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Abstract

Glyoxal is a physiological metabolite formed by lipid peroxidation, ascorbate autoxidation, oxidative degradation of glucose, and degradation of glycated proteins. Glyoxal has been linked to oxidative stress and can cause a number of cellular damages, including covalent modification of amino and thiol groups of proteins to form advanced glycation end products. However, the mechanism of glyoxal toxicity has not been fully understood. In this study, we have focused on glyoxal toxicity in isolated rat liver mitochondria. Isolated mitochondria (0.5 mg protein per milliliter) were prepared from the Wistar rat liver using differential centrifugation and incubated with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM) for 30 min. The activity of mitochondrial complex II was determined by measurement of MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) conversion. The mitochondrial membrane potential (MMP), lipid peroxidation (MDA), reactive oxygen species (ROS) formation, glutathione (GSH) content, and protein carbonylation were also assessed. After an incubation of isolated liver mitochondria with glyoxal, disrupted electron transport chain, increased mitochondrial ROS formation, lipid peroxidation, mitochondrial membrane damage, GSH oxidation, and protein carbonylation ensued as compared to the control group (p < 0.05). Glyoxal toxicity in isolated rat liver mitochondria was dose-dependent. In conclusion, glyoxal impaired the electron transport chain, which is the cause of increased ROS and MDA production, depletion of GSH, and disruption of MMP. Mitotoxicity of glyoxal might be related to the pathomechanisms involved in diabetes and its complications.

Keywords

Glyoxal toxicity mitochondria liver rat

Introduction

Currently, diabetes mellitus affects 1–2% of the worldwide population. Patients with diabetes have a higher tendency to develop long-term complications, including nephropathy, neuropathy, atherosclerosis, retinopathy, and cataracts.¹ In diabetes mellitus, abnormal glucose metabolism leads to glucose intolerance and also hyperglycemia. Hyperglycemia plays a critical role in the pathogenesis of upcoming complications, and diabetic patients with poor maintenance of blood glucose levels are predominantly at higher risk.²

Under hyperglycemic conditions, reducing sugars can be autoxidized to form reactive dicarbonyls including glyoxal and methylglyoxal (MG).^3
–5 These reactive dicarbonyls promote oxidative stress and can cause a number of cellular damages, including covalent modification of amino and thiol groups of proteins to form advanced glycation end products (AGEs).⁶ AGEs have been implicated in the pathogenesis of diabetic complications such as atherogenesis, nephropathy, and cataractogenesis.⁷

Glyoxal and its derivative, MG, are reactive oxoaldehydes that originate endogenously from pathways linked to various pathologies such as glucose autoxidation, DNA oxidation, and lipid peroxidation.⁸ Glyoxal can also be readily formed from the autoxidation of glyceraldehyde, glycoaldehyde, and hydroxypyruvate in the absence of enzymes. Additionally, cooking food at high temperatures also results in the formation of glyoxal.⁹ Glyoxal and MG react nonenzymatically with amino and thiol groups of biomolecules by the Maillard reaction.¹⁰ The resulting Schiff base undergoes rearrangement to form relatively stable ketoamines known as Amadori products. Glycated biomolecules then undergo progressive dehydration, cyclization, and oxidation reactions to form AGEs.

Reaction of proteins with reactive dicarbonyls can result in the inactivation of essential cellular proteins that can potentially lead to cytotoxicity.¹¹ Although glyoxal and MG have not been classified as carcinogens, they appear to be tumor promoters and have been shown to be direct mutagens in several cellular models. Previous studies showed that glyoxal cytotoxicity occurred following the loss of hepatocytes mitochondrial membrane potential (MMP), glutathione (GSH) oxidation, and reactive oxygen species (ROS) formation.⁵ Glycolytic substrates, for example, fructose, sorbitol and xylitol, or ROS scavengers inhibited glyoxal-induced cytotoxicity and prevented the MMP collapse, suggesting that mitochondrial toxicity and oxidative stress contribute to the cytotoxic mechanism. This mitotoxicity could primarily be the result of oxidative stress and excess ROS production, not the direct influence of glyoxals and the direct mitochondrial effect of glyoxal toxicity remains to be elucidated. So, in this study, we have focused on the toxic mechanisms of glyoxal in isolated rat liver mitochondria.

Materials and methods

Chemicals

Glyoxal (40%, w/v), dithiobis-2-nitrobenzoic acid (DTNB), reduced GSH, sucrose, 2′,7′-dichlorofluorescin diacetate (DCF-DA), rhodamine 123 (Rh 123), Coomassie blue G, ethylene glycol-bis (2-aminoethylether)-N,N,N′,N′-tetraacetic acid (EGTA), D-mannitol, tetraethoxypropane (TEP), MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide), dinitrophenylhydrazine (DNPH), thiobarbituric acid (TBA), bovine serum albumin (BSA), and 2-[4-(2-hydroxyethyl) piperazin-1-yl] ethanesulfonic acid (HEPES) were all purchased from Sigma Chemical Co. (St. Louis, Missouri, USA). All other chemicals used in the experiments were of analytical grade.

Animals

Male Wistar rats (200–250 g) were obtained from Ahvaz Jundishapur University of Medical Sciences. All rats were kept in polypropylene cages and given standard rat chow and drinking water ad libitum. The animals were maintained in a controlled condition of temperature (25 ± 2°C) with a 12-h light:12-h dark cycle. All experimental procedures were conducted according to the ethical standards and protocols approved by the Committee of Animal Experimentation of Ahvaz Jundishapur University of Medical Sciences (ethics approval number: 1393.11.17).

Mitochondrial preparation

Mitochondria were prepared from Wistar rat liver by differential centrifugation.¹² The liver was removed and minced with a small scissor in a cold mannitol solution containing 200 mM d-mannitol, 70 mM sucrose, 1 mM EGTA, 0.1% (w/v) BSA, 10 mM HEPES-potassium hydroxide, and pH 7.4 prepared freshly. The minced liver was gently homogenized in a glass homogenizer and then nuclei and broken cell debris were sedimented through centrifuging at 600 × g for 10 min at 4°C, and the supernatants were centrifuged at 10,000 × g for 15 min. The upper layer was carefully discarded. The pellet was washed by gentle suspension in the isolation buffer and centrifuged again at 10,000 × g for 10 min. Final mitochondrial pellets were suspended in the mannitol solution. Protein concentrations were determined through the Coomassie blue protein-binding method as explained by Bradford.¹³ Mitochondrial samples (0.5 mg protein per milliliter) were incubated with various concentrations of glyoxal for 30 min.

Complex II activity

Mitochondrial complex II activity is mediated by succinate dehydrogenase enzymes. The activity of mitochondrial complex II was assayed through the measurement of MTT reduction. Briefly, 1 mL of mitochondrial suspensions (0.5 mg protein per milliliter) was centrifuged at 10,000 × g for 1 min. Pellet was resuspended in 970 µL of isolation medium and 500 µL of 0.5-µM MTT and incubated at 37°C for 45 min. Formed purple formazan crystals were dissolved in 800 µL DMSO, and the absorbance at 570 nm was measured with a spectrophotometer (UV-1650 PC, Shimadzu, Japan).¹⁴

Mitochondrial ROS level assay

The mitochondrial ROS measurement was performed using the fluorescent probe DCF-DA. Briefly, isolated mitochondria (0.5 mg protein per milliliter) were incubated with 1.6 µM DCF-DA at 37°C for 10 min. The fluorescence was measured using Perkin Elmer LS-50B Luminescence fluorescence spectrophotometer (California, USA) at the excitation and emission wavelength of 500 and 520 nm, respectively.¹⁵

Lipid peroxidation assay

Lipid peroxidation (MDA) was determined using the method of Zhang et al.¹⁶ Mitochondrial suspensions (0.5 mg protein per milliliter) were incubated with 0.25 ml sulfuric acid (0.05 M) and 0.3 ml 0.2% TBA. The tubes were placed in a boiling water bath for 30 min. At the end, the tubes were moved to an ice-bath and 0.4 ml n-butanol was added to each tube. Then, they were centrifuged at 3500 × g for 10 min. The amount of MDA formed in each of the samples was assessed through measuring the absorbance of the supernatant at 532 nm using a spectrophotometer (UV-1650 PC). TEP was used as the standard and MDA content was expressed as nanomoles per milligram of protein.

MMP assay

The mitochondrial uptake of the cationic fluorescent dye, Rh 123, has been used for the determination of MMP. The mitochondrial suspensions (0.5 mg protein per milliliter) were incubated with 1.5 µM Rh 123 at 37°C for 10 min with gentle shaking. The fluorescence was measured using Perkin Elmer LS-50B Luminescence fluorescence spectrophotometer at the excitation and emission wavelength of 490 and 535 nm, respectively.¹⁷

Measurement of GSH content

GSH contents were determined using DTNB by spectrophotometric method in isolated mitochondria.¹⁸ Mitochondrial suspensions (0.5 mg protein/ml) were added into 0.04% DTNB in 0.1 mol/l of phosphate buffers (pH 7.4). The yellow color development was read at 412 nm using a spectrophotometer (UV-1650 PC). GSH content was expressed as micrograms per milligram of protein.

Determination of protein carbonyl content

The total protein-bound carbonyl content was measured by derivatizing the protein carbonyl adducts with DNPH. Briefly, an aliquot of the suspension of mitochondria (0.5 mg protein/ml) was added to an equivalent volume (0.5 mL) of 0.1% DNPH (w/v) in 2 N hydrogen chloride (HCl) and allowed to incubate for 1 h at room temperature. This reaction was terminated and the total protein precipitated by the addition of an equivalent of 1 mL volume of 20% TCA (w/v). Protein was pelleted by centrifugation at 10,000 r min⁻¹, and the supernatant was discarded. Excess unincorporated DNPH was extracted three times using an excess volume (0.5 mL) of ethanol:ethyl acetate (1:1) solution. Following extraction, the recovered cellular protein was dried under a stream of nitrogen and dissolved in 1 mL of tris-buffered 8.0 M guanidine–HCl, pH 7.2. The resulting solubilized hydrazones were measured at 370 nm. The concentration of 2,4-DNPH-derivatized protein carbonyls was determined using an extinction coefficient of 22,000 M⁻¹ cm⁻¹.¹⁹

Statistical analyses

Results were reported as mean ± SD. The statistical analyses were conducted using one-way analysis of variance by SPSS (version 20). The group differences were calculated by post hoc analysis using Tukey test.

Results

Effect of glyoxal on the mitochondrial complex II

The activity of succinate dehydrogenase (complex II) was determined using the MTT assay at different concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Figure 1 shows a significant concentration-dependent decrease in the mitochondrial metabolism of MTT to formazan (p < 0.05).

Figure 1.

Effect of glyoxal on succinate dehydrogenase (complex II) activity. Liver mitochondria (0.5 mg/mL) were incubated for 30 min with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Values are represented as mean ± SD (n = 6). ***p < 0.001: significant as compared with control.

Effect of glyoxal on the mitochondrial ROS production

As shown in Figure 2, following the glyoxal exposure in the concentrations of 2.5, 5, 7.5, and 10 mM, ROS generation was significantly increased in isolated rat liver mitochondria (p < 0.001).

Figure 2.

Effect of glyoxal on mitochondrial ROS formation. Liver mitochondria (0.5 mg/mL) were incubated for 30 min with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Values are represented as mean ± SD (n = 6). *p < 0.05; **p < 0.01: significant as compared with control.

Effect of glyoxal on the mitochondrial lipid peroxidation

MDA is the final product of lipid peroxidation that is often used as a marker of oxidative damage. As shown in Figure 3, in isolated rat liver mitochondria, MDA was markedly increased following the glyoxal exposure in the concentrations of 5, 7.5, and 10 mM (p < 0.001). However, low concentrations of glyoxal (1 and 2.5 mM) did not significantly increase MDA. The amounts of MDA formation in liver mitochondria were 3.3 ± 0.3, 3.5 ± 0.4, 4.1 ± 0.3, 4.3 ± 0.5, and 4.5 ± 0.6 nanomoles per milligram of protein at 1, 2.5, 5, 7.5, and 10mM glyoxal concentrations, respectively, whereas that of the control group was 2.8 ± 0.3 nanomoles per milligram of protein.

Figure 3.

Effect of glyoxal on mitochondrial lipid peroxidation. Liver mitochondria (0.5 mg/mL) were incubated for 30 min with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Values are represented as mean ± SD (n = 6). ***p < 0.001: significant as compared with control.

Effect of glyoxal on MMP

MMP is an electrochemical potential that consists of a transmembrane electrical potential and a proton gradient. MMP collapse is an early sign of mitochondrial dysfunction. The mitochondrial membrane damage was found to be significantly higher (p < 0.05) in the glyoxal-treated isolated mitochondria compared with control mitochondria (Figure 4).

Figure 4.

The effect of glyoxal on the mitochondrial membrane damage in liver mitochondria. Liver mitochondria (0.5 mg/mL) were incubated for 30 min with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Values are represented as mean ± SD (n = 6). *p < 0.05; **p < 0.01; and ***p < 0.001: significant as compared with control.

Effect of glyoxal on mitochondrial GSH content

The GSH concentration in liver mitochondria after a 30-min exposure to glyoxal was evaluated to determine the extent of oxidative stress induced by glyoxal. At 1, 2.5, 5, 7.5, and 10 mM glyoxal concentrations, GSH levels decreased to 55.8 ± 7.1, 40.8 ± 8.2, 33.4 ± 5.3, 29.5 ± 5.1, and 23.7 ± 4.5 (micrograms per milligram of protein), respectively, as compared to 65.37 ± 6.4 (micrograms per milligram of protein) for control mitochondria (Figure 5).

Figure 5.

Effect of glyoxal on mitochondrial GSH content. Liver mitochondria (0.5 mg/mL) were incubated for 30 min with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Values are represented as mean ± SD (n = 6). ***p < 0.001: significant as compared with control.

Effect of glyoxal on protein carbonylation

As shown in Figure 6, protein carbonylation after a 30-min exposure to glyoxal was significantly increased in isolated rat liver mitochondria (p < 0.01).

Figure 6.

Effect of glyoxal on protein carbonylation. Liver mitochondria (0.5 mg/mL) were incubated for 30 min with various concentrations of glyoxal (1, 2.5, 5, 7.5, and 10 mM). Values are represented as mean ± SD (n = 6). **p < 0.01; ***p < 0.001: significant as compared with control.

At 1, 2.5, 5, 7.5, and 10 mM glyoxal concentrations, total protein-bound carbonyl content increased to 26.7 ± 5.6, 44.6 ± 7.3, 49.0 ± 5.9, 57.5 ± 6.1, and 64.0 ± 11.1 (nanomoles per milligram of protein), respectively, that are comparable to 9.1 ± 3.6 (nanomoles per milligram of protein) for control mitochondria.

Discussion

Glyoxal generates via enzymatic and nonenzymatic degradation of glucose, lipid, and protein catabolism and also lipid peroxidation. In hyperglycemic condition in which oxidative stress and lipid peroxidation are common, glyoxal maybe over produced so that when combined with proteins form AGEs that are implicated in diabetic complications.²⁰ Interestingly, this oxoaldehyde was reported to form in early glycation from the degradation of glucose and Schiff’s base adduct and even a short period of hyperglycemia, as occurred in impaired glucose tolerance, may be sufficient to increase the concentrations of oxoaldehydes in vivo.²¹ Glyoxal amount is particularly important in the evaluation of the possible effect of oxidative stress.²² Currently, subclinical hyperglycemia and diabetes complications are among the most important worldwide public health problems. As a result, in this study, we mainly focused on and discussed the effect of glyoxal on mitochondria in the context of diabetes.

Hepatocytes of rats were previously reported to be protected from glyoxal cytotoxicity when polyphenols were used after the toxic insult was applied.²³ Many rat hepatocyte models were extensively used to evaluate the toxic effect of glyoxal and from them it was suggested that the glyoxal toxic effects upon rat hepatocytes were attributed to potential radical formation and ROS overproduction.²⁴ On the other hand, mitochondria are well known as the main sources for intracellular ROS production.^25,26 So, to elucidate the mechanism of glyoxal toxicity on rat hepatocytes, in this study, their mitochondria were isolated and evaluated under the toxic concentrations of glyoxal.

The range of glyoxal concentrations used in this study were much higher than those found under physiological (approximately 12.5 µg/ml) and diabetic (approximately 27.2 µg/ml) conditions.²² Other estimates of the concentration of glyoxal in human blood plasma were in the range of 100–120 nM and cellular concentrations of glyoxal 0.1–1 μM. However, insufficient control of interferences during analysis, estimates of 10–1000 fold higher than this may be easily determined.²⁷ All data obtained via in vitro models questioned how extrapolatable they are to the normal physiological condition. In toxicology and for demonstrating a toxic outcome during a short period (acute toxicity) in in vitro and in vivo models, usually higher concentrations than normal values are used to determine a toxic effect. For evaluating toxic insults on isolated mitochondria model, there is a limitation of short period of mitochondrial maintenance and stability upon in vitro conditions (3–4 h). At the same time, we must be careful to the pattern of concentration–response curve which may, at lower concentrations of toxicant, achieve an inverse outcome (J shape). In this regard, MG when used at millimolar concentrations to demonstrate an impairment in insulin signaling is physiologically irrelevant.²⁷ Mitochondria-associated chronic diseases, particularly diabetes and cancer rates, are rising, and an efficient in vitro method for evaluation of mitochondrial maintenance and also its protection has already been investigated.^15,28 In this study, the main objective is to establish a rapid in vitro method for glyoxal mitotoxicity and then to evaluate mitochondrial protection against this toxicity.

Mitochondria are important sources of ROS generation besides being the powerhouse of cells that provides over 90% of Adenosine triphosphate (ATP) consumed by the body. Mitochondria also play an important role in both apoptotic and necrotic cell death. So, mitochondrial dysfunction not only leads to oxidative stress but also results in cell death and finally tissue damage.^29,30

Succinate dehydrogenase (succinate: ubiquinone oxidoreductase; mitochondrial complex II) has been reported to play a central role at high respiration rates; therefore, the activity of this enzyme has been considered as a good marker for the mitochondrial oxidative capacity.³¹ We investigated the effect of glyoxal in the activity of complex II of mitochondrial electron transport chain. The effect of glyoxal on succinate dehydrogenase activity was determined through the reduction of the MTT dye to formazan metabolite. Our results showed a significant reduction in function of complex II activity following glyoxal, which contributes to glyoxal toxicity. Previous studies showed that mitochondrial complexes I, II, and III are the main sources for ROS production in the respiratory chain.^15,32

The results of the present study indicated that mitochondrial ROS production was significantly induced by glyoxal in a concentration-dependent manner that, with regard to the mechanism described above, can be caused by impaired electron transport chain and mitochondrial membrane.

Incubation of mitochondrial suspension with glyoxal significantly increased lipid peroxidation, which is comparable to the previous studies in hepatocyte.¹¹ This suggests that mitochondrial lipids are early targets of oxygen free radicals, due to their high content of unsaturated fatty acids and their location in the inner mitochondrial membrane, near the site of ROS production. On the other hand, oxidation of lipid membrane results in disruption of mitochondrial membrane and consequent collapse of MMP and cytochrome c release.³³ Mitochondrial membrane is an electrochemical potential that consists of a transmembrane electrical potential and a proton gradient. In this study, glyoxal significantly induced MMP collapse.

Numerous enzymatic and nonenzymatic defense mechanisms such as GSH peroxidase and total-SH proteins have an important role in regulation of ROS effects. GSH is one of the primary nonenzymatic antioxidant systems against hydrogen peroxide and other ROS, of which nearly 10–15% of total cellular GSH is present in the mitochondria.³⁴ Therefore, depletion of reduced GSH contents in the mitochondria could cause severe deficiency in their defense system against oxidative damage, leading to a further lipid peroxidation. GSH depletion has markedly enhanced the sensitivity of the mitochondrial structure to the ROS-mediated damage.³⁴ In the current study, glyoxal also significantly decreased the GSH content as compared to the control group (p < 0.05). This effect was in a concentration-dependent manner. Previous studies also reported the GSH depletion in glyoxal-exposed isolated rat hepatocytes.^5,35

Glyoxal and MG are detoxified by the cytosolic glyoxalase I and glyoxalase II systems to form glycolate and d-lactate, respectively.^11,35 The rate of detoxification of glyoxal and MG by the cell is dependent on the cytosolic GSH concentration.³⁶ Under conditions of oxidative stress, the subsequent decrease in cellular GSH concentration impairs glyoxal and MG metabolism.¹¹

ROS-induced oxidation of protein lysine, arginine, histidine, threonine, proline, and cysteine amino acids to form reactive protein carbonyl functional groups was generally termed “protein carbonylation.” In addition to carbonyl stress, oxidative stress has also been implicated in the pathogenesis of diabetic complications.

Mitochondrial dysfunction can occur at the beginning of toxicity and has been implicated in the pathogenesis of various conditions such as cancer, diabetes, cardiovascular disease, and age-related neurodegenerative diseases and oxidative stress.³⁷ Understanding the function of mitochondria and their role in toxicity induced by various toxicants provides the important target for the treatment and prevention of related diseases. The distinctive nature of mitochondrial genome and its sensitivity to mutations, enzyme activation, and also detoxification of different toxicants and xenobiotics in mitochondria and its membrane swelling due to the diverse injuries have been evaluated in many studies.³¹

Mitochondrion not only plays a key role in generating energy through the process called oxidative phosphorylation (OXPHOS), it also plays a central role in apoptosis, cellular stress responses, and genetic diseases.³⁸ Disruption of OXPHOS leads to changes in the intracellular redox states and intracellular organelles, ATP production, the formation of ROS, and eventually leads to cell death, including apoptosis.

Mitochondria could be considered a target for environmental pollutants such as heavy metals. In this context, mitochondrial dysfunction leads to the oxidative stress, a noxious process that is believed to be involved in the development of many diseases. Glyoxal is one of the key targets for therapeutic intervention in pathological conditions involving carbonyl toxicity, including diabetes.

In conclusion, glyoxal impaired the electron transport chain in isolated rat liver mitochondria, which is the cause of the increased ROS production. Mitochondrial ROS production in turn contributes to the mitochondrial lipid peroxidation, depletion of GSH content, and MMP disruption. This study supports the role of mitochondria in the oxidative injury resulting from glyoxal exposure and implies a vicious cycle that is the ROS-induced ROS generation.

Footnotes

Authors’ note

This article is issued from the thesis of Mehdi Goudarzi.

Declaration of Conflicting Interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work was supported by a grant (grant no MPRC-069) from Deputy of Research, Ahvaz Jundishapur University of Medical Sciences, Ahvaz, Iran.

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