Nlrc5^-/- mice in C57BL/6N background, generated by crossing Nlrc5-floxed mice with CMV-Cre mice, were a generous gift from Dr. Dana Philpott (38). Wildtype C57Bl/6N mice were used as controls. Both groups of mice were bred and housed in ventilated cages on the same housing unit throughout the experiment. The experiments were done as and when the knockout mice became available. Therefore, the numbers of mice used per group in different experiments was variable and are indicated in the corresponding figure legends. All experimental protocols on animals were carried out with the approval of the Université de Sherbrooke Animal Ethics Committee (Protocol # 2018-2083, 359-18C).

Liver fibrosis was induced as we have described previously (39). Male mice were used for liver fibrosis induction as female sex hormones diminish inflammatory cytokine production in the liver (40). Briefly, CCl₄ (Sigma-Aldrich, Oakville, ON) diluted in corn oil (1:3) was injected via intraperitoneal (i.p) route (0.5μl CCl₄ per gram body weight) twice a week for five weeks. Three days after the last treatment, mice were euthanized, blood collected by cardiac puncture and liver tissues resected. Serum was separated and kept frozen at -80°C. Liver pieces were snap frozen and stored at -80°C for gene and protein expression studies and hydroxyproline assay. For histopathology analyses, 3-4 cubic mm size liver pieces from 4-5 different locations of the same liver were fixed for 12-16 hours in 4% paraformaldehyde solution and embedded in paraffin on the same tissue block.

Serum alanine transaminase (ALT) levels were measured using a kinetic assay (Pointe Scientific Inc, Brussels, Belgium) following manufacturer’s instructions. Hydroxyproline content was measured as described previously (39). Ten mg of liver tissue, homogenized in 1 mL of 6N HCl using the bead mill MM 400 (Retsch, Hann, Germany), was transferred to glass tubes, topped up with 2 mL of 6N HCl and the tubes were kept on a heat block for 16h at 110°C to hydrolyze proteins. After filtering the hydrolysate through Whatman #1 filter paper, aliquots were evaporated on a heat block and the residues were dissolved in 50% 2-propanol. Hydroxyproline standards and samples, distributed in a 96-well microtiter plate, were oxidized by adding chloramine T (Sigma-Aldrich; dissolved in 50% isopropanol and adjusted to pH 6.5 with acetate/citrate buffer). Following incubation at room temperature for 25 min, Ehrlich reagent [p-dimethylaminobenzaldehyde dissolved in n-propanol/perchloric acid (2:1)], was added and the samples incubated at 50°C for 10 min for color development. Absorbance at 550 nm was measured using the SPECTROstar Nano (BMG Labtech, Germany) spectrophotometer.

Liver sections were deparaffinized, rehydrated, and stained with hematoxylin and eosin (H&E) or Sirius red following standard procedures. For immunohistochemical detection of αSMA, rehydrated liver sections immersed in citrate buffer (pH 6.0) were given intermittent microwave treatment to retrieve antigenic epitopes. Following incubation in 3% hydrogen peroxide for 10 min to inhibit endogenous peroxidase activity, sections were blocked with 5% BSA in Tris-buffered saline (TBS) containing 20% Tween-20 (TBS-T). Slides were incubated overnight at 4°C with a rabbit mAb against mouse αSMA (Cell Signaling Technology, Cat #19245S) diluted in blocking buffer, washed and then incubated with horseradish peroxidase (HRP)-conjugated secondary Ab for 1 h. After thorough washing, a substrate solution containing 3,3’-diaminobenzidine (DAB; Sigma-Aldrich; 30 μL chromogen diluted in 1 mL of DAB liquid buffer) was added for 10 min. The sections were counterstained with hematoxylin and mounted with a coverslip. Images of the stained sections, digitized using the NanoZoomer Slide Scanner (Hamamatsu Photonics, Japan), were analyzed by the NanoZoomer Digital Pathology software NDPview2.0. Sirius red staining and αSMA-positive areas were quantified using the NIH ImageJ software (version 1.53e). Data from six randomly selected fields from different liver pieces for each of the three mice per group were used for quantification.

Total RNA from frozen tissues was extracted using QIAzol Lysis Reagent (Qiagen, Toronto, Ontario, Canada), according to the manufacturer’s instructions. cDNA was synthetized from 1µg of purified RNA using QuantiTect^® reverse transcription kit (Qiagen, Toronto, Ontario, Canada). Quantitative RT-PCR amplification reactions were carried out in CFX Connect Real-Time PCR Detection System (Bio-Rad, Canada) or QuantStudio 3 Real-Time PCR System (Thermo Fisher Scientific, Canada) using SYBR Green Supermix (Bio-Rad, Mississauga, Ontario, Canada). The expression of indicated genes was measured using primers listed in Supplementary Table S1 . Gene expression levels between samples were normalized based on the Cycle threshold (Ct) values compared to housekeeping gene 36B4 and the fold induction was calculated using the vehicle (oil)-treated wildtype mice as controls.

Serum TNF protein levels were quantified using a sandwich ELISA kit from eBioscience (Cat # 88-7324) following manufacturer’s instructions. Capture Ab diluted in coating buffer was added to high protein-binding 96-well plates (Nunc Maxisorp^®) and incubated overnight at 4°C. After washing with PBS-0.05% Tween-20 (wash buffer), the plates were blocked with assay diluent for 1 h at room temperature. Serum samples diluted 1:1 in assay diluent and serial dilutions of recombinant TNF standard were added in duplicates, and plates were incubated at room temperature for 2 h. After thorough washing, biotinylated detection antibody was added for 1 h followed by the addition of avidin-HRP for 30 min. After thorough washing, tetramethylbenzidine substrate solution was added for 15 min and color development was measured at 450 nm using SPECTROstar Nano. The values were plotted against the standard curve to calculate TNF protein levels in serum.

Mice liver tissue samples were taken in a 2 mL round bottom tube and homogenized using bead mill MM 400 (Retsch, Hann, Germany) containing TNE buffer (50mM Tris-HCl, 150mM NaCl, 1mM EDTA; pH 8.0) supplemented with phosphatase and protease inhibitor cocktails (Roche, Indianapolis, IN). TNE buffer containing detergents (0.2% SDS, 1% sodium deoxycholate and 1% Triton-X) was added in equal volumes into the homogenates and kept on rocker for 30 min at 4°C. Lysate was centrifuged for 20 min at 15,000 ×g and the supernatant collected. Protein concentration was determined using RC-DC Protein Assay Kit (Bio-Rad, Mississauga, ON). Protein samples containing 30-50 µg proteins were electrophoresed on SDS-PAGE gels and analysed by Western Blot. Primary Ab used are listed in Supplementary Table S2 . HRP-conjugated anti-mouse or anti-rabbit secondary antibodies and enhanced chemiluminescence reagents (ECL) were from GE Healthcare Life Sciences (Pittsburg, PA). Images of western blot were captured by the VersaDOC 5000 imaging system (Bio-Rad).

The numbers of mice in experimental and control groups for the two genotypes of mice in each experiment are indicated in corresponding figure legends. Data were analyzed using the GraphPad Prism9 (San Diego, CA). Statistical significance was calculated by two-way ANOVA with Tukey’s post-hoc test. p values <0.05 were considered significant.

TNFα, expressed by macrophages and hepatocytes in response to toll-like receptor signaling, contributes to liver fibrosis by activating HSC and immune cells (12). Loss of TNF receptor TNFR1 attenuates liver fibrosis induced by CCl₄ or bile duct ligation, accompanied by reduced expression of Col1a1 and Il6 genes and decreased NF-κB activation in liver tissues as well as in isolated HSCs (41, 42). NF-κB signaling promotes cell survival and proliferation of not only hepatocytes but also HSCs (42–44). As NLRC5 knockdown in HSCs was shown to increase NF-κB signaling (35), we examined whether NLRC5 deficiency promoted liver fibrosis in vivo. To this end, we induced liver fibrosis by intraperitoneal administration of CCl₄ in NLRC5-deficient and control mice for five weeks. Alterations in liver function were evaluated and histological and molecular changes were assessed. As shown in Figure 1A , both wildtype and Nlrc5^-/- mice showed comparable levels of liver damage as revealed by elevated serum levels of alanine transaminase (ALT). Hematoxylin and eosin-stained liver sections showed similar features of hepatocyte damage and mononuclear cell infiltration in both wildtype and Nlrc5^-/- mice ( Figure 1B ). Together these results indicated that loss of NLRC5 does not increase hepatocyte damage induced by chronic chemical injury.

Next, we compared the extent of liver fibrosis in CCl₄-treated Nlrc5^-/- and control mice. Sirius red staining of collagen fibers revealed comparable pattern and distribution of fibrotic areas in Nlrc5^-/- and wildtype mice that was also confirmed by quantification of the stained areas ( Figures 2A, B ). Moreover, measurement of hydroxyproline, which is enriched in connective tissue collagen fibers (45), was increased in CCl₄-treated wildtype mice ( Figure 2C ). Interestingly, Nlrc5^-/- mice treated with vehicle (corn oil, control) showed significantly elevated hydroxyproline content compared to wildtype mice. Because of such elevated hydroxyproline content in Nlrc5^-/- mice, the CCl₄-mediated increase in this group was not statistically significant, even though these levels are appreciably higher than in CCl₄-treated wildtype mice ( Figure 2C ). These observations suggested that NLRC5 deficiency may augment certain aspects of the hepatic fibrogenic response that is not discernible in the presence of strong fibrogenic inducers such as CCl₄.

As fibrogenesis is mainly driven by HSCs activation and their differentiation to myofibroblasts (13), we evaluated the expression of the Acta2 gene coding for αSMA and that of Pdgfb coding for platelet-derived growth factor beta, a mitogen for HSC. The induction of Acta2 was significantly high in CCl₄-treated wildtype mice livers but showed only marginal increase in Nlrc5^-/- mice. On the other hand, Pdgfb upregulation was significantly elevated in the livers of CCl₄-treated Nlrc5^-/- mice but less prominently in control mice ( Figure 3A ). However, the upregulation of Acta2 and Pdgfb genes was not significantly different between CCl₄-treated wildtype and Nlrc5^-/- mice. Moreover, immunohistochemical staining of αSMA in the liver sections from vehicle- or CCl₄- treated mice showed a comparable increase in pattern and staining of myofibroblast distribution in CCl₄- treated wildtype and Nlrc5^-/- mice that was also confirmed by digital quantification of the stained areas ( Figures 3B, C ). These findings indicated that NLRC5 deficiency does not markedly affect myofibroblast differentiation during chemically induced liver fibrosis.

Consistent with the comparable levels of myofibroblast differentiation in Nlrc5^-/- and wildtype mice livers following CCL₄ treatment, genes encoding the fibrillar collagens, collagen 1α1 and collagen 3α1 (46) were strongly induced in both groups ( Figure 4A ). Similarly, the gene coding for the ECM modifying enzyme MMP2 and tissue inhibitor of MMPs-2 (Mmp2, Timp2), which respectively exert anti- and pro-fibrogenic roles in liver fibrosis (47–49), were strongly upregulated by CCL₄ treatment in both Nlrc5^-/- and control mice livers ( Figure 4B ). However, Mmp3 and Timp1 genes, whose impact on liver fibrosis is controversial or unclear (49), were strongly induced in wildtype mice livers but showed significantly lower or negligible induction in NLRC5-deficient livers ( Figure 4B ). These findings indicate that NLRC5 deficiency does not appreciably affect the induction of many fibrogenic response genes and that the observed differences caused by NLRC5 deficiency are not strong enough to influence the severity of liver fibrosis.

Liver fibrosis establishes feed forward loops involving pro-inflammatory and profibrogenic cytokine gene expression by immune cells and their recruitment by chemokines (50, 51). To determine how NLRC5 deficiency affects these processes, we first evaluated the expression of candidate genes implicated in these processes. NLRC5-deficient livers displayed a significantly higher induction of the pro-fibrogenic tumor necrosis factor gene Tnf ( Figure 5A ). Serum TNF levels were elevated in both control and Nlrc5^-/- mice following CCl₄ treatment ( Figure 5B ). Notably, vehicle-treated Nlrc5^-/- mice displayed appreciably higher levels of TNF than control mice. The interleukin-1β gene Il1b did not show appreciable induction following CCl₄ treatment in control livers but was significantly elevated in NLRC5-deficient livers due to lower expression in the oil-treated group ( Figure 5A ). The transcript levels of IL-6, a survival cytokine, was appreciably lower in Nlrc5^-/- livers ( Figure 5A ). The Tgfb gene coding for the key fibrogenic cytokine transforming growth factor beta showed comparable upregulation in both groups following CCl₄ treatment ( Figure 5C ). On the other hand, the antifibrogenic interferon gamma gene Ifng was appreciably reduced in wildtype livers following CCl₄ treatment, whereas Nlrc5^-/- livers showed a significant upregulation ( Figure 5C ). These findings indicate that NLRC5 deficiency did cause an upregulation of hepatic Tnf gene expression and systemic TNF protein levels, but this did not result in increased liver fibrosis.

The key producer cells of TNF during liver fibrosis are activated liver-resident Kupffer cells and monocyte-derived macrophages, which are recruited by chemokines expressed in the inflamed liver (51). As NLRC5-deficient mice showed elevated TNF expression, we evaluated the gene expression of the macrophage recruiting chemokine CCL2 (macrophage chemoattractant protein-1) and the T cell chemoattractant CCL5, as well as CX3CR1, the receptor for CX3CL1 (fractalkine) expressed on monocyte-derived macrophages and required for their homeostasis (52, 53). Whereas the expression of Ccl2 and Ccl5 showed only marginal induction in both wildtype and Nlrc5^-/- livers, Cx3cr1 was strongly upregulated in both groups ( Figure 6A ). Next, we examined the gene expression of macrophage markers CD68 and F4/80 (ADGRE1) and T lymphocytes markers CD3ϵ and CD8α. As shown in Figure 6B , the fibrotic livers of both control and NLRC5-deficient mice showed increased expression of Cd68 and Adgre1, and the latter was significantly higher in Nlrc5^-/- livers. Whereas F4/80 is highly expressed in tissue-resident macrophages, CD68 is expressed in both tissue-resident and infiltrating macrophages (54, 55). The T cell marker transcript levels were not markedly altered by CCl₄ treatment in both groups of mice. These findings suggest that NLRC5 deficiency increases the activation of liver-resident macrophages, which presumably contributes to elevated Tnf expression.

Finally, we examined the protein expression of molecules associated with fibrosis and signaling events reported to be regulated by NLRC5 in whole liver homogenates. CCl₄-treated wildtype and Nlrc5^-/- mice livers showed increased levels of αSMA and MMP2 compared to vehicle-treated control groups ( Figure 7A ), reflecting the increased transcript levels of Acta2 and Mmp2 genes in the fibrotic livers ( Figures 3A , 4B ). Notably, phosphorylation of the p65 subunit of NF-κB, which occurs downstream of diverse inflammatory signaling pathways including TNF (56), was found to be elevated in vehicle-treated Nlrc5^-/- mice livers compared to wildtype control mice and this p65 phosphorylation was sustained following CCl₄ treatment, with a concomitant decrease in total IκB ( Figure 7B ). This observation is consistent with the findings in the HSC cell line LX-2 following NLRC5 knockdown (37). However, phosphorylation of SMAD3, which occurs downstream of TGFβ signaling and reported to be reduced by NLRC5 knockdown in LX-2 cells (36), was reduced in Nlrc5^-/- mice livers with or without CCl₄ treatment, whereas phosphorylation of SMAD2 was comparable to control mice livers ( Figure 7C ). These results indicate that NLRC5 deficiency deregulates NF-κB activation and may also modulate the SMAD signaling pathway in the liver.