Introduction
Manganese (Mn) is a trace element incorporated in several metabolic pathways and in the structures of some vital enzymes (1, 2). However, it has also been found that Mn overexposure is associated with several deleterious adverse effects, such as neurotoxicity (1, 3–5). Renal injury and disturbances of serum electrolytes are the other adverse effects associated with Mn overexposure (6–8). There is no precise mechanism for Mn-induced nephrotoxicity.
The mechanism of Mn neurotoxicity is widely investigated (9). It has been found that Mn-induced oxidative stress plays a central role in the adverse effects of this metal on the nervous system (9–12). It has been reported that Mn accumulates in cellular mitochondria through calcium (Ca2+) channels (2). Hence, cellular mitochondria are critical targets for Mn cytotoxicity. Mitochondrial depolarization, mitochondria swelling, increased mitochondria-mediated reactive oxygen species (ROS) formation, and mitochondria-mediated cell death have been reported in different experimental models that investigated Mn neurotoxicity (9–13).
Renal tissue contains numerous mitochondria, the proper functioning of which guarantee appropriate energy (ATP) level required for the reabsorption process of chemicals (14–16). The mechanism of nephrotoxicity induced by several xenobiotics relies on mitochondrial impairment and mitochondria-mediated cell death (17). Oxidative stress and mitochondrial injury are two mechanistically related events (18). Hence, mitochondrial impairment could deteriorate oxidative stress and vice versa.
As already mentioned, there is no precise mechanism for Mn-induced renal injury. The current study was designed to evaluate the role of oxidative stress and mitochondrial impairment in the pathogenesis of Mn nephrotoxicity. Rats were exposed to Mn for 30 consecutive days. Several biomarkers in serum and urine, as well as histopathological alterations and oxidative stress markers in renal tissue, were evaluated. Moreover, kidney tissue mitochondria were isolated and assessed.
Materials and Methods
Reagents
3-[4,5dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT), iodoacetic acid, potassium hydroxide, bovine serum albumin (BSA), methanol high performance liquid chromatography (HPLC) grade, 3-(N-morpholino) propane sulfonic acid (MOPS), 4,2Hydroxyethyl,1-piperazineethanesulfonic acid (HEPES), dimethyl sulfoxide (DMSO), thiobarbituric acid (TBA), glacial acetic acid, glutathione (GSH), malondialdehyde (MDA), 2′,7′ Dichlorofluorescein diacetate (DCFH-DA), acetonitrile HPLC grade, ethylene glycol-bis (2-aminoethyl ether)-N, N, N′, N′-tetraacetic acid (EGTA), sucrose, dithiothreitol (DTT), sodium chloride, and Rhodamine 123 were obtained from Sigma Chemical Co. (St. Louis, MO, USA). Manganese chloride (MnCl2), trichloroacetic acid (TCA), and hydroxymethyl aminomethane hydrochloride (Tris-HCl) were purchased from Merck (Darmstadt, Germany).
Animals and treatments
Mature male Sprague–Dawley rats (200–250 g, live weight; n = 24) were obtained from Shiraz University of Medicine, Shiraz, Iran. Rats were maintained under standard conditions (22–24°C; 12:12 h, photo schedule; appropriate ventilation; and 40 ± 2% relative humidity). Animals had free access to tap water and a commercial rodents chow diet (RoyanFeed®, Isfahan, Iran). All procedures on experimental animals were performed in compliance with the ethical guidelines approved by the Shiraz University of Medical Sciences ethics committee (#95-01-36-11290). Animals were allotted to three groups (n = 8 in each group), and treated as follows: (i) Control (vehicle-treated); (ii) MnCl2 (20 mg/mL in drinking water); and (iii) MnCl2 (40 mg/mL in drinking water). Animals were treated for 42 consecutive days. On day 43, rats were anesthetized, and serum and kidney tissue samples were collected.
Sample collection
Animals were anesthetized (Thiopental 80 mg/kg, i.p). Blood was collected from the abdominal aorta and transferred to standard tubes (VACUSERA®, Serum gel, and clot activator tubes) for serum preparation. The kidney tissue was washed in ice-cooled (4°C) normal saline and used for further assessments. Kidney weight index (WI) was determined as WI = [wet weight of organ (g)/body weight (g)] × 100.
Reactive oxygen species
Renal tissue ROS levels were measured using dichlorofluorescein diacetate (DCFH-DA) as a fluorescent probe (19). Briefly, 10 μL of DCFH-DA (10 μM final concentration) was added to 990 μL of tissue homogenate (10% w: v in KCl buffer). Samples were incubated for 15 min at 37°C in the dark. Finally, the DCF fluorescence intensity was assessed using a fluorimeter (FLUOstar Omega®, BMG LABTECH, Germany; λexcit = 485 nm and λem = 525 nm) (19).
Renal tissue lipid peroxidation
Thiobarbituric acid reactive substances (TBARS) in renal tissue were measured as an index of lipid peroxidation in the renal tissue (12, 20). Briefly, renal tissue (500 mg) was homogenized in ice-cooled KCl buffer (1.15% w: v; 4°C). Then, 500 µL of tissue homogenate was added to a reaction mixture consisting of 1 mL TBA (0.375%, w: v), 1 mL trichloroacetic acid (20% w: v), and meta-phosphoric acid (3 mL f 1% w: v solution, pH = 2) (21). The mixture was incubated in a water bath (100°C) for 45 min (22). Then, 4 mL of n-butanol was added and vortexed (1 min). Finally, samples were centrifuged (17,000 × g for 10 min) and the absorbance of the developed color in upper phase (n-butanol) was measured at λ = 532 nm (EPOCH® plate reader, Highland Park, USA) (21).
Renal tissue and mitochondria glutathione content
Renal glutathione levels (oxidized and reduced; GSSG and GSH) were measured by the HPLC method (23). The HPLC system consisted of an NH2 column as the stationary phase (25 cm, Bischoff chromatography, Leonberg, Germany) (24). The mobile phases consisted of buffer A (Water: Methanol; 1: 4 v: v) and buffer B (Acetate buffer: Buffer A; 1: 4 v: v), and a gradient method with a steady increase of buffer B to 95% in 25 min (24). The flow rate of the mentioned mobile phase was 1 mL/min, and the UV detector (UV) detector was set at λ = 254 nm. Tissue samples were homogenized in Tris-HCl buffer (250 mM; pH = 7.4; 4°C), and 500 µL of TCA (50% w: v) was added. Isolated mitochondria (1 mL, 1 mg protein/mL) were also treated with 100 µL of TCA 50% w: v. Samples were mixed well, incubated on ice (10 min), and centrifuged (17,000 g, 30 min, 4°C). Afterward, 1 mL of the supernatant was collected in 5 mL tubes and 300 µL of the NaOH: NaHCO3 (2 M: 2 M) solution was added. Then, 100 µL of iodoacetic acid (1.5% w: v in deionized water) was added, and samples were incubated for 1 h (4°C, in the dark). Afterward, 2, 4-dinitrofluorobenzene (DNFB, 500 µL of 1.5% w: v in absolute ethanol) was added and mixed well. Samples were incubated in the dark (25°C, 24 h). Finally, samples were centrifuged (17,000 g, 30 min) and injected (25 µL) into the described HPLC system (23).
Protein carbonylation
The oxidative damage of kidney tissue proteins was assessed based on the reaction with dinitrophenyl hydrazine (DNPH) (25). Briefly, 1 mL of the tissue homogenate (10% w: v) was treated with 100 µL of the triton X-100 (0.1% w: v) and centrifuged (10 min, 700 g, 4°C). Then, 500 µL aliquots of the resulting supernatant were treated with 300 µL of DNPH (10 mM in HCl). Samples were then incubated for 1 h (25°C, vortexing every 10 min). Afterward, 100 µL of trichloroacetic acid (20% w: v) was added and centrifuged (12,000 g, 5 min) (26). The pellet was washed three times with ethanol: ethyl acetate (1 mL of 1: 1 v: v solution). Finally, the precipitate was re-dissolved in guanidine hydrochloride solution (600 µL of 6 M solution), and the absorbance at λ = 370 nm was measured (EPOCH® plate reader, Highland Park, USA) (25).
Ferric reducing antioxidant power
The ferric reducing antioxidant power (FRAP) of kidney tissue was measured (27). The FRAP assay measures change in the absorbance at λ = 593 nm due to the formation of a blue-colored ferrous (Fe2+)-4, 6-tripyridyl-s-triazine (TPTZ) complex from the colorless oxidized ferric form (Fe3+) by the action of tissue electron-donating antioxidants. The working FRAP solution was freshly prepared by mixing 25 mL of acetate buffer (300 mmol/L; pH = 3.6) with 2.5 mL of TPTZ ( 10 mmol/L in 40 mmol/L HCl) and 2.5 mL of ferric chloride (FeCl2, 20 mmol/L). Tissue samples (200 mg) were homogenized in 5 mL of 250 mM Tris-HCl buffer (pH = 7.4; 4°C). Afterward, 50 µL of tissue homogenate was added to 900 µL of the FRAP reagent and incubated in the dark (37°C, 5 min). The intensity of the resultant blue color was measured at λ = 593 nm using an EPOCH plate reader (BioTek® Instruments, Highland Park, US) (27).
Histopathological assessment
For histopathological evaluations, renal specimens were fixed in a buffered formalin solution prepared from NaH2PO4 (0.4% w: v), Na2HPO4 (0.64% w: v), and formaldehyde (10%) in double-distilled water (pH = 7.4). Paraffin-embedded tissue specimens were cut (5 µm) using a microtome and stained with hematoxylin and eosin. Kidney histopathological changes were evaluated using a light microscope (Olympus BX41; Olympus Optical Co. Ltd, Japan).
Renal and serum Mn levels
Serum and kidney Mn levels were measured using an inductively coupled plasma mass spectrometry (ICP-MS) method (28). Briefly, serum (500 µL) and kidney tissue samples (500 µL of 10% w: v tissue homogenate) were treated with 100 µL of nitric acid (HNO3; 2.5% w: v) and centrifuged (17,000 g, 30 min). The supernatants were collected and used for Mn determination (28).
Kidney mitochondria isolation
Rats’ kidneys were washed (NaCl 0.9% w: v, 4°C) and minced in the ice-cold isolation buffer containing 0.5 mM EGTA, 2 mM HEPES, 220 mM sucrose, 70 mM mannitol, and BSA (0.1% w: v) (pH = 7.4). Minced tissue was transported into fresh isolation buffer (5 mL buffer: 1 g tissue) and homogenized. Kidney mitochondria were isolated based on the differential centrifugation method (29). For this purpose, unbroken cells and nuclei were pelleted at the first round of centrifugation (1000 g for 10 min at 4°C). Afterward, the supernatant was centrifuged at 10,000 g (10 min at 4°C) to pellet the mitochondria fraction (brown-colored). The second centrifugation step was repeated at least thrice using a fresh buffer medium. Finally, mitochondrial pellets were resuspended in a buffer (5 mL buffer/g tissue) containing 70 mM mannitol, 220 mM sucrose, and 2 mM HEPES (pH = 7.4). The mitochondria fractions used to measure mitochondrial permeabilization and mitochondrial depolarization were suspended in mitochondria permeabilization buffer (65 mM KCl, 10 mM HEPES, 125 mM Sucrose, pH = 7.2) and depolarization assay buffer (220 mM Sucrose, 10 mM KCl, 68 mM Mannitol, 5 mM KH2PO4, 2 mM MgCl2, 50 μM EGTA, and 10 mM HEPES, pH = 7.2) (29). Sample protein concentrations were determined based on the Bradford method to standardize the obtained data.
Mitochondrial dehydrogenases activity
A colorimetric technique based on the production of purple formazan crystals from the 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide was used for the estimation of mitochondrial dehydrogenases activity (30). Briefly, a mitochondrial suspension (0.5 mg protein/mL) was incubated with 40 µL of MTT (0.4% w: v) and incubated in the dark (37°C, 30 min). Samples were centrifuged (15,000 g, 15 min), and the pellet was dissolved in dimethyl sulfoxide (DMSO; 1000 µL). The optical density (OD) at λ = 570 nm was measured with an EPOCH® plate reader (Highland Park, USA) (30).
Mitochondrial ATP levels
Based on a previously reported protocol, mitochondrial ATP level was assessed by HPLC (31, 32). Briefly, isolated mitochondria (1 mg protein/mL) were mixed with 100 µL ice-cooled meta-phosphoric acid solution (50% w: v, 4°C) and centrifuged (30 min, 17,000 g, 4°C). Afterward, the supernatant (100 µL) was treated with 15 µL of ice-cooled potassium hydroxide solution (KOH, 1 M). Samples were centrifuged (30 min, 17,000 g, 4°C) again, and 25 µL of the supernatant was injected into an HPLC system composed of an LC-18 column (µ-Bondapak, 25 cm). The mobile phase was composed of potassium hydrogen phosphate mono-basic (100 mM KH2PO4, pH = 7 adjusted with KOH), acetonitrile (2.5% v: v), and tetrabutylammonium hydroxide (1 mM). The flow rate was 1 mL/min, and the UV detector was set at λ = 254 nm (31).
Mitochondrial depolarization
Mitochondrial uptake of the cationic dye rhodamine 123 was applied for the evaluation of mitochondrial depolarization (33, 34). Rhodamine 123 accumulates in the mitochondrial matrix by facilitated diffusion. When the mitochondrion is depolarized, there is no facilitated diffusion, and the amount of rhodamine 123 in the supernatant will be increased (35). In the current investigation, the mitochondrial fractions (0.5 mg protein/mL; in the depolarization assay buffer) were incubated with 10 µM of rhodamine 123 (30 min, 37°C, in the dark). Afterward, samples were centrifuged (15,000 g, 10 min, 4°C) and the fluorescence intensity of the supernatant was monitored with a fluorimeter (FLUOstar Omega®; BMG, Germany; λexcit = 485 nm and λem = 525 nm) (33, 36).
Mitochondrial permeabilization and swelling
Mitochondrial swelling was estimated by analyzing the changes in optical density at λ = 540 nm (34, 37). Briefly, isolated mitochondria (0.5 mg protein/ml) were suspended in the mitochondria permeabilization buffer (65 mM KCl, 125 mM Sucrose, 10 mM HEPES, pH = 7.2), and the absorbance was monitored (30°C, during 30 min of incubation) using an EPOCH® plate reader (Highland Park, USA). An increase in mitochondrial swelling is associated with a decrease in absorbance. The results are reported as maximal mitochondrial swelling amplitude (ΔOD 540 nm) (37).
Lipid peroxidation in kidney mitochondria
TBARS were measured in isolated kidney mitochondria. Previous studies mentioned that sucrose interferes with the lipid peroxidation test in isolated mitochondria preparations (37). Therefore, mitochondria preparations were washed once (to remove sucrose) in ice-cooled MOPS-KCl buffer (100 mM KCl, 50 mM MOPS, 4°C, pH = 7.4). For this purpose, isolated kidney mitochondria were suspended in 5 mL of MOPS-KCl buffer and centrifuged (15,000 g, 4°C, 20 min). The pellet was re-suspended in MOPS-KCl buffer and used for TBARs assay. The mitochondrial suspension (1 mg protein/mL) was mixed with 1 mL of a solution containing trichloroacetic acid (15% w: v), HCl (0.24 N), TBA (0.375% w: v), and Trolox (500 µM). Samples were heated for 15 min at 100°C (37). Then n-butanol (1 mL) was added and vortexed (5 min). Samples were centrifuged (15,000 g, 10 min), and the absorbance of the n-butanol phase (upper phase) was measured (EPOCH® plate reader, Highland Park, USA, λ = 532 nm) (37).
Statistical methods
Data are represented as mean ± SD. Data analysis was accomplished by the one-way analysis of variance (ANOVA) and the Tukey’s multiple comparison test as the post hoc test. A P < 0.05 was considered as a statistically significant difference.
Results
Animal weight gain was significantly lower in the Mn-treated group (40 mg/mL) in comparison with control rats (Figure 1A). The kidney WI was also significantly lower in Mn 40 mg/mL group (Figure 1A). Serum and kidney tissue Mn levels were also significantly higher in Mn-treated animals (Figure 1B).
Figure 1. Effect of manganese treatment (20 and 40 mg/mL for 30 consecutive days) on animals’ weight gain, renal weight index (Panel A), and serum and kidney tissue manganese (Mn) (Panel B).
Data are shown as mean ± SD (n = 8).
#Indicates significantly different as compared with the control group (P < 0.05).
Significant deterioration in serum biochemical measurements indicates renal injury in Mn-treated rats (Table 1). Signs of hypophosphatemia were evident in the Mn group (Table 1). On the other hand, serum BUN and creatinine levels were significantly higher in Mn-exposed animals (20 and 40 mg/mL) (Table 1). Significant elevation in urine protein, alkaline phosphatase (ALP), γ-glutamyl transferase (γ-GT), and glucose level was also detected in Mn-treated rats (Table 2).
Table 1. Serum biochemical measurements in manganese (Mn)-treated rats.
| Control | Mn 20 mg/mL |
Mn 40 mg/mL | |
|---|---|---|---|
| Ca2+ (mg/dL) | 5.0 ± 0.20 | 4.9 ± 0.40 | 5.0 ± 0.50 |
| K+ (mmol/L) | 5.5 ± 0.40 | 4.6 ± 0.32 | 4.5 ± 0.50 |
| Na+ (mmol/L) | 84.0 ± 4.00 | 76 ± 3.00 | 71.0 ± 2.00 |
| Glucose (mg/dL) | 113.0 ± 9.00 | 105 ± 5.00 | 97.0 ± 11.00 |
| Phosphate (mg/dL) | 3.4 ± 0.40 | 2.33 ± 0.40 | 2.1 ± 0.11# |
| Total protein (mg/dL) | 6.8 ± 0.44 | 7.1 ± 0.9 | 6.9 ± 0.30 |
| Blood urea nitrogen (mg/dL) | 43.0 ± 2.00 | 66.0 ± 5.00# | 68 ± 4.00# |
| Creatinine (mg/dL) | 0.3 ± 0.02 | 0.6 ± 0.03# | 0.8 ± 0.03# |
Data are represented as mean ± SD (n = 8).
#Indicates significantly different as compared with the control group (P < 0.01).
Table 2. Urinalysis of manganese (Mn)-treated animals
| Control | Mn 20 mg/mL |
Mn 40 mg/mL | |
|---|---|---|---|
| Protein (mg/dL) | 0.4 ± 0.1 | 1.1 ± 0.10# | 1.2 ± 0.20# |
| ALP (U/L) | 2122 ± 115 | 2380 ± 193 | 2670 ± 390# |
| γ-GT (U/L) | 2569 ± 166 | 3134 ± 113 | 3897 ± 575# |
| Glucose (mg/dL) | 80.0 ± 4.00 | 96.0 ± 19.00 | 132.0 ± 3.00# |
Data are represented as mean ± SD (n = 8).
ALP: alkaline phosphatase; γ-GT: γ-glutamyl transferase.
#Indicates significantly different as compared with the control group (P < 0.01)
Significant ROS formation, lipid peroxidation, and protein carbonylation were detected in Mn groups (Figure 2). Moreover, kidney tissue GSH was depleted, and the GSSG level was significantly increased in Mn-exposed animals (Figure 2). Tissue antioxidant capacity was also dose-dependently decreased in the kidneys of Mn-treated rats (Figure 2).
Figure 2. Markers of oxidative stress in the kidney tissue of manganese-treated rats.
Data are given as mean ± SD (n = 8).
Asterisks indicate significantly different from the control group (* P < 0.05, *** P < 0.001).
Several mitochondrial indices were assessed in the kidney tissue of Mn-treated rats (Figure 3). It was found that Mn exposure significantly decreased mitochondrial dehydrogenases activity and ATP levels in a dose-dependent manner (Figure 3). Moreover, a significant increment of mitochondrial depolarization and swelling was detected in the kidney mitochondria of Mn-treated animals (Figure 3). Lipid peroxidation was also dose-dependently increased in the kidney mitochondria isolated from Mn-exposed animals (Figure 3).
Figure 3. Markers of mitochondrial impairment in the kidneys of manganese-treated animals.
Data are presented as mean ± SD (n = 8).
*** Significantly different from the control group (P < 0.001).
Significant interstitial inflammation and tubular atrophy were evident in the kidneys of Mn 20 and 40 mg/mL groups (Figure 4 and Table 3). Moreover, Mn 40 mg/mL caused renal tissue necrosis (Figure 4 and Table 3).
Figure 4. Kidney tissue histopathological alterations in manganese-exposed animals. Hematoxylin and eosin staining. The grades of histopathological changes are given in Table 3.
Table 3. Renal tissue histopathological alterations in manganese-exposed rats.
| Interstitial inflammation | Tubular atrophy | Necrosis | |
|---|---|---|---|
| Control | − | − | − |
| Manganese 20 mg/mL | + | + | + |
| Manganese 40 mg/mL | ++ | ++ | +++ |
+: Mild; ++: Moderate; and +++: Severe histopathological alterations.
Discussion
Mn is a trace element that plays a fundamental role in several metabolic pathways and enzyme structures (2). However, overexposure to this metal is associated with a wide range of adverse effects, including renal injury (6–8, 38). Acute tubular necrosis, proteinuria, oliguria, and significant elevation in serum creatinine levels have been reported in human cases of Mn-induced nephrotoxicity (6, 38–40). No precise mechanism for Mn-induced nephrotoxicity has been identified so far. In the current investigation, it was found that Mn caused significant oxidative stress as well as mitochondrial impairment in the kidney tissue. The results might help in the development of therapeutic options against renal failure and serum electrolyte imbalance observed in Mn-intoxicated patients (6, 38–41).
Neurotoxicity is a well-described adverse effect of Mn (9, 13, 42–45). Oxidative stress and its consequences, such as disruption of biomembrane lipids and protein carbonylation, seem to play a fundamental role in Mn-induced toxicity in different organs such as the brain (9, 13, 42–45). Severe elevation in brain tissue ROS level and lipid peroxidation has been documented in Mn-induced neurotoxicity (9, 13, 42–45). Moreover, it has been found that brain tissue antioxidant systems are hampered upon Mn overexposure (9, 13, 42–45). The mechanism(s) of nephrotoxicity induced by Mn is less understood. Previous studies mentioned the occurrence of oxidative stress in the renal tissue of Mn-exposed animals (7). Mn-induced oxidative stress could affect several cellular targets, including biomembrane lipids, proteins, as well as deoxyribonucleic acid (7). In the current study, significant ROS formation, protein carbonylation, lipid peroxidation, and depletion of kidney tissue antioxidant capacity was evident in Mn-exposed rats. These results are consistent with previous investigations indicating Mn-induced oxidative stress in the kidney (7). Moreover, we found that kidney mitochondria could also be affected by Mn overexposure.
Cellular mitochondria are critical targets affected by Mn. (13). It has been found that Mn is accumulated in the mitochondrial matrix through Ca2+ channels (13). Induction of mitochondrial permeabilization, enhancement of mitochondria-facilitated ROS formation, a decrease of cellular ATP levels, and mitochondria-mediated cell death and apoptosis are associated with Mn-induced mitochondrial impairment (13).
Mitochondria play a fundamental role in kidney tissue (14–16). The reabsorption of chemicals (e.g., amino acids, glucose, and minerals) from nephrons to the bloodstream is an energy-dependent activity (14–16). Kidney tissue contains numerous mitochondria, the proper functioning of which guarantee enough ATP required for the reabsorption process of chemicals (14–16). Hence, Mn-induced mitochondrial impairment leads to an energy crisis and defect in the reabsorption of many chemicals in the renal tubules. Consequently, serum electrolyte disturbances could occur. Cellular mitochondria are also important sites of ROS production (18). It has been repeatedly reported that xenobiotics-induced mitochondrial impairment could facilitate mitochondria-mediated ROS formation (18). Based on the data obtained from the current study, we might be able to speculate that Mn-induced mitotoxicity could serve as a major cause of oxidative stress in the renal tissue.
Conclusion
Collectively, our results indicate the fundamental role of oxidative stress and mitochondrial impairment in the pathogenesis of Mn-induced renal injury. Therefore, targeting cellular mitochondria might serve as a therapeutic point against Mn-induced nephrotoxicity.