Protective effects of retinoic acid on cadmium toxicity in human proximal tubular epithelial cells

Abstract

Cadmium (Cd) is a toxic heavy metal that induces proximal tubular cell damage. Previously, we identified that retinoic acid receptor (RAR) activity was suppressed in the kidney of Cd-exposed mice. In addition, peroxisome proliferator-activated receptor δ, PPARδ, contributed to the modification of Cd toxicity in HK-2 human proximal tubular cells. In this study, we investigated the protective effects of retinoic acid (RA) and its precursor, retinol, against Cd-induced cytotoxicity in HK-2 cells. Pretreatment with RA or retinol significantly reduced Cd toxicity. Knockdown of RARA, RARG, or PPARD did not alter the protective effects of RA; moreover, the double knockdown of RARA and RARG partly suppressed the RA-reduced Cd toxicity. This suggested that RA may reduce Cd toxicity by a receptor-independent mechanism. Furthermore, RA did not affect the expression of metallothionein genes (MT-1X and MT-2A) or the intracellular accumulation of Cd after Cd treatment. RA pretreatment suppressed Cd-induced apoptosis, partly by inhibiting caspase-3 activation. These findings suggest that RA prevents Cd toxicity via a novel, receptor-independent mechanism involving the suppression of apoptosis.

INTRODUCTION

Cadmium (Cd) is a pervasive environmental contaminant that induces severe toxic effects in multiple tissues, with the kidney being particularly susceptible (Järup et al., 1998). Following its long-term oral ingestion, Cd predominantly accumulates in the kidney and exhibits an exceptionally long biological half-life, ranging from 15 to 30 years (Järup, 2002; Järup and Akesson, 2009). The excessive accumulation of Cd in the kidney leads to proximal tubular damage, a hallmark of Cd-induced nephrotoxicity (Järup et al., 1998). Although Cd is bound to the metal-binding protein metallothionein (MT), a primary detoxification factor, elevated levels of unbound Cd disrupt cellular functions and trigger toxic responses (Järup et al., 1998; Klaassen et al., 2009). At the molecular level, Cd exposure has been associated with a broad spectrum of cytotoxic events, including the induction of oxidative stress, inhibition of key enzymatic activities, and dysregulation of gene expression (Fujiwara et al., 2012; Lee et al., 2025). Despite extensive research, the precise molecular mechanisms underlying Cd-induced renal toxicity remain incompletely understood.

Recently, we reported that the activity of retinoic acid receptor/direct repeat-5 (RAR/DR-5) was suppressed among a number of transcription factors whose activities were markedly suppressed in the kidney of mice chronically exposed to Cd (Lee et al., 2021). The reduced activity of RAR/DR-5 was also shown in rat proximal tubular cells (NRK-52E cells) upon Cd exposure (Tokumoto et al., 2014). RAR belongs to a nuclear receptor superfamily and functions as a ligand-dependent transcription factor. In humans, three isoforms—RARα, RARβ, and RARγ—have been characterized (Das et al., 2014). RAR typically forms heterodimers with retinoid X receptors (RXRs) and binds to specific DNA sequences known as retinoic acid response elements located in the promoter regions of target genes to regulate their transcriptional activity (Das et al., 2014). One of the principal ligands for RAR is retinoic acid (RA), a derivative of vitamin A (Yamamoto et al., 2021), which is also a ligand of peroxisome proliferator-activated receptor δ (PPARδ) (Rieck et al., 2008; Wolf, 2010). Our previous study demonstrated that PPARδ is one of a number of modification factors of Cd renal toxicity (Mori et al., 2022). In humans and other vertebrates, the dietary intake of retinoids—including vitamin A and its derivatives—is essential for maintaining normal physiological functions (Brun et al., 2015). Retinoids play crucial roles in diverse biological processes by regulating cellular proliferation, differentiation, and apoptosis, and are therefore indispensable for immune regulation, reproduction, and embryonic development (Chambon, 1996; Gudas, 2012).

Despite these findings, the relationship between Cd-induced renal toxicity and the RA-RAR signaling axis remains largely unexplored. Therefore, we examined whether RA could attenuate Cd-induced cytotoxicity in human proximal tubular epithelial cells (HK-2), and whether this protective effect is mediated via RARs or alternative signaling pathways.

MATERIALS AND METHODS

Cell culture and Cd treatment

HK-2 cells (ATCC, Manassas, VA, USA) were maintained in DMEM/Ham’s F-12 (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% fetal bovine serum (FBS) (Gibco; Thermo Fisher Scientific, Waltham, MA, USA), penicillin (25 U/mL), streptomycin (25 µg/mL), 1% Insulin–Transferrin–Selenium-X (Gibco), epidermal growth factor (EGF; 10 ng/mL), and hydrocortisone (5 ng/mL) at 37°C in a humidified atmosphere of 5% CO₂. For exposure experiments, cells were seeded into 96-well plates at 2.5 × 10⁴ cells/cm² and grown to confluence. The medium was then replaced with serum-free medium containing cadmium chloride (CdCl₂; Wako Pure Chemical Industries, Osaka, Japan), and cells were incubated for the indicated times.

Cell viability

After treatment, culture medium was changed to fresh 10% FBS-DMEM containing 10% alamarBlue™ (Invitrogen; Thermo Fisher Scientific) and incubated for another 4 hr at 37°C. Fluorescence was measured at an excitation wavelength of 540 nm and emission wavelength of 595 nm with a SpectraMax® iD3 microplate reader (Molecular Devices, San Jose, CA, USA).

siRNA transfection

Silencer Select siRNAs (Ambion; Thermo Fisher Scientific) were used. Transfections were performed with Lipofectamine RNAiMAX (Invitrogen) following the manufacturer’s instructions. Briefly, siRNA complexes were prepared by incubating siRNAs with RNAiMAX in Opti-MEM (Gibco) for 15 min at room temperature and then added to HK-2 cells to yield final conditions of 1 nM or 0.05 nM per sequence, 0.2% (v/v) RNAiMAX, and 10% (v/v) Opti-MEM. Cells were incubated for 48 hr before downstream analyses.

RNA extraction and real-time RT-PCR

Cd-treated or siRNA-transfected HK-2 cells were washed twice with ice-cold phosphate-buffered saline [PBS(–); Nissui, Tokyo, Japan]. Total RNA was extracted using a PureLink™ RNA Mini Kit (Ambion) in accordance with the manufacturer’s instructions. RNA concentration and purity were determined with a BioSpec-nano spectrophotometer (Shimadzu Biotech, Kyoto, Japan). First-strand cDNA was synthesized from total RNA using the PrimeScript RT Reagent Kit (Perfect Real Time; Takara Bio, Shiga, Japan). Quantitative real-time PCR was performed with SYBR Premix Ex Taq II (Perfect Real Time; Takara Bio) on a Thermal Cycler Dice Real Time System (Takara Bio). Cycling conditions were 95°C for 10 sec, followed by 40 cycles of 95°C for 5 sec and 60°C for 30 sec. Gene expression levels were normalized to GAPDH mRNA. The oligonucleotide sequences of the primers were as follows: sense, 5′-GCACCGTCAAGGCTGAGAAC-3′, and antisense, 5′-TGGTGAAGACGCCAGTGGA-3′, for human GAPDH; sense, 5′-GTGTCACCGGGACAAGAACT-3′, and antisense, 5′-GGGCACCTCCTTCTTCTTCT-3′, for human RARA; sense, 5′-GGATCAATGCCACCTCTCAT-3′, and antisense, 5′-GGTGACTGACTGACCCCACT-3′ for human RARB; sense, 5′-AACAAGGTGACCAGGAA-3′, and antisense, 5′-TGTCAGGTGACCCTTCT-3′ for human RARG; sense, 5′-GAAACAGGCCTTCTCAGTGC-3′, and antisense, 5′-TTGCTGGGTCGTCTTTTTCT-3′ for human PPARD; sense, 5′-TCTAAGCGTCACCACGACTTCA-3′, and antisense, 5′-GTGCACTTGCAGTTCTTGCAG-3′ for human MT-1X; sense, 5′-CCTGCAATGCAAACAACAATGC-3′, and antisense, 5′-AGCTGCACTTGTCGGAAGC-3′ for human MT-2A.

Cd contents

After treatment, cells were washed twice with ice-cold PBS(–) and then three times with PBS(–) containing 2 nM ethylene glycol tetraacetic acid (EGTA; Nacalai Tesque, Kyoto, Japan). Cells were harvested in 1 mL RIPA buffer [25 mM Tris-HCl (pH 7.6), 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS; Thermo Fisher Scientific] and subjected to acid digestion with nitric acid and hydrogen peroxide. Cd was quantified by atomic absorption spectrometry (200 Series AA; Agilent Technologies, Santa Clara, CA, USA). Protein concentrations were determined using the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific) and used to normalize cellular Cd content.

Western blotting

HK-2 cells were treated with the indicated RA and Cd in 6-cm dishes. Following treatment, cells were washed twice with ice-cold PBS(–) and lysed in RIPA buffer. Protein concentrations were determined using the Pierce™ BCA Protein Assay Kit. Equal amounts of protein were resolved by SDS-PAGE and transferred to polyvinylidene difluoride (PVDF) membranes. Membranes were incubated with primary antibodies followed by horseradish peroxidase-conjugated secondary antibodies (1:10,000; GE Healthcare, Little Chalfont, UK). Immunoreactive bands were visualized by enhanced chemiluminescence (Chemi-Lumi One Super; Nacalai Tesque) and imaged with a ChemiDoc™ system (Bio-Rad, Hercules, CA, USA). Primary antibodies were anti-GAPDH (1:1,000; American Research Products, Waltham, MA, USA), anti-caspase-3 (1:1,000; Cell Signaling Technology, Danvers, MA, USA), and anti-cleaved caspase-3 (1:1,000; Cell Signaling Technology).

Apoptosis

Apoptosis was quantified using the Cell Death Detection ELISA^PLUS kit (Roche, Basel, Switzerland) according to the manufacturer’s instructions (Mori et al., 2022). The absorbance was measured at 405 nm with 490 nm as reference using a SpectraMax® iD3 microplate reader. Staurosporine (STS; Sigma-Aldrich) was used as a positive control. Cd-induced apoptosis was normalized to the apoptosis level induced by 0.1 µM STS.

Statistical analysis

Statistical analyses were performed using one-way or two-way ANOVA. When the omnibus F-test was significant (P < 0.05), post hoc pairwise comparisons were conducted using Bonferroni-adjusted t-tests (P < 0.05). All analyses were carried out with SPSS Statistics (IBM, Tokyo, Japan).

RESULTS

Effects of RA and retinol on Cd toxicity

To investigate the influence of RA on Cd sensitivity, HK-2 cells were pretreated with RA and its metabolic precursor, retinol, 24 hr prior to Cd exposure. As a result, pretreatment with RA or retinol significantly attenuated Cd-induced toxicity in a concentration-dependent manner (Fig. 1). These findings demonstrated that RA and retinol function as protective factors against Cd toxicity.

Fig. 1

Effects of RA and retinol on Cd toxicity. HK-2 cells were pretreated with RA or retinol for 24 hr. After washing out RA or retinol, HK-2 cells were treated with Cd for 24 hr. Cell viability was examined with an alamrBlue assay. Values are the means ± S.D. (n = 3). (A) *RA 5 μM group significantly different from RA 0 μM group, P < 0.05. ^†RA 10 μM group significantly different from RA 0 μM group, P < 0.05. (B) *Retinol 10 μM group significantly different from retinol 0 μM group, P < 0.05. ^†Retinol 20 μM group significantly different from retinol 0 μM group, P < 0.05.

Effects of knockdown of RAR on RA-reduced Cd toxicity

As RA is involved in gene transcription by binding to RAR in the nuclei (Yamamoto et al., 2021), we assessed the involvement of RAR isoforms in Cd sensitivity. Of the three RAR isoforms, RARA and RARG are mainly expressed in HK-2 cells (Fig. 2A). Therefore, we examined the involvement of RARα and RARγ in RA-suppressed Cd toxicity. Treatment with RARA or RARG siRNA significantly decreased the expressions of RARA and RARG, respectively (Figs. 2B and 2C). However, the siRNA-mediated knockdown of RARA or RARG did not affect the expression of the other gene (Figs. 2B and 2C). Furthermore, the knockdown of RARA did not significantly alter Cd toxicity compared with the control cells (Fig. 2D), nor did knockdown of RARG (Fig. 2E).

Fig. 2

Effects of RAR siRNA on RA-reduced Cd toxicity. (A) The gene expression levels of RARs in HK-2 cells. RARA, RARB, and RARG mRNA levels were examined by real-time RT-PCR and calculated using the double delta Ct method. Values are the means ± S.D. (n = 3). *P < 0.05. (B, C) The gene expression levels of RARA and RARG in HK-2 cells treated with RARA or RARB siRNA for 48 hr. mRNA levels were examined by real-time RT-PCR and normalized to GAPDH mRNA levels. *Significantly different from the control group, P < 0.05. (D, E) HK-2 cells were transfected with RARA or RARG siRNA for 48 hr, treated with RA for 24 hr, washed, and then exposed to Cd for 24 hr. Cell viability was examined with an alamarBlue assay. (D) ^#Control siRNA + RA 10 μM group significantly different from control siRNA + RA 0 μM group, ^$RARA siRNA + RA 10 μM group significantly different from RARA siRNA + RA 0 μM group, P < 0.05. (E) *Control siRNA + RA 10 μM group significantly different from control siRNA + RA 0 μM group, ^#RARG siRNA + RA 0 μM group significantly different from control siRNA + RA 0 μM group, ^†RARG siRNA + RA 10 μM group significantly different from RARG siRNA + RA 0 μM group, P < 0.05.

The single knockdown of RARA or RARG did not affect the RA-induced resistance to Cd toxicity. Next, the effects of double knockdown of RARA and RARG on RA-reduced Cd toxicity were examined and we found that 1.0 nM RARG siRNA induced cell toxicity by approximately 20% (Fig. 2E). To prevent the marked reduction in cell death caused by siRNA treatment, we transfected HK-2 cells with siRNAs targeting RARA and RARG at 0.05 nM each for the double knockdown. The gene expressions of RARA and RARG were decreased significantly by the complexed siRNA treatment (Figs. 3A and 3B). Although the gene expressions of RARA and RARG were suppressed simultaneously, the protective effect of RA against Cd toxicity was significant even with simultaneous knockdown of RARA and RARG (P < 0.05) (Fig. 3C). Only treatment with 20 µM Cd weakened RA-reduced Cd toxicity by the double knockdown (Fig. 3C). Therefore, although the RAR transcriptional pathway is partly involved, RA suppresses Cd toxicity predominantly through RAR-independent mechanisms.

Fig. 3

Effects of the double knockdown of RARA and RARG on RA-reduced Cd toxicity. (A, B) The gene expression levels of RARA and RARG knockdown in HK-2 cells treated with RARA and RARG siRNAs for 48 hr. mRNA levels were examined by real-time RT-PCR and normalized to GAPDH mRNA levels. Values are the means ± S.D. (n = 6). *Significantly different from the control group, P < 0.05. (C) HK-2 cells were transfected with RARA and RARG siRNAs for 48 hr, treated with RA for 24 hr, washed, and then exposed to Cd for 24 hr. Cell viability was examined with an alamarBlue assay. Values are the means ± S.D. (n = 4 – 5). ^#Control siRNA + RA 10 μM group significantly different from control siRNA + RA 0 μM group, ^$RAR siRNAs + RA 10 μM group significantly different from RAR siRNAs + RA 0 μM group, P < 0.05.

Effects of PPARD siRNA on RA-reduced Cd toxicity

Previously, PPARδ was identified as a modulatory factor in Cd toxicity, as the knockdown of PPARD rendered HK-2 cells resistant to Cd-induced cytotoxicity (Mori et al., 2022). RA is known to act as a ligand for PPARδ (Rieck et al., 2008; Wolf, 2010). If the protective effect of RA against Cd toxicity is mediated through PPARδ, it is expected that the knockdown of PPARD would attenuate the effect of RA. Unexpectedly, however, RA-decreased Cd toxicity was not altered by PPARD knockdown (Fig. 4). Despite RA being reported as a PPARδ ligand, PPARD knockdown does not change RA’s protective effect against Cd toxicity.

Fig. 4

Effects of PPARD siRNA on RA-reduced Cd toxicity. (A) The gene expression level of PPARD in HK-2 cells treated with PPARD siRNA for 48 hr. mRNA levels were examined by real-time RT-PCR and normalized to GAPDH mRNA levels. *Significantly different from the control group, P < 0.05. (B) HK-2 cells were transfected with PPARD siRNAs for 48 hr, treated with RA for 24 hr, washed, and then exposed to Cd for 24 hr. Cell viability was examined with an alamarBlue assay. *Control siRNA + RA 10 μM group significantly different from the control siRNA + RA 0 μM group; ^#PPARD siRNA + RA 0 μM group significantly different from the control siRNA + RA 0 μM; ^†PPARD siRNA + RA 10 μM group significantly different from the PPARD siRNA + RA 0 μM group, P < 0.05.

Effect of RA on intracellular Cd levels

Changes in intracellular Cd levels following RA pretreatment may contribute to its protective effect, and therefore we examined the influence of RA pretreatment on the cellular accumulation of Cd. After HK-2 cells were exposed to Cd, intracellular Cd concentrations were measured and were increased with increasing concentrations of Cd. However, RA pretreatment did not significantly alter intracellular Cd levels compared with Cd treatment alone (Fig. 5). Consequently, the protective effect of RA against Cd toxicity does not occur via the changes in intracellular Cd accumulation.

Fig. 5

Effect of RA pretreatment on the intracellular Cd concentration. HK-2 cells were pretreated with RA for 24 hr. After washing out RA, HK-2 cells were treated with Cd for 12 hr. The Cd concentration was measured by an atomic absorption spectrometer. Values are the means ± S.D. (n = 3). *Significantly different from the Cd 10 μM group, P < 0.05.

Effect of RA treatment on MT gene expression

MT, a metal-binding protein, protects against Cd toxicity, with MT-1X and MT-2A identified as the major isoforms most closely associated with Cd exposure (Miura and Koizumi, 2007). To examine whether RA modulated the expression of these isoforms, we analyzed mRNA levels of MT-1X and MT-2A following RA treatment. RA treatment did not significantly alter the expressions of MT-1X or MT-2A at the mRNA level (Fig. 6). Taken together, the protective effect of RA pretreatment against Cd toxicity is not mediated through the induction of MT expression.

Fig. 6

Effect of RA on MT mRNA levels. HK-2 cells were pretreated with RA for 24 hr. MT-1X and MT-2A mRNA levels were examined by real-time RT-PCR and normalized to GAPDH mRNA levels. Values are the means ± S.D. (n = 3).

Effect of RA on Cd-induced apoptosis

As Cd is known to induce apoptosis and contribute to renal toxicity (Fujiwara et al., 2012; Lee et al., 2025), we examined whether RA pretreatment attenuated Cd-induced apoptosis. Exposure to 20 μM Cd for 12 hr significantly increased apoptosis in HK-2 cells (Fig. 7A). Notably, RA pretreatment reduced apoptotic cells from 32% to 18% (P < 0.05), indicating a significant anti-apoptotic effect (Fig. 7A). Cd-induced apoptosis has been previously shown to involve the activation of caspase-3, a key executioner of the apoptotic pathway (Fujiwara et al., 2012; Lee et al., 2025; Li et al., 2000). Therefore, we examined the effect of RA pretreatment on caspase-3 activation in Cd-exposed HK-2 cells. After 6 hr of Cd exposure, RA pretreatment attenuated the increase in cleaved caspase-3 levels observed after treatment with 20 μM or 30 μM Cd (Fig. 7B). Similarly, following 12 hr of exposure, the elevation of cleaved caspase-3 induced by 10 μM Cd was reduced by RA pretreatment (Fig. 7C). Collectively, these findings demonstrated that RA pretreatment suppresses Cd-induced apoptosis in HK-2 cells, at least in part, by inhibiting the activation of caspase-3.

Fig. 7

Effect of RA pretreatment on apoptosis induced by Cd. (A) After pretreatment with RA for 24 hr, HK-2 cells were treated with Cd for 12 hr. Apoptosis was examined by a Cell Death Detection ELISA^PLUS Kit. Values are the means ± S.D. (n = 3). * Significantly different from the control group, P < 0.05. ^#P < 0.05 (B, C) After pretreatment with RA for 24 hr, HK-2 cells were treated with Cd for 6 hr (B) or 12 hr (C). Protein levels of caspase-3 and cleaved caspase-3 were examined by western blotting. GAPDH was used as the loading control.

DISCUSSION

In the present study, we demonstrated that RA functions as a protective factor against Cd toxicity. Importantly, the protective effect of RA was implicated to involve the suppression of apoptosis by inhibiting caspase-3 activation. Although RA primarily functions as a ligand for RAR and PPARδ to regulate gene transcription, our findings revealed that RAR and PPARδ were mostly not involved in the protective effects of RA against Cd-induced cytotoxicity. Moreover, RA was demonstrated as one of the modification factors of Cd toxicity in addition to PPARδ; however, RA and PPARδ affect Cd toxicity through separate pathways.

Within the cytoplasm, RA bound to cellular retinoic acid-binding protein (CRABP) to direct RAR-dependent transcription, whereas RA bound to fatty-acid–binding protein 5 (FABP5) engages RAR-independent transcriptional routes (Connolly et al., 2013; Costantini et al., 2020). Accordingly, a variety of RA-binding proteins might participate in the RA-mediated reduction of Cd toxicity. Previous studies have reported that RA alleviated apoptosis in mesangial cells by inhibiting the JNK-AP-1 (c-Jun N-terminal kinase–activator protein-1) signaling pathway, including suppression of JNK activation, downregulation of c-fos and c-jun expressions, and inhibition of AP-1 transcriptional activity (Moreno-Manzano et al., 1999). Furthermore, in cardiac tissues, RA mitigated apoptosis by partially suppressing ROS production and inhibiting the phosphorylation of MAPKs (mitogen-activated protein kinases), including p38, JNK, and ERK (extracellular signal-regulated kinase) (Zhu et al., 2015). Notably, mitochondrial ROS generation and MAPK pathways involving p38 and JNK have been implicated in Cd-induced toxicity (Gobe and Crane, 2010; Jiang et al., 2015). These findings raise the possibility that RA may suppress Cd-induced apoptosis through multiple intracellular signaling pathways beyond RAR-mediated transcription. Thus, for fully understanding the apoptosis-related mechanism, it is necessary to test the direct involvement of upstream pathways (e.g., mitochondrial ROS, JNK/p38/ERK MAPKs, AP-1). Additionally, RA has been reported to enhance the function of the mannose-6-phosphate/insulin-like growth factor II (M6P/IGF-II) receptor, which is involved in the regulation of cell proliferation and apoptosis (Kang et al., 1997). Taken together, these findings support the notion that RA may exert its anti-apoptotic effects via diverse pathways and receptor systems, including mechanisms independent of classical RAR and RXR signaling.

Our experiments were conducted exclusively in HK-2 cells, an immortalized proximal tubular epithelial cell line. Further studies are required for the replicate key findings using mice or other in vivo models.

Vitamin A, an essential fat-soluble micronutrient that exists in multiple chemical forms (retinol, retinal, RA, retinyl esters, and provitamin A carotenoids), supports diverse physiological processes (Carazo et al., 2021). In addition to sustaining the visual cycle, vitamin A supports normal cell development and metabolic functions (Carazo et al., 2021; Yamamoto et al., 2021). Retinol functions as a cofactor in multiple enzymatic reactions; retinal serves as a chromophore in vision; and RA mediates diverse effects by binding to RARs to regulate gene expression (Carazo et al., 2021; Yamamoto et al., 2021). Several groups reported that RA has preventive and therapeutic effects on kidney diseases in experimental models (Chen et al., 2021; Mallipattu and He, 2015). However, RA can cause significant adverse effects, including liver toxicity, hyperlipidemia, and central nervous system abnormalities (Chen et al., 2021; Orfanos et al., 1987). Thus, RA may produce opposite outcomes through the activation of different downstream and tissue-restricted pathways. Clarifying these cascades and their gene targets will facilitate the design of agents that promote the protective effect of RA against Cd renal toxicity.

Funding

This work was supported partly by the Study of the Health Effects of Heavy Metals, organized by the Ministry of the Environment, Japan.

Conflict of interest

The authors declare that they have no conflicts of interest with regard to this study.

Data availability

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

Author contributions

Conceptualization: J.Y.L., M.S.

Data curation: J.Y.L., C.M., M.T., M.S.

Funding acquisition: M.S.

Formal analysis: J.Y.L., C.M., M.T., M.S.

Investigation: J.Y.L., C.M.

Project administration: M.S.

Supervision: M.S.

Writing – original draft: J.Y.L.

Writing – review and editing: M.T., M.S.

Ethical approval and consent to participate

Not applicable.

Patient consent for publication

Not applicable.

REFERENCES

•  Brun,  P.J.,  Grijalva,  A.,  Rausch,  R.,  Watson,  E.,  Yuen,  J.J.,  Das,  B.C.,  Shudo,  K.,  Kagechika,  H.,  Leibel,  R.L. and  Blaner,  W.S. (2015): Retinoic acid receptor signaling is required to maintain glucose-stimulated insulin secretion and β-cell mass. FASEB J., 29, 671-683.
•  Carazo,  A.,  Macáková,  K.,  Matoušová,  K.,  Krčmová,  L.K.,  Protti,  M. and  Mladěnka,  P. (2021): Vitamin A Update: Forms, Sources, Kinetics, Detection, Function, Deficiency, Therapeutic Use and Toxicity. Nutrients, 13.
•  Chambon,  P. (1996): A decade of molecular biology of retinoic acid receptors. FASEB J., 10, 940-954.
•  Chen,  A.,  Liu,  Y.,  Lu,  Y.,  Lee,  K. and  He,  J.C. (2021): Disparate roles of retinoid acid signaling molecules in kidney disease. Am. J. Physiol. Renal Physiol., 320, F683-F692.
•  Connolly,  R.M.,  Nguyen,  N.K. and  Sukumar,  S. (2013): Molecular pathways: current role and future directions of the retinoic acid pathway in cancer prevention and treatment. Clin. Cancer Res., 19, 1651-1659.
•  Costantini,  L.,  Molinari,  R.,  Farinon,  B. and  Merendino,  N. (2020): Retinoic Acids in the Treatment of Most Lethal Solid Cancers. J. Clin. Med., 9.
•  Das,  B.C.,  Thapa,  P.,  Karki,  R.,  Das,  S.,  Mahapatra,  S.,  Liu,  T.C.,  Torregroza,  I.,  Wallace,  D.P.,  Kambhampati,  S.,  Van Veldhuizen,  P.,  Verma,  A.,  Ray,  S.K. and  Evans,  T. (2014): Retinoic acid signaling pathways in development and diseases. Bioorg. Med. Chem., 22, 673-683.
•  Fujiwara,  Y.,  Lee,  J.Y.,  Tokumoto,  M. and  Satoh,  M. (2012): Cadmium renal toxicity via apoptotic pathways. Biol. Pharm. Bull., 35, 1892-1897.
•  Gobe,  G. and  Crane,  D. (2010): Mitochondria, reactive oxygen species and cadmium toxicity in the kidney. Toxicol. Lett., 198, 49-55.
•  Gudas,  L.J. (2012): Emerging roles for retinoids in regeneration and differentiation in normal and disease states. Biochim. Biophys. Acta, 1821, 213-221.
•  Järup,  L. (2002): Cadmium overload and toxicity. Nephrol. Dial. Transplant., 17 (Suppl 2), 35-39.
•  Järup,  L. and  Akesson,  A. (2009): Current status of cadmium as an environmental health problem. Toxicol. Appl. Pharmacol., 238, 201-208.
•  Järup,  L.,  Berglund,  M.,  Elinder,  C.G.,  Nordberg,  G. and  Vahter,  M. (1998): Health effects of cadmium exposure--a review of the literature and a risk estimate. Scand. J. Work Environ. Health, 24 (Suppl 1), 1-51.
•  Jiang,  J.H.,  Ge,  G.,  Gao,  K.,  Pang,  Y.,  Chai,  R.C.,  Jia,  X.H.,  Kong,  J.G. and  Yu,  A.C. (2015): Calcium Signaling Involvement in Cadmium-Induced Astrocyte Cytotoxicity and Cell Death Through Activation of MAPK and PI3K/Akt Signaling Pathways. Neurochem. Res., 40, 1929-1944.
•  Kang,  J.X.,  Li,  Y. and  Leaf,  A. (1997): Mannose-6-phosphate/insulin-like growth factor-II receptor is a receptor for retinoic acid. Proc. Natl. Acad. Sci. USA, 94, 13671-13676.
•  Klaassen,  C.D.,  Liu,  J. and  Diwan,  B.A. (2009): Metallothionein protection of cadmium toxicity. Toxicol. Appl. Pharmacol., 238, 215-220.
•  Lee,  J.Y.,  Mori,  C.,  Tokumoto,  M. and  Satoh,  M. (2021): Changes in DNA-binding activity of transcription factors in the kidney of mice exposed to cadmium. J. Toxicol. Sci., 46, 125-129.
•  Lee,  J.Y.,  Tokumoto,  M. and  Satoh,  M. (2025): Molecular Mechanisms of Cadmium-Induced Toxicity and Its Modification. Int. J. Mol. Sci., 26.
•  Li,  M.,  Kondo,  T.,  Zhao,  Q.L.,  Li,  F.J.,  Tanabe,  K.,  Arai,  Y.,  Zhou,  Z.C. and  Kasuya,  M. (2000): Apoptosis induced by cadmium in human lymphoma U937 cells through Ca2+-calpain and caspase-mitochondria- dependent pathways. J. Biol. Chem., 275, 39702-39709.
•  Mallipattu,  S.K. and  He,  J.C. (2015): The beneficial role of retinoids in glomerular disease. Front. Med. (Lausanne), 2, 16.
•  Miura,  N. and  Koizumi,  S. (2007): Heavy metal responses of the human metallothionein isoform genes. Yakugaku Zasshi, 127, 665-673.
•  Moreno-Manzano,  V.,  Ishikawa,  Y.,  Lucio-Cazana,  J. and  Kitamura,  M. (1999): Suppression of apoptosis by all-trans-retinoic acid. Dual intervention in the c-Jun n-terminal kinase-AP-1 pathway. J. Biol. Chem., 274, 20251-20258.
•  Mori,  C.,  Lee,  J.Y.,  Tokumoto,  M. and  Satoh,  M. (2022): Cadmium Toxicity Is Regulated by Peroxisome Proliferator-Activated Receptor δ in Human Proximal Tubular Cells. Int. J. Mol. Sci., 23.
•  Orfanos,  C.E.,  Ehlert,  R. and  Gollnick,  H. (1987): The retinoids. A review of their clinical pharmacology and therapeutic use. Drugs, 34, 459-503.
•  Rieck,  M.,  Meissner,  W.,  Ries,  S.,  Müller-Brüsselbach,  S. and  Müller,  R. (2008): Ligand-mediated regulation of peroxisome proliferator-activated receptor (PPAR) beta/delta: a comparative analysis of PPAR-selective agonists and all-trans retinoic acid. Mol. Pharmacol., 74, 1269-1277.
•  Tokumoto,  M.,  Lee,  J.Y.,  Fujiwara,  Y. and  Satoh,  M. (2014): Alteration of DNA binding activity of transcription factors in NRK-52E rat proximal tubular cells treated with cadmium. J. Toxicol. Sci., 39, 735-738.
•  Wolf,  G. (2010): Retinoic acid activation of peroxisome proliferation-activated receptor delta represses obesity and insulin resistance. Nutr. Rev., 68, 67-70.
•  Yamamoto,  Y.,  Matsumoto,  Y.,  Suzuki,  T. and  Inoue,  J. (2021): Study of Control Mechanism of Vitamin A Metabolism through its Specific Binding Proteins. VITAMINS, 95, 257-265.
•  Zhu,  Z.,  Zhu,  J.,  Zhao,  X.,  Yang,  K.,  Lu,  L.,  Zhang,  F.,  Shen,  W. and  Zhang,  R. (2015): All-Trans Retinoic Acid Ameliorates Myocardial Ischemia/Reperfusion Injury by Reducing Cardiomyocyte Apoptosis. PLoS One, 10, e0133414.