The major cellular event in the development and progression of liver fibrosis is the activation of hepatic stellate cells (HSCs). Activated HSCs proliferate and produce excess collagen, leading to accumulation of scar matrix and fibrotic liver. As such, the induction of activated HSC death has been proposed as a means to achieve resolution of liver fibrosis. Here we demonstrate that cannabidiol (CBD), a major non-psychoactive component of the plantCannabis sativa, induces apoptosis in activated HSCs through a cannabinoid receptor-independent mechanism. CBD elicits an endoplasmic reticulum (ER) stress response, characterized by changes in ER morphology and the initiation of RNA-dependent protein kinase-like ER kinase-, activating transcription factor-6-, and inositol-requiring ER-to-nucleus signal kinase-1 (IRE1)-mediated signaling cascades. Furthermore, CBD induces downstream activation of the pro-apoptotic IRE1/ASK1/c-Jun N-terminal kinase pathway, leading to HSC death. Importantly, we show that this mechanism of CBD-induced ER stress-mediated apoptosis is specific to activated HSCs, as it occurs in activated human and rat HSC lines, and in primary in vivo-activated mouse HSCs, but not in quiescent HSCs or primary hepatocytes from rat. Finally, we provide evidence that the elevated basal level of ER stress in activated HSCs has a role in their susceptibility to the pro-apoptotic effect of CBD. We propose that CBD, by selectively inducing death of activated HSCs, represents a potential therapeutic agent for the treatment of liver fibrosis.

CBD induces activated HSC apoptosis in a cannabinoid receptor-independent manner

To investigate the potential of CBD and other cannabinoid ligands to induce activated HSC death, we treated cultured activated HSCs, isolated from livers of rats that were fed an 8-month ethanol diet¹³ (see details in Materials and Methods), with increasing concentrations of different cannabinoids for 8h, and measured cell viability using an acid phosphatase assay. Incubation of cells with up to 10μM of either AM630 (CB₂R antagonist), JWH133 (CB₂R agonist), PF514273 (CB₁R antagonist), AM251 (CB₁R antagonist), or ACEA (CB₁R agonist) had little or no effect, whereas 5μM of either CBD or CP55940 (CB₁R/CB₂R agonist) was sufficient to cause ~60% and 90% loss of viability, respectively. In contrast to exogenous ligands, 20μM AEA (endogenous CB₁R/CB₂R agonist) was required to achieve a similar effect, whereas 2-arachydonoyl glycerol (2-AG, endogenous CB₁R/CB₂R agonist) treatment resulted in no significant change in the survival of these cells (Figure 1a).

Figure 1

CBD induces apoptosis in activated HSCs. (a) Cell viability of activated rat HSCs was determined using an acid phosphatase assay following serum starvation and 8-h treatment with indicated concentrations of cannabinoid ligands. Cell survival is presented …

In parallel, we investigated the ability of these cannabinoids to kill activated HSCs by examining the presence of cleaved poly-(ADP ribose)-polymerase (PARP), a marker for cell death, following 5μM treatment of cells. Among these compounds, only AEA and CBD were able to promote activated HSC death, as indicated by western blot analysis of PARP cleavage (Figure 1b), in agreement with the results obtained with the cell viability assay. Upon further examination, we found that CBD induced activated HSC death in a dose- and time-dependent manner, detectable after 6h with 5μM treatment (Figure 1c). We then confirmed the induction of apoptosis by CBD using flow cytometric analysis of FITC-Annexin V staining (Figure 1d).

In order to characterize the mechanism through which CBD causes apoptosis, we first investigated whether CBD was acting through cannabinoid receptors CB₁R or CB₂R. We found that blocking CB₁R or CB₂R had no effect on CBD-induced PARP cleavage (Figure 2a) and activated HSC viability (Figure 2b), in contrast to the death induced by CP55940, which appears to be mediated at least, in part, by CB₂R. Co-treatment with the CB₂R antagonist AM630 decreased CP55940-induced cell death by over 50% (data not shown). These results support a cannabinoid receptor-independent mechanism for CBD-induced apoptosis.

Figure 2

CBD induces ER dilation and expression of ER stress markers. (a) Activated rat HSCs were incubated for 6h in serum-free media with 5μM CBD in the presence or absence of either 1μM AM251 or 1μ …

CBD causes a change in morphology of the ER in activated HSCs through induction of ER stress

During the course of our experiments, we noticed marked changes in the morphology of CBD-treated activated HSCs. The presence of distinct structures surrounding the nucleus suggested an effect of CBD on the ER (Figure 2c). This was supported by an alteration in the distribution and localization of the ER chaperone calnexin, with the disappearance of uniform network-like ER structure and the formation of perinuclear vacuole-like structures (Figure 2d). This was further confirmed by electron microscopy showing that CBD treatment caused ER dilation (Figure 2e). These changes in ER morphology suggested that CBD induces ER stress.

We further investigated the effect of CBD on the ER in the activated HSCs by examining changes in the expression of calnexin, as well as the transcription factor C/EBP (CCAAT/enhancer-binding protein) homologous protein (CHOP), a major marker of prolonged ER stress. Western blot analysis showed that CBD treatment led to increased expression of both calnexin and CHOP (Figure 2f). Furthermore, upregulation of CHOP, an important potentiator of pro-apoptotic signaling following ER stress, provided evidence that ER stress may mediate CBD-induced apoptosis of activated HSCs.

CBD promotes apoptosis and ER stress in activated HSCs, but not in control HSCs or primary hepatocytes

In order to assess the specificity of CBD-induced apoptosis and ER stress, we examined the effect of CBD on the expression of PARP, calnexin, and CHOP in different cell lines, as well as in primary cells. We first compared HSCs from ethanol-treated rats with HSCs from control rats.¹³ HSCs from control rats exhibit a markedly lower activation state as compared with HSCs from ethanol-treated rats, indicated by 20-fold lower expression of alpha smooth muscle actin (αSMA; see Figure 3a, inset). Upon comparing the effect of treatment with 5μM CBD in these cells, we found that CBD did not alter PARP, calnexin, and CHOP expression in control HSCs, suggesting that the pro-apoptotic and ER stress effects of CBD were specific to the activated HSC phenotype (Figure 3a).

Figure 3

Pro-apoptotic effect of CBD is specific to activated HSCs. (a) HSCs from control or ethanol-fed rats, (b) primary mouse in vivo-activated HSCs, (c) LX-2 cells (human HSC line derived from cirrhotic patient), and (d) primary rat hepatocytes were incubated …

In primary in vivo-activated mouse HSCs and in LX-2 cells (immortalized human HSC line derived from cirrhotic patient¹⁴), CBD led to PARP cleavage, upregulation of calnexin, and induction of CHOP expression, consistent with what we observed in activated HSCs from rats (Figures 3b and c). In contrast, CBD treatment did not induce PARP cleavage or upregulation of calnexin and CHOP in primary rat hepatocytes (Figure 3d), confirming the specificity of the pro-apoptotic effect of CBD to activated HSCs.

To further examine the effect of CBD across different cell types, we compared the viability of HSCs from control rats, HSCs from ethanol-treated rats, and primary rat hepatocytes under 0–20μM range of CBD treatment. As shown before, 5μM CBD was sufficient to cause ~60% decrease in viability of HSCs from ethanol-treated rats, but not in HSCs from control rats, for which 10μM CBD was required to induce a significant loss that occurs to an even lesser extent (~40% reduction). In contrast to HSCs, primary rat hepatocytes maintained maximal viability under treatment with up to 20μM CBD (Figure 3e). Altogether, these findings suggest that CBD, in a dose-dependent manner, specifically targets activated HSCs to cause death.

CBD induces PERK-, ATF6-, and IRE1-mediated ER stress response in activated HSCs

We further investigated the mechanism for CBD-induced ER stress and apoptosis by characterizing the CBD-induced changes in the major mediators of ER stress signaling as described below. Cells respond to ER stress through a protective mechanism termed the unfolded protein response (UPR) in order to restore normal ER function. Upon stressful stimulus, this adaptive response is initiated by the repression release of three ER transmembrane proteins – PRKR (RNA-dependent protein kinase)-like ER kinase (PERK), ATF6, and inositol-requiring ER-to-nucleus signal kinase-1 (IRE1) – from the ER chaperone glucose-regulated protein 78kDa (GRP78). The subsequent activation of these ER stress sensors initiates a cascade of signaling events that result in the expression of genes that are critical for overcoming ER stress, including transcription factors and molecular chaperones^15, 16 (Figure 4a).

Figure 4

CBD induces ER stress response. (a) Schematic representation of the signaling pathways characterizing the ER stress response. In (b) through (e), activated rat HSCs were incubated in serum-free media with 5μM CBD, unless indicated otherwise, …

Release of PERK from GRP78 allows PERK activation through receptor autophosphorylation. Activated PERK can phosphorylate and inhibit eukaryotic translation initiation factor 2α (eIF2α), potentiating the eIF2α-independent translation and nuclear translocation of activating transcription factor (ATF)-4 to induce gene expression of chaperones and transcription factors such as CHOP. In activated HSCs, we found that CBD induced phosphorylation of PERK at 2h, followed by phosphorylation of eIF2α at 4h (Figure 4b). In line with this, we also detected a time-dependent accumulation of ATF4 in the nucleus (Figure 4c).

Upon release from GRP78, ATF6 is cleaved and translocated to the nucleus to induce CHOP and X-box-binding protein-1 (XBP1) expression. Western blot analysis showed that CBD induced accumulation of cleaved ATF6α in the nucleus (Figure 4c), indicating activation of the ATF6-mediated ER stress response.

Release of IRE1 from GRP78 during the UPR is followed by IRE1 autophosphorylation and phospho-IRE1-mediated excision of a 26-nucleotide intron from XBP1 mRNA, facilitating nuclear translocation of spliced XBP1 (XBP1s) protein and induction of expression of genes such as P58^IPK.¹⁷ CBD treatment of activated HSCs led to an increase in phosphorylated IRE1, accompanied by slight induction of P58^IPK expression (Figure 4b). We also found that CBD induced splicing of XBP1 mRNA in a time- and dose-dependent manner (Figure 4d). Moreover, a sharp increase in nuclear XBP1s protein was detected from 2–8h of treatment (Figure 4c), supporting CBD-induced activation of the IRE1-mediated ER stress-signaling cascade.

Induction of the ER stress response by CBD was further confirmed by an increase in expression of the UPR genes ATF4, ATF6, CHOP, and XBP1. Although 2μM CBD treatment for 6h induced significant increase in transcript levels of ATF4, ATF6, and CHOP, 5μM CBD led to even greater upregulation of these genes at earlier time points (Figure 4e).

Taken together, these results demonstrate that CBD treatment leads to the induction of all three signaling arms of the ER stress response in activated HSCs, resulting in significant upregulation of UPR genes and transcription factors. Along with our observation that CBD leads to apoptosis in activated HSCs, this provided strong evidence for ER stress as the underlying mechanism mediating the CBD-induced apoptotic response.

CBD promotes ER stress-induced apoptosis through activation of the IRE1/ASK1/JNK pathway in activated HSCs

In order to link the induction of ER stress to CBD-mediated cell death, we examined signaling events downstream of the UPR that trigger apoptosis. IRE1 has been shown to be pro-apoptotic by activating apoptosis signal-regulating kinase 1 (ASK1) through dephosphorylation of its Ser967 residue. Activated ASK1 leads to phosphorylation of c-Jun N-terminal kinase (JNK), which then promotes a series of downstream pro-apoptotic events.^18, 19, 20 As shown before (Figure 4b), we found that CBD leads to IRE1 phosphorylation. In addition, CBD induced dephosphorylation (activation) of ASK1, followed by an increase in phosphorylation (activation) of JNK. This cascade was activated in LX-2 cells, in cultured activated rat HSCs, and in primary in vivo-activated mouse HSCs (Figures 5a–c). Furthermore, overexpression of dominant-negative (DN) IRE1α in activated mouse HSCs (Figure 5d) prevented JNK phosphorylation and protected cells from apoptosis induced by CBD (Figure 5e). Expression of the DN IRE1α also led to a marked decrease in XBP1s mRNA in DN IRE1α-expressing activated HSCs after CBD treatment (Figure 5f), confirming the decreased activity of IRE1α. In addition, we found that JNK inhibitor protected activated HSCs from the apoptotic effect of CBD, decreasing PARP cleavage by over 50% (Figure 5g), and restoring cell viability to near maximal (Figure 5h). These findings indicate that CBD triggers apoptosis through ER stress-mediated activation of the IRE1/ASK1/JNK pathway.

Figure 5

CBD-induced cell death is mediated by the IRE1/ASK1/JNK pathway. Expression levels of phospho-ASK1, phospho-JNK, and JNK were determined by western blot analysis in (a) LX-2 cells, (b) HSCs from ethanol-fed rats, and (c) primary mouse in vivo-activated …

Blocking ER stress protects against CBD-induced apoptosis in activated HSCs

In order to investigate further the role of ER stress in CBD-induced apoptosis, we downregulated ATF4, a transcription factor induced upon activation of PERK in the UPR, in activated mouse HSCs. This resulted in a significant decrease in CHOP induction and cell death caused by CBD (Figure 6a), suggesting the importance of the PERK-mediated arm of the ER stress response in the pro-apoptotic effect of CBD. Using another approach to block the ER stress response, we treated LX-2 cells with CBD in the presence of imatinib, which has been shown to attenuate ER stress.²¹ Imatinib led to a significant decrease in CHOP induction, JNK phosphorylation, and PARP cleavage induced by CBD (Figure 6b), and protected against CBD-mediated activated HSC death (Figure 6c), providing strong evidence that CBD causes apoptosis by promoting ER stress.

Figure 6

Inhibition of ER stress response protects against CBD-induced cell death. (a) JS1 cells were transfected with siRNA against ATF4. After 16h, cells were incubated with 5μM CBD in serum-free media for 8h. Western blot …

Basal level of ER stress determines apoptotic effect of CBD

We then assessed the ER stress-mediated apoptotic effect of CBD by examining its action in the presence or absence of the ER stress inducer tunicamycin. We found that CBD and tunicamycin worked in synergy to increase LX-2 cell death in a dose-dependent manner (Figure 7a), supporting our finding that the apoptotic effect of CBD is mediated through an ER stress mechanism.

Figure 7

Basal ER stress determines sensitivity to CBD. (a) LX-2 cells were incubated in serum-free media with 0–4μM CBD for 6h in the presence or absence of tunicamycin (1μg/mL) to induce ER stress. Cell death …

In order to further elucidate the role of CBD-induced ER stress as a mediator for apoptosis, we investigated the basis for specificity of this occurrence in activated HSCs. We first examined basal ER stress levels in activated versuscontrol HSCs and found that expression levels of the ER stress response genes ATF4, ATF6, and CHOP were increased in activated HSCs from ethanol-treated rats as compared to control HSCs from untreated rats (Figure 7b). This led us to hypothesize that the elevated basal ER stress level of activated HSCs rendered these cells more susceptible to the ER stress-mediated apoptotic effect of CBD. To test this, we treated control HSCs with tunicamycin and examined CHOP expression and PARP cleavage in the presence or absence of CBD. Interestingly, we found that increasing the basal level of ER stress with tunicamycin in control HSCs led to an increase in CBD-induced CHOP expression and HSC death (Figure 7c). Taken together, this provided evidence that the increased basal level of ER stress in activated HSCs is an underlying factor, determining sensitivity to CBD and its induction of ER stress-mediated apoptosis.

Materials

The following primary antibodies were used in this study: rabbit monoclonal anti-phospho-JNK, rabbit polyclonal anti-JNK, anti-PARP, anti-P58IKP, anti-eIF2α, anti-phospho-PERK, mouse monoclonal anti-CHOP (Cell Signaling Technology, Danvers, MA, USA), polyclonal anti-calnexin, monoclonal anti-β-actin (Sigma, St. Louis, MO, USA), rabbit polyclonal anti-ATF4, anti-ATF6, anti-XBP1, anti-pASK1, phospho-c-Jun, anti-CHOP (Santa Cruz Biotechnology, Santa Cruz, CA, USA), monoclonal anti-α-SMA (Abcam, Cambridge, MA, USA), and polyclonal anti-phospho-IRE1 (Novus Biological, Littleton, CO, USA). The following secondary antibodies were used in this study: IRDye 680-labeled anti-rabbit, and IRDye 800-labeled anti-mouse and anti-goat (Rockland Immunochemicals, Gilbertsville, PA, USA). The following compounds were used in this study: CBD, AM630, JWH133, PF514273, AM251, ACEA, AEA, 2-AG, CP55940, tunicamycin (Tocris Bioscience, Ellisville, MO, USA), JNK inhibitor 2 (Calbiochem, Gibbstown, NJ, USA), imatinib (Selleck Chemicals, Houston, TX, USA), and bis(p-nitrophenyl) phosphate (Sigma).

Cell culture

HSCs were maintained in DMEM/F12 50% v/v+10% FBS. LX-2 cells¹⁴ were a gift from Dr. Scott Friedman. HSCs from ethanol-treated and control rats^13, 34were a gift from Dr. Natalia Nieto. JS1 cells (mouse activated HSCs) and primary HSCs from mice that were chronically administered CCl₄ for 1 month were a gift from Drs. Scott Friedman, Virginia Hernandez-Gea and Zahra Ghiassi-Nejad. Primary rat hepatocytes were obtained from Invitrogen (Carlsbad, CA, USA) and maintained according to manufacturer’s protocol.

Cell viability assay

Cell viability was determined using an acid phosphatase assay as described.³⁵Briefly, cells were treated with vehicle (DMSO), CBD, or the indicated ligands and drugs for indicated times, then washed with PBS. 10mM bis(p-nitrophenyl) phosphate (substrate) in 165mM Na acetate buffer, pH 5.5, containing 0.1% Triton X-100, was added to cells and incubated at room temperature for 16–24h. Reaction was terminated by adding 1N NaOH, and the absorbance at 410nM was measured using a spectrophotometer to determine acid phosphatase activity.

Western blot

Western blot analysis was carried out as described.³⁶ Briefly, cells starved for 16h were treated with CBD or with the indicated compounds. Cells were lysed in RIPA buffer and lysates were subjected to western blot analysis and probed for the indicated proteins using specific antibodies, according to manufacturer’s protocols. Both blotting and imaging with the Odyssey imaging system (LI-COR, Lincoln, NE, USA) were performed following the manufacturer’s protocols.

Flow cytometry

Cells were treated with vehicle or CBD for indicated times, and stained with FITC-AnnexinV and propidium iodide (BD Biosciences, San Jose, CA, USA), according to manufacturer’s protocol. Flow cytometric analyses were performed on a three-laser LSRII with DiVa software (BD Biosciences). Data were analyzed with FlowJo software (Tree Star Inc., Ashland, OR, USA).

Electron microscopy

Cells were fixed with 3% glutaraldehyde, pH 7.4. The specimens were then treated with 1% osmium tetroxide in 0.2M sodium cacodylate for 1h, followed by ethanol dehydration in graded steps through propylene oxide, and then embedded in Embed 812 (Electron Microscopy Sciences, Hatfield, PA, USA). Images were acquired on a Hitachi H7000 transmission electron microscope (Hitachi, Tokyo, Japan).

Immunofluorescence and confocal microscopy

This was carried out as described.³⁶ Slides were visualized with a Leica TCS SP5 confocal microscope (Leica Microsystems, Heidelberg, Germany). Images were acquired with an x63/1.32 PL APO objective lens (Leica Microsystems) and analyzed in sequential scanning mode.

Reverse transcription-polymerase chain reaction (RT-PCR)

Cells were treated with vehicle (DMSO) or CBD for indicated times, then washed with PBS. Total RNA was isolated using the RNeasy Mini Kit (QIAGEN, Valencia, CA, USA) and reverse transcribed using the High Capacity RNA-to-cDNA Kit (Applied Biosystems, Carlsbad, CA, USA). cDNA from activated HSCs was subjected either to standard PCR with Taq Polymerase (Invitrogen) to detect splicing, or to real-time PCR with SYBR Green (Applied Biosystems) to detect relative mRNA levels. For spliced Xbp1 detection by standard PCR, 500nM of each rat Xbp1-specific primers,³⁷ sense 5′-AAACAGAGTAGCAGCACAGACTGC-3′ and antisense 5′-TCCTTCTGGGTAGACCTCTGGGAG-3′ were used with the following program: (1) 94°C for 4min, (2) 35 cycles of 94°C for 45s, 63°C for 30s, and 72°C for 30s, and (3) 72°C for 10min. PCR products were separated by agarose gel electrophoresis to resolve the 473bp (unspliced) and 428bp (spliced) amplicons. Real-time PCR was performed using the following primers (500nM) for rat: ATF4 sense 5′-TATGGATGGGTTGGTCAGTG-3′ and antisense 5′-CTCATCTGGCATGGTTTCC-3′ ATF6 sense 5′-GATTTGATGCCTTGGGAGTC-3′ and antisense 5′-GGACCGAGGAGAAGAGACAG-3′ CHOP sense 5′-CCACACCTGAAAGCAGAAAC-3′ and antisense 5′-CACTGTCTCAAAGGCGAAAG-3′ XBP1 sense 5′-GATGAATGCCCTGGTTACTG-3′ and antisense 5′-AGATGTTCTGGGGAGGTGAC-3′ glyceraldehyde 3-phosphate dehydrogenase (GAPDH) sense 5′-TCAAGAAGGTGGTGAAGCAG-3′ and antisense 5′-AGGTGGAAGAATGGGAGTTG-3′ RNA polymerase II sense 5′-GAAGATCGGGAAGTGCTCAG-3′ and antisense 5′-ATCCGCAGAAGGTTACAAGG-3′ hypoxanthine-guanine phosphoribosyltransferase sense 5′-CCCAAAATGGTTAAGGTTGC-3′ and antisense 5′-TCAAGGGCATATCCAACAAC-3′. Quantitative analysis was performed using the ΔΔC_T method and REST software (http://www.gene-quantification.de).³⁸

DN IRE1α transfection

JS1 cells (mouse activated HSCs) were transfected with Addgene plasmid 20745 (Addgene, Cambridge, MA, USA) coding DN IRE1α (K599A, dominant negative kinase mutant), generated by Dr. Fumihiko Urano,³⁹ using Lipofectamine (Invitrogen). Stable lines were established by puromycin selection and clonal growth of resistant cells. Similar results were observed with three different clones. DN IRE1α expression was verified by RT-PCR using the human/mouse IRE1-specific primers, sense 5′-CTCTATGCCTCTCCCTCAATG-3′ and antisense 5′-AAGTCCTTCTGCTCCACATACTC-3′ (spanning the mutation site), in JS1 wild-type cells, JS1 cells expressing DN IRE1α, and LX-2 cells, according to the RT-PCR methods as described above. PCR products were analyzed and purified by agarose gel electrophoresis, then sequenced using the IRE1 antisense primer. GAPDH-specific primers for either human (sense 5′-GAAGGTGAAGGTCGGAGTC-3′, antisense 5′-GAAGATGGTGATGGGATTTC-3′) or mouse (sense 5′-TGAAGGTCGGTGTGAACG-3′, antisense 5′-CAATCTCCACTTTGCCACTG-3′) were used in parallel as internal controls.

siATF4 transfection

JS1 cells in six-well plates were transfected with Stealth RNAi siRNA to ATF4 (Invitrogen) for 6h using Lipofectamine (Invitrogen). Following transfection, cells were serum-starved for 16h, and then treated with CBD for the indicated times.

Statistical analyses

Statistical significance and P-values were determined by the Student’s t-test when comparing two groups, or by one-way ANOVA with Bonferroni post test when comparing multiple groups, using the GraphPad Prism software (GraphPad Software, San Diego, CA, USA). P-value (^*P<0.05; ^**P<0.01;^***P<0.001) of <0.05 was considered statistically significant.

We thank Dr. Scott Friedman for constructive discussions, comments, and the gift of the LX-2 cells, Drs. Virginia Hernandez-Gea and Zahra Ghiassi-Nejad for the primary mouse HSCs and JS1 cells, Dr. Natalia Nieto for the rat HSCs, and Dr. Serine Avagyan for assistance with flow cytometry. This work was supported by NIH grants DA08863, DA019521 to LAD, AA017067 to RR, and R24 CA095823 to MSSM-Microscopy Shared Resource Facility.

2-AG

2-arachydonoyl glycerol

AEA

anandamide

ASK1

apoptosis signal-regulating kinase 1

αSMA

alpha smooth muscle actin

ATF4

activating transcription factor 4

ATF6

activating transcription factor 6

CB₁R

type I cannabinoid receptor

CB₂R

type II cannabinoid receptor

CBD

cannabidiol

DN

dominant negative

ECM

extracellular matrix

eIF2α

eukaryotic translation initiation factor 2α

ER

endoplasmic reticulum

CHOP

C/EBP (CCAAT/enhancer-binding protein) homologous protein

GAPDH

glyceraldehyde 3-phosphate dehydrogenase

GRP78

glucose-regulated protein 78kDa

HSC

hepatic stellate cell

IRE1α

inositol-requiring ER-to-nucleus signal kinase-1α

JNK

c-Jun N-terminal kinase

PARP

poly-(ADP ribose) polymerase

PERK

PRKR (RNA-dependent protein kinase)-like ER kinase

UPR

unfolded protein response

XBP-1

X-box-binding protein-1

Edited by M Piacentini

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