in HK‐2 cells was evaluated using qRT–PCR. (E) After macrophages were cocultured with HK‐2‐NC or HK‐2‐shKMT5a cells, their chemotaxis was evaluated in vitro. (F) The concentrations of CCL2, CCL3, and CCL7 in the culture supernatants of HK‐2‐NC and HK‐2‐shKMT5a cells were measured using ELISA. (G) The expression of CCL2, CCL3, and CCL7 in HK‐2‐NC and HK‐2‐shKMT5a cells was measured by qRT–PCR. (H) The concentrations of CCL2, CCL3, CCL7, and TGF‐β in the homogenate of renal tissues from the UUO model were measured using ELISA. The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, ***p < 0.001, and ns, not significant, according to unpaired Student's t‐tests (B, D, E, F, G and H).

3.4. KMT5a Stabilized IRF3 to Mediate Macrophage Chemotaxis

To identify the potential mechanism underlying KMT5a‐mediated macrophage chemotaxis, we overexpressed Flag‐KMT5a in HK‐2 cells and performed immunoprecipitation assays with an anti‐Flag antibody. The immunoprecipitates were screened by LC–MS. Among the candidates that interacted with KMT5a, we detected IRF3, which is an important transcription factor for inflammation [15, 16]. Thus, we hypothesized that IRF3 is an important mediator of KMT5a‐induced macrophage chemotaxis (Figure S2A). Immunoprecipitation assays using anti‐KMT5a and anti‐IRF3 antibodies in HK‐2 cells suggested that KMT5a and IRF3 interact with each other (Figure 4A). Moreover, western blotting revealed that KMT5a knockdown clearly decreased the protein level of IRF3, with no apparent decrease in its mRNA level, indicating that KMT5a regulated the protein level of IRF3 through posttranslational modification (Figure 4B,C). The coimmunoprecipitation assays using anti‐IRF3 and anti‐Lys‐Methylation antibodies confirmed decreased IRF3 methylation level in HK‐2‐shKMT5a cells (Figure 4D). After treating HK‐2‐shKMT5a cells with cycloheximide (CHX), western blotting revealed that IRF3 protein stability was impaired (Figure 4E). In the UUO model, IHC staining also confirmed that the level of IRF3 was significantly decreased in KMT5a^KO mice (Figure 4F). To determine whether IRF3 rescues the KMT5a knockdown‐induced impairment of macrophage chemotaxis, IRF3 was overexpressed in HK‐2‐KMT5a^sh cells. Macrophage chemotaxis dramatically increased after IRF3 overexpression (Figure 4G). Furthermore, ELISA and qRT–PCR analysis revealed upregulated CCL2, CCL3 and CCL7 expression in HK‐2‐KMT5a^sh cells overexpressing IRF3 (Figure 4H,I). The ChIP‐PCR with IRF3 antibodies presented significant enrichment in the promoter region of CCL2, CCL3 and CCL7, revealing direct transcription regulation of theses cytokines by IRF3 (Figure S2B). We also performed AAV‐IRF3 injection in UUO model. The data presented that IRF3 overexpression restored renal fibrosis progression in KMT5KO mice (Figure S2C,D). These results indicate that KMT5a stabilizes IRF3 to mediate macrophage chemotaxis.

FIGURE 4.

KMT5a stabilized IRF3 to mediate macrophage chemotaxis. (A) Co‐IP assays were performed using HK cells and anti‐KMT5a and anti‐IRF3 antibodies. (B) Differences in IRF3 mRNA levels in HK‐2‐NC and HK‐2‐shKMT5a cells were assessed using qRT–PCR. (C) Changes in IRF3 protein levels in HK‐2‐NC and HK‐2‐shKMT5a cells were assessed via western blotting. (D) Co‐IP assays were performed in HK‐2‐NC and HK‐2‐shKMT5a cells using anti‐IRF3 antibodies to detect the IRF3 methylation level. (E) After cycloheximide (CHX) treatment, the protein levels of IRF3 in HK‐2‐NC and HK‐2‐shKMT5a cells at 0 h, 2 h, 4 h, and 8 h were assessed via western blotting. (F) IHC staining for IRF3 in renal tissues from WT and KMT5a^KO mice with UUO for 28 days. (G) HK‐2 cells in the KMT5a^NC, KMT5a^sh + IRF3^vector and KMT5a^sh + IRF3^OE groups were cocultured with macrophages, and macrophage chemotaxis was assessed. (H) The concentrations of CCL2, CCL3, and CCL7 in the culture supernatants from the KMT5a^NC, KMT5a^sh + IRF3^vector, and KMT5a^sh + IRF3^OE groups were evaluated using ELISA. (I) CCL2, CCL3, and CCL7 expression in KMT5a^NC, KMT5a^sh + IRF3^vector, and KMT5a^sh + IRF3^OE HK‐2 cells was assessed via qRT–PCR. The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, ***p < 0.001, and ns, not significant, according to unpaired Student's t‐tests (B, C and F) or one‐way ANOVA (G, H and I).

3.5. The KMT5a Inhibitor UNC0379 Impeded Macrophage Chemotaxis In Vitro

UNC0379 is a classical small‐molecule inhibitor of KMT5a [7]. We investigated whether UNC0379 impedes KMT5a‐mediated renal fibrosis progression. After UNC0379 treatment, western blotting revealed that the IRF3 level in HK‐2 cells decreased in a dose‐dependent manner (Figure 5A). However, the transcription level of IRF3 was not affected (Figure 5B). In addition, the IRF3 protein stability was greatly impaired in HK‐2 cells with UNCO379 treatment (Figure S3A). Macrophage chemotaxis was also inhibited by UNC0379 in a dose‐dependent manner (Figure 5C). ELISA and qRT–PCR results demonstrated that the concentrations and mRNA levels of CCL2, CCL3, and CCL7 in HK‐2 cells gradually decreased after UNC0379 treatment (Figure 5D,E). Thus, these data suggest that UNC0379 impedes macrophage chemotaxis in vitro.

FIGURE 5.

The KMT5a inhibitor UNC0379 impeded macrophage chemotaxis in vitro. (A) After UNC0379 treatment (DMSO, 1 μM and 10 μM) for 48 h, the protein level of IRF3 in HK‐2 cells was assessed using western blotting. (B) After UNC0379 treatment (0 μM, 1 μM and 10 μM) for 48 h, the protein level of IRF3 in HK‐2 cells was assessed via qRT–PCR. (C) HK‐2 cell‐mediated macrophage chemotaxis was detected after treatment with UNC0379 (0 μM, 1 μM, and 10 μM). (D) After UNC0379 treatment (0 μM, 1 μM and 10 μM) for 48 h, the concentrations of CCL2, CCL3 and CCL7 in the culture supernatant of HK‐2 cells were evaluated via ELISA. (E) After UNC0379 treatment (0 μM, 1 μM and 10 μM) for 48 h, the expression of CCL2, CCL3 and CCL7 in HK‐2 cells was evaluated using qRT–PCR. The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, ***p < 0.001, and ns, not significant, according to one‐way ANOVA (A, B, C, D and E).

3.6. UNC0379 Suppressed Renal Fibrosis Progression In Vivo

Considering the promising data showing that UNC0379 inhibits macrophage chemotaxis in vitro, we investigated the efficacy of UNC0379 in UUO mice. UNC0379 (1 mg/kg every three days) was administered for 4 weeks, and renal tissues were dissected for evaluation. Masson and Sirius red staining revealed that UNC0379 significantly inhibited collagen deposition in the renal tissues of UUO mice. IHC staining also revealed that α‐SMA and IRF3 levels were decreased in the UNC0379 treatment group (Figure 6A,B). Consistent with those findings, western blotting revealed decreased levels of fibronectin, α‐SMA, and IRF3 in the UNC0379 treatment group (Figure 6C). IHC staining for CD206 and CD86 in renal tissues from mice in the control and UNC0379 groups suggested that UNC0379 suppressed macrophage infiltration in renal fibrosis in vivo (Figure 6D). Taken together, these data prove that UNC0379 suppresses renal fibrosis progression in vivo.

FIGURE 6.

UNC0379 suppressed renal fibrosis progression in vivo. (A) Representative images of Masson, Sirius red, and IHC staining for α‐SMA and IRF3 in renal tissues from UUO mice treated with DMSO or UNC0379 for 28 days. (B) Statistical analyses of Masson, Sirius red‐ and α‐SMA‐positive areas and the proportion of IRF3‐positive renal tissues in UUO mice treated with DMSO or UNC0379 for 28 days. (C) The fibronectin, α‐SMA, and IRF3 protein levels in renal tissues from UUO mice treated with DMSO or UNC0379 for 28 days were assessed via western blotting. (D) IHC staining for CD206⁺ macrophages and CD86⁺ macrophages in renal tissues from UUO mice treated with DMSO or UNC0379 for 28 days. The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, ***p < 0.001, and ns, not significant, according to unpaired Student's t‐tests (B and D).

4. Discussion

This study reveals a novel role for the histone methyltransferase KMT5a in the pathogenesis of renal fibrosis. We demonstrate that KMT5a expression is upregulated during fibrogenesis and drives renal fibrosis progression by orchestrating a profibrotic inflammatory response. Mechanistically, we revealed that KMT5a posttranslationally stabilizes the transcription factor IRF3 in renal tubular cells, leading to the increased secretion of macrophage chemotaxis‐related cytokines (CCL2, CCL3, and CCL7) and consequent infiltration of macrophages, a key promoter of renal fibrosis. The functional significance of this axis was firmly established through genetic knockout and pharmacological inhibition, both of which robustly attenuated fibrosis in vivo. Our findings thus position KMT5a as a critical epigenetic‐immune nexus in chronic kidney disease and a promising therapeutic target.

In chronic kidney disease, macrophage recruitment within glomerular and tubulointerstitial compartments is a unique pathological feature [17]. Macrophage enrichment can be a predictor of renal fibrosis progression and disease prognosis, including renal dysfunction, tubular dilatation, glomerular damage, and crescent formation [18]. Acute kidney injury is characterized by the enrichment of M1‐type macrophages. In contrast, chronic kidney injury involves a phenotypic shift, with a substantial proportion of M1‐type macrophages transitioning to M2 macrophages [19, 20]. Moreover, macrophages can also differentiate into myofibroblasts, which contribute to renal fibrosis progression [21]. In this study, we investigated the potential relationship between KMT5a and macrophage recruitment in renal fibrosis. KMT5a enhanced macrophage chemotaxis via IRF3 stabilization.

The discovery that KMT5a stabilizes the IRF3 protein in HK‐2 cells represents a significant conceptual advancement. Conventionally, KMT5a has been shown to play a role in histone H4K20 methylation, influencing gene silencing and genome stability [22, 23]. Our data reveal a noncanonical function in which KMT5a directly interacts with IRF3 and regulates its stabilization. IRF3 is a central hub in innate immune responses and is classically activated by viral infection via TBK1‐mediated phosphorylation [24]. Our findings reveal a novel, epigenetically linked activation pathway for IRF3 in sterile inflammation (fibrosis). The KMT5a‐mediated stabilization of IRF3 potentially lowers the threshold for its activation or prolongs its transcriptional activity for chemokine genes; this provides a direct molecular conduit from an epigenetic modifier to sustained inflammatory gene expression. Our results broaden the function of KMT5a from being a chromatin modulator to also being a direct regulator of inflammatory signaling complexes.

In this study, the novel small‐molecule inhibitor UNC0379 was used to treat renal fibrosis. This inhibitor has been widely investigated in the treatment of cancer, including colon cancer [7], bladder cancer [25] and Ewing sarcoma [26]. In this study, we confirmed that UNC0379 slows renal fibrosis progression. UNC0379, by inhibiting KMT5a methyltransferase activity, successfully decreased the protein expression of IRF3 and the expression of the chemokines CCL2, CCL3, and CCL7 (at both the protein level and the mRNA level), impaired macrophage chemotaxis in vitro, and depleted infiltrating macrophages in vivo, ultimately reducing collagen deposition. Given the challenges associated with macrophage depletion therapies, targeting specific recruitment signals from injured parenchyma (via KMT5a inhibition) offers a more precise and potentially safer alternative.

Our study has several limitations. First, while the UUO model is robust for proof‐of‐concept, validation in more chronic, clinically relevant models (e.g., diabetic nephropathy and hypertensive nephrosclerosis) is essential to confirm the broad role of this axis. Second, the precise molecular mechanism of IRF3 stabilization needs to be elucidated further: which lysine residue on IRF3 is methylated by KMT5a, and how this modification inhibits IRF3 degradation (e.g., by blocking ubiquitination)?

In summary, we identified a novel KMT5a‐IRF3‐chemokine signaling axis that serves as a critical amplifier loop in renal fibrosis by driving pathogenic macrophage infiltration. This work redefines KMT5a as a central epigenetic‐immune regulator in sterile inflammation and provides a compelling preclinical rationale for the development of KMT5a inhibitors as a new class of antifibrotic agents aimed at disrupting the vicious cycle of tubular injury and immune cell recruitment.

Author Contributions

M.Y.: Data curation, Writing Original draft preparation, Investigation; P.X.: Data curation, Visualization, Investigation; J.L.: Software, Validation; H.Y.: Supervision; Y.J.: Software, Validation; M.W.: Conceptualization, Methodology, Software, Writing Reviewing and Editing, Funding acquisition.

Funding

This study was supported by the author(s) declare that financial support was received for the research, authorship, and/or publication of this article. This work was supported by the Jiangsu Province Leading Talents Cultivation Project for Traditional Chinese Medicine (SLJ0330), the Jiangsu Province Sixth Phase 333 High‐Level Talents Project, the Jiangsu Provincial Medical Innovation Center (CXZX202233), the 2024 Suzhou Science and Education Driven Healthcare Enhancement Project (MSXM2024006), and the 2024 Research Projects of the Jiangsu Association of Traditional Chinese Medicine (PDJH2024040, CYTF2024042).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: A. The kidney function including glomerular filtration, Scr and BUN in the peripheral blood of mice from Sham and UUO group was measured. B. The KMT5a level in 5 human CKD samples and 5 human normal renal samples was detected using western blotting. C. The KMT5a level in 5 human CKD samples and 5 human normal renal samples was detected using qRT‐PCR.

The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, and ***p < 0.001; unpaired Student's t‐test (A and C).

Figure S2: A. Candidates immunoprecipitated from HK‐2 cells were identified via LC–MS; the top ten candidates based on the Sequest HT score are shown. B. The ChIP‐PCR data or IRF3 enrichment in the promoter of CCL2, CCL3 and CCL7. C. Representative images of Masson, Sirius red and IHC staining for α‐SMA in renal tissues from KMT5a^WT and KMT5a^KO UUO mice model accepting AAV‐IRF3 administration. D. Statistical analyses of Masson, Sirius red‐ and α‐SMA‐positive areas in renal tissues from KMT5a^WT and KMT5a^KO UUO mice model accepting AAV‐IRF3 administration.

The results are representative of at least three independent experiments; *p < 0.05, **p < 0.01, and ***p < 0.001; unpaired Student's t‐test (B) and one‐way ANOVA (D).

Figure S3: A. After UNC0379 (10 μM) treatment, the protein stability profile of IRF3 in HK‐2 cells at 0 h, 2 h, 4 h and 8 h were assessed via western blotting.

Data Availability Statement

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.