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Inhibition of RAC1 activator DOCK2 ameliorates cholestatic liver injury via regulating macrophage polarisation and hepatic stellate cell activation

Biology Direct volume 20, Article number: 21 (2025) Cite this article

Abstract

Background

The Rho GTPase Rac family small GTPase 1 (RAC1) is considered a promising fibrotic therapeutic target, but the role of its activator, dedicator of cytokinesis 2 (DOCK2), in liver fibrosis is largely unknown. This study aimed to investigate the expression and role of DOCK2 in cholestasis-induced liver fibrosis and to further explore the potential mechanisms.

Results

Cholestasis was induced in male C57BL/6 mice by bile duct ligation (BDL). DOCK2 knockdown was achieved by tail vein injection of adenovirus containing DOCK2-targeting shRNA. The effect of DOCK2 knockdown on cholestatic liver injury was evaluated at different time points after BDL. Hepatic DOCK2 expression gradually increased after BDL. Knockdown of DOCK2 reduced the necrotic area in BDL liver and downregulated serum levels of liver injury indicators. At 3d post-BDL (acute phase), DOCK2 knockdown alleviated M1 macrophage inflammation in the liver, as evidenced by reduced infiltrating iNOS + macrophages and inflammatory cytokines and mitigated NLRP3 inflammasome activation. At 14d post-BDL (chronic phase), DOCK2 knockdown suppressed hepatic stellate cell (HSC) activation and liver fibrosis as indicated by decreased α-SMA + HSCs and extracellular matrix deposition. In vitro experiments further demonstrated that DOCK2 knockdown suppressed M1 macrophage polarisation and HSC to myofibroblast transition, accompanied by inhibition of RAC1 activation.

Conclusions

In summary, this study demonstrates for the first time that the RAC1 activator DOCK2 regulates M1 macrophage polarisation and hepatic stellate cell activation to promote cholestasis-induced liver inflammation and fibrosis, suggesting that DOCK2 may be a potential therapeutic target in cholestatic liver injury.

Graphical Abstract

Background

Cholestasis is a condition characterised by impaired bile formation and/or flow to the bile ducts, resulting in the retention of toxic hydrophobic bile salts and other bile constituents in hepatocytes, triggering inflammatory responses and compromising cell membrane integrity. These elevated metabolites in the systemic circulation will further contribute to the dysfunction of other organs [1]. Cholestatic liver injury can progress to hepatic fibrosis, cirrhosis and liver failure. In cases of end-stage cholestatic diseases, liver transplantation is the only available therapeutic option [2]. Currently, the pathophysiology of the progression of cholestatic liver injury is largely undefined.

The liver is composed of different cell types and these cells cooperate to maintain liver function and homeostasis. Macrophages are characterised by their diversity and plasticity and can be polarised into M1 and M2 phenotypes. M1 macrophage inflammation is known to promote liver fibrosis, and suppressing macrophage infiltration and M1 polarisation has been shown to reduce liver inflammation and promote regression of liver fibrosis [3,4,5]. Hepatic stellate cells (HSCs) are located in the perisinusoidal space of Disse, which lies between liver sinusoidal endothelial cells and hepatocytes. The quiescent HSCs are responsible for the storage, transport and metabolism of vitamin A [6]. Upon chronic stimuli such as cholestasis, HSCs are activated to transdifferentiate into proliferative and pro-fibrogenic myofibroblasts that produce fibrous collagens and extracellular matrix (ECM) remodelling factors, leading to excessive ECM production and deposition, termed fibrosis [6].

Rac family small GTPase 1 (RAC1) is a member of the Rac family of Rho GTPases that becomes active when bound to GTP and is considered a potential fibrotic target [7, 8]. Persistent activation of RAC1 in HSCs has been shown to exacerbate carbon tetrachloride (CCl4)-induced liver injury and facilitate fibrosis in mice [8], and targeting RAC1 has been shown to prevent biliary duct ligation (BDL)-induced liver fibrosis in rats [9, 10]. Dedicator of cytokinesis 2 (DOCK2) is an atypical RAC activator that controls RAC1 activation by catalysing the exchange of GDP for GTP [11, 12]. DOCK2 and engulfment and cell motility protein 1 (ELMO1) typically form a closed, auto-inhibited binary complex. The conformational change of the ELMO1 subunit promotes the auto-inhibited state to an active state, resulting in the exposure of binding sites for RAC1 on DOCK2 [13].

As an activator of RAC1, DOCK2 has been implicated in inflammatory diseases [14]. DOCK2 activates pro-inflammatory macrophages to promote the expression of pro-inflammatory genes [15]. Targeting DOCK2 impairs pulmonary macrophage recruitment and infiltration [14, 16], as well as reduces RAC1 activation and reactive oxygen species (ROS) production [17]. In addition, DOCK2 facilitates the transition from lung fibroblast to myofibroblast, thereby promoting pulmonary fibrosis [18]. DOCK2 also mediates mesothelial to mesenchymal transition and promotes matrix deposition, thereby contributing to pleural fibrosis [19]. However, the role of DOCK2 in hepatic macrophages and HSCs and the contribution of DOCK2 to cholestasis-induced liver fibrosis are poorly characterised.

In this study, we measured liver DOCK2 expression during the acute and chronic phases of cholestasis-induced liver injury and evaluated the effects and underlying mechanisms of adenovirus-mediated DOCK2 knockdown on liver inflammation and fibrosis. The results of this work would contribute to a better understanding of the underlying molecular and cellular features during the progression of cholestatic liver injury.

Methods

Animal experiments

Male C57BL/6 mice (RRID: MGI:2159769) aged 6 and 8 weeks were used for modelling. Cholestasis was induced in mice by BDL, which can induce highly reproducible liver fibrosis in mice [20]. BDL surgery was performed as previously reported [21]. In brief, after midline laparotomy, the common bile duct was exposed, double-ligated with 4 − 0 silk and transected between the ligations. Mice in the sham group underwent similar surgery without ligation and transection of the common bile duct. Mice were injected with adenovirus containing DOCK2-targeting shRNA (Ad_shDOCK2) or non-targeting shRNA (Ad_shNC) via the tail vein (30 µL of 1 × 109 PFU/mL) two days before BDL, and injections were maintained once a week for two weeks, as previously reported with slight modifications [22, 23]. At 3d, 1w and 2w after BDL, mice were sacrificed and serum and liver samples were collected and processed for subsequent experiments. The ratio of liver weight to body weight (LW/BW) was measured. All animal experiments were approved by the Laboratory Animal Welfare and Ethics Committee of Henan University of Traditional Chinese Medicine and conducted following the US National Research Council’s Guide for the Care and Use of Laboratory Animals (8th edition) and the ARRIVE guidelines.

Cell culture and treatment

RAW264.7 mouse macrophages (Cat. #: iCell-m047) and immortalised mouse HSCs (Cat. #: iCell-0094a) were purchased from Shanghai iCell Bioscience Inc. RAW264.7 cells were cultured in DMDM medium supplemented with 10% fetal bovine serum. HSCs were cultured in an HSC culture system purchased from iCell (Cat. #: PriMed-iCell-009). To induce M1 polarisation, RAW264.7 cells were incubated with 100 ng/mL lipopolysaccharide (LPS) for 6 h. To induce the transition to myofibroblasts, HSCs were incubated with 10 ng/mL transforming growth factor (TGF)-β1 for 24 h. To knock down DOCK2, RAW264.7 cells and HSCs were transduced with Ad_shDOCK2 or Ad_shNC.

Real-time quantitative PCR (qPCR)

Total RNA was isolated from liver homogenates or cells using TRIpure reagent (BioTeke, Beijing, China). The RNA concentrations were determined using Nanodrop 2000 (ThermoFisher Scientific). Complementary DNA synthesis was carried out using BeyoRT™ II M-MLV reverse transcriptase (Beyotime Biotechnology, Shanghai, China) and RNase inhibitor (Biosharp Life Sciences, Hefei, China). Real-time qPCR was conducted on an Exicycler™ 96 instrument (Bioneer Company, Daejeon, Republic of Korea) using PCR MasterMix supplemented with SYBR Green (Solarbio, Beijing, China). β-actin (Actb) primers were CATCCGTAAAGACCTCTATGCC (forward) and ATGGAGCCACCGATCCACA (reverse). DOCK2 primers were GTGGCTGTCCCTATTGAA (forward) and TCTTGCTGTCTCCCTTGA (reverse). Alpha smooth muscle actin (α-SMA) primers were GAGCGTGAGATTGTCCG (forward) and GCTGTTATAGGTGGTTTCG (reverse). Collagen type I alpha 1 chain (Col1a1) primers were GACGCCATCAAGGTCTACTG (forward) and AATCCATCGGTCATGCTCT (reverse). Collagen type III alpha 1 chain (Col3a1) primers were TGCCCACAGCCTTCTACAC (forward) and GCCAGGGTCACCATTTCTC (reverse). Interleukin 6 (IL-6) primers were TAACAGATAAGCTGGAGTC (forward) and TAGGTTTGCCGAGTAGA (reverse). Tumour necrosis factor (TNF)-α primers were ACAGAAAGCATGATCCGCGA (forward) and TTGCTACGACGTGGGCTAC (reverse). Inducible nitric oxide synthase (iNOS) primers were CACCACCCTCCTCGTTC (forward) and CAATCCACAACTCGCTCC (reverse).

Biochemical analysis and assessment of pro-inflammatory cytokines

Serum levels of total bilirubin (T-Bil), aspartate aminotransferase (AST) and alanine aminotransferase (ALT) were determined using commercially available detection kits from Beijing Solarbio Science & Technology Co., Ltd. (T-Bil Cat. #: BC5180) and Nanjing Jiancheng Bioengineering Institute (AST Cat. #: C010; ALT Cat. #: C009). Levels of pro-inflammatory cytokines (IL-6, TNF-α and iNOS) in liver tissue were measured using enzyme-linked immunosorbent assay kits purchased from Wuhan FineTest Biotech Co., Ltd. (iNOS Cat. #: EM0272) and Hangzhou Multisciences (Lianke) Biotech, Co., Ltd. (IL-6 Cat. #: EK206; TNF-α Cat. #: EK282).

Histopathology, immunofluorescence (IF) and immunohistochemistry (IHC)

Paraffin-embedded liver specimens were sectioned into 5-µm thick slices, deparaffinised and rehydrated. To assess pathological changes such as necrosis, and collagen fibre deposition, the slices were stained with haematoxylin and eosin (H&E), Sirius red and Masson’s trichrome. After dehydration and permeabilization, the slices were sealed with neutral resin and photographed at 40 and 200 magnifications. For IHC and IF staining, after antigen retrieval and blocking, the slices were incubated with primary antibodies against DOCK2 (Bioss Corporation, Beijing, China; Cat. #: bs-12431R), NLR family pyrin domain containing 3 (NLRP3; Affinity; Cat. #: DF7438, RRID: AB_2839376), cleaved caspase-1 (Affinity; Cat. #: AF4005, RRID: AB_2845463), iNOS (Affinity; Cat. #: AF0199, RRID: AB_2833391), α-SMA (Novus Biologicals, Littleton, CO, USA; Cat. #: NBP2-33006, RRID: AB_3094464), F4/80 (Santa; Cat. #: Sc-377009, RRID: AB_2927461), Rac1 (ProteinTech Ltd., Wuhan, China; Cat. #: 66122-1-Ig, RRID: AB_2881521), and GTP-RAC1 (Neweast Biosciences, PA, USA; Cat. #: 26903, RRID: AB_1961793) overnight at 4 °C. The slices were then incubated with the corresponding horseradish peroxidase-conjugated (Solarbio; Cat. #: SE134, RRID: AB_2797593) or fluorescence dye-labelled secondary antibodies (Proteintech; Cat. #: SA00009-2 and SA00003-1, RRID: AB_2890957 and RRID: AB_2890896) and photographed at 200, 400 or 600 magnification.

Western blot and active RAC1 pull-down assay

Cell or tissue samples were lysed. The BCA protein assay kit (ProteinTech) was used to determine protein concentrations. Equal amounts of proteins were separated in 5-15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and then transferred onto polyvinylidene fluoride membranes (ThermoFisher Scientific, Pittsburgh, PA, USA). The membranes were blocked in 5% skimmed milk/TBST and then incubated with primary antibodies and horseradish peroxidase-conjugated secondary antibodies, followed by visualization using the ECL method.

RAC1 activation was evaluated using the Rac1 Pull-down Activation Assay Biochem Kit (Cytoskeleton, Inc., CO, USA; Cat. #: BK035) following the manufacturer’s instructions. The kit provided PAK-PBD protein beads that specifically pull down the GTP-bound form of RAC proteins. The pull-down GTP-RAC1 proteins were visualized through the western blot using the anti-RAC1 monoclonal antibody supplied with the kit.

Gene expression omnibus (GEO) datasets

To find differentially expressed DOCK members between control and BDL livers, we analysed three mRNA microarray datasets from the Gene Expression Omnibus (GEO) database (https://www.ncbi.nlm.nih.gov/gds/). The GSE152494 [24] series contains 6 control livers and 6 BDL livers; the GSE140498 [25] and GSE40041 [26] series each contain 3 control livers and 3 BDL livers. Differentially expressed genes (DEGs) were identified using GEO2R online tools (https://www.ncbi.nlm.nih.gov/geo/geo2r/) with|Log2FC|≥0.6 and p < 0.05. Circular heatmaps were generated using the ComplexHeatmap (version 2.18.0) and circlize (version 0.4.16) packages in R (version 4.3.2). Box plots were generated using the ggplot2 (version 3.5.0) and ggpubr (version 0.6.0) packages.

High throughput RNA sequencing (RNA-seq)

DOCK2-silenced RAW264.7 cells and HSCs and control cells were treated with LPS for 6 h or TGF-β1 for 24 h. Total RNA was extracted from the cells for RNA-seq. Genes that met the criteria of|Log2FC|>1 and p < 0.05 were identified as DEGs. DEGs were then subjected to Gene Ontology (GO) Biological Process (BP) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis. Venn diagrams were generated using the eulerr package (version 7.0.1) in R (version 4.3.2). Volcano plots were generated using the ggplot2 package (version 3.5.0). Heat maps were generated using the pheatmap package (version 1.0.12). Enriched GO terms and KEGG pathways were visualised using the package ggplot2 (version 3.5.0).

Statistical analysis

GraphPad Prism 9.0 (RRID: SCR_002798) was used for statistical analysis and graphing. One-way analysis of variance (ANOVA) with post-hoc multiple tests was applied for comparisons among groups. A p-value less than 0.05 was considered statistically significant.

Results

DOCK2 is upregulated in the liver of BDL mice

RAC1 is a known target in liver fibrosis [8,9,10], but the role of its activator, DOCK, in liver fibrosis remains unknown. To identify candidate DOCK members potentially involved in the progression of cholestasis, we used publicly available BDL mouse liver transcriptome datasets, GSE152494 [24], GSE140498 [25], and GSE40041 [26]. We found that five DOCK members (1, 2, 7, 8, 10 and 11) are differentially expressed (Fig. 1A-C;|Log2FC|≥0.6 and p < 0.05). Notably, DOCK2 is the common differential gene among the three datasets and its expression was significantly higher in BDL liver than in control liver (Fig. 1D).

Fig. 1
figure 1

DOCK2 is increased in the liver of BDL mice. (A-C) Circular heatmaps demonstrating the expression of DOCK family genes in three BDL mouse liver transcriptome datasets. (D) Box plot showing DOCK2 expression in three BDL mouse liver transcriptome datasets. **P < 0.01 and *P < 0.05. (E) Schematic illustration of experimental protocols. (F) Relative mRNA levels of DOCK2 in livers. (G) Representative microphotographs of IHC staining of DOCK2 in liver sections. Magnification: 200 fold. Scale bar: 100 μm

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BDL mice develop time-dependent pathological and molecular changes, making them suitable for studying the pathogenesis of cholestatic liver injury that progresses over time. However, the liver samples in the above datasets were all collected at the chronic stage of the disease (more than 4 weeks after BDL), so it was unclear whether DOCK2 expression had changed in the acute stage. Therefore, we evaluated the hepatic expression of DOCK2 at different time points (3d, 1w and 2w) after BDL (Fig. 1E). As determined by real-time PCR, DOCK2 expression was gradually upregulated over the disease progression (Fig. 1F), which was further confirmed by IHC results (Fig. 1G). These results suggest a potentially important role for DOCK2 in the progression of BDL-induced liver injury.

Knockdown of DOCK2 alleviates BDL-induced liver injury

To determine the effect of DOCK2 knockdown on BDL-induced liver injury, we injected mice with Ad-shDOCK2 or Ad-shNC two days before BDL surgery. The experimental protocol is shown in Fig. 2A. IHC staining showed that DOCK2 staining was weaker after Ad-shDOCK2 injection, confirming adenoviral vector-mediated interference with DOCK2 expression (Fig. 2B). Compared to the sham group, BDL livers exhibited significant fibrosis, whereas Ad-shDOCK2 caused an obvious restoration of fibrotic liver in BDL mice (Fig. 2C) and decreased the LW/BW ratio of BDL mice (Fig. 2D). Upon histopathological observation, DOCK2 knockdown reduced the necrotic area in the liver after BDL (Fig. 2E). Furthermore, the elevated serum levels of liver injury indicators (T-Bil, ALT and AST) in BDL mice were reduced after DOCK2 knockdown (Fig. 2F). Taken together, these results suggest that DOCK2 knockdown alleviates BDL-induced liver injury.

Fig. 2
figure 2

Silencing DOCK2 alleviates BDL-induced liver injury. (A) Schematic illustration of experimental protocols. (B) Representative microphotographs of IHC staining of DOCK2 in liver sections. Magnification: 200 fold. Scale bar: 100 μm. (C) Representative photographs of mouse livers. (D) Liver weight (LW)/body weight (BW) of mice. (E) Representative microphotographs of H&E staining of liver sections. Magnification: 40 fold. Scale bar: 500 μm. (F) Serum levels of total bilirubin (T-Bil), ALT (alanine aminotransferase), and AST (aspartate aminotransferase)

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Knockdown of DOCK2 suppresses liver inflammation 3d post-BDL

Since hepatic DOCK2 expression gradually increased after BDL, we first tested whether DOCK2 was involved in hepatic macrophage inflammation in the acute phase (3d post-BDL). IF staining showed that the majority of F4/80 + macrophages in BDL livers were also positive for DOCK2, and the infiltrated F4/80+/DOCK2 + macrophages in BDL livers were reduced by DOCK2 knockdown (Fig. 3A). Similarly, DOCK2 knockdown reduced the elevated pro-inflammatory M1 (F4/80+iNOS+) macrophages in BDL livers (Fig. 3B). These results indicate that DOCK2 is associated with M1 macrophage infiltration in BDL livers. Consistently, the elevated levels of M1 macrophage-related pro-inflammatory cytokines (IL-6, TNF-α and iNOS) in liver tissues were significantly reduced by DOCK2 knockdown (Fig. 3C). Furthermore, BDL livers showed strong staining for NLRP3 and cleaved caspase 1, suggesting NLRP3 inflammasome activation, which was mitigated in Ad-shDOCK2-infected mice (Fig. 3D and E).

Fig. 3
figure 3

Silencing DOCK2 mitigates liver inflammation 3d post-BDL. (A) Representative microphotographs of double IF staining of DOCK2 and F4/80 or (B) iNOS and F4/80. Magnification: 400 fold. Scale bar: 50 μm. (C) Levels of M1 macrophage-related pro-inflammatory cytokines (IL-6, TNF-α, and iNOS) in liver tissues. (D) Representative microphotographs of IHC staining of NLRP3 (E) Caspase 1. Magnification: 200 fold. Scale bar: 100 μm

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Knockdown of DOCK2 ameliorates liver fibrosis 2w post-BDL

We next investigated the effects of DOCK2 knockdown on long-term BDL-induced liver fibrosis. As shown in Figs. 2w and 4A BDL caused an increase in the number of DOCK2+/α-SMA + cells in the liver, indicating that DOCK2 expression was increased in activated HSCs, whereas Ad_shDOCK2-mediated silencing of DOCK2 reduced the number of DOCK2+/α-SMA + cells. As indicated by Sirius Red and Masson’s trichrome staining, the ECM deposition in BDL livers was attenuated by DOCK2 knockdown (Fig. 4B and C). Additionally, the increased mRNA expression of Acta2 (α-SMA encoding gene) and major fibre genes (Col1a1 and Col3a1) in 2w BDL livers was reduced by DOCK2 knockdown (Fig. 4D). Taken together, these results suggest that DOCK2 deficiency ameliorates long-term BDL-induced liver fibrosis.

Fig. 4
figure 4

Knockdown of DOCK2 attenuates liver fibrosis after 2w post-BDL. (A) Representative microphotographs of double IF staining of DOCK2 and α-SMA. Magnification: 200 fold and 600 fold. Scale bar: 100 μm and 50 μm. (B) Representative microphotographs of Sirius Red staining. Magnification: 200 fold. Scale bar: 100 μm. (C) Representative microphotographs of Masson’s trichrome staining. Magnification: 200 fold. Scale bar: 100 μm. (D) Relative mRNA levels of fibrotic markers (Acta2, Col1a1 and Col3a1) in livers

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DOCK2 activates RAC1 to promote progression of BDL-induced liver injury

We next investigated whether RAC1 was involved in the role of DOCK2 in the progression of BDL-induced liver injury. As shown by double IF staining, DOCK2 and RAC1 were co-localised in the liver, and this co-localisation was strong in both 3d and 2w BDL livers and was reduced in livers harvested from Ad_shDOCK2-infected mice (Fig. 5A and B). Additionally, DOCK2 silencing caused a marked reduction in activated RAC1 (GTP-RAC1) in both 3d and 2w BDL livers (Fig. 5C and D). Taken together, these results demonstrate that DOCK2 silencing inhibits the activation of RAC1 during the progression of BDL-induced liver injury.

Fig. 5
figure 5

DOCK2 activates RAC1 to participate in BDL-induced liver injury and its deficiency suppressed LPS-induced M1 macrophage polarisation. (A and B) Representative microphotographs of double IF staining of DOCK2 and RAC1 in 3d and 2w BDL livers. Magnification: 600 fold. Scale bar: 50 μm. (C and D) Representative microphotographs of IF staining of GTP-RAC1 in 3d and 2w BDL livers. The quantification of mean fluorescence intensity (MFI) is shown in the right panels. Magnification: 400 fold. Scale bar: 50 μm

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DOCK2 knockdown suppressed LPS-induced M1 polarisation of mouse macrophages

Next, we treated RAW264.7 mouse macrophages with LPS to induce M1 polarisation. The experimental protocol is shown in Fig. 6A. Compared to control cells, DOCK2 expression was increased upon LPS stimulation (Fig. 6B). Real-time PCR confirmed Ad_shDOCK2-mediated silencing of DOCK2 in macrophages (Fig. 6C). In parallel, the number of iNOS-positive M1 macrophages was increased upon LPS stimulation but was reduced in the DOCK2-silenced group (Fig. 6D). Consistently, LPS-induced mRNA expression of IL-6, TNF-α and iNOS was also downregulated by DOCK2 silencing (Fig. 6E). In addition, as evidenced by IF staining (Fig. 6F) and pull-down (Fig. 6G) of GTP-RAC1, LPS-induced activation of RAC1 was mitigated by Ad_shDOCK2-mediated knockdown of DOCK2 (Fig. 6H).

Fig. 6
figure 6

Knockdown of DOCK2 suppressed LPS-induced M1 polarisation of mouse macrophages. (A) Schematic illustration of experimental protocols. (B) Protein levels of DOCK2 6 h after 100 ng/mL LPS stimulation. (C) Relative mRNA level of DOCK2 in RAW264.7 cells 48 h after adenovirus transduction. (D) 48 h after adenovirus transduction, cells were stimulated with 100 ng/mL LPS. Representative microphotographs of IF staining of iNOS 6 h later. Magnification: 400 fold. Scale bar: 50 μm. (E) Relative mRNA levels of M1 macrophage cytokines (IL-6, TNF-α, and iNOS). (F) Representative microphotographs of IF staining of GTP-RAC1 in cells. Magnification: 400 fold. Scale bar: 50 μm. The quantification of mean fluorescence intensity (MFI) is shown in the right panel. (G) The pull-down assay of GTP-RAC1 by PAK-PBD protein beads. (H) Protein levels of DOCK2 in cells

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Furthermore, we performed mRNA-seq to illustrate the changes in the expression profile in DOCK2-silenced M1 macrophages. The volcano plot and heat map of DEGs are shown in Figure S1A and B. A total of 522 downregulated and 314 upregulated DEGs were identified. GO enrichment and KEGG pathway analysis revealed that these DEGs were enriched in biological processes and pathways related to immune response and ECM organisation (Figure S1C).

DOCK2 knockdown suppressed the TGF-β1-induced transition of mouse HSCs to myofibroblasts

Finally, we investigated the role of DOCK2 in the TGF-β1-induced transition of mouse HSCs to myofibroblasts. The experimental protocol is shown in Fig. 7A. Similarly, DOCK2 expression was increased in TGF-β1-induced myofibroblasts (Fig. 7B). Therefore, we knocked down DOCK2 in mouse HSCs by Ad_shDOCK2 transduction (Fig. 7C). We found that knockdown of DOCK2 reduced fibrotic α-SMA expression in TGF-β1-induced myofibroblasts (Fig. 7D and E). IF staining (Fig. 7F and G) and pull-down assay (Fig. 7H) of GTP-RAC1 showed that TGF-β1-induced activation of RAC1 was also mitigated by DOCK2 knockdown (Fig. 7I).

Fig. 7
figure 7

Knockdown of DOCK2 inhibits TGF-β1-induced activation of HSC cells. (A) Schematic illustration of experimental protocols. (B) Protein levels of DOCK2 24 h after 10 ng/mL TGF-β1 stimulation. (C) Relative mRNA levels of DOCK2 in HSCs 48 h after adenovirus transduction. (D) 24 h after adenovirus transduction, cells were stimulated with 10 ng/mL TGF-β1. Representative microphotographs of IF staining of α-SMA 24 h later. Magnification: 400 fold. Scale bar: 50 μm. (E) Protein levels ofα-SMA in cells. (F) Representative microphotographs of IF staining of GTP-RAC1 in cells. Magnification: 400 fold. Scale bar: 50 μm. (G) The quantification of MFI. (H) The pull-down assay of GTP-RAC1 by PAK-PBD protein beads. (I) Protein levels of DOCK2 in cells. (J) Venn analysis showing the common differentially expressed genes (DEGs;|Log2FC|>1 and p < 0.05) between DOCK2-silenced M1 macrophages and myofibroblasts. (K) GO biological processes and KEGG pathways enriched for these common DEGs

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Next, we performed mRNA-seq to illustrate the changes in the expression profile in DOCK2-silenced myofibroblasts. The volcano plot and heat map of DEGs are shown in Figure S1D and E. A total of 508 downregulated and 213 upregulated DEGs were identified. GO enrichment and KEGG pathway analysis revealed that these DEGs were mainly enriched in biological processes and pathways related to cell growth regulation and ECM organisation (Figure S1F).

By performing Venn analysis, we obtained 377 common DEGs in both M1 macrophages and myofibroblasts (Fig. 7J). Interestingly, in addition to the previously indicated biological processes and pathways related to immune response, cell growth regulation and ECM organisation, these common DEGs were also enriched in PI3K-Akt, PPAR, and TGF-β pathways, suggesting their involvement in the molecular mechanisms underlying DOCK2 in M1 macrophage polarisation and HSC activation (Fig. 7K).

Discussion

The morphological, biochemical and molecular changes in the liver after BDL surgery are complex and time-dependent. Typically, the acute phase of injury (up to 3 days) is characterised by an inflammatory response manifested by inflammatory gene expression and immune cell infiltration, and the chronic phase of injury (more than 10 days) often shows persistent inflammation and progressive fibrosis [20]. In the early phase of cholestatic liver injury (3d post-BDL), fibrosis is not yet evident, but a significant increase in hepatic DOCK2 can be observed and this increase persists until 2w post-BDL. This finding suggests that increased DOCK2 contributes to or mediates liver fibrosis rather than being the result of fibrosis.

Impaired liver function is marked by elevated serum enzymes (ALT, AST, alkaline phosphatase and γ-glutamyltranspeptidase) and total bilirubin concentration. As previously reported, bile acids modulate the release of alkaline phosphatase from the canalicular membrane of rat hepatocytes [27]. Hepatic and serum bile acids are elevated within the first 6 h after BDL surgery and this elevation persists for up to 14 days [28], which may contribute to the sustained increase in serum enzymes and total bilirubin concentration after BDL, along with the steady increase in liver infarct areas [29]. In this study, we found that the elevated serum enzymes and total bilirubin in BDL mice were significantly reversed by DOCK2 knockdown. In parallel, DOCK2 knockdown ameliorated the morphological and histopathological alterations in BDL livers, supporting the hepatoprotective effects of DOCK2 silencing.

Bile acids can act as inflammatory initiators, especially in BDL models [28, 30]. As previously determined, the proportion of liver-infiltrating CD11b + F4/80 + macrophages in the liver was markedly increased 3d and 7d after BDL [31]. M1 macrophages produce multiple inflammatory cytokines such as IL-1β and TNF-α, which promote the survival of activated HSCs [32]. Besides, macrophages also produce various pro-fibrogenic factors such as TGF-β and platelet-derived growth factor, thereby contributing to the activation of quiescent HSCs [33, 34]. iNOS is an enzyme responsible for catalysing the production of reactive nitrogen species. iNOS-producing hepatic macrophages exert pro-inflammatory and pro-fibrogenic functions, producing TGF-β to promote HSC activation [35]. In contrast, genetic or pharmacological inhibition of iNOS markedly mitigated liver fibrosis [36, 37].

The accumulated bile acids can act as endogenous danger-associated molecular patterns, thereby activating signals of the NLRP3 inflammasome in macrophages [38]. The activated NLRP3 inflammasome will further induce inflammatory cell death, contributing to liver fibrosis [39, 40]. As early as 2d after BDL, the activated NLRP3 inflammasome can be observed in the liver [41], and the levels of NLRP3 and cleaved caspase 1 were gradually increased during the progression of liver injury [42]. Hou et al. reported that the majority (approximately 80%) of NLRP3 + cells in BDL liver were F4/80 + macrophages; blockade of macrophage-specific NLRP3 inflammasome activation can attenuate liver inflammation and fibrosis [42]. In this study, we observed that in the acute phase (3d after BDL), DOCK2 knockdown decreased the number of infiltrating F4/80 + and iNOS + macrophages in BDL livers, mitigated the activation of NLRP3 inflammasome, and reduced the levels of M1 inflammatory cytokines, suggesting that targeting DOCK2 contributes to the inhibition of the M1 macrophage-mediated inflammatory response in the acute phase of cholestatic liver injury.

Liver fibrosis also follows a time-dependent process. At 3d after BDL, α-SMA + activated HSCs first appeared around the infarcted bile duct and their number gradually increased until 14d after BDL; in parallel, the accumulation of collagen also gradually increased until 14 d, with no further increase thereafter [43]. Importantly, sustained activation of HSCs leads to excessive production and deposition of ECM and secretion of chemokines to recruit inflammatory cells, which in turn activate HSCs through direct interactions or indirect cytokines, leading to the progression of fibrosis [6, 44]. In this study, we demonstrated that DOCK2 knockdown had an anti-fibrotic effect at 14d post-BDL, suggesting that targeting DOCK2 has a significant therapeutic effect in cholestatic liver fibrosis.

Mounting studies have shown that RAC1 is closely involved in liver inflammation and fibrosis. Activation of RAC1 causes a significant increase in the expression of pro-inflammatory cytokines by macrophages [45,46,47]. In contrast, interfering with RAC1 activation by genetic ablation or pharmacological inhibition would suppress macrophage M1 polarisation and potently reduce the production of pro-inflammatory cytokines (IL-6 and TNF-α) [48,49,50], and interestingly, without affecting macrophage recruitment [48]. Additionally, RAC1 activated the Hedgehog pathway to stimulate the myofibroblastic transition of HSCs and increased the viability of myofibroblastic HSCs, thereby facilitating the fibrosis process [51]. Here in this study, we observed that the activation of RAC1 in macrophages and HSCs in the liver was suppressed with DOCK2 knockdown, in both the acute and the chronic phases of disease; therefore, we proposed that DOCK2 participates in BDL-induced liver injury through the activation of RAC1. Considering that DOCK2-mediated activation of RAC1 involves the conformational change of ELMO1, the role of ELMO1 in the context of BDL-induced liver injury needs to be identified.

We also noticed an important point in this work. In addition to infiltrating monocyte-derived macrophages, resident Kupffer cells are also important components of hepatic macrophages and are often involved in the progression of liver fibrosis [52]. More complexly, Kupffer cells have been found to play a binary role in cholestatic liver disease, either protecting hepatocytes from damage or promoting NLRP3 inflammasome-mediated inflammation and fibrosis [53, 54]. As both Kupffer cells and monocyte-derived macrophages were positive for F4/80, we cannot distinguish these two cell types in F4/80 + macrophage populations. Therefore, it is difficult to distinguish the contribution of monocyte-derived macrophages and Kupffer cells in DOCK2-mediated macrophage inflammation, which requires further investigation.

Conclusions

Summing up, in this work we observed that hepatic DOCK2 expression was gradually increased during the progression of BDL-induced cholestatic liver injury in mice. Adenovirus vector-mediated knockdown of DOCK2 rescued the impaired liver function of BDL mice, reduced acute inflammatory responses and ameliorated the chronic fibrotic phenotype. Mechanistically, DOCK2 regulated M1 macrophage polarisation and HSC-myofibroblast transition by promoting RAC1 activation. These findings suggest that DOCK2 may play an important role in the development and progression of cholestatic liver fibrosis.

Data availability

Data and material will be provided on request to the corresponding authors.

Abbreviations

ALT:

Alanine aminotransferase

AST:

Aspartate aminotransferase

α-SMA:

alpha smooth muscle actin

BDL:

Biliary duct ligation

BP:

Biological process

CCl4 :

Carbon tetrachloride

Col1a1:

Collagen type I alpha 1 chain

Col3a1:

Collagen type III alpha 1 chain

DEGs:

Differentially expressed genes

DOCK2:

Dedicator of cytokinesis 2

ECM:

Extracellular matrix

ELMO1:

Engulfment and cell motility protein 1

GO:

Gene ontology

H&E:

Haematoxylin and eosin

HSCs:

Hepatic stellate cells

IF:

Immunofluorescence

IHC:

Immunohistochemistry

IL-6:

Interleukin 6

iNOS:

Inducible nitric oxide synthase

KEGG:

Kyoto encyclopedia of genes and genomes

LPS:

Lipopolysaccharide

NLRP3:

NLR family pyrin domain containing 3

qPCR:

real-time quantitative PCR

RAC1:

Rac family small GTPase 1

ROS:

Reactive oxygen species

T-Bil:

Total bilirubin

TGF:

Transforming growth factor

TNF:

Tumour necrosis factor

References

  1. Pollheimer MJ, Fickert P, Stieger B. Chronic cholestatic liver diseases: clues from histopathology for pathogenesis. Mol Aspects Med. 2014;37:35–56.

    Article  CAS  PubMed  Google Scholar 

  2. Allen K, Jaeschke H, Copple BL. Bile acids induce inflammatory genes in hepatocytes: a novel mechanism of inflammation during obstructive cholestasis. Am J Pathol. 2011;178(1):175–86.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. She S, Wu X, Zheng D, Pei X, Ma J, Sun Y, et al. PSMP/MSMP promotes hepatic fibrosis through CCR2 and represents a novel therapeutic target. J Hepatol. 2020;72(3):506–18.

    Article  CAS  PubMed  Google Scholar 

  4. Rao J, Wang H, Ni M, Wang Z, Wang Z, Wei S, et al. FSTL1 promotes liver fibrosis by reprogramming macrophage function through modulating the intracellular function of PKM2. Gut. 2022;71(12):2539–50.

    Article  CAS  PubMed  Google Scholar 

  5. Miura K, Yang L, van Rooijen N, Ohnishi H, Seki E. Hepatic recruitment of macrophages promotes nonalcoholic steatohepatitis through CCR2. Am J Physiol Gastrointest Liver Physiol. 2012;302(11):G1310–21.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  6. Tsuchida T, Friedman SL. Mechanisms of hepatic stellate cell activation. Nat Rev Gastroenterol Hepatol. 2017;14(7):397–411.

    Article  CAS  PubMed  Google Scholar 

  7. Heasman SJ, Ridley AJ. Mammalian rho GTPases: new insights into their functions from in vivo studies. Nat Rev Mol Cell Biol. 2008;9(9):690–701.

    Article  CAS  PubMed  Google Scholar 

  8. Choi SS, Sicklick JK, Ma Q, Yang L, Huang J, Qi Y, et al. Sustained activation of Rac1 in hepatic stellate cells promotes liver injury and fibrosis in mice. Hepatology. 2006;44(5):1267–77.

    Article  CAS  PubMed  Google Scholar 

  9. Ghoreshi ZA, Kabirifar R, Safari F, Karimollah A, Moradi A, Eskandari-Nasab E. Hepatoprotective effects of curcumin in rats after bile duct ligation via downregulation of Rac1 and NOX1. Nutrition. 2017;36:72–8.

    Article  CAS  PubMed  Google Scholar 

  10. Ghoreshi ZA, Kabirifar R, Khodarahmi A, Karimollah A, Moradi A. The preventive effect of atorvastatin on liver fibrosis in the bile duct ligation rats via antioxidant activity and down-regulation of Rac1 and NOX1. Iran J Basic Med Sci. 2020;23(1):30–5.

    PubMed  PubMed Central  Google Scholar 

  11. Gotoh K, Tanaka Y, Nishikimi A, Nakamura R, Yamada H, Maeda N, et al. Selective control of type I IFN induction by the rac activator DOCK2 during TLR-mediated plasmacytoid dendritic cell activation. J Exp Med. 2010;207(4):721–30.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Cherfils J, Zeghouf M. Regulation of small GTPases by GEFs, GAPs, and GDIs. Physiol Rev. 2013;93(1):269–309.

    Article  CAS  PubMed  Google Scholar 

  13. Chang L, Yang J, Jo CH, Boland A, Zhang Z, McLaughlin SH, et al. Structure of the DOCK2-ELMO1 complex provides insights into regulation of the auto-inhibited state. Nat Commun. 2020;11(1):3464.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Qian G, Adeyanju O, Sunil C, Huang SK, Chen SY, Tucker TA, et al. Dedicator of cytokinesis 2 (DOCK2) Deficiency attenuates Lung Injury Associated with Chronic High-Fat and High-Fructose Diet-Induced obesity. Am J Pathol. 2022;192(2):226–38.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Xu X, Su Y, Wu K, Pan F, Wang A. DOCK2 contributes to endotoxemia-induced acute lung injury in mice by activating proinflammatory macrophages. Biochem Pharmacol. 2021;184:114399.

    Article  CAS  PubMed  Google Scholar 

  16. Namkoong H, Edahiro R, Takano T, Nishihara H, Shirai Y, Sonehara K, et al. DOCK2 is involved in the host genetics and biology of severe COVID-19. Nature. 2022;609(7928):754–60.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. Ma X, Tan X, Yu B, Sun W, Wang H, Hu H, et al. DOCK2 regulates antifungal immunity by regulating RAC GTPase activity. Cell Mol Immunol. 2022;19(5):602–18.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Guo X, Adeyanju O, Sunil C, Mandlem V, Olajuyin A, Huang S, et al. DOCK2 contributes to pulmonary fibrosis by promoting lung fibroblast to myofibroblast transition. Am J Physiol Cell Physiol. 2022;323(1):C133–44.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Qian G, Adeyanju O, Roy S, Sunil C, Jeffers A, Guo X, et al. DOCK2 promotes Pleural Fibrosis by modulating Mesothelial to Mesenchymal Transition. Am J Respir Cell Mol Biol. 2022;66(2):171–82.

    Article  CAS  PubMed  Google Scholar 

  20. Tag CG, Sauer-Lehnen S, Weiskirchen S, Borkham-Kamphorst E, Tolba RH, Tacke F et al. Bile duct ligation in mice: induction of inflammatory liver injury and fibrosis by obstructive cholestasis. J Vis Exp. 2015(96).

  21. Buryova H, Chalupsky K, Zbodakova O, Kanchev I, Jirouskova M, Gregor M, et al. Liver protective effect of ursodeoxycholic acid includes regulation of ADAM17 activity. BMC Gastroenterol. 2013;13:155.

    Article  PubMed  PubMed Central  Google Scholar 

  22. Borkham-Kamphorst E, Huss S, Van de Leur E, Haas U, Weiskirchen R. Adenoviral CCN3/NOV gene transfer fails to mitigate liver fibrosis in an experimental bile duct ligation model because of hepatocyte apoptosis. Liver Int. 2012;32(9):1342–53.

    Article  CAS  PubMed  Google Scholar 

  23. Chung YH, Huang YH, Chu TH, Chen CL, Lin PR, Huang SC, et al. BMP-2 restoration aids in recovery from liver fibrosis by attenuating TGF-β1 signaling. Lab Invest. 2018;98(8):999–1013.

    Article  CAS  PubMed  Google Scholar 

  24. Gijbels E, Vilas-Boas V, Annaert P, Vanhaecke T, Devisscher L, Vinken M. Robustness testing and optimization of an adverse outcome pathway on cholestatic liver injury. Arch Toxicol. 2020;94(4):1151–72.

    Article  CAS  PubMed  Google Scholar 

  25. Cubero FJ, Mohamed MR, Woitok MM, Zhao G, Hatting M, Nevzorova YA, et al. Loss of c-Jun N-terminal kinase 1 and 2 function in liver epithelial cells triggers biliary hyperproliferation resembling Cholangiocarcinoma. Hepatol Commun. 2020;4(6):834–51.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Zhao G, Hatting M, Nevzorova YA, Peng J, Hu W, Boekschoten MV, et al. Jnk1 in murine hepatic stellate cells is a crucial mediator of liver fibrogenesis. Gut. 2014;63(7):1159–72.

    Article  CAS  PubMed  Google Scholar 

  27. Hatoff DE, Hardison WG. Bile acid-dependent secretion of alkaline phosphatase in rat bile. Hepatology. 1982;2(4):433–9.

    Article  CAS  PubMed  Google Scholar 

  28. Zhang Y, Hong JY, Rockwell CE, Copple BL, Jaeschke H, Klaassen CD. Effect of bile duct ligation on bile acid composition in mouse serum and liver. Liver Int. 2012;32(1):58–69.

    Article  PubMed  Google Scholar 

  29. Abshagen K, König M, Hoppe A, Müller I, Ebert M, Weng H, et al. Pathobiochemical signatures of cholestatic liver disease in bile duct ligated mice. BMC Syst Biol. 2015;9:83.

    Article  PubMed  PubMed Central  Google Scholar 

  30. Cai SY, Ouyang X, Chen Y, Soroka CJ, Wang J, Mennone A, et al. Bile acids initiate cholestatic liver injury by triggering a hepatocyte-specific inflammatory response. JCI Insight. 2017;2(5):e90780.

    Article  PubMed  PubMed Central  Google Scholar 

  31. Cabrera-Rubio R, Patterson AM, Cotter PD, Beraza N. Cholestasis induced by bile duct ligation promotes changes in the intestinal microbiome in mice. Sci Rep. 2019;9(1):12324.

    Article  PubMed  PubMed Central  Google Scholar 

  32. Pradere JP, Kluwe J, De Minicis S, Jiao JJ, Gwak GY, Dapito DH, et al. Hepatic macrophages but not dendritic cells contribute to liver fibrosis by promoting the survival of activated hepatic stellate cells in mice. Hepatology. 2013;58(4):1461–73.

    Article  CAS  PubMed  Google Scholar 

  33. Dooley S, ten Dijke P. TGF-β in progression of liver disease. Cell Tissue Res. 2012;347(1):245–56.

    Article  CAS  PubMed  Google Scholar 

  34. Kocabayoglu P, Lade A, Lee YA, Dragomir AC, Sun X, Fiel MI, et al. β-PDGF receptor expressed by hepatic stellate cells regulates fibrosis in murine liver injury, but not carcinogenesis. J Hepatol. 2015;63(1):141–7.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. Karlmark KR, Weiskirchen R, Zimmermann HW, Gassler N, Ginhoux F, Weber C, et al. Hepatic recruitment of the inflammatory Gr1 + monocyte subset upon liver injury promotes hepatic fibrosis. Hepatology. 2009;50(1):261–74.

    Article  CAS  PubMed  Google Scholar 

  36. Aram G, Potter JJ, Liu X, Torbenson MS, Mezey E. Lack of inducible nitric oxide synthase leads to increased hepatic apoptosis and decreased fibrosis in mice after chronic carbon tetrachloride administration. Hepatology. 2008;47(6):2051–8.

    Article  CAS  PubMed  Google Scholar 

  37. Cinar R, Iyer MR, Liu Z, Cao Z, Jourdan T, Erdelyi K et al. Hybrid inhibitor of peripheral cannabinoid-1 receptors and inducible nitric oxide synthase mitigates liver fibrosis. JCI Insight. 2016;1(11).

  38. Hao H, Cao L, Jiang C, Che Y, Zhang S, Takahashi S, et al. Farnesoid X receptor regulation of the NLRP3 Inflammasome underlies Cholestasis-Associated Sepsis. Cell Metab. 2017;25(4):856–e675.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Wree A, Eguchi A, McGeough MD, Pena CA, Johnson CD, Canbay A, et al. NLRP3 inflammasome activation results in hepatocyte pyroptosis, liver inflammation, and fibrosis in mice. Hepatology. 2014;59(3):898–910.

    Article  CAS  PubMed  Google Scholar 

  40. Qu J, Yuan Z, Wang G, Wang X, Li K. The selective NLRP3 inflammasome inhibitor MCC950 alleviates cholestatic liver injury and fibrosis in mice. Int Immunopharmacol. 2019;70:147–55.

    Article  CAS  PubMed  Google Scholar 

  41. Boaru SG, Borkham-Kamphorst E, Tihaa L, Haas U, Weiskirchen R. Expression analysis of inflammasomes in experimental models of inflammatory and fibrotic liver disease. J Inflamm. 2012;9:1–16.

    Article  Google Scholar 

  42. Hou L, Yang L, Chang N, Zhao X, Zhou X, Dong C, et al. Macrophage sphingosine 1-Phosphate receptor 2 Blockade attenuates liver inflammation and fibrogenesis triggered by NLRP3 inflammasome. Front Immunol. 2020;11:1149.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Georgiev P, Jochum W, Heinrich S, Jang JH, Nocito A, Dahm F, et al. Characterization of time-related changes after experimental bile duct ligation. Br J Surg. 2008;95(5):646–56.

    Article  CAS  PubMed  Google Scholar 

  44. Chang J, Hisamatsu T, Shimamura K, Yoneno K, Adachi M, Naruse H, et al. Activated hepatic stellate cells mediate the differentiation of macrophages. Hepatol Res. 2013;43(6):658–69.

    Article  PubMed  Google Scholar 

  45. Yi YS, Kim HG, Kim JH, Yang WS, Kim E, Jeong D, et al. Syk-MyD88 Axis is a critical determinant of inflammatory-response in activated macrophages. Front Immunol. 2021;12:767366.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  46. Jin J, Qian H, Wan B, Zhou L, Chen C, Lv Y, et al. Geranylgeranyl diphosphate synthase deficiency hyperactivates macrophages and aggravates lipopolysaccharide-induced acute lung injury. Am J Physiol Lung Cell Mol Physiol. 2021;320(6):L1011–24.

    Article  CAS  PubMed  Google Scholar 

  47. Fu H, Alabdullah M, Großmann J, Spieler F, Abdosh R, Lutz V, et al. The differential statin effect on cytokine production of monocytes or macrophages is mediated by differential geranylgeranylation-dependent Rac1 activation. Cell Death Dis. 2019;10(12):880.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  48. Nagase M, Kurihara H, Aiba A, Young MJ, Sakai T. Deletion of Rac1GTPase in the myeloid lineage protects against inflammation-mediated kidney Injury in mice. PLoS ONE. 2016;11(3):e0150886.

    Article  PubMed  PubMed Central  Google Scholar 

  49. Bandaru S, Ala C, Ekstrand M, Akula MK, Pedrelli M, Liu X, et al. Lack of RAC1 in macrophages protects against atherosclerosis. PLoS ONE. 2020;15(9):e0239284.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  50. Fu H, Zhang P, Zhao XD, Zhong XY. Interfering with Rac1-activation during neonatal monocyte-macrophage differentiation influences the inflammatory responses of M1 macrophages. Cell Death Dis. 2023;14(9):619.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Choi SS, Witek RP, Yang L, Omenetti A, Syn WK, Moylan CA, et al. Activation of Rac1 promotes hedgehog-mediated acquisition of the myofibroblastic phenotype in rat and human hepatic stellate cells. Hepatology. 2010;52(1):278–90.

    Article  CAS  PubMed  Google Scholar 

  52. Tacke F. Targeting hepatic macrophages to treat liver diseases. J Hepatol. 2017;66(6):1300–12.

    Article  CAS  PubMed  Google Scholar 

  53. Osawa Y, Seki E, Adachi M, Suetsugu A, Ito H, Moriwaki H, et al. Role of acid sphingomyelinase of Kupffer cells in cholestatic liver injury in mice. Hepatology. 2010;51(1):237–45.

    Article  CAS  PubMed  Google Scholar 

  54. Holtmann TM, Inzaugarat ME, Knorr J, Geisler L, Schulz M, Bieghs V et al. Bile acids activate NLRP3 inflammasome, promoting Murine Liver inflammation or fibrosis in a cell type-specific manner. Cells. 2021;10(10).

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Acknowledgements

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Funding

This study was funded by the Natural Science Foundation of Henan Province (232300420057), China Postdoctoral Science Foundation (2020M670026ZX) and the National Natural Science Foundation of China (82474570).

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Author notes

  1. Jianli Qiu, Yitong Qu and Yinli Li contributed equally to this work.

Authors and Affiliations

  1. Department of Pediatrics, The First Affiliated Hospital of Henan University of Chinese Medicine, Zhengzhou, Henan, China

    Jianli Qiu, Yitong Qu, Yinli Li, Cancan Li, Junling Wang, Lu Meng, Xiaojin Jing, Jiangping Fu & Yan Xu

  2. Department of Pediatrics, Henan University of Chinese Medicine, Zhengzhou, Henan, China

    Jianli Qiu, Yitong Qu, Yinli Li, Cancan Li, Junling Wang, Lu Meng, Xiaojin Jing, Jiangping Fu & Yan Xu

  3. Department of Pharmacy, The First Affiliated Hospital of Zhengzhou University, Zhengzhou, China

    Yuna Chai

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Contributions

J.L.Q., Y.T.Q., and Y.L.L. contributed to Methodology, Investigation, Formal analysis, Writing– original draft. C.C.L. and J.L.W. contributed to Investigation, Data curation, Formal analysis. L.M., X.J.J., and J.P.F. contributed to Visualization, Validation. Y.X. and Y.N.C. contributed to Conceptualization, Supervision, Resources, Writing– review and editing. J.L.Q. contributed to Funding acquisition. All authors have read and approved the final manuscript.

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Correspondence to Jianli Qiu, Yan Xu or Yuna Chai.

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Qiu, J., Qu, Y., Li, Y. et al. Inhibition of RAC1 activator DOCK2 ameliorates cholestatic liver injury via regulating macrophage polarisation and hepatic stellate cell activation. Biol Direct 20, 21 (2025). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s13062-025-00612-3

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