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. 2023 Mar 10;46(3):149–159. doi: 10.1007/s12272-023-01438-1

Anti-fibrotic effect of aurocyanide, the active metabolite of auranofin

Hyun Young Kim 1, Undarmaa Otgontenger 2, Jun-Woo Kim 3, Young Joo Lee 1, Sang-Bum Kim 3, Sung Chul Lim 4, Young-Mi Kim 2,✉, Keon Wook Kang 1,✉
PMCID: PMC9998255  PMID: 36894745

Abstract

Drug repositioning has gained significant attention over the past several years. The anti-rheumatoid arthritis drug auranofin has been investigated for the treatment of other diseases, including liver fibrosis. Because auranofin is rapidly metabolized, it is necessary to identify the active metabolites of auranofin that have detectable levels in the blood and reflect its therapeutic effects. In the present study, we investigated whether aurocyanide as an active metabolite of auranofin, can be used to evaluate the anti-fibrotic effects of auranofin. Incubation of auranofin with liver microsomes showed that auranofin was susceptible to hepatic metabolism. Previously, we found that the anti-fibrotic effects of auranofin are mediated via system xc–-dependent inhibition of the NOD-, LRR-, and pyrin domain-containing protein 3 (NLRP3) inflammasome. Therefore, we tried to identify active metabolites of auranofin based on their inhibitory effects on system xc– and NLRP3 inflammasome in bone marrow-derived macrophages. Among the seven candidate metabolites, 1-thio-β-D-glycopyrano-sato-S-(triethyl-phosphine)-gold(I) and aurocyanide potently inhibited system xc– and NLRP3 inflammasome. A pharmacokinetics study on mice detected significant plasma levels of aurocyanide after auranofin administration. Oral administration of aurocyanide significantly prevented thioacetamide-induced liver fibrosis in mice. Moreover, the in vitro anti-fibrotic effects of aurocyanide were assessed in LX-2 cells, where aurocyanide significantly decreased the migratory ability of the cells. In conclusion, aurocyanide is metabolically stable and detectable in plasma, and has inhibitory effects on liver fibrosis, suggesting that it is a potential marker of the therapeutic effects of auranofin.

Supplementary Information

The online version contains supplementary material available at 10.1007/s12272-023-01438-1.

Keywords: Active metabolite, Auranofin, Aurocyanide, Liver fibrosis

Introduction

Auranofin is a gold(I) complex that contains thioglucose tetraacetate and triethylphosphine (Walz et al. 1983). It was initially approved for the treatment of rheumatoid arthritis in 1985 (Bombardier et al. 1986). Although it is not commonly used because it is less effective than other disease‐modifying anti-rheumatic drugs, such as methotrexate (Kean et al. 1997), recent studies have evaluated its use for the treatment of diseases other than rheumatoid arthritis (Roder and Thomson 2015), such as cancer, bacterial and parasitic infections, SARS-CoV-2, and liver fibrosis (Capparelli et al. 2017; Stratton et al. 2020; Abdalbari and Telleria 2021; Kim et al. 2021; Sonzogni-Desautels and Ndao 2021; Liu et al. 2022; Lu et al. 2022).

The main advantages of auranofin are its safety and oral availability (Glennas et al. 1997). In particular, it is safer than other injectable gold compounds, gold sodium thiomalate and aurothioglucose (Blocka 1983). Auranofin undergoes rapid ligand exchange and maintains a very low concentration in blood. Due to its rapid metabolism, the pharmacokinetic parameters of auranofin are determined on the basis of gold (Au) levels in blood (Messori and Marcon 2004; Sonzogni-Desautels and Ndao 2021). Therefore, it is necessary to identify the active metabolites of auranofin that have detectable levels in the blood and reflect its therapeutic effects. In the present study, we evaluated whether aurocyanide, an active metabolite of auranofin, can be used to evaluate the therapeutic effects of auranofin.

Although the precise mechanism underlying the effects of auranofin is unclear, most previous studies have found that it involves cellular oxidative stress. A recent omics analysis of auranofin-resistant cancer cells showed that changes in antioxidant gene expression indicate a response to auranofin (Falchetti et al. 2023). The mechanism underlying oxidative stress associated with auranofin probably involves inhibition of thioredoxin reductase (TrxR) (Zhang et al. 2019). In ovarian cancer cells, auranofin targets both the cytosolic (TrxR1) and mitochondrial (TrxR2) subtypes of TrxR (Gandin et al. 2010). The cysteine/selenocysteine redox active sequence of TrxR is involved in the interactions with its electrophilic chemical inhibitors. However, auranofin mainly binds to physiological thiols to form stable thiol gold(I) adducts before binding to intracellular TrxR (Sadler and Guo 1998). Therefore, sequential thiol-exchange mechanisms and membrane-located targets may explain the pharmacological actions of auranofin (Snyder et al. 1986; Kim et al. 2021).

Liver fibrosis progresses due to multiple events, involving macrophages, hepatic stellate cells (HSCs), and hepatocytes (Matsuda and Seki 2020; Kisseleva and Brenner 2021). Previously, we found that auranofin inhibits system xc–, a cysteine/glutamate antiporter, and induces oxidative stress in hepatic nonparenchymal cells (Kim et al. 2021). The oxidative bursts in hepatic macrophages inhibit the NOD-, LRR-, and pyrin domain-containing protein 3 (NLRP3) inflammasome and block interleukin (IL)-1β secretion (Hwangbo et al. 2020; Kim et al. 2021). NLRP3 inflammasome regulates immune responses to fibrosis progression (Davis et al. 2011) and auranofin inhibits liver fibrosis.

In the present study, we evaluated the inhibitory effects of auranofin metabolites on system xc– and NLRP3 inflammasome activity in bone marrow-derived macrophages (BMDMs) and identified aurocyanide as a potent active metabolite of auranofin. Furthermore, we evaluated the pharmacokinetic profile of aurocyanide in mice following a single oral administration of auranofin or aurocyanide. Finally, we demonstrated the anti-fibrotic effect of aurocyanide using a thioacetamide (TAA)-induced mouse model of liver fibrosis.

Materials and methods

Materials

Aurocyanide (Gold(I) potassium cyanide) was purchased from Alfa Aesar (Ward Hill, MA, USA). l-thio-3-D-glucopyranosato-(triethylphosphine) gold(I) (M1) was synthesized from chloro(triethylphosphine) gold(I) and the thiosugar. Other proposed metabolites of auranofin (M2-M6) were obtained from Sigma (St. Louis, MO, USA). Recombinant human transforming growth factor-β1 (TGF-β1) was supplied by PeproTech (Cranbury, NJ, USA). TAA, lipopolysaccharide (LPS), and adenosine triphosphate (ATP) were purchased from Sigma (St. Louis, MO, USA).

Animal experiments

Male BALB/c mice were purchased from SLC Inc. (Kotoh-cho, Japan). BALB/c mice were intraperitoneally injected with 100 mg/kg of TAA for 8 weeks, twice a week to induce hepatic fibrosis. The mice were acclimatized for one week prior to experiment and were housed in groups of 3–5 at a temperature of 21 ± 2 °C and 50 ± 5% humidity with a 12 h light/dark cycle.

Seven or eight-week-old ICR mice were purchased from OrientBio Inc. (Siheung, Korea) for the pharmacokinetic study. Mice had free access to food and water. On the experiment day, the mice were fasted for 16 h before oral administration of auranofin or aurocyanide. The animal handlings and experimental procedures were approved by IACUC (Institutional Animal Care and Use Committee, SNU-190924-6 and DGMIF-19071401-00).

Assessment of liver injury in blood and fibrotic liver

Serum aspartate transaminase (AST) and alanine transferase (ALT) levels were determined using Spectrum® (Abbott Laboratories, Abbott Park, IL, USA). Liver tissue samples were fixed using 10% neutral-buffered formalin solution (Sigma, St. Louis, MO, USA), embedded in paraffin blocks, and stained for histological assessment. Masson’s trichrome stained liver sections were assessed by a liver pathologist in a blind manner. Fibrosis extent was graded as 0, absent; 1, enlarged, fibrotic portal areas; 2, periportal or portal-portal septa but intact architecture; 3, fibrosis with architectural distortion but no obvious cirrhosis; and 4, probable or definite cirrhosis.

Isolation and culture of BMDMs

Bone marrow cells were isolated by flushing the femur and tibia of 8-week-old male C57BL/6 mice. The isolated bone marrow cells were incubated with macrophage colony-stimulating factor (M-CSF; Peprotech, Cranbury, NJ, USA), and were differentiated (7 days) into BMDMs. Bone marrow cells were incubated in RPMI medium containing 30 ng/ml M-CSF, 10% fetal bovine serum (FBS), 1% penicillin/streptomycin, and 25 mM HEPES. The media were changed every 3 days.

Quantification of aurocyanide in plasma and pharmacokinetic analysis

Blood samples were taken at 0.25, 0.5, 1, 2, 4, 6, and 8 h after oral administration of auranofin or aurocyanide (Ott Joslin 2009). 20 μl of plasma samples were added to 180 μl of acetonitrile containing internal standard, vortexed for 5 min at 15,000 rpm. The supernatants were subjected to LC–MS/MS (Triple Quad 5500, Applied Biosystems, Foster City, CA, USA). Phoenix WinNonlin 6.4 version program (Pharsight Corporation, Mountain View, CA, USA) and non-compartmental analysis model were used for the calculation of pharmacokinetic parameters.

In vitro microsomal stability test

Auranofin (1 μM) was incubated with liver microsomes from human or mouse (0.5 mg/ml) in 0.1 M phosphate buffer. The amount of auranofin remaining after 30 min incubation at 37 °C was determined using LC–MS/MS (Triple Quad 5500, Applied Biosystems, Foster City, CA, USA).

Glutamate assay

BMDMs were incubated in phenol red free-medium. The glutamate level was measured after 3 h incubation with auranofin metabolites, using the Amplex® Red glutamate assay kit (Thermo Fisher Scientific, Waltham, MA, USA). According to the manufacturer’s instruction, the glutamate concentrations of the medium were measured by a fluorescence microplate reader (SpectraMAXi3x, Molecular Devices, Sunnyvale, CA, USA).

Cell culture

LX-2 cells, an immortalized human hepatic stellate cell line were kindly provided by Dr. Friedman (Mount Sinai School of Medicine, New York, NY, USA) (Yang et al. 2014). LX-2 cells were cultured in high-glucose Dulbecco's modified Eagle's medium (DMEM; HyClone, Logan, UT, USA) supplemented with 2% FBS (Gibco, Waltham, MA, USA), 100 U/ml penicillin, and 100 μg/ml streptomycin at 37 °C in a humidified atmosphere containing 5% CO2. LX-2 cells were seeded at a density of 5 × 105 cells/well in a 6-well plate for western blot. After the cells reach 70–80% confluence, they were serum-starved overnight and treated with the compounds.

Real-time monitoring of cell proliferation

LX-2 cell proliferation was monitored by phase-based quantification of cell confluences using an IncuCyte® S3 Live-Cell Analysis System (Sartorius, Ann Arbor, MI, USA). LX-2 cells were plated at 2 × 104 cells/well in a 48-well plate. After 24 h incubation, cells were treated with various concentrations of aurocyanide in the presence of 2% FBS and the phase confluences of the cells were measured every 3 h over 2 days. The relative cell confluence was calculated by setting the phase confluence at 0 h to be 1.

Transwell migration assay

The 24-well transwell unit with 8-μm pore-sized polycarbonate membrane (Costar, Corning Inc., Corning, NY, USA) was used. The undersurface of upper chamber was coated with 0.5 mg/ml of type I collagen (Sigma, St. Louis, MO, USA). LX-2 cells serum-starved overnight in serum-free DMEM were plated in the upper chamber (2 × 104 cells/well) and the lower chamber was filled with DMEM containing 2% FBS with or without aurocyanide. The cells were allowed to migrate for 15 h, fixed with methanol, and stained with hematoxylin and eosin. After removing the non-migrated cells in the upper chamber with a cotton swab, migratory ability was assessed by counting the number of cells moved to the undersurface of the membrane in 8 high-power fields captured with microscopy. The representative images were taken using a Nikon Eclipse Ti inverted microscope (Nikon, Tokyo, Japan).

Western blot analysis

Liver tissues were lysed using RIPA lysis buffer and centrifuged at 16,000g for 15 min. LX-2 cells were lysed in RIPA buffer (Cell Signaling Technology, Beverly, MA, USA) containing a cocktail of protease inhibitors (Calbiochem part of Merck KGaA, Darmstadt, Germany) on ice. The supernatants were then obtained after centrifugation at 16,200g for 15 min at 4℃. The protein concentrations of the cell lysates or liver tissues were measured by Bradford assay (Pro-Measure, iNtRON Biotechnology, Seoul, Korea) and the proteins were dissolved in the SDS sample buffer. Western blot analysis was conducted as previously described (Song et al. 2016). Samples were separated using sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE). The separated proteins were transferred to nitrocellulose membranes (GE Healthcare, Madison, WI, USA) and blocked with 5% skim milk. The proteins on the membrane were incubated with the primary antibodies (Supplementary Table 1) overnight. Horseradish peroxidase-conjugated IgG antibodies (Cell Signaling Technology, Beverly, MA, USA) were used as the secondary antibodies. After 2 h incubation with the secondary antibody, immune complexes were detected using the Immobilon Western Chemiluminescent HRP Substrate (Merck Millipore, Billerica, MA, USA).

RNA preparation and real-time quantitative polymerase chain reaction

Total RNA was extracted from liver tissue using Trizol® reagent (Thermo Fisher Scientific, Waltham, MA, USA). Oligo (dT) primer sequences used in this study are shown in Supplementary Table 2. Transcript levels were normalized to 18s ribosomal RNA levels. Expression levels of the mRNA obtained from liver tissue were quantified by real-time qPCR, using Bio-Rad CFX Manager™ Software (Bio-Rad, Hercules, CA, USA) with SYBR Select Master Mix (Applied Biosystems, Foster City, CA, USA).

Statistical analysis

Data are presented as mean ± S.D. Statistically significant differences were assessed by the unpaired Student’s t-test or one-way ANOVA followed by Tukey’s multiple comparison test. GraphPad PRISM 7.0 software (GraphPad Software, Inc., San Diego, CA, USA) was used for the statistical analysis. P-value < 0.05 was considered statistically significant.

Results

Identification of aurocyanide as an active metabolite of auranofin

Before evaluating the metabolites of auranofin, we determined the microsomal stability of auranofin. Liver microsomal assays indicate the hepatic clearance of drugs (Obach 1999). Auranofin was incubated with microsomes from the livers of humans or mice and the remaining quantity of auranofin was compared to that of verapamil (Fig. 1A). Verapamil was used as the reference compound because it undergoes extensive first-pass hepatic metabolism (Vogelgesang et al. 1984). Microsomal metabolism was faster for auranofin than for verapamil. Only 6.9% and 1.5% of auranofin remained after 30 min of incubation with human and mouse microsomes, respectively, suggesting that auranofin exhibits significant first-pass metabolism. Therefore, we hypothesized that the pharmacological effects of auranofin are mainly mediated via its metabolites. Based on previous studies (Tepperman et al. 1984; Graham et al. 2008; Madeira et al. 2012), we identified seven candidate metabolites generated by the hepatic metabolism of auranofin (Fig. 1B, C).

Fig. 1.

Fig. 1

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Proposed metabolites of auranofin. A 1 μM of auranofin or verapamil was incubated in 0.1 M phosphate buffer (pH 7.4) containing liver microsomes from human or mouse (0.5 mg/ml). The percentage of remaining auranofin or verapamil was determined using LC–MS/MS. Molecular structures of B auranofin and C its metabolites. 7 candidates were proposed as metabolites of auranofin. M, metabolites of auranofin; ND, not determined

In a previous study, we found that auranofin prevents liver fibrosis by inhibiting the NLRP3 inflammasome in macrophages and fibrotic liver tissues (Kim et al. 2021). The NLRP3 inflammasome mediates liver fibrosis progression (Kaufmann et al. 2022). Therefore, we investigated the effects of the candidate metabolites on the NLRP3 inflammasome based on IL-1β release in LPS-primed BMDMs. Because gold is essential for the effects of auranofin (Snyder et al. 1987), gold-containing metabolites (M1, M2, M4, and M7) were tested first. Treatment with 0.1 µM auranofin or its gold-containing metabolites inhibited ATP-induced IL-1β secretion in LPS-primed BMDMs. Among the metabolites tested, M1 (1-thio-β-D-glycopyrano-sato-S-[triethyl-phosphine]-gold[I]) and M7 (aurocyanide) exhibited the most potent anti-inflammasome effect (Fig. 2A).

Fig. 2.

Fig. 2

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Screening for identifying active metabolites of auranofin. A BMDMs were primed with LPS (100 ng/ml) for 4 h, followed by 1 h exposure to ATP (1 mM) plus 0.1 μM of auranofin or its metabolites. The effects of auranofin or its metabolites on NLRP3 inflammasome were determined by measuring IL-1β secreted to the supernatants (n = 3). Data are presented as mean ± S.D. ***P < 0.005, compared with the LPS-primed group. B BMDMs were treated with 1 μM of auranofin or proposed metabolites of auranofin (M1-M7) for 3 h in serum free media. Extracellular glutamate levels were determined using glutamate assay kit in the supernatant of BMDMs (n = 3). Data are presented as mean ± S.D. *P < 0.05, **P < 0.01, ***P < 0.005, compared with the control group. C NLRP3 inflammasome activities were compared by measuring IL-1β in the supernatants of LPS-primed BMDMs, which were subsequently exposed to 1 mM of ATP with or without auranofin or its metabolite aurocyanide (M7) for 1 h (n = 3). Data are presented as mean ± SD ***P < 0.005, compared with the LPS-primed group; ##P < 0.01, ###P < 0.005 compared with the LPS-primed group exposed to ATP. AC. aurocyanide; AF, auranofin; BMDM, bone marrow-derived macrophage; M, metabolite

Auranofin inhibits the NLRP3 inflammasome by blocking system xc– activity (Kim et al. 2021). System xc– is a cystine/glutamate antiporter that regulates oxidative stress by importing cystine, the major source of intracellular cysteine and glutathione (Kobayashi et al. 2018). Inhibition of system xc– induces a short-term increase in reactive oxygen species and blocks the NLRP3 inflammasome in macrophages during liver fibrogenesis (Kim et al. 2021). To evaluate the role of system xc– in metabolite-induced inhibition of NLRP3 inflammasome, we measured its activity in BMDMs after treatment with metabolites. We determined the glutamate level in the supernatant after incubation with 1 µM each metabolite. In comparison to auranofin, all of the metabolites except M3 showed more potent inhibitory effects on system xc− activity (Fig. 2B). Considering the inhibitory effects of auranofin metabolites on system xc− and inflammsome, we propose that M1 and M7 may be potential active metabolites of auranofin. Because the tetraethylthioglucose moiety in M1 is rapidly modified and aurocyanide (M7) is detected in the blood and urine of patients who have received auranofin (Tepperman et al. 1984; Elder et al. 1993), we further evaluated aurocyanide as an active metabolite of auranofin. Similar to auranofin, aurocyanide significantly inhibited NLRP3 inflammasome activity in a concentration-dependent manner (Fig. 2C).

Pharmacokinetics of aurocyanide, an active metabolite of auranofin

The plasma concentration–time curve of aurocyanide following a single dose of auranofin in mice is shown in Fig. 3A. Auranofin was administered orally at a dose of 10 mg/kg, which showed anti-fibrotic effects on liver fibrosis in mice (Kim et al. 2021). The area under the plasma concentration–time curve from time 0 to the last measured concentration in plasma (AUClast) was 513.5 ng⋅h/mL, and the maximum plasma concentration (Cmax) was 119.3 ng/mL. Because aurocyanide inhibited the NLRP3 inflammasome from 30 nM (Fig. 2C), the plasma level of aurocyanide was physiologically reliable up to 8 h after auranofin administration.

Fig. 3.

Fig. 3

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Plasma concentration–time profiles of aurocyanide, an active metabolite of auranofin, in mice. Mean plasma concentration–time profiles of aurocyanide A after a single oral administration of auranofin at a dose of 10 mg/kg (n = 4) and B following an oral dose of 1, 3, or 10 mg/kg aurocyanide in mice (n = 4)

The plasma concentration–time curves and pharmacokinetic parameters of aurocyanide after a single oral dose of aurocyanide (1, 3, or 10 mg/kg) in mice are presented in Fig. 3B and Table 1, respectively. The Cmax and AUClast of aurocyanide after administration of 10 mg/kg auranofin were similar to those after administration of 3 mg/kg aurocyanide (Table 1), suggesting that > 3 mg/kg aurocyanide prevents liver fibrosis.

Table 1.

Pharmacokinetic parameters of aurocyanide after a single dose of aurocyanide in mice

Parameter AC 1 mg/kg AC 3 mg/kg AC 10 mg/kg
T1/2 (h) 7.4 ± 3.7 3.6 ± 0.6 2.6 ± 1.3
Cmax (ng/ml) 77.3 ± 19.0 199.8 ± 75.3 638.5 ± 102.9
Tmax (h) 0.4 ± 0.1 0.6 ± 0.4 1.3 ± 0.9
AUClast (ng⋅h/ml) 220.6 ± 22.4 583.6 ± 103.0 2324.4 ± 310.7
AUCINF_obs (ng⋅h/ml) 412.7 ± 83.6 750.7 ± 117.7 2857.8 ± 504.4
%AUCExtrap (%) 44.6 ± 14.1 22.4 ± 4.6 17.2 ± 15.3
MRTINF_obs (h) 10.7 ± 4.9 5.1 ± 0.8 4.6 ± 1.6
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Data are presented as mean ± SD (n = 4). AC, aurocyanide; T1/2, terminal half-life; Cmax, maximum plasma concentration; Tmax, time to reach Cmax; AUClast, the total area under the plasma concentration–time curve from time 0 to the last measured concentration in plasma; AUCINF_obs, AUC from time of dosing extrapolated to infinity, based on the last observed concentration; %AUCExtrap, percentage of the area extrapolated for calculation of area under the plasma concentration–time curve from time 0 to infinity; MRTINF_obs, mean residence time extrapolated to infinity

Inhibition of liver fibrosis by aurocyanide

To evaluate the anti-fibrotic effects of aurocyanide in vivo, we used a TAA-induced liver fibrosis model. BALB/c mice were injected with TAA (100 mg/kg) twice per week and with 10 mg/kg auranofin or aurocyanide six times a week for 8 weeks. Masson’s trichrome staining revealed significantly decreased fibrotic score and area in mice treated with aurocyanide (Fig. 4A–C). Aurocyanide administration significantly attenuated the TAA-induced increase in ALT and AST levels (Fig. 4D). Furthermore, the mRNA expression levels of profibrotic markers, including Timp1, Col1a1, Acta2, and Ctgf in the liver tissues were significantly attenuated by aurocyanide administration (Fig. 4E). We also evaluated the expression of α-smooth muscle actin (αSMA) protein (Fig. 4F). The protein expression levels of the inflammasome markers, mature IL-1β and caspase-1 (p20) were mildly decreased in the aurocyanide-treated group compared to the TAA-treated group (Fig. S1A).

Fig. 4.

Fig. 4

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Inhibitory effects of aurocyanide on thioacetamide-induced liver fibrosis in mice. A Liver sections were subjected to Masson’s Trichrome staining to evaluate collagen deposition in the liver and representative images are shown. B, C Fibrotic scores and fibrotic area were assessed by a liver pathologist in a blind manner. D Effects of auranofin and its active metabolite aurocyanide on the liver injury markers, serum ALT and AST levels, were determined. E Profibrotic markers including Timp1, Col1a1, Acta2, and Ctgf were measured in liver tissues using real-time PCR analysis. mRNA expressions were presented as relative ratio to the CTRL whose value was taken as 1.0. F Protein expression levels of αSMA in liver tissues were determined using western blot analysis (left). Bands were quantified by densitometry (right). Data B–F are presented as mean ± SD. n = 6–9 per group. ***P < 0.005 compared with the vehicle-treated control group; #P < 0.05, ##P < 0.01, ###P < 0.005 compared with the group treated with thioacetamide alone; +++P < 0.005 compared with the group treated with thioacetamide and auranofin. AC, aurocyanide; AF, auranofin; CTRL, control; NS, normal saline; TAA, thioacetamide; ns, not significant

Although aurocyanide includes cyanide, no adverse events related to cyanide administration were observed (Wright and Vesey 1986). Furthermore, none of the mice died or had significant weight loss after aurocyanide administration (Fig. S1B). The serum levels of AST, ALT, and creatinine, and urea nitrogen in blood were normal in mice treated with aurocyanide, indicating no significant hepatic or renal toxicity (Fig. 4D and Fig. S1C). These results suggest that aurocyanide is an active metabolite of auranofin that prevents liver fibrosis.

Inhibition of cell proliferation and migration of hepatic stellate cells (HSCs) by aurocyanide

During liver fibrogenesis, activated HSCs migrate to the perisinusoidal space, expand, and produce extracellular matrix (Kisseleva and Brenner 2021). Considering the crucial role of HSCs in the pathogenesis of liver fibrosis, we investigated the effects of aurocyanide on HSCs to elucidate its anti-fibrotic effects at the cellular level. First, we assessed its effect on the proliferation of LX-2 cells, a human hepatic stellate cell line, using a live-cell monitoring system. Aurocyanide significantly suppressed LX-2 cell proliferation at concentrations higher than 0.5 µM (Fig. 5A). Next, we evaluated its effect on the TGF-β1 signaling, a key inducer of HSC activation. Aurocyanide did not affect canonical TGF-β1 signaling which promotes Smad2/3 phosphorylation (Fig. 5B). However, it significantly inhibited the migratory ability of HSCs, which is also involved in the attenuation of liver fibrosis by auranofin (Kim et al. 2021). The number of migrated LX-2 cells decreased by 23% and 48% at 0.1 and 0.3 µM aurocyanide, respectively (Fig. 5C).

Fig. 5.

Fig. 5

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In vitro effects of aurocyanide in LX-2 human hepatic stellate cells. A LX-2 cells were incubated with increasing concentrations of aurocyanide (0.01–1 μM) in the presence of serum and cell proliferation was monitored using a live-cell imaging system over 2 days. The phase confluence at 0 h was set to 1 (n = 4). B Immunoblotting for the phosphorylated Smad2/3 was performed in the lysates of LX-2 cells exposed to 10 ng/ml of TGF-β1 in the presence or absence of aurocyanide for 1 h (left). Bands were quantified by densitometry (right; n = 3). C The migratory ability of LX-2 cells co-incubated with 2% FBS and aurocyanide for 15 h was assessed by a transwell migration assay. Migrated cells were stained with hematoxylin and eosin. Representative images are shown (left) and number of migrated cells was counted (right; n = 3). Scale bar = 100 μm. Data are presented as mean ± S.D. *P < 0.05, **P < 0.01, ***P < 0.005 compared with the vehicle-treated control group; ##P < 0.01, ###P < 0.005 compared with the group treated with 2% FBS alone. AC, aurocyanide; FBS, fetal bovine serum; ns, not significant

Discussion

Although an increasing number of studies are being conducted on liver fibrosis, its first-line treatment is not clear (Manka et al. 2019). In a previous study, we proposed auranofin, a clinically approved anti-rheumatoid drug, as a treatment for liver fibrosis (Kim et al. 2021). Auranofin inhibited the pathogenesis of liver fibrosis, inflammatory signals of macrophages, and migration of HSCs (Kim et al. 2021). The clinical efficacy of auranofin in liver fibrosis is being evaluated in a phase II clinical trial. However, repositioning auranofin for the treatment of liver fibrosis is challenging because its rapid and extensive hepatic metabolism makes it difficult to evaluate its therapeutic effects on the basis of its plasma concentration (Madeira et al. 2012). Plasma concentrations of gold drugs are measured using atomic absorption techniques, which only detect gold and cannot detect its ligands. Similarly, most pharmacokinetic studies of auranofin have only evaluated the kinetics of gold and not intact auranofin (Messori and Marcon 2004). Based on the plasma concentration of gold after auranofin administration, 15–25% of auranofin is absorbed through the gastrointestinal tract, and most of the gold is bound to albumin (Roder and Thomson 2015). Moreover, the absorbed auranofin undergoes extensive first-pass metabolism (Fig. 1A), which makes it difficult to predict the clinical outcome. Therefore, in the present study, we attempted to identify a metabolically stable active metabolite of auranofin.

Based on previous studies (Graham et al. 2008; Madeira et al. 2012), we proposed seven metabolites of auranofin, of which many are produced by oxidation and hydrolysis (Kupiec et al. 2019). Based on the previously reported mechanism of auranofin, we assessed the effects of the candidate metabolites on system xc–-mediated glutamate efflux and LPS/ATP-stimulated NLRP3 inflammasome activation in BMDMs. Aurocyanide potently inhibited the activation of NLRP3 inflammasome. Similar to auranofin, aurocyanide reduced cell proliferation and migration of HSCs. Aurocyanide produced from anti-rheumatic gold complexes also has anti-arthritic and anti-inflammatory effects (Graham et al. 2008). However, to the best of our knowledge, the present study is the first to show the anti-fibrotic effect of aurocyanide (Fig. 6).

Fig. 6.

Fig. 6

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Involvement of aurocyanide formation in anti-liver fibrotic actions of auranofin

Aurocyanide is also a metabolite of other anti-rheumatic gold complexes (Graham and Kettle 1998). It is produced from aurothiomalate due to the action of myeloperoxidase. Under physiological conditions, gold is present in plasma mainly as complexes with albumin and endogenous thiols. Aurothiomalate or the albumin complexes are converted into aurocyanide following myeloperoxidase-mediated oxidation of thiocyanate (Graham and Pile 2016).

The plasma level of aurocyanide after administration of auranofin and aurocyanide in mice suggested a relatively high metabolic stability of aurocyanide. Auranofin undergoes ligand exchange with cyanides and forms dicyanoaurate complex, aurocyanide. Cyanide is a very strong ligand for gold(I) and exists in small quantities in the blood; it is produced endogenously or derived from cigarette smoke (Graham et al. 1984; Messori and Marcon 2004). Unlike auranofin, aurocyanide is resistant to metabolism, remarkably stable, and has an estimated formation constant of 1038. In our study, 10 mg/kg aurocyanide prevented TAA-induced liver fibrosis in mice, suggesting that the plasma level of aurocyanide predicts the anti-fibrotic effect of auranofin. Moreover, there were no significant adverse events following oral administration of aurocyanide for 8 weeks. In fact, barely no cyanide is liberated from aurocyanide under the physiological conditions, and no adverse events were noted (Wu et al. 2001).

There were some limitations to the present study. Aurocyanide may not be the only active metabolite of auranofin. Based on the pharmacokinetic parameters, the AUClast value of aurocyanide after administration of 10 mg/kg auranofin (513.5 ng⋅h/mL) was similar to that after administration of 3 mg/kg aurocyanide (583.6 ng⋅h/mL). However, the efficacy of 10 mg/kg auranofin was similar to that of 10 mg/kg aurocyanide. This suggests that other metabolites may also mediate the protective effect of auranofin on liver fibrosis. Although further studies are needed to identify additional active metabolites, aurocyanide is a potent active metabolite that inhibits liver fibrosis. Another important factor to consider is the possibility that aurocyanide's inhibitory effect on liver fibrosis may be attributed to effects beyond the inhibition of inflammasome. In our preliminary experiments, M7 demonstrated a protective effect against apoptosis induced by reactive oxygen species. Therefore, it is plausible that the antioxidant effect of M7 could contribute to its inhibition of liver fibrosis and should also be taken into consideration.

In our in vitro assay using BMDMs, metabolites containing gold (M1, M4, M6, and M7) significantly reduced IL-1β secretion, consistent with a previous study that found that gold is essential for the activity of auranofin (Snyder et al. 1987). In future studies, active gold compounds may be chemically modified to develop more potent anti-fibrotic agents.

Based on our results, aurocyanide is an active metabolite of auranofin. It is metabolically stable and detectable in plasma, and has anti-fibrotic effects. Although additional metabolites may also mediate the pharmacological effects of auranofin, aurocyanide is a potential marker of the therapeutic effects of auranofin.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOCX 231 KB) (229.9KB, docx)

Acknowledgements

This work was supported by the National Research Foundation of Korea (NRF) grants funded by the Korean Government [NRF-2022M3A9B6017654 and NRF-2020M3A9C8015798 (K.W. Kang) and NRF-2020R1A6A1A03042854 and NRF-2022R1A2C1092649 (Y.-M. Kim)]. Figure 6 was created using biorender.com.

Data availability statement

The data used to support the findings of this study are available from the corresponding author upon request.

Declarations

Conflict of interest

The authors declare no conflict of interest.

Footnotes

Publisher's Note

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Contributor Information

Young-Mi Kim, Email: ymikim12@hanyang.ac.kr.

Keon Wook Kang, Email: kwkang@snu.ac.kr.

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Associated Data

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Supplementary Materials

Supplementary file1 (DOCX 231 KB) (229.9KB, docx)

Data Availability Statement

The data used to support the findings of this study are available from the corresponding author upon request.

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