Experimental Diabetes ResearchVolume 2012 (2012), Article ID 430176, 8 pagesdoi:10.1155/2012/430176

Research Article

Hepatic Mitochondrial Alterations and Increased Oxidative Stress in Nutritional Diabetes-Prone Psammomys obesus Model

Saida Bouderba,¹ M. Nieves Sanz,² Carlos Sánchez-Martín,² M. Yehia El-Mir,² Gloria R. Villanueva,² Dominique Detaille,3,4 and E. Ahmed Koceïr¹

¹USTHB, FSB, Equipe de Bioénergétique et Métabolisme Intermédiaire, Alger 16111, Algeria²Departamento de Fisiología y Farmacología, Faculad de Farmacia, Universidad de Salamanca, 37007 Salamanca, Spain³Laboratoire de Bioénergétique Fondamentale et Appliquée (LBFA), Université Joseph Fourier, Grenoble 38041, France⁴INSERM U1055, Grenoble cedex 9, Grenoble 38041, France

Received 10 January 2012; Accepted 16 March 2012

Academic Editor: Pietro Galassetti

Copyright © 2012 Saida Bouderba et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Mitochondrial dysfunction is considered to be a pivotal component of insulin resistance and associated metabolic diseases. Psammomys obesus is a relevant model of nutritional diabetes since these adult animals exhibit a state of insulin resistance when fed a standard laboratory chow, hypercaloric for them as compared to their natural food. In this context, alterations in bioenergetics were studied. Using liver mitochondria isolated from these rats fed such a diet for 18 weeks, oxygen consumption rates, activities of respiratory complexes, and content in cytochromes were examined. Levels of malondialdehyde (MDA) and gluthatione (GSH) were measured in tissue homogenates. Diabetic Psammomys showed a serious liver deterioration (hepatic mass accretion, lipids accumulation), accompanied by an enhanced oxidative stress (MDA increased, GSH depleted). On the other hand, both ADP-dependent and uncoupled respirations greatly diminished below control values, and the respiratory flux to cytochrome oxydase was mildly lowered. Furthermore, an inhibition of complexes I and III together with an activation of complex II were found. With emergence of oxidative stress, possibly related to a defect in oxidative phosphorylation, some molecular adjustments could contribute to alleviate, at least in part, the deleterious outcomes of insulin resistance in this gerbil species.

1. Introduction

The pathophysiology of type 2 diabetes mellitus (T2DM) is varied and very complex but the association of T2DM with obesity and inactivity indicates a potentially pathogenic link between fuel homeostasis, emergence of insulin resistance, and disease progression. Given the central role for mitochondria in energy production, dysregulated mitochondrial function at the cellular level can impact whole-body metabolism. Three major players are believed to be involved in such a disordered context: hepatocytes, insulin-dependent tissues (skeletal muscle, fat), and β-cells. Evidence pointing to defects in mitochondrial oxidative capacity for all these cell types demonstrates that each of them contributes to glucose imbalance [1]. Nevertheless, the nature, origin, and extent of this dysfunctioning remain controversial. Reduced expression of oxidative phosphorylation genes was observed in muscle and adipose tissue of humans with T2DM [2], while other studies reported on increases in respiration intensity of liver mitochondria from Goto-Kakizaki (GK) rats, a rodent model of diabetes [3], or diabetic patients [4]. In any case, it is important to realize that mitochondrial abnormalities could add to hyperglycemia once the insulin resistance is in place and lead to worsening of the diabetic state.

Due to its strategic position, between the intestinal bed and the systemic circulation, the liver was regarded as buffer organ for the regulation of metabolic fluxes [5, 6]. As glucose and fatty acid metabolisms are largely dependent on mitochondria to generate energy in cells, any impairment in nutrients oxidation, together with a reduced mitochondrial content, could thereby establish a vicious cycle of metabolic alterations involved in the pathogenesis of T2DM, leading to an increased generation of free radicals. Oxidative stress is recognized to contribute to many pathological processes, and a number of works point to the role of hyperglycaemia in promoting overproduction of mitochondria-derived reactive oxygen species (ROS) [7]. On the other hand, oxidative stress might also result from diminution of the antioxidative capacity in plasma and within the cells of diabetic subjects. Indeed, and in spite of more uncertain data, various reports shown that liver concentrations of reduced glutathione (GSH) decreased in different rodents models of diabetes [8, 9].

Several rodents models have been used to investigate the pathogenesis of metabolic syndrome but they do not reflect the human disease sufficiently. Psammomys obesus, a desert gerbil species, is able to subsist on a halophilic plants-based hypocaloric diet. However, in the presence of a relatively high energy regimen such as the standard laboratory diet, this sand rat develops T2DM [10, 11]. Indeed, the potential toxicity of glucose from this exogenous source (high but normal if compared to Wistar rats) will be drastically amplified, leading to a rapid evolution of the disease. Interestingly, prevention of this hyperglycemic state together with an enhancement of hepatic insulin sensitivity was found in diabetes-prone Psammomys obesus after exercise training [12]. Through this peculiar behavior, this kind of rodent represents an appropriate biological tool to uncover the features of nutritional diabetes. In addition, a previous experiment with perifused hepatocytes of Psammomys reported oxygen consumption rates smaller than those measured from Wistar rats. As well, levels of ATP and ADP were markedly lower in these gerbils [13].

In the light of these above considerations, the aim of the present work was to monitor in parallel the mitochondrial functioning and oxidative damage in diabetes-prone Psammomys obesus, by measuring the respiration intensity related to electron transfer chain complexes activities as well as some oxidative stress parameters after 18 weeks of treatment.

2. Materials and Methods

2.1. Animals and Diet

The Algerian sand rats Psammomys obesus used for this investigation were housed in suitable cages under controlled temperature and light conditions. Adult animals of both sexes (80–100 g) were divided into two groups: the control group consuming their plants-(Salsola-foetida-) based natural food, with a low energy diet (20 kcal/day) but rich in water and minerals—and the group fed the standard laboratory diet of high caloric value (32.5 kcal/day). Food and water were supplied during 18 weeks. Each animal was monitored for body weight and blood glucose in order to select Psammomys having a glycaemia superior to 100 mg/dL. All experimental procedures were authorized by the Institutional Animal Care Committee.

2.2. Biochemical Analysis

Fasted Psammomys rats were killed by cervical dislocation at the end of treatments, without anesthesia to avoid any further stress, and blood samples were collected in EDTA tubes. Plasma glucose and lipids (triglycerides, total cholesterol, HDL-cholesterol, LDL-c), hepatic and renal function markers (alanine transaminase (ALT), aspartate transaminase (AST), creatinine, urea) were measured by a spectrophotometric method adapted on a Cobas Mira automatic analyser. Plasma insulin was determined by radioimmunoassay. Extraction of hepatic total lipids was carried out using a process earlier described [14], and they were gravimetrically measured.

2.3. Oxidative Stress Assessment

Plasma total antioxidant status (TAS) was analysed in blood samples, using a commercial kit (Randox Laboratories LTD, UK). The principle of this assay is based on the reaction of peroxydase and H₂O₂ with the substrate azinodiethyl-benzothiazoline sulfonic acid (ABTS) to produce a radical cation of stable blue-green colour which was detected at 600 nm. The antioxidant capacity was inversely proportional to this coloration intensity and was expressed as mmol/L.

Lipid peroxidation was estimated from liver homogenates by measuring levels of malondialdehyde (MDA) through the thiobarbituric acid reactive substances method [15].

Liver GSH content was assayed using a commercially available kit (Cayman Chemical Company), based on the reaction with the thiol-specific reagent dithionitrobenzoic acid (DTNB). In this procedure, the sulfhydryl group of GSH reacts with DTNB to form a product which is reduced by glutathione reductase for recycling GSH and producing more thionitro-benzoic acid (TNB). The rate of TNB production was directly proportional to GSH levels, which were expressed as μmoles/mg protein.

The protein content of hepatic samples was determined according to the Lowry method [16], using bovine serum albumin (BSA) solution as a standard.

2.4. Mitochondria Isolation

Liver mitochondria from both Psammomys groups were prepared according to a standard differential centrifugation procedure, with all steps carried out at 4°C. After killing the animals, livers were quickly excised, rinsed, and chopped into an isolation medium (250 mM sucrose, 20 mM Tris-HCl, 1 mM EGTA, pH 7.4). The homogenates were centrifuged at 800 g for 10 min to remove nuclei and cell debris. Mitochondria were obtained from the supernatant by spinning twice at 8000 g for 10 min, and the pellet was resuspended in 0.5 mL of isolation buffer, then kept on ice. After measuring protein concentrations as above described, final mitochondrial suspensions were used immediately for respiration or stored at –80°C until enzyme analysis.

2.5. Oxidative Phosphorylation Measurement

Mitochondrial respiration was recorded polarographically, using a sealed oxygraphy chamber equipped with a Clark oxygen electrode and magnetic stirring. Oxygen consumption rates (JO₂) were determined at 37°C in a respiration buffer (125 mM KCl, 20 mM Tris-HCl, 1 mM EGTA, 5 mM Pi) with FFA-BSA to avoid the presence of uncoupled mitochondria. The non-phosphorylating state 2 was initiated by the addition of either 5 mM glutamate/2.5 mM malate (G/M) or 5 mM succinate/0.5 mM malate (S/M) in presence of 1.25 μM rotenone. The phosphorylating state 3 was obtained after addition of 1 mM ADP while state 4 was measured with 1.25 μg/mL oligomycin, a specific inhibitor of ATP synthase. For uncoupled respiration, 75 μM of dinitrophenol (DNP) was added, while cytochrome c oxidase activity was indirectly evaluated with 1 mM TMPD/5 mM ascorbate. The efficiency of oxidative phosphorylation was then assessed by the state 3-to-state 4 ratio, also called respiratory control index (RCR).

2.6. Electron Transfer Chain Activity and Cytochromes Content

Activities of respiratory complexes were assayed via slight changes of the protocols described by Malgat et al., using liver mitochondrial particles resulting from freezing-thawing cycles [17]. Complex I was assayed as the rate of NADH oxidation at 340 nm in 50 mM KPi buffer containing 3.75 mg/mL BSA, 100 μM decylubiquinone, 100 μM NADH. Rotenone (10 μM) was specifically used to inhibit this complex whose real activity was deduced from difference between NADH oxidation without and with rotenone. Complex II was measured as the rate of 2,6-dichloro-indophenol (DCIP) reduction at 600 nm in 50 mM KPi buffer supplemented with 2.5 mg/mL BSA, 9.3 μM antimycin A, 5 μM rotenone, 100 μM DCIP, 30 mM succinate. Enzyme activity was calculated between the difference before and after addition of 50 μM decylubiquinone. Complex III was assayed by measuring the reduction of cytochrome c at 550 nm, with and without 9.1 μM antimycin A. Isolated mitochondria were incubated in 100 mM KPi medium with 1 mg/mL BSA, 50 μM EDTA, 1 mM KCN, 100 μM oxidized cytochrome c, and the reaction was started by addition of 105.6 μM decylubiquinol.

In parallel, the content in different cytochromes of the electron transfer chain was measured by dual-wavelength spectrophotometry, comparing the spectra of fully oxidized versus fully reduced cytochromes [18].

2.7. Statistical Analysis

All data were reported as mean ± SEM. Differences between both rat groups were determined by Student’s t-tests, with a P value of either <0.05, <0.01, or <0.001 considered as statistically significant.

3. Results

3.1. Long-Term Metabolic Effects of High Caloric Diet and Impact on the Liver Redox State

Psammomys rats fed a high caloric chow for 18 weeks developed a metabolic syndrome, with significant changes in their body weight (P < 0.05), glycemia (P < 0.01), and insulinemia (P < 0.001) as compared with those of control group (Table 1). Plasma lipids levels, in particular triglyceridemia and cholesterolemia, were also altered. Furthermore, diabetic Psammomys exhibited a severe liver deterioration, as evidenced by a substantial increase of transaminases activity (P < 0.05), the hepatic mass accretion together with tissue accumulation of triglyce-rides. A renal injury, characterized by a rise in uremia and creatininemia, was besides showed. On the other hand, such harmful conditions markedly decreased plasma antioxidant capacity (Figure 1), whereas index of lipid peroxidation simultaneously increased (Figure 1). In addition, a depletion in hepatic GSH was found (Figure 1), and the resulting oxidative stress status in the liver, as the GSH/GSSG ratio, was largely diminished by the hypercaloric diet (1.07 ± 0.82 versus 4.29 ± 0.69 for control animals, P < 0.01).

Table 1: Body weight and biochemical parameters of control (n = 10) and high caloric diet-fed Psammomys rats (n = 15). Each run was performed in duplicate. *P < 0.05, **P < 0.01, ***P < 0.001 versus control rat group (natural food).

Parameter
Control (18 weeks)
Diabetic (18 weeks)

Initial body wt (g)
87.8 ± 3.3
85 ± 3.9

Final body wt (g)
121.5 ± 4.1
138.5 ± 8.7*

Hepatic mass/body wt ratio (%)
3.4 ± 0.09
4.05 ± 0.12*

Glucose (mg/dL)
65.2 ± 0.03
267 ± 0.5**

Insulin (pmol/L)
166.5 ± 1.8
1439 ± 3.3***

Triglycerides (mg/dL)
74.6 ± 19
253 ± 45.4**

Cholesterol (mg/dL)
61 ± 4.2
145.7 ± 4.9**

HDL-C (mg/dL)
44.2 ± 3.3
48.5 ± 7.2

LDL-C (mg/dL)
17.8 ± 1.3
29.5 ± 3.2*

Hepatic total lipids (mg/100 g wet wt)
3720 ± 90
4900 ± 190*

Hepatic triglycerides (mg/100 g wet wt)
291 ± 19
654 ± 79**

Hepatic cholesterol (mg/100 g wet wt)
273 ± 11
322 ± 21*

Urea (mg/dL)
53.8 ± 4.6
73.6 ± 6.9*

Creatinine (mg/dL)
0.28 ± 0.05
0.45 ± 0.02*

ALT (U/L)
76 ± 8.4
136 ± 19**

AST (U/L)
86 ± 6.7
167 ± 25**

(a)

(b)

(c)

Figure 1: Oxidative stress in plasma and liver tissue. Total antioxidant defences or TAS (a), as well as the intrahepatic contents in MDA (b) and GSH (c), was, respectively, measured in fresh plasma and frozen liver homogenate of control or treated Psammomys. *P < 0.05 versus control group.

3.2. Oxygen Consumption

With mitochondria isolated from diabetic Psammomys livers, we noticed a net decline of the respiratory chain activity (Figure 2). Indeed, both basal state 2 and ADP-stimulated state 3 were significantly lower in mitochondria respiring on G/M (−24 and −31%, resp.) but barely decreased with S/M plus rotenone (−8 and −7.5%, resp.). The assessment of oligomycin-induced state 4 showed no modification whatever the substrates. These values led to a decreased RCR (state 3-to-state 4 ratio) for mitochondria only energized with G/M (Table 2). An inhibition of either DNP-uncoupled or TMPD/ascorbate-activated respirations (−25 and −19% resp.) was still observed under this particular condition, suggesting an alteration of some respiratory fluxes which could alter the oxidative phosphorylation machinery.

Table 2: Effect of dietary treatment on respiratory control ratios (RCR) of liver mitochondria from both Psammomys groups. *P < 0.05 versus control group.

Oxygen consumption rate (natoms O/min/mg protein)

Glutamate/Malate (GM)
Succinate/Malate (SM) + rotenone

State 3
State 4
RCR
State 3
State 4
RCR

Control
138.8 ± 9.9
14.8 ± 0.6
9.4 ± 0.9
143.1 ± 5.3
39.3 ± 3.0
3.7 ± 0.3

Diabetic
96.1 ± 7.3*
14.5 ± 0.6
6.6 ± 0.2*
133.0 ± 12.3
35.4 ± 3.5
3.8 ± 0.2

(a)

(b)

Figure 2: Mitochondrial respiration in Psammomys. Oxygen consumption rates (JO₂) were assayed on mitochondria freshly isolated from control (open bars) or treated Psammomys (black bars), in the presence of glutamate/malate (a) or succinate/malate with rotenone (b) as energizing substrates. Various respiratory states were next assessed following the addition of different drugs. *P < 0.05 versus control group.

3.3. Mitochondrial Complexes Activity

To assess whether the above respiration data were directly linked to some defects inside the electron transfer chain, enzyme activities of complexes I, II, III, combined with the level in different cytochromes, were measured in broken liver mitochondria. Complexes I and III were substantially decreased (−32 and −40% resp.) in organelles from diabetic animals as compared to control group, yet complex II unexpectedly increased by 42.4% (Figure 3). Interestingly, a smaller content in cytochrome aa3 was found in diabetic Psammomys liver mitochondria (Table 3), a result rather consistent with a reduced activity of cytochrome oxidase that we had above seen through the TMPD-dependent respiration.

Table 3: Effect of dietary treatment on levels in different cytochromes of liver mitochondria from both Psammomys groups.*P < 0.05 versus from control group.

Cytochromes content (pmols/mg protein)

a + a3
b
c1
c

Control
66.1 ± 3.1
184.2 ± 6.0
95.0 ± 9.2
63.2 ± 21.0

Diabetic
51.3 ± 2.8*
219.4 ± 12.0
79.8 ± 7.9
58.2 ± 7.2

Figure 3: Activities of respiratory complexes I, II, III in liver mitochondria freshly isolated from control (open bars) or treated Psammomys (black bars). *P < 0.05 versus control group.

4. Discussion

In this work, we have confirmed that Psammomys obesus is a reliable biological support for the study of metabolic disorders such as insulin resistance or T2DM, and whose the etiology is similar to its manifestation in humans. Our results are in accordance with studies using the Israeli Psammomys [19] or the Nile rat [20], even though the latter displayed less pronounced metabolic disturbances than Psammomys obesus after feeding a calorie-rich diet for 18 weeks. It may be underlined that the increment in hepatic mass-body weight ratio was positively correlated with hyperinsulinemia and tissue accumulation of lipids, meaning a profound liver injury similar to that reported by others [21]. A high proportion of soluble transaminases was also seen. Knowing that these enzymes are released when hepatocellular damage occurs [22] and, on the other side, that an incomplete oxidation due to chronic fuel excess can be linked to the inability for mitochondria to maintain sufficient ATP levels [23, 24], it seems that these deleterious metabolic defects are consistently associated with a drastic endogenous oxidative stress and subsequent mitochondrial dysfunction, mainly at the inner membrane activity level.

The current work studied oxidative phosphorylation capacity using liver mitochondria isolated from Psammomys obesus. Because there existed no or few relevant data about bioenergetics in these rats, it became quite difficult to compare our findings with those of recent literature. We, therefore, tried to discuss them at best with respect to other experimental models involving a mitochondrial dysfunction. Mitochondria from diabetes-prone Psammomys obesus showed a lower respiratory rate than that of control group: the decreases in both state 3 (with ADP) and uncoupled state indicate a loss of oxidative capacity. Similar results were obtained from Wistar rats fed a high-fat diet [25, 26], while no modifications or even higher respiration intensities were found with Zucker rats [27] or GK rats of 6 months age [3]. Cytochrome oxidase, last component of the respiratory chain, is well recognized as a controlling step of nonphosphorylating oxygen consumption [28]. Its activity, when indirectly determined in the presence of TMPD/ascorbate, was lower in diabetic Psammomys obesus compared to control animals, and this response was moreover accompanied by a significant loss in cytochrome aa3. Such an observation could reflect a smaller mitochondrial efficiency as evidenced by the reduction of RCR though this fact concerned more particularly the NAD-driven respiration (with glutamate/malate). These findings would be also partially explained by diet-induced changes in membrane lipids composition but this phenomenon merits to be further explored.

High glucose or free fatty acids flux or both impairs metabolic flexibility, which may enhance mitochondrial substrate supply and ROS production [7]. As the harmful effects of ROS on tissues are widely appreciated, the severity of diabetic state for Psammomys obesus is likely related to an exacerbated oxidative stress. An increase in lipoperoxidation together with a decline in TAS were reported. In line with these alterations, a decrease of GSH/GSSG ratio was clearly indicative of an impaired liver antioxidant system. Our data are in agreement with several studies using streptozotocin-treated rats [29, 30], rats fed a high fructose diet [31], or diabetic mice [32, 33]. In view of that an increased oxidative stress is also consistent with a slowing-down of oxidative phosphorylation, our bioenergetics results suggest the existence of multiple damaged sites along the electron transfer chain. Indeed, mitochondria from diabetic Psammomys revealed an inhibition of complex I and complex III, attributable to an excess in mitochondrial ROS as earlier suggested [34]. Another important finding of this work is that Psammomys fed a calorie-rich diet exhibited a net activation of complex II despite a nearly decrease of JO₂ in presence of succinate (FAD-linked substrate), but with comparable RCR between diabetic and control rats. In support of our data, Cunningham et al. observed decreases in all respiratory complexes, except complex II, using liver biopsies from patients with steatosis [35]. Otherwise, complexes II and IV activities were augmented in GK rats or streptozotocin-induced diabetic rats [36].

A large number of studies invoked the involvement of ROS in the pathogenesis of mitochondrial DNA-related disorders [37, 38], and lipid peroxidation products can damage the mitochondrial genome [39]. One may infer that an increasing oxidative stress linked to the diabetic state should lead to mitochondrial DNA damages, altering the function of complexes I and III. Taken into account that the proteins of complex II encoded by nuclear genes are probably spared, any improvement of its activity can alleviate the abnormality or loss of other respiratory parameters, and determine the overall oxidative capacity. Interestingly enough, another kind of adaptative response of mitochondrial metabolism to a high glucose milieu was revealed in pancreatic islets from diabetic Psammomys obesus [40]. In this regard, the desert gerbil model could set up metabolic and/or molecular adjustments to circumvent some of the deleterious outcomes of insulin resistance.

It is concluded that a high-caloric diet causes metabolic troubles at the hepatic level in the Psammomys obesus rat after an 18-week treatment, and these changes are likely associated with a vast mitochondrial dysfunctioning. The present work is in agreement with an altering expression of oxidative phosphorylation genes, possibly resulting from aggravated oxidative stress, which justifies the performance of further studies to identify some molecular processes responsible for that mitochondrial impairment, and to quantify their relative influence for the liver.

Conflict of Interests

The authors state no conflict of interests.

Acknowledgments

This paper was supported by the Algerian Agency for the Research & Development in Health (ANDRS; Grant no. 03/03/03/10/008) the Program of Interuniversity Cooperation (PCI; Grant no. A/026422/09) as well as by the National Research Program (PNR no. 208/ANDRS/2011 and PNR no. 41/ANDRS/2011) from the National Administration of Algerian Higher Education and Scientific Research (DG-RSDT). The authores would like to dedicate this paper to our friend and colleague Professor Yehia El-Mir, recently deceased, with all their love and gratitude.

References

J. A. Kim, Y. Wei, and J. R. Sowers, “Role of mitochondrial dysfunction in insulin resistance,” Circulation Research, vol. 102, no. 4, pp. 401–414, 2008.

M. E. Patti, A. J. Butte, S. Crunkhorn et al., “Coordinated reduction of genes of oxidative metabolism in humans with insulin resistance and diabetes: potential role of PGC1 and NRF1,” Proceedings of the National Academy of Sciences of the United States of America, vol. 100, no. 14, pp. 8466–8471, 2003.

F. M. L. Ferreira, C. M. Palmeira, M. J. Matos, R. Seiç, and M. S. Santos, “Decreased susceptibility to lipid peroxidation of Goto-Kakizaki rats: relationship to mitochondrial antioxidant capacity,” Life Sciences, vol. 65, no. 10, pp. 1013–1025, 1999.

T. Takamura, H. Misu, N. Matsuzawa-Nagata et al., “Obesity upregulates genes involved in oxidative phosphorylation in livers of diabetic patients,” Obesity, vol. 16, no. 12, pp. 2601–2609, 2008.

J. Radziuk and S. Pye, “The liver, glucose homeostasis, and insulin action in type 2 diabetes mellitus,” in Contempory Endocrinology: The Metabolic Syndrome: Epidemiology, Clinical Treatment and Underlying Mechanisms, B. C. Hansen and G. A. Bray, Eds., pp. 343–371, Humana Press, 2007.

G. A. Rutter, “Diabetes: the importance of the liver,” Current Biology, vol. 10, no. 20, pp. R736–R738, 2000.

M. Brownlee, “Biochemistry and molecular cell biology of diabetic complications,” Nature, vol. 414, no. 6865, pp. 813–820, 2001.

S. S. Melo, M. R. Arantes, M. S. Meirelles, A. A. Jordao Jr., and H. Vannucchi, “Lipid peroxidation in nicotinamide-deficient and nicotinamide-supplemented rats with streptozotocin-induced diabetes,” Acta Diabetologica, vol. 37, no. 1, pp. 33–39, 2000.

F. M. Rauscher, R. A. Sanders, and J. B. Watkins, “Effects of coenzyme Q10 treatment on antioxidant pathways in normal and streptozotocin-induced diabetic rats,” Journal of Biochemical and Molecular Toxicology, vol. 15, no. 1, pp. 41–46, 2001.

G. Marquié, J. Duhault, and B. Jacotot, “Diabetes mellitus in sand rats (Psammomys obesus). Metabolic pattern during development of the diabetic syndrome,” Diabetes, vol. 33, no. 5 I, pp. 438–443, 1984.

E. Ziv, E. Shafrir, R. Kalman, S. Galer, and H. Bar-On, “Changing pattern of prevalence of insulin resistance in Psammomys obesus, a model of nutritionally induced type 2 diabetes,” Metabolism, vol. 48, no. 12, pp. 1549–1554, 1999.

Y. Heled, Y. Shapiro, Y. Shani et al., “Physical exercise enhances hepatic insulin signaling and inhibits phosphoenolpyruvate carboxykinase activity in diabetes-prone Psammomys obesus,” Metabolism, vol. 53, no. 7, pp. 836–841, 2004.

E. A. Koceir, Y. Dahmani, and X. Leverve, “Low rate of glucose 6-phosphate hydrolysis in liver cells is a physiological feature of non-diabetic wild sand rats (Psammomys obesus),” Diabetes and Metabolism, vol. 29, no. 4, pp. 363–374, 2003.

J. Folch, M. Lees, and G. H. Sloane Stanley, “A simple method for the isolation and purification of total lipides from animal tissues,” The Journal of Biological Chemistry, vol. 226, no. 1, pp. 497–509, 1957.

R. O. Recknagel and E. A. Glende Jr., “Spectrophotometric detection of lipid conjugated dienes,” Methods in Enzymology, vol. 105, pp. 331–337, 1984.

O. H. Lowry, N. J. Rosebrough, A. L. Farr, and R. J. Randall, “Protein measurement with the Folin phenol reagent,” The Journal of Biological Chemistry, vol. 193, no. 1, pp. 265–275, 1951.

M. Malgat, T. Letellier, G. Durrieu, and J. P. Mazat, “Enzymatic and polarographic measurements of the respiratory chain complexes,” in Mitochondrial Diseases: Models and Methods, P. Lestienne, Ed., pp. 357–377, Springer, Berlin, Germany, 1999.

J. N. Williams Jr., “A method for the simultaneous quantitative estimation of cytochromes a, b, c1, and c in mitochondria,” Archives of Biochemistry and Biophysics, vol. 107, no. 3, pp. 537–543, 1964.

E. Shafrir and E. Ziv, “Cellular mechanism of nutritionally induced insulin resistance: the desert rodent Psammomys obesus and other animals in which insulin resistance leads to detrimental outcome,” Journal of Basic and Clinical Physiology and Pharmacology, vol. 9, no. 2–4, pp. 347–385, 1998.

F. Chaabo, A. Pronczuk, E. Maslova, and K. Hayes, “Nutritional correlates and dynamics of diabetes in the Nile rat (Arvicanthis niloticus): a novel model for diet-induced type 2 diabetes and the metabolic syndrome,” Nutrition & Metabolism, vol. 7, article 29, 2010.

M. Maislos, V. Medvedovskv, I. Sztarkier, A. Yaari, and E. Sikuler, “Psammomys obesus (sand rat), a new animal model of non-alcoholic fatty liver disease,” Diabetes Research and Clinical Practice, vol. 72, no. 1, pp. 1–5, 2006.

B. J. Song, K. H. Moon, N. U. Olsson, and N. Salem Jr., “Prevention of alcoholic fatty liver and mitochondrial dysfunction in the rat by long-chain polyunsaturated fatty acids,” Journal of Hepatology, vol. 49, no. 2, pp. 262–273, 2008.

B. Fromenty and D. Pessayre, “Inhibition of mitochondrial beta-oxidation as a mechanism of hepatotoxicity,” Pharmacology and Therapeutics, vol. 67, no. 1, pp. 101–154, 1995.

S. K. Mantena, D. P. Vaughn, K. K. Andringa et al., “High fat diet induces dysregulation of hepatic oxygen gradients and mitochondrial function in vivo,” Biochemical Journal, vol. 417, no. 1, pp. 183–193, 2009.

G. Vial, H. Dubouchaud, K. Couturier et al., “Effects of a high-fat diet on energy metabolism and ROS production in rat liver,” Journal of Hepatology, vol. 54, no. 2, pp. 348–356, 2011.

J. Ciapaite, S. J. L. Bakker, G. van Eikenhorst et al., “Functioning of oxidative phosphorylation in liver mitochondria of high-fat diet fed rats,” Biochimica et Biophysica Acta, vol. 1772, no. 3, pp. 307–316, 2007.

M. Flamment, M. Arvier, Y. Gallois et al., “Fatty liver and insulin resistance in obese Zucker rats: no role for mitochondrial dysfunction,” Biochimie, vol. 90, no. 9, pp. 1407–1413, 2008.

V. Nogueira, M. Rigoulet, M. A. Piquet, A. Devin, E. Fontaine, and X. M. Leverve, “Mitochondrial respiratory chain adjustment to cellular energy demand,” The Journal of Biological Chemistry, vol. 276, no. 49, pp. 46104–46110, 2001.

J. A. Herlein, B. D. Fink, Y. O'Malley, and W. I. Sivitz, “Superoxide and respiratory coupling in mitochondria of insulin-deficient diabetic rats,” Endocrinology, vol. 150, no. 1, pp. 46–55, 2009.

K. R. Shanmugam, K. Mallikarjuna, N. Kesireddy, C. H. Kuo, and K. S. Reddy, “Protective effect of dietary ginger on antioxidant enzymes and oxidative damage in experi-mental diabetic rat tissues,” Food and Chemical Toxicology, vol. 124, no. 4, pp. 1436–1442, 2011.

P. Rajasekar and C. V. Anuradha, “Effect of L-carnitine on skeletal muscle lipids and oxidative stress in rats fed high-fructose diet,” Experimental Diabetes Research, vol. 2007, Article ID 72741, 8 pages, 2007.

I. Garcia-Ruiz, C. Rodriguez-Juan, T. Diaz-Sanjuan, et al., “Uric acid and anti-TNF antibody improve mitochondrial dysfunction in ob/ob mice,” Hepatology, vol. 44, no. 3, pp. 581–591, 2006.

D. Du, Y. H. Shi, and G. W. Le, “Oxidative stress induced by high-glucose diet in liver of C57BL/6J mice and its underlying mechanism,” Molecular Biology Reports, vol. 37, no. 8, pp. 3833–3839, 2010.

T. A. Young, C. C. Cunningham, and S. M. Bailey, “Reactive oxygen species production by the mitochondrial respiratory chain in isolated rat hepatocytes and liver mitochondria: studies using myxothiazol,” Archives of Biochemistry and Biophysics, vol. 405, no. 1, pp. 65–72, 2002.

C. C. Cunningham, W. B. Coleman, and P. I. Spach, “The effects of chronic ethanol consumption on hepatic mitochondrial energy metabolism,” Alcohol and Alcoholism, vol. 25, no. 2-3, pp. 127–136, 1990.

F. M. Ferreira, C. M. Palmeira, R. Seiça, A. J. Moreno, and M. S. Santos, “Diabetes and mitochondrial bioenergetics: alterations with age,” Journal of Biochemical and Molecular Toxicology, vol. 17, no. 4, pp. 214–222, 2003.

A. P. Rolo and C. M. Palmeira, “Diabetes and mitochondrial function: role of hyperglycemia and oxidative stress,” Toxicology and Applied Pharmacology, vol. 212, no. 2, pp. 167–178, 2006.

C. Vives-Bauza, R. Gonzalo, G. Manfredi, E. Garcia-Arumi, and A. L. Andreu, “Enhanced ROS production and antioxidant defenses in cybrids harbouring mutations in mtDNA,” Neuroscience Letters, vol. 391, no. 3, pp. 136–141, 2006.

C. Demeilliers, C. Maisonneuve, A. Grodet et al., “Impaired adaptive resynthesis and prolonged depletion of hepatic mitochondrial DNA after repeated alcohol binges in mice,” Gastroenterology, vol. 123, no. 4, pp. 1278–1290, 2002.

S. M. Katzman, M. A. Messerli, D. T. Barry et al., “Mitochondrial metabolism reveals a functional architecture in intact islets of Langerhans from normal and diabetic Psammomys obesus,” American Journal of Physiology. Endocrinology and Metabolism, vol. 287, no. 6, pp. E1090–E1099, 2004.