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Impact of fluoride on antioxidant activity and mitochondrial homeostasis in fetal rat kidney.
Abstract
Full-text study online at
https://www.sciencedirect.com/science/article/pii/S0300483X25003014?via%3Dihub
Fluoride exposure harms human health. Additionally, recent attention has focused on understanding the impact of an adverse in utero environment on the programming of chronic kidney disease in adult offspring. A previous study has described that fluoride alters kidney development, inducing oligonephronia and promoting the premature maturation of remnant nephrons. Therefore, it is essential to continue researching the potential mechanisms involved in fluoride-induced alterations. Female Wistar rats were administered two doses of fluoride (2.5 or 5.0 mg/kg body weight/day) for 20 days before and after mating. The results showed that fluoride diminished catalase and superoxide dismutase activities while increasing the markers of nuclear factor erythroid 2-related factor 2 (Nrf2) activation, mitochondrial dynamics, autophagy, and biogenesis. Morphometric analysis revealed an increase in mitochondrial size and content. Although the levels of Bcl-2-associated X protein (Bax) and B-cell lymphoma 2 (Bcl-2) increased, the activity of caspase-3 and the immunodetection in the TUNEL assay decreased. Several of these changes were mainly observed with the high fluoride dose. In conclusion, the results suggest that fluoride alters the redox status and mitochondrial homeostasis while decreasing apoptosis. It is essential to recognize that, under normal conditions, apoptosis plays a crucial role in proper kidney development, and its occurrence decreases as the structures mature. Therefore, collectively, these alterations could lead to undesirable effects on postnatal life.
Abbreviations
AR, Aspect ratio; ATP, Adenosine triphosphate; Bax, Bcl-2-associated X protein; Bcl-2, B-cell lymphoma 2; CAT, Catalase; CTRL, Control; DRP1, Dynamin-related protein 1; F2.5, 2.5 mg fluoride/kg body weight; F5.0, 5.0 mg fluoride/kg body weight; GD, Gestation day; yGCS, Gamma-glutamylcysteine synthetase; GPx, Glutathione peroxidase; GRx, Glutathione reductase; GSH, Glutathione; GSS, glutathione synthetase; HO-1, Heme oxygenase 1; LC3-I, Microtubule-associated protein 1 light chain 3; LC3-II, LC3-phosphatidylethanolamine conjugate; MDA, Malondialdehyde; MFN2, Mitofusin 2; MtDNA, Mitochondrial DNA; NADPH, Nicotinamide adenine dinucleotide phosphate; NBT, Nitro blue tetrazolium; NRF1, Nuclear respiratory factor 1; NRF2, Nuclear respiratory factor 2.; Nrf2, Nuclear factor erythroid 2-related factor 2; PNrf2, Phosphorylated Nrf2; P62, Ubiquitin-binding protein p62; PGC-1a, Peroxisome proliferator-activated receptor gamma coactivator 1-alpha; PINK1, PTEN-induced putative kinase 1; PPINK1, Phosphorylated PINK1; PTEN, Phosphatase and tensin homolog; RFU, Relative fluorescence unit; ROS, Reactive oxygen species; S40, Serine 40; SEM, Standard error of the mean; SOD, Superoxide dismutase; TFAM, Mitochondrial transcription factor A; TUNEL, Terminal deoxynucleotidyl transferase TdT-mediated nick-end labeling; VDAC, Voltage-dependent anion channel.
Keywords
Fluoride
Kidney
Mitochondrial
Apoptosis
Oxidative stress
Autophagy
1. Introduction
Fluoride is an inorganic anion widely distributed globally (Kimambo et al., 2019). Humans can be exposed to fluoride from various sources, including anthropogenic activities, fluoride-containing dental products, and water fluoridation (Johnston and Strobel, 2020). However, fluoride’s impact on human health can vary, exhibiting both beneficial and harmful effects, depending on its concentration and the quantity ingested over time (Ahmad et al., 2022).
Fluoride’s detrimental effects impact hard and soft tissues, including the liver, lungs, heart, brain, reproductive organs, and kidneys (Hamza et al., 2015, Sharma et al., 2023). The mechanisms underlying fluoride-induced toxicity involve oxidative stress (Zuo et al., 2018), mitochondrial dysfunction (Zhao et al., 2020), and an imbalance in autophagy (Tian et al., 2020, Zhou et al., 2019).
Regarding the kidney, although there is conflicting information on fluoride’s effects in adult, adolescent, and child populations (Jiménez-Córdova et al., 2019, Jiménez-Córdova et al., 2018), its impact during the last pregnancy stage (Montañez-Rodriguez et al., 2024) and in early life (Niu et al., 2016) remains limited. Previously, our research group found that fluoride exposure during gestation disrupts normal kidney development and accelerates the maturation of tubular segments (Montañez-Rodriguez et al., 2024).
The maturation of antioxidant capacity in the kidneys is a critical factor during the late gestation stage. Sprague-Dawley rat fetuses from gestation day 18 (GD18) to GD22 show a significant increase in levels of superoxide dismutase 1 and 2 (SOD1 and SOD2, respectively), glutathione peroxidase (GPx), and catalase (CAT) (Hayashibe et al., 1990). These adaptations seem to align with the changes in the nephron’s metabolic demand (Diniz et al., 2023, Pejznochova et al., 2010), whereby at GD13.5, nephrons exhibit high glycolysis rates and low oxidative phosphorylation levels; although this profile switches between the end of the late gestation and the onset of the postnatal stage (Liu et al., 2017). During late gestation, mitochondria undergo significant changes, including increased mitochondrial DNA (mtDNA) levels, oxygen consumption, and the content and activity of adenosine triphosphate (ATP) synthase and cytochrome c oxidase, indicating an increase in the number of mitochondria (Prieur et al., 1998, Prieur et al., 1995). Yet, an increase in ATP production by mitochondria is also indirectly associated with a rise in reactive oxygen species (ROS) generation (Murphy, 2009).
Briefly, mitochondria can undergo reshaping through fusion-fission cycles, without changing their total mass (Eisner et al., 2018). Fission helps segregate damaged mitochondria for degradation. It is mainly regulated by dynamin-related protein 1 (DRP1) (Tang et al., 2021). Conversely, fusion allows the exchange of proteins, metabolites, and substrates, ensuring the optimal function of the mitochondrial network. It is facilitated by mitofusins, among other proteins (Bhargava and Schnellmann, 2017).
On the other hand, the response to stressors may also involve changes in mitochondrial mass through biogenesis and/or mitophagy (Eisner et al., 2018). Briefly, mitochondrial transcription factor A (TFAM) is involved in initiating mtDNA transcription and replicating its genome (Rubalcava-Gracia et al., 2023). TFAM is regulated by the nuclear respiratory factors 1 and 2 (NRF1 and NRF2, respectively), which are influenced by the peroxisome proliferator-activated receptor gamma coactivator 1-alpha (PGC-1a) (Ploumi et al., 2017).
Mitophagy enables the selective degradation of damaged mitochondria by the autophagy machinery (Doke and Susztak, 2022). The phosphatase and tensin homolog (PTEN)-induced putative kinase 1 (PINK1) and Parkin are the main proteins involved in the ubiquitin-dependent mitophagy pathway (Cho and Sun, 2020). Additionally, mitophagy activation depends on the loss of mitochondrial membrane potential (Xian and Liou, 2021).
As mentioned, fluoride alters the redox status and mitochondrial homeostasis in postnatal life. Therefore, we conducted a study using female Wistar rats exposed to environmentally relevant fluoride concentrations. The primary aim of this study was to assess the impact of fluoride on the redox status and processes involved in mitochondrial homeostasis. In addition, we investigated whether these alterations affect the induction of apoptosis in the fetal kidney.
2. Material and methods
2.1. Chemicals
All details about the chemicals and antibodies used in this study are available in the supplementary material.
2.2. Experimental design
The experimental protocol was reviewed and approved by the Institutional Committee for the Care and Use of Laboratory Animals of CINVESTAV (Protocol No. 0041–13) and adhered to the Mexican Official Standard NOM-062-ZOO-1999. In addition, all animal experiments were conducted in strict compliance with the ARRIVE guidelines.
Female Wistar rats (200–250 g) were housed in polycarbonate cages. Animals had free access to water and food throughout the study. After one week of acclimatization, the rats were randomly assigned to three groups (n = 8/group): (i) control group (CTRL); (ii) F2.5 and (iii) F5.0 group, animals treated by oral gavage with fluoride (2.5 or 5.0 F–/kg body weight/day, respectively). Animals were administered daily for 20 days, and their estrous cycles were monitored simultaneously. Additionally, on day 20, rats in the proestrus stage were mated with fertile male rats to induce pregnancy. The next day, the conception was confirmed by the presence of sperm in vaginal smears. Once conception was confirmed, this was considered GD0. The rats continued to receive treatment for the next 19 days. On the GD20, the animals were anesthetized with ketamine/xylazine (80 and 8 mg/kg, respectively), and the fetuses were obtained by cesarean (Fig. 1). Immediately, fetal kidneys were acquired. One kidney was immersed in a fixing solution for electron microscopy analysis. The remaining kidneys of the litter were stored at -70°C until the preparation of the homogenates.
Fig. 1. Schematic experimental design.
2.3. Antioxidant activity
Total protein extraction required a pool of 5 fetal kidneys per litter. The tissue was homogenized in 50 mM phosphate buffer, pH 7.0, containing 0.05 % Triton X-100 and immediately centrifuged at 10,000 g for 15 min at 4°C to obtain the supernatant. Immediately, total protein content in the supernatant was determined using the Lowry method. CAT activity was evaluated by determining the disappearance of 30 H2O2 at 240 nm (Aebi, 1984). Glutathione reductase (GRx) activity was determined by evaluating nicotinamide adenine dinucleotide phosphate (NADPH) disappearance at 340 nm coupled with the reduction of glutathione disulfide to glutathione (GSH) (Carlberg and Mannervik, 1975). The oxidation of 1 µmol of NADPH/min was used as one unit of GRx activity. GPx activity was measured at 340 nm using GRx and NADPH in a coupled reaction with H2O2 decomposition (Lawrence and Burk, 1976). The oxidation of 1 µmol of NADPH/min was used as one unit of enzyme activity for both assays. Total SOD activity was assayed at 560n m using the xanthine/xanthine oxidase system as a generator of superoxide anion radical and nitro blue tetrazolium (NBT) as the indicator reagent (Weydert and Cullen, 2010). The amount of protein that inhibited NBT reduction to 50 % was defined as one unit of SOD activity. SOD2 activity was determined using the same method; however, it was differentiated by adding 2 mM potassium cyanide, which explicitly inhibits SOD1. All data were expressed as Units per milligram protein (U/mg protein), except for CAT, which was expressed as k/mg protein.
2.4. Antioxidant status
Malondialdehyde (MDA) concentrations were determined according to described by Gérard-Monnier et al. (1998). Briefly, at least six fetal kidneys per litter were homogenized in phosphate buffer (50 mM, pH 7.0), supplemented with 5 mM butylated hydroxytoluene. Samples were centrifuged at 3000 g for 10 min at 4°C. 100 L of supernatant was added to 650 uL of a solution of 15.4 mM 1-methyl-2-phenylindole in a mixture of acetonitrile/methanol (3:1). The reaction was started by adding 150 uL of concentrated HCl. The 586 nm absorbance was measured upon incubating the mixture reaction at 45°C for 40 min (Gérard-Monnier et al., 1998). The results were calculated by using the tetramethoxypropane standard curve. MDA concentrations were expressed as umol/mg of protein.
2.5. Transmission electron microscopy
For the transmission electron microscopy, one kidney per litter was fixed with Karnovsky’s fixative (a mixture of 4 % paraformaldehyde and 1.5 % glutaraldehyde) in 0.2 M Sörensen buffer pH 7.2. Then, the tissue was post-fixed with 2 % osmium tetraoxide, dehydrated with ethyl alcohol, and embedded in epoxy resins. 70–90 nm sections were placed on copper grips, contrasted with uranyl acetate and lead citrate Reynol’s, and inspected with a transmission electron microscope (Tecnai Spirit BioTwin, FEI, Hillsboro, OR, USA). The mitochondrial size (um2), aspect ratio (AR), and mitochondrial area/field area ratio were determined using the ImageJ software provided by the National Institutes of Health (NIH; Bethesda, MD, USA).
2.6. Western blot
Total protein extraction required a pool of 5 fetal kidneys per litter. The tissue was homogenized in RIPA buffer containing protease and phosphatase inhibitors (1 mM NaF, 1 mM Na3VO4, and 1X protease inhibitor cocktail). The samples were centrifuged at 10,000 g for 15 min at 4°C. Immediately, protein concentration in the supernatant was determined using the Bradford method. Later, 25 µg of protein was loaded per well into 10–17 % polyacrylamide gels for separation. The proteins were immediately transferred onto a nitrocellulose membrane and stained with Ponceau red to verify protein transfer and equal loading among samples. Nonspecific binding sites on the membranes were blocked with 5 % non-fat milk or 5 % non-fat milk plus 2.5 % BSA in phosphate buffer saline with 0.1 % Tween 20, for 1 h at room temperature. The membranes were incubated overnight at 4°C with a specific primary antibody (Supplementary Table 1).
The next day, the membranes were incubated with the appropriate secondary antibody for 2 h at room temperature. The bands were visualized by chemiluminescence (Immobilon Forte Western horseradish peroxidase substrate) using autoradiography films. The bands were quantified by densitometric analysis using ImageJ software (Bethesda, MD, USA). Data were normalized with data of B-actin (1:3500).
2.7. DNA fragmentation (TUNEL)
DNA fragmentation in apoptotic cells was detected using the terminal deoxynucleotidyl transferase (TdT)-mediated nick-end labelling (TUNEL) method. For this, one kidney from each of three different litters were used to identify TUNEL-positive cells. Briefly, semi-thin sections were obtained from all kidneys. Thus, tissue sections were deparaffinized in xylene, rehydrated, and treated with proteinase K (20 µg/mL). Endogenous peroxidase activity was blocked by incubation in 3 % H2O2. After quenching, sections were incubated with Sniper Background to reduce nonspecific binding. Next, sections were incubated with the terminal deoxynucleotidyl transferase enzyme in a humidified chamber at 37°C for 1 h. Following a wash step, sections were incubated with anti-digoxigenin conjugate for 30 min at room temperature. Finally, after additional washing, sections were developed with peroxidase substrate, rinsed, and counterstained with hematoxylin. Finally, sections were examined with a light microscope equipped with a Nikon digital camera.
2.8. Caspase-3 activity
Caspase-3 activity was determined using the caspase-3 assay kit according to the manufacturer’s instructions. Briefly, 5–6 kidneys per litter were homogenized in lysis buffer, and the protein concentration was determined using the Bradford method. One hundred and fifty µg of proteins were placed in a 96-well plate and incubated for 30 min at 37°C. Caspase activity was expressed as relative fluorescence units (RFU)/µg protein.
2.9. Statistical analysis
All results are expressed as mean ± standard error of the mean (SEM). Additionally, normality was tested using the Kolmogorov-Smirnov test. The data were analyzed by one-way ANOVA followed by the Dunnett post hoc test using GraphPad Prism 10 (GraphPad Software, Boston, MA, USA). A p < 0.05 was considered statistically significant.
3. Results
3.1. Fluoride alters the antioxidant status
To determine if fluoride exposure evokes an antioxidant response, we evaluated the activity of several antioxidant enzymes. Firstly, the GRx activity remained consistent across all groups exposed to fluoride (Fig. 2A). Notably, the GPx activity showed a significant increase only in the F5.0 group compared to the CTRL group (Fig. 2B). Furthermore, both groups exposed to fluoride decreased CAT (Fig. 2C) and total SOD (Fig. 2D) activities relative to the CTRL group. In contrast, SOD-2 activity remained unchanged following fluoride exposure (Fig. 2E). The overproduction of ROS can result in oxidative damage. Therefore, we assessed whether fluoride exposure induces lipid peroxidation by quantifying MDA. However, the levels of MDA significantly decreased in the F2.5 group compared to the CTRL group (Fig. 2F). Since the nuclear factor erythroid 2-related factor 2 (Nrf2) transcription factor is considered a master regulator of the antioxidant response, we determined both the levels of Nrf2 and its phosphorylated form (pNrf2) (Fig. 2G), finding that fluoride exposure did not alter the Nrf2 levels (Fig. 2H). Conversely, both groups exposed to fluoride showed a significantly elevated level of pNrf2 compared to the CTRL group (Figs. 2G and 2I). By determining the pNrf2/Nrf2 ratio, it was observed that this pathway was only activated with the treatment of F2.5 (Fig. 2J). Additionally, we assessed the levels of gamma-glutamylcysteine synthetase (yGCS) and SOD2, which are regulated by Nrf2. Both proteins remained unchanged across all groups (Fig. 2K and L).
Fig. 2. Impact of fluoride on antioxidant enzymes activity, Nrf2 activation, and its related targets. Determination of antioxidant activities of A) glutathione reductase, B) glutathione peroxidase, C) catalase, D) total superoxide dismutase (SOD), and E) superoxide dismutase 2 (SOD2). F) Quantification of malondialdehyde (MDA). G) Representative immunoblots and densitometric analysis of H) nuclear factor erythroid 2-related factor 2 (Nrf2), and I) its phosphorylated form (pNrf2), as well as, the J) pNrf2/Nrf2 ratio. Representatives of immunoblot and densitometric analysis of K) gamma glutamylcysteine synthetase (yGCS), and L) SOD2. Data are mean (n = 7–8) ± SEM and were analyzed by using one-way ANOVA followed by the Dunnett test. *p < 0.05; **p < 0.01; ***p < 0.001.
3.2. Fluoride shifts mitochondrial dynamics and enhances biogenesis
To determine whether fluoride exposure affects mitochondrial dynamics, we examined key markers of fission and fusion. Regarding fission, both groups exposed to fluoride significantly increased the DRP1 protein levels relative to the CTRL group (Fig. 3A). Concerning fusion, mitofusin 2 (MFN2) protein levels only increased in the F5.0 group compared to the CTRL group (Fig. 3B).
Fig. 3. Effects of fluoride on mitochondrial dynamics and biogenesis. Representative immunoblots and densitometric analysis of A) dynamin-related protein 1 (DRP1), B) mitofusin 2 (MFN2), C) peroxisome proliferator-activated receptor coactivator 1-alpha (PGC-1a), D) nuclear respiratory factor 1 (NRF1), E) mitochondrial transcription factor A (TFAM), and F) voltage-dependent anion channel (VDAC). Data are mean (n = 6–7) ± SEM and were analyzed by using one-way ANOVA followed by the Dunnett test. * p < 0.05; **p < 0.01.
3.3. Fluoride promotes mitophagy activation
Beyond alterations in its structure and synthesis, mitochondria can also undergo degradation through a process known as mitophagy. So, we assessed markers of autophagic flux and mitophagy to examine the potential effects of fluoride. The levels of Beclin-1 (Fig. 4A), pPINK1 (Fig. 4C), LC3-I (Fig. 4D), and LC3-II (Fig. 4D and E) increased significantly in the F5.0 group compared to the CTRL group. Furthermore, the F2.5 group also showed a considerable increase in LC3-I levels. On the other hand, fluoride exposure did not affect the ubiquitin-binding protein p62 (p62) levels (Fig. 4B) or the LC3-II/LC3-I ratio (Fig. 4F).
Fig. 4. Effects of fluoride on autophagy-mitophagy induction. Representative immunoblots and densitometric analysis of A) Beclin-1, B) ubiquitin-binding protein p62 (p62), C) phosphorylated PTEN-induced putative kinase 1 (pPINK1), D) microtubule-associated protein 1 light chain 3 (LC3-I), E) LC3-phosphatidylethanolamine conjugate (LC3-II), as well as, F) the LC3-II/LC3-I ratio. Data are mean (n = 6–7) ± SEM and were analyzed by using one-way ANOVA followed by the Dunnett test. * p < 0.05; **p < 0.01; ***p < 0.001.
3.4. Fluoride alters the mitochondrial ultrastructure
The observed alterations in mitochondrial dynamics necessitated a detailed evaluation of the mitochondrial ultrastructure in tubular epithelial cells by electron microscopy. The CTRL group predominantly displayed oval mitochondria, with only a tiny population of bean-shaped mitochondria. Additionally, the mitochondria in this group exhibited diffuse cristae (Fig. 5A). By contrast, the groups exposed to fluoride showed a markedly higher quantity of mitochondria than the CTRL group. Furthermore, these mitochondria displayed several morphologic changes, including both bean-shaped and elongated forms. In addition, the cristae of mitochondria are shown in greater detail in both groups (Fig. 5B and C). Curiously, some fields within the fluoride-exposed groups revealed the presence of donut-shaped mitochondria and lysosomes (Supplementary Fig. 1).
Fig. 5. Effects of fluoride on mitochondrial ultrastructure. Representative micrographs of A) Control, B) F2.5, and C) F5.0 groups. BB, Border brush; N, nucleus; (*) mitochondria. Quantitative analysis of the D) mitochondrial size, E) aspect ratio (AR), and F) relative mitochondrial area/field area. Data are mean ± SEM and were analyzed by using one-way ANOVA followed by the Dunnett test. **p < 0.01; ***p < 0.001; ****p < 0.0001.
3.5. Fluoride modifies the induction of apoptosis
The imbalance of mitochondrial homeostasis may lead to cell death. Given the close relationship between mitochondrial alterations and the mitochondrial apoptosis pathway, we initially assessed the levels of pro-apoptotic and anti-apoptotic factors (Fig. 6A). Our findings indicate that both groups exposed to fluoride significantly increased Bcl-2-associated X protein (Bax) levels (Fig. 6B). By contrast, B-cell lymphoma (Bcl-2) levels only increased in the F5.0 group (Fig. 6C). Furthermore, the Bax/Bcl-2 ratio exhibited a significant increase solely in the F2.5 group compared to the CTRL group (Fig. 6D).
Fig. 6. Effects of fluoride on cell apoptosis. A) Representative immunoblots and densitometric analysis of B) Bcl-2-associated X protein (Bax), C) B-cell lymphoma 2 (Bcl-2), as well as, D) Bax/Bcl-2 ratio. E) Representative micrographs of cell apoptosis detection in the control, F2.5, and F5.0 groups, using TUNEL staining. F) Caspase-3 activity. Relative fluorescence units (RFU). Data are mean (n = 6–7 and 5–6, for the densitometric analysis and Caspase-3 activity, respectively) ± SEM and were analyzed by using one-way ANOVA followed by the Dunnett test. * p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.
4. Discussion
So far, it is known that fluoride exposure during gestation has an adverse effect on kidney development. Since oxidative stress, mitochondrial dysfunction, and apoptosis contribute to impairing kidney development in other models (Ho et al., 2023, Stewart et al., 2019), we investigated these potential mechanisms in fluoride-induced kidney alterations during the gestation stage.
Unlike what happens in the postnatal life, where it has been described that fluoride exposure reduces the activity of CAT, GPx, SOD, and GRx, GSH levels, and the Nrf2 activation (Owumi et al., 2024, Sharma et al., 2023), the response during gestation was partially different. Firstly, based on the abundance of SOD isoforms during this stage (Hayashibe et al., 1990), as well as the negative impact of fluoride on SOD1 mRNA expression and its cofactors (Cu, Zn) (Bouaziz et al., 2007, Luo et al., 2017), it can be inferred that SOD1 may be more susceptible than SOD2 to fluoride exposure during gestation. Concerning the activities of GPx and CAT, these enzymes operate in concert (Lushchak, 2012), which could explain the changes in GPx activity in response to the fall in CAT activity. Meanwhile, the fact that fluoride did not alter the GRx activity could be linked to its activity at the end of gestation, which significantly surpasses that seen in adult life (Vlessis and Mela-Riker, 1989).
Overall, these changes can compromise the ability to neutralize the ROS production. Consequently, it can favor cellular damage, such as lipid peroxidation. Unexpectedly, MDA levels decreased in the F2.5 group, possibly due to apparent Nrf2 activation. Although we did not confirm its activation by quantifying Nrf2 nuclear levels, it is well-known that Nrf2 nuclear translocation requires the phosphorylation of S40 (Huang et al., 2002), which was positively identified in our research. Additionally, based on the results on yGCS levels, a Nrf2 target, we dismiss the notion that MDA changes are associated with GSH novo synthesis. Conversely, the potential involvement of the Nrf2/heme oxygenase 1 (HO-1)/ferritin pathway cannot be ruled out, since HO-1 contributes to maintaining intracellular homeostasis by promoting ferritin expression (Kajarabille and Latunde-Dada, 2019), thereby limiting free Fe²+ availability and preventing hydroxyl radical formation and subsequent MDA production.
Beyond toxicity, ROS also regulates transcription factors involved in proliferation, differentiation, migration, and apoptosis (Spinelli and Haigis, 2018). Therefore, we cannot rule out the possibility that fluoride-induced redox status imbalance may be involved in the changes in kidney development.
Regarding mitochondrial dynamics, both morphometric changes and modifications in MFN2 and DRP1 partially support the proposal by Meyer et al. indicating that exposure to low-to-moderate levels of stressors enhances mitochondrial fusion and fission, thereby resulting in a greater interconnection of mitochondria and augmented turnover of damaged components (Meyer et al., 2017).
Similarly, recent research shows that enhancing fission mitigates fluoride’s harmful effects by elevating mitophagy and inhibiting apoptosis (Zhao et al., 2019). This fragmentation is crucial for initiating mitophagy because autophagosomes cannot engulf mitochondrial fragments longer than a specific size (Cho and Sun, 2020). Additionally, we also detected donut-shaped mitochondria, which help them resist mitophagy, and they can even recover their original shape when the metabolic stress disappears (Liu and Hajnóczky, 2011, Zhou et al., 2020).
Autophagy is acknowledged as an adaptive mechanism that protects cells against various stressors. However, when the stress level exceeds a critical threshold, it may trigger the activation of apoptosis (Kroemer et al., 2010). Monitoring autophagic flux requires assessing proteins at different stages, including Beclin-1 (initiating autophagosome formation), LC3-II (facilitating the extension and closure of the autophagosome), and p62 (a polyubiquitin-binding protein that integrates into the autophagosome) (Tian et al., 2020). Our results showed that all autophagic flux markers increased, except for p62. This response is consistent with observations in the kidneys of ICR mouse pups exposed to fluoride in drinking water before and during gestation, supporting the activation of autophagy (Guo et al., 2020). Conversely, another study found that fluoride exposure from gestation to puberty leads to inhibition of autophagy (Tian et al., 2020). Furthermore, although p62 levels remained unchanged after fluoride exposure, other autophagy adaptors may be involved (Zellner et al., 2021). In fact, p62 could remain unchanged due to Nrf2 activation, given that antioxidant response element sequences are present within the p62 promoter, thereby facilitating positive feedback (Silva-Islas and Maldonado, 2018). Ultimately, the changes in pPINK1 ensure the selective degradation of mitochondria, thereby preventing cellular damage (Palikaras et al., 2018).
Beyond its protective or harmful roles, autophagy also plays a role in cellular structure differentiation. Specifically, a study in C57BL/6 mice showed a correlation between the degree of podocyte differentiation and the expression of autophagy markers, while the inhibition of autophagy had a negative impact (Zhang et al., 2017). Therefore, activating autophagy may provide valuable insights into how fluoride exposure promotes podocyte maturation.
Our findings suggest that fluoride exposure promotes mitochondrial biogenesis despite the diminished TFAM levels. Meanwhile, TFAM was detected as a dimer in the fetal kidney. TFAM dimerization is necessary for the compactation of mtDNA into nucleoids (Ngo et al., 2014). Regardless of whether TFAM exists as a monomer or dimer, small changes in TFAM levels can significantly affect the available DNA for replication and transcription (Farge et al., 2014). Therefore, we hypothesize that a reduction in TFAM dimerization leads to the relaxation of mtDNA, facilitating its transcription.
Bcl-2 and Bax are well-established proteins that play a significant role in regulating apoptosis (Doke and Susztak, 2022). Although the increase in Bax levels is consistent with other reports, our finding regarding Bcl-2 levels contrasts with much of the existing literature (Song et al., 2017). These discrepancies may be attributed to the fact that these kidneys are still developing. In this context, Bcl-2 is expressed in various tissues during development, including the kidney, where it is detected in areas characterized by inductive interactions between epithelial and mesenchymal structures (LeBrun et al., 1993). In the same line, a study in mouse fetuses found that Bax and Bcl-2 are strongly expressed in mature renal structures, while their expression is weak in immature proximal and distal tubules, supporting that Bax and Bcl-2 have a role in kidney morphogenesis (Song et al., 2012). Conversely, the loss of Bcl-2 has a dramatic effect on kidney development, hindering differentiation and maturation (Sorenson, 2004). Interestingly, the changes in Bax and Bcl-2 during kidney development also coincide with changes in the expression of cytochrome c, establishing a relationship between them (Song et al., 2017a). Still, whether this relationship is linked to energetic requirements or ROS production as a signaling pathway is unknown.
Finally, beyond the induction of apoptosis by fluoride exposure, it is important to highlight that during development, apoptosis itself plays a crucial role in establishing tissue architecture (Woolf and Welham, 2002). So, it facilitates the remodeling of developing structures into their mature forms and removes excess cells, ensuring optimal tissue functionality (Carev et al., 2006). In this context, animal models have shown that both caspase activation and TUNEL staining increase toward the end of gestation and decrease during the postnatal period (Song et al., 2017a). By contrast, both markers associated with apoptosis induction diminished with the fluoride exposure, which may be counterproductive since caspase-3 inhibition or inactivation can harm kidney development, resulting in decreased nephron formation (Hayashi and Araki, 2002).
Our findings so far may suggest a potential shift in the metabolic profile, which is known to contribute to abnormal development and even kidney diseases (Cargill and Sims-Lucas, 2020). Furthermore, inhibiting glycolysis can promote nephrogenesis, leading to premature nephron differentiation (Diniz et al., 2023, Liu et al., 2017). Additionally, research utilizing organoids has shown that exposure to other environmental pollutants, such as microplastics, also involves metabolic modifications, reducing glycolysis while increasing activity in the tricarboxylic acid cycle, which subsequently affects development (Zhou et al., 2025).
Regarding the limitations of our study, evaluating mitochondrial bioenergetics, encompassing oxygen consumption, ATPase activity, ATP production, and membrane potential, would have been ideal; however, the limited sample availability for mitochondrial isolation made this impossible. Despite these limitations, the analysis of mitochondrial morphology and the levels of several mitochondrial biomarkers provides valuable insight into mitochondrial bioenergetics, given the close relationship among these components.
Taken together, our results suggest that fluoride exposure alters the redox status and mitochondrial homeostasis. At the same time, it decreased the apoptosis induction (Fig. 7). These changes may be related to early kidney maturation, as previously described by our research group. Our findings may provoke controversy regarding the safety of fluoride exposure during pregnancy; therefore, it is crucial to continue in-depth studies on the effects of fluoride in this specific condition.
Fig. 7. Integrative scheme. Several parameters within the fetal kidney were affected following fluoride exposure: (1) the activities of superoxide dismutase 1 (SOD1) and catalase (CAT) declined, whilst glutathione peroxidase (GPx) increased. Additionally, (2) total nuclear factor erythroid 2-related factor 2 (Nrf2) levels remained unchanged; however, its phosphorylated form (pNrf2) was significantly enhanced. Regarding mitochondrial dynamics, (3) mitofusin 2 showed a slight increase, whereas (4) dynamin-related protein 1 (DRP1) levels were markedly elevated. On the other hand, (5) proteins involved in biogenesis, such as peroxisome proliferator-activated receptor gamma coactivator 1-alpha (PGC-1a) and nuclear respiratory factor 1 (NRF1), increased following fluoride exposure; meanwhile, the mitochondrial transcription factor A (TFAM) dimer decreased, likely resulting in diminished mitochondrial DNA compaction. Similarly, (6) proteins associated with mitophagy, including beclin-1 (BCN1), microtubule-associated protein 1 light chain 3 (LC3), and phosphorylated PTEN-induced putative kinase 1 (pPINK1), also showed elevated levels. Ultimately, mitochondrial alterations affect apoptosis; thus, (7) Bax levels increased significantly, while Bcl-2 levels exhibited a slight elevation. Despite the changes in Bax, the caspase-3 activity decreased in those fetal kidneys exposed to fluoride.
CRediT authorship contribution statement
Esaú Montañez-Rodriguez: Writing – review & editing, Methodology, Investigation. Olivier Christophe Barbier: Writing – review & editing, Supervision, Resources, Project administration, Funding acquisition, Conceptualization. Sabino Hazael Avila-Rojas: Writing – original draft, Visualization, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. José Pedraza-Chaverri: Writing – review & editing, Resources, Methodology. Casimiro Gerarduzzi: Writing – review & editing. Ana Karen Pantaleón-Gómez: Writing – review & editing, Visualization, Methodology, Investigation. Iliana Angélica Tostado-Fernández: Writing – review & editing, Methodology. Juan Carlos León-Contreras: Writing – review & editing, Methodology, Investigation. Rogelio Hernández-Pando: Writing – review & editing, Methodology, Investigation. Omar Noel Medina-Campos: Writing – review & editing, Methodology. Brenda Marquina-Castillo: Writing – review & editing, Methodology.
Funding
SHAR is grateful for the postdoctoral scholarship from CONAHCyT (CVU: 487583). This work was partially supported by PAPIIT (DGAPA, UNAM) IN202725 and Federal Funds from Cinvestav (Presupuesto Federal).
Declaration of Competing Interest
The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Sabino Hazael AVILA ROJAS reports financial support was provided by Conahcyt. Jose PEDRAZZA CHAVERRI reports financial support was provided by National Autonomous University of Mexico. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
Acknowledgments
We sincerely appreciate the technical assistance of MVZ Rafael Muñoz Leyva, Rene Panfilo Morales, and Maria Antonieta Lopez Lopez for animal care.
Appendix A. Supplementary material
Supplementary material
Data availability
Data will be made available on request.
References
Aebi, 1984
H. Aebi
Catalase in vitro
Methods Enzym., 105 (1984), pp. 121-126, 10.1016/S0076-6879(84)05016-3
View PDFView articleGoogle ScholarAhmad et al., 2022
Environ. Sci. Adv. (2022), 10.1039/d1va00039j
Bhargava and Schnellmann, 2017
Nat. Rev. Nephrol., 13 (2017), pp. 629-646, 10.1038/NRNEPH.2017.107
View in ScopusGoogle ScholarBouaziz et al., 2007
Exp. Toxicol. Pathol., 58 (2007), pp. 339-349, 10.1016/J.ETP.2006.11.004
View PDFView articleView in ScopusGoogle ScholarCárdenas-González et al., 2013
Toxicol. Appl. Pharm., 272 (2013), pp. 888-894, 10.1016/J.TAAP.2013.07.026
View PDFView articleView in ScopusGoogle ScholarCarev et al., 2006
Pedia Nephrol., 21 (2006), pp. 627-636, 10.1007/S00467-006-0057-Y
View in ScopusGoogle ScholarCargill and Sims-Lucas, 2020
Pedia Nephrol., 35 (2020), 10.1007/S00467-018-4157-2
J. Biol. Chem., 250 (1975), pp. 5475-5480
View PDFView articleView in ScopusGoogle ScholarCho and Sun, 2020
Exp. Mol. Med., 52 (2020), pp. 730-737, 10.1038/S12276-020-0434-9
View in ScopusGoogle ScholarDiniz et al., 2023
Nat. Commun., 14 (2023), 10.1038/S41467-023-43513-7
Trends Cell Biol., 32 (2022), pp. 841-853, 10.1016/J.TCB.2022.03.012
View PDFView articleView in ScopusGoogle ScholarEisner et al., 2018
Nat. Cell Biol., 20 (2018), pp. 755-765, 10.1038/S41556-018-0133-0
View in ScopusGoogle ScholarFarge et al., 2014
Cell Rep., 8 (2014), pp. 66-74, 10.1016/J.CELREP.2014.05.046
View PDFView articleView in ScopusGoogle ScholarGérard-Monnier et al., 1998
Chem. Res. Toxicol., 11 (1998), pp. 1176-1183, 10.1021/TX9701790
Chemosphere, 241 (2020), 10.1016/J.CHEMOSPHERE.2019.124861
J. Basic Clin. Physiol. Pharm., 26 (2015), pp. 237-251, 10.1515/JBCPP-2014-0065
View in ScopusGoogle ScholarHayashi and Araki, 2002
Nephrol. Dial. Transpl., 17 (.9) (2002), pp. 8-10, 10.1093/NDT/17.SUPPL_9.8
View in ScopusGoogle ScholarHayashibe et al., 1990
Pedia Res., 27 (1990), pp. 472-475, 10.1203/00006450-199005000-00011
View in ScopusGoogle ScholarHo et al., 2023
Nutr. Metab., 20 (2023), 10.1186/S12986-023-00769-6
J. Biol. Chem., 277 (2002), pp. 42769-42774, 10.1074/JBC.M206911200
View PDFView articleView in ScopusGoogle ScholarJiménez-Córdova et al., 2018
Toxicol. Appl. Pharm., 352 (2018), pp. 97-106, 10.1016/J.TAAP.2018.05.027
View PDFView articleView in ScopusGoogle ScholarJiménez-Córdova et al., 2019
Environ. Res., 169 (2019), pp. 220-228, 10.1016/J.ENVRES.2018.10.028
View PDFView articleView in ScopusGoogle ScholarJohnston and Strobel, 2020
Arch. Toxicol., 94 (2020), pp. 1051-1069, 10.1007/S00204-020-02687-5
View in ScopusGoogle ScholarKajarabille and Latunde-Dada, 2019
Int J. Mol. Sci., 20 (2019), 10.3390/IJMS20194968
Ground Sustain Dev., 9 (2019), Article 100223, 10.1016/J.GSD.2019.100223
View PDFView articleView in ScopusGoogle ScholarKroemer et al., 2010
Mol. Cell, 40 (2010), pp. 280-293, 10.1016/J.MOLCEL.2010.09.023
View PDFView articleView in ScopusGoogle ScholarLawrence and Burk, 1976
Biochem Biophys. Res. Commun., 71 (1976), pp. 952-958, 10.1016/0006-291X(76)90747-6
View PDFView articleView in ScopusGoogle ScholarLeBrun et al., 1993
Am. J. Pathol., 142 (1993), pp. 743-753
Google ScholarLiu et al., 2017
J. Am. Soc. Nephrol., 28 (2017), pp. 3323-3335, 10.1681/ASN.2016111246/-/DCSUPPLEMENTAL
View in ScopusGoogle ScholarLiu and Hajnóczky, 2011
Cell Death Differ., 18 (2011), pp. 1561-1572, 10.1038/CDD.2011.13
View in ScopusGoogle ScholarLuo et al., 2017
Oncotarget, 8 (2017), pp. 50430-50446, 10.18632/ONCOTARGET.17365
View in ScopusGoogle ScholarLushchak, 2012
J. Amino Acids, 2012 (2012), pp. 1-26, 10.1155/2012/736837
Toxicology, 391 (2017), pp. 42-53, 10.1016/J.TOX.2017.07.019
View PDFView articleView in ScopusGoogle ScholarMontañez-Rodriguez et al., 2024
Environ. Toxicol. Pharm., 110 (2024), 10.1016/J.ETAP.2024.104545
Biochem J., 417 (2009), pp. 1-13, 10.1042/BJ20081386
View in ScopusGoogle ScholarNgo et al., 2014
Nat. Commun., 5 (2014), 10.1038/NCOMMS4077
Fluoride (2016), pp. 5-12
Google ScholarOwumi et al., 2024
Naunyn Schmiede Arch. Pharm., 397 (2024), 10.1007/S00210-024-03133-4
Nat. Cell Biol., 20 (2018), pp. 1013-1022, 10.1038/S41556-018-0176-2
View in ScopusGoogle ScholarPejznochova et al., 2010
Mitochondrion, 10 (2010), pp. 321-329, 10.1016/J.MITO.2010.01.006
View PDFView articleView in ScopusGoogle ScholarPloumi et al., 2017
FEBS J., 284 (2017), pp. 183-195, 10.1111/FEBS.13820
View in ScopusGoogle ScholarPrieur et al., 1998
Eur. J. Biochem, 252 (1998), pp. 194-199, 10.1046/J.1432-1327.1998.2520194.X
View in ScopusGoogle ScholarPrieur et al., 1995
Biochem J., 305 (Pt 2) (1995), pp. 675-680, 10.1042/BJ3050675
View in ScopusGoogle ScholarRubalcava-Gracia et al., 2023
Mol. Cell, 83 (2023), pp. 832-842, 10.1016/J.MOLCEL.2022.09.010
View PDFView articleView in ScopusGoogle ScholarSharma et al., 2023
Biol. Trace Elem. Res, 201 (2023), pp. 229-241, 10.1007/S12011-022-03113-W
View in ScopusGoogle ScholarSilva-Islas and Maldonado, 2018
Pharm. Res, 134 (2018), pp. 92-99, 10.1016/J.PHRS.2018.06.013
View PDFView articleView in ScopusGoogle ScholarSong et al., 2017
Sci. Rep., 7 (2017), 10.1038/S41598-017-00796-3
PLoS One, 7 (2012), 10.1371/JOURNAL.PONE.0032771
Nephron, 135 (2017), pp. 77-86, 10.1159/000450585
View in ScopusGoogle ScholarSorenson, 2004
Biochim Biophys. Acta Mol. Cell Res, 1644 (2004), pp. 169-177, 10.1016/j.bbamcr.2003.08.010
View PDFView articleView in ScopusGoogle ScholarSpinelli and Haigis, 2018
Nat. Cell Biol., 20 (2018), pp. 745-754, 10.1038/S41556-018-0124-1
View in ScopusGoogle ScholarStewart et al., 2019
Fetal Pedia Pathol., 38 (2019), pp. 185-194, 10.1080/15513815.2018.1564160
View in ScopusGoogle ScholarTang et al., 2021
Nat. Rev. Nephrol., 17 (2021), pp. 299-318, 10.1038/S41581-020-00369-0
View in ScopusGoogle ScholarTian et al., 2020
Arch. Toxicol., 94 (2020), pp. 749-760, 10.1007/S00204-019-02651-Y
View in ScopusGoogle ScholarVlessis and Mela-Riker, 1989
Pedia Res, 26 (1989), pp. 220-226, 10.1203/00006450-198909000-00013
View in ScopusGoogle ScholarWeydert and Cullen, 2010
Nat. Protoc., 5 (2010), pp. 51-66, 10.1038/NPROT.2009.197
View in ScopusGoogle ScholarWoolf and Welham, 2002
Nephrol. Dial. Transpl., 17 (9) (2002), pp. 2-4, 10.1093/NDT/17.SUPPL_9.2
View in ScopusGoogle ScholarXian and Liou, 2021
Cell Death Differ., 28 (2021), pp. 827-842, 10.1038/S41418-020-00657-Z
View in ScopusGoogle ScholarZellner et al., 2021
Mol. Cell, 81 (2021), pp. 1337-1354.e8, 10.1016/J.MOLCEL.2021.01.009
View PDFView articleView in ScopusGoogle ScholarZhang et al., 2017
J. Cell Mol. Med, 21 (2017), pp. 1315-1328, 10.1111/JCMM.13061
View in ScopusGoogle ScholarZhao et al., 2019
Arch. Toxicol., 93 (2019), pp. 709-726, 10.1007/S00204-019-02390-0
View in ScopusGoogle ScholarZhao et al., 2020
Theranostics, 10 (2020), pp. 4822-4838, 10.7150/THNO.42387
View in ScopusGoogle ScholarZhou et al., 2020
Autophagy, 16 (2020), pp. 562-574, 10.1080/15548627.2019.1634944
View in ScopusGoogle ScholarZhou et al., 2019
Toxicol. Appl. Pharm., 378 (2019), 10.1016/J.TAAP.2019.114608
J. Hazard Mater., 486 (2025), 10.1016/J.JHAZMAT.2024.137002
View at publisherGoogle Scholar
Life Sci., 198 (2018), pp. 18-24, 10.1016/J.LFS.2018.02.001
View PDFView articleView in ScopusGoogle Scholar