Human recombinant TGFβ1 was purchased from Peprotech (Rocky Hill, CT, United States). Dulbecco’s modified Eagle’s Medium (DMEM), penicillin, streptomycin, and L-glutamine were obtained from Corning (NYC, NY, United States). Fetal bovine serum (FBS) was purchased from Invitrogen (NYC, NY, United States), and rottlerin and paxilline were from Sigma–Aldrich (Saint Louis, MO, United States). Antibodies against SMAD3, p-SMAD3, JAK2, p-JAK2, and β-actin were obtained from Cell Signaling Technology (Danvers, MA, United States), the anti-α-SMA (ACTA2) antibody was from Novus Biology (Centennial, CO, United States), and the anti-BK-Slo1 was purchased from Alomone Labs (Jerusalem, Israel). Alexa Fluor 680-conjugated anti-mouse and anti-rabbit secondary antibodies were purchased from Life Technologies (Eugene, OR, United States). Alanine transaminase (ALT) and aspartate transaminase (AST) detection reagents were obtained from Shanghai Shensuo UNF Medical Diagnostic Articles Corporation (Shanghai, China).

Isolation and culture of liver HSCs from adult male Sprague–Dawley rats (body weight, 500–600 g) were performed as previously described (Seki et al., 2007). Briefly, rats were anesthetized with 1 mL of ketamine (10%) and 50 μL of xylazin (2%) per kg body weight according to an institutionally approved protocol (Shanghai Jiao Tong University Institutional Animal Care and Use Committee). The peritoneum was then exposed and the liver quickly perfused via the portal vein with 40 mL of Hank’s balanced salt solution (HBSS) without Ca²⁺ and Mg²⁺ at a flow rate of 10 mL/min. Next, the liver was perfused with 30 mL (1 mg/mL) of pronase E followed by 30 mL (0.25 mg/mL) of collagenase H. The liver was carefully separated and transferred to HBSS (with Ca²⁺ and Mg²⁺) containing pronase E (0.2 mg/mL) and collagenase H (0.25 mg/mL), and the liver lobes were cut into small pieces and digested for 20 min at 37°C with continuous stirring. The hepatocytes were centrifuged at 50 × g and the cell suspension was filtered through 100-μm cell strainers, washed four times with HBSS (with Ca²⁺ and Mg²⁺), and subjected to Nycodenz (13.2%) gradient centrifugation at 1400 × g for 20 min at 4°C. HSCs were concentrated in the interphase as a white, ring-like structure. After harvesting and washing with HBSS, the HSCs were seeded on uncoated plastic tissue culture dishes in DMEM supplemented with 10% FBS, 4 mM L-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin. The first change of medium was performed 24 h after seeding. Quiescent HSCs were used for experiments 2–3 days after seeding, and activated HSCs were used 9–10 days after seeding. The human hepatic stellate cell line LX2 was cultured under the same conditions as for the rat HSCs. TGFβ1 (4 ng/mL) was added to the culture medium to test protein phosphorylation of HSCs at different time points or to differentiate LX2 cells into activated myofibroblast-like cells. All the cells were grown and maintained at 37°C and 5% CO₂ in a humidified incubator.

Total RNA was extracted from primary rat HSCs and LX2 cells using Trizol reagent (Invitrogen, Carlsbad, CA, United States). cDNA was synthesized using a reverse transcription kit (TaKaRa, Dalian, China). Real-time PCR was performed using 2 × SYBR Green qPCR Master Mix (Bimake, Houston, TX, United States). RNA was amplified using the primers indicated in Table 1. Relative mRNA expression levels were calculated and normalized to the levels of endogenous beta-actin in the same samples. Real-time PCR was performed in triplicate for each experiment and repeated at least three times and data was analyzed by 2^–△△CT method.

Cells were seeded in 96-well plates at a density of 5 × 10³ cells per well overnight, and then exposed to chemicals at the indicated concentrations for 48 h. Subsequently, 10 μL of MTS was added to each well and incubated for a minimum of 1 h. Measurements were made in accordance with the manufacturer’s instructions (Promega Corp., Madison, WI, United States). Metabolic activity was recorded as the relative colorimetric change measured at 492 nm. Each experiment was performed at least three times.

The EdU cell proliferation assay was performed according to the manufacturer’s instructions. Briefly, LX2 cells were transfected with KCNMA1-expressing plasmids or KCNMA1 siRNA for 48 h, and then plated in 96-well plates and incubated for 24 h. After incubation, the EdU solution (10 mM) was added directly to the culture medium and incubated for a further 30 min to ensure capture of most of the proliferating cells. The cells were subsequently fixed in 4% paraformaldehyde. Incorporation of EdU was performed by incubating the fixed cells with 2% bovine serum albumin (BSA) in PBS for 30 min and Alexa Fluor 488 for a further 30 min under Cu(I)-catalyzed click reaction conditions. The cells were washed with PBS and counterstained with Hoechst 33342 (1:1000) in PBS before detection by fluorescence microscopy using ImageXpress^® Micro 4 (Molecular Devices, San Jose, CA, United States).

Cells were lysed on ice with lysis buffer (50 mM Tris–HCl, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, and 0.1% SDS, pH 7.5) in the presence of a protease inhibitor mixture and phosphatase inhibitors (both from Roche, Mannheim, Germany). Protein concentrations were measured using the BCA protein assay kit (Thermo Fisher Scientific, Rockford, IL, United States). Samples were added to 5 × SDS loading buffer, boiled for 10 min, and loaded onto 10% polyacrylamide-SDS gels. The proteins were then transferred to PVDF membranes (Millipore, Billerica, MA, United States) at 300 mA for different durations, and the membranes were incubated in 5% BSA for 1 h at room temperature. After incubation with primary antibodies (Tables 2, 3) overnight at 4°C, the membranes were washed three times (10 min each) in TBS containing 0.1% Tween 20 (TBST) and then incubated with peroxidase- or fluorophore-conjugated secondary antibodies for 1 h in TBST at room temperature. The blots were detected by the ProteinSimple system (San Jose, CA, United States) with an ECL substrate or a LiCor imaging system (Lincoln, NE, United States).

Macroscopic currents were recorded from HSCs in a whole-cell configuration using an EPC-10 amplifier controlled by PatchMaster software (HEKA Electronics, Ludwigshafen/Rhein, Rheinland-Pfalz, Germany) as previously described (Han et al., 2016). Patch pipettes were prepared from borosilicate glass (Warner, Hamden, CT, United States) and had a resistance of 2–5 M Ω when filled with a pipette solution containing 140 mM K-aspartate, 4.3 mM CaCl₂, 2.06 mM MgCl₂, 5 mM EGTA, and 10 mM HEPES, pH 7.2. Free Ca²⁺ concentrations were calculated to be 1 μM using the Maxchelator program (Stanford University). The standard external recording solution comprised 150 mM sodium aspartate, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 10 mM glucose, and 10 mM HEPES, pH 7.4. Currents were elicited every 5 s by 200 ms voltage ramps from −100 to 80 mV. Data analysis and curve fitting were performed using FitMaster (HEKA Electronics, Ludwigshafen/Rhein, Rheinland-Pfalz, Germany) and figures were prepared with Origin 8.5 (OriginLab, Northampton, MA, United States).

Overexpression or downregulation of human KCNMA1 (AAB65837) was accomplished by transient transfection of plasmids containing the KCNMA1 gene or siRNA targeting KCNMA1 into LX2 cells for 48–72 h. Lipofectamine^TM 2000 (Thermo Fisher Scientific, Carlsbad, CA, United States) was used for transient transfection according to the manufacturer’s instructions.

The migration capacity of human LX2 cells and activated primary rat HSCs was measured by transwell assay (Corning, NY, United States) according to the manufacturer’s instructions. Briefly, 5 × 10⁴ cells were seeded into the upper chamber of transwell inserts. DMEM supplemented with 10% (v/v) FBS was added to the bottom chamber. Cells were incubated at 37°C and allowed to migrate for 16 h for LX2 cells and 12 h for activated rat HSCs, respectively. At the designated time points, cells were fixed in 4% paraformaldehyde and stained with 0.2% crystal violet, and non-migrating cells on the upper chamber surface were removed with cotton swabs. The number of cells on the lower surface was counted in five random fields under a light microscope.

For immunostaining, cells were fixed in 4% (w/v) paraformaldehyde in PBS for 1 h at room temperature. After fixation, the cells were rinsed with PBS and then permeabilized with 0.3% Triton X-100. Non-specific binding was blocked with 10% (w/v) BSA for 1 h at room temperature. The cells were then incubated with the respective primary antibodies (Table 3) overnight at 4°C. After extensive washing, cells were incubated with secondary antibodies for 1 h at room temperature, and nuclei were stained with DAPI. Confocal microscopy and image acquisition were carried out using a Nikon A1Si laser scanning confocal microscope. For liver section staining, paraffin sections (5 μm) were deparaffinized, rehydrated through a graded ethanol series, incubated with 0.3% hydrogen peroxide for 30 min, and blocked with goat serum. The sections were then incubated with the anti-ACTA2 antibody overnight at 4°C, followed by labeling with a horseradish peroxidase (HRP)-conjugated antibody (Thermo Fisher Scientific, Eugene, OR, United States) at room temperature for 1 h. Staining was visualized with DAB (Gene Tech, Shanghai, China). All sections were observed and imaged under a microscope (Carl Zeiss, Jena, Germany).

Eighteen pathogen-free male Sprague–Dawley rats (∼200 g) were randomly assigned to three groups, namely, control, fibrosis (CCl₄-treated), and rottlerin-treated groups. The animals in the fibrosis group were injected intraperitoneally with 0.2 mL/100 g of a CCl₄/olive oil mix (1:1) twice weekly, at equal intervals, for 8 weeks, as previously described (Constandinou et al., 2005). The rats in the rottlerin-treated group were injected intraperitoneally with rottlerin (2.5 mg/kg body weight) 2 weeks after the first CCl₄ injection. All the animal experiments were conducted according to protocols approved by the Shanghai Jiao Tong University Institutional Animal Care and Use Committee.

LX2 cells at 60–70% confluence were plated in 96-well plates and transiently transfected with 0.15 μg of the pGL4.48[luc2P/SBE/Hygro] vector (Promega, Madison, WI, United States) and 0.015 μg of the Renilla luciferase control reporter vector (Promega) using FuGENE 6 transfection reagent (Promega) according to the manufacturer’s instructions. SBE means SMAD binding element. Co-transfection with a vector expressing Renilla luciferase under the control of the SV40 early enhancer/promoter was used to normalize transfection efficiency. After 24 h of transfection, LX2 cells were treated with TGFβ1 (4 ng/mL) for 24 h. In the rottlerin-treated group, LX2 cells were treated with TGFβ1 (4 ng/mL) for 12 h and then co-treated with rottlerin (3 μM) for 12 h. The cells were collected and subjected to a dual-luciferase reporter assay (E1910, Promega). For each sample, firefly luciferase activity was normalized to that of Renilla luciferase.

Comparative microarray analysis was performed using the Affymetrix GeneChip^® Rat Genome 230 2.0 platform. Each treatment had three replicates, resulting in three independent microarrays for each treatment and controls (9 arrays in total). Genes showing differences in probe intensity of 1.5-fold or more were further analyzed. Differentially expressed genes were classified into their Gene Ontology (GO)¹ functional categories using R/Bioconductor (Bioconductor).² Cellular pathway association was analyzed according to the Kyoto Encyclopedia of Genes and Genomes (KEGG) database³ and pathway maps were analyzed according to BioCarta.⁴ Microarray data are available via the GEO accession number GSE139994.

Data are expressed as means ± SEM. Significance was evaluated using either paired or unpaired Student t-test or one-way AONVA analysis if the data showed normal distribution and homogeneity of variance which were evaluated by Shapiro-Wilk test and Levene’s test, respectively. Otherwise, Wilcoxon signed-rank test was used to analyze the paired data and Mann-Whitney U test and Kruskal-Wallis H test were used to analyze unpaired data for two and multiple groups, respectively. A p-value < 0.05 was considered significant.

To determine whether BK channels were expressed in activated HSCs, we examined the mRNA and protein expression of the BK channel pore-forming alpha subunit KCNMA1 in the LX2 cell line (Xu et al., 2005), a partially activated human HSC line that can be fully activated by TGFβ1 (Tsuchida and Friedman, 2017). We found that KCNMA1 was expressed at both the mRNA and protein level in TGFβ1-activated LX2 cells (Figures 1A,B) that showed robust expression of ACTA2, a marker for activated HSCs (Tsuchida and Friedman, 2017; Shang et al., 2018). To investigate whether KCNMA1 is also expressed in activated native HSCs, rat primary HSCs were isolated and spontaneously activated in serum-containing medium following 8–10 days of continuous culture on uncoated plastic. These spontaneously activated HSCs can faithfully recapitulate the major in vivo characteristic phenotypes of activated HSCs (Friedman et al., 1989). We found that KCNMA1 was expressed in primary activated HSCs at the level of both mRNA and protein (Figures 1A,B), while immunostaining indicated that KCNMA1 was distributed throughout the cell membrane of activated HSCs (Figure 1C).

To investigate whether BK channels in activated HSCs were functional, we performed whole-cell patch-clamp recordings combined with pharmacological evaluation. We found that whole-cell K⁺ currents from activated HSCs showed apparent voltage-dependent outward rectification properties. Extracellular application of rottlerin, a BK channel-specific activator, markedly increased the outward K⁺ current (Figures 1D,E), whereas treatment with paxilline, a BK channel-specific blocker, elicited the opposite effect, demonstrating that BK channels were functional in activated HSCs.

BK channels are associated with cell migration in multiple cell types (Toro et al., 2014). To explore the potential roles of BK channels in migration, a prominent feature of activated HSCs (Tsuchida and Friedman, 2017), we overexpressed the BK channel pore-forming alpha subunit, KCNMA1, by transfecting KCNMA1-expressing vectors into LX2 cells. The results showed that overexpression of KCNMA1 (Figure 2A) decreased LX2 cell migration (Figure 2B). Furthermore, ACTA2, collagen type I alpha 1 chain (COL1A1), COL1A2, and C-C motif chemokine ligand 2 (CCL2) mRNA expression levels were also decreased (Figure 2C). In contrast, downregulating KCNMA1 expression (Figure 2D) through siRNA transfection increased the migration rates of LX2 cells (Figure 2E), as well as the expression of ACTA2, COL1A1, COL1A2, and CCL2 (Figure 2F), indicating that enhancement of BK channel function could reverse HSC activation. However, neither overexpression nor knockdown of KCNMA1 affected HSC proliferation, as evidenced by the results of EdU staining (Figure 2G).

We then examined the effects of rottlerin treatment on phenotypic changes of activated HSCs. The results showed that rottlerin decreased the migration of both human LX2 cells (Figure 3A) and spontaneously activated rat HSCs (Figure 3B) at micromolar concentrations. In addition, rottlerin treatment also suppressed TGFβ1-induced expression of ACTA2, COL1A1, COL1A2, and CCL2 (Figure 3C), the expression of which is greatly increased in HSCs of liver fibrosis patients. Accordingly, rottlerin-induced BK channel activation did not affect the proliferation of LX2 cells (Figure 3D).

The results of the in vitro experiments suggested that augmentation of BK channel activity likely facilitates the reversion of activated HSCs to a quiescent state, which may have a protective effect against liver fibrosis. To assess whether increasing BK channel function elicits antifibrotic effects in vivo, we tested the effect of rottlerin administration on CCl₄-induced liver fibrosis in rats, in which myofibroblasts are mainly derived from activated HSCs (Iwaisako et al., 2014). As expected, the levels of ALT and AST, two typical serum markers of liver injury (Figure 4A), the expression of ACTA2, and levels of collagen deposition all were increased in rats with CCl₄-induced liver fibrosis (Figures 4B–D). Interestingly, rottlerin treatment markedly suppressed these CCl₄-induced effects (Figures 4A–D), indicating that rottlerin can elicit a significant improvement in the pathology of fibrosis, as well as in liver function.

Several studies have shown that TGFβ1 plays pivotal roles in HSC activation and hepatic fibrosis (Koyama and Brenner, 2017; Tsuchida and Friedman, 2017). To illustrate the underlying molecular mechanisms of the antifibrotic effect of BK channel activation, we first examined whether the canonical SMAD3-dependent TGFβ1 signaling pathway was involved in these processes. We found that rottlerin treatment completely blocked the TGFβ1-stimulated increase in SMAD transcriptional activity in LX2 cells, as determined by a luciferase reporter assay (Figure 5A). Western blotting results further showed that rottlerin treatment partially reduced the enhanced levels of phosphorylated, but not total, SMAD3 in TGFβ1-treated HSCs (Figure 5B), indicating that rottlerin specifically affected the phosphorylated form of SMAD3. In CCl₄-treated rats, rottlerin administration also greatly suppressed the increased levels of total and phosphorylated SMAD3, as well as the phosphorylation of TGFβ receptor types I and II (Figure 5C), suggesting that rottlerin exerts its antifibrotic effects through the suppression of the canonical TGFβ1 signaling pathway in vivo. To address whether other mechanisms were also involved, we used mRNA microarray to identify differentially expressed genes in the liver tissue of rats administered vehicle, CCl₄, or Rottlerin + CCl₄ treatments. Pathway analysis indicated that multiple signaling pathways, including the JAK/STAT3 pathway, were involved in the rottlerin-associated antifibrotic effects in vivo (Figures 6A,B). As crosstalk between SMAD3 and STAT3 in numerous biological functions is well documented, and because TGFβ1 can activate STAT3, which also has a role in liver fibrosis (Tang et al., 2017; Itoh et al., 2018; Xiang et al., 2018), we hypothesized that STAT3 may contribute to the antifibrotic effects of rottlerin. Western blot results showed that rottlerin treatment could reverse the increased levels of phosphorylated JAK2, as well as the levels of total and phosphorylated STAT3, in rats with CCl₄-induced fibrosis (Figure 6C). In addition, rottlerin treatment also attenuated the levels of phosphorylated JAK2 and STAT3 in primary activated HSCs stimulated by TGFβ1 (Figure 6D), as well as the mRNA expression of Il6, Cxcl2, and Ifng (Figure 6E), downstream target genes of STAT3. Combined, these results provide strong evidence that rottlerin treatment inhibits the TGFβ1/SMAD3 and JAK/STAT3 signaling pathways in vivo.