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Antimicrobial Agents and Chemotherapy, February 2007, p. 583-590, Vol. 51, No. 2
0066-4804/07/$08.00+0     doi:10.1128/AAC.01078-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Site-Specific Reduction of Oxidative and Lipid Metabolism in Adipose Tissue of 3'-Azido-3'-Deoxythymidine-Treated Rats{triangledown}

Catherine Deveaud,* Bertrand Beauvoit, Annabel Reynaud, and Jacques Bonnet

INSERM U441, Avenue du Haut Lévêque, 33600 Pessac, France

Received 26 August 2006/ Returned for modification 9 November 2006/ Accepted 27 November 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Although it is well accepted that treatment with some nucleoside reverse transcriptase inhibitors modifies both fat metabolism and fat distribution in humans, the mechanisms underlying these modifications are not yet known. The present investigation examined whether a decrease in oxidative capacity, induced by a chronic oral administration of 3'-azido-3'-deoxythymidine (AZT) in rats, could be associated with an alteration of the lipogenic capacity of white adipose tissues. The impact of obesity as a factor was then evaluated. Results showed that AZT treatment induced differential effects depending on anatomical localization. Indeed, in the inguinal adipose tissue, the specific activities of cytochrome c oxidase and fatty acid synthase, two rate-controlling enzymes in energy and lipogenic metabolisms, respectively, both decreased under AZT treatment, thus leading to a lowered cell lipid accumulation. Moreover, the AMP-activated protein kinase phosphorylation level tended to increase, thus implying that AZT causes an energy imbalance. Furthermore, the inguinal tissue of obese rats presented a sensitivity to AZT treatment that was higher than that of lean rats. In contrast, for epididymal tissue, no significant change in all these parameters could be detected under AZT treatment, regardless of the nutritional status of the animals. Taken together, these data demonstrate differential effects of AZT on subcutaneous adipose tissue and visceral white adipose tissue. It could be considered that the chronic decreases in energy and lipogenic metabolism of inguinal adipocyte, consecutive to AZT treatment, may lead, in the long term, to adipose tissue atrophy.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Protease inhibitors (PIs) in combination with nucleoside reverse transcriptase inhibitors (NRTIs) and/or nonnucleoside reverse transcriptase inhibitors are considered to be the standard treatment for optimal antiretroviral therapy for human immunodeficiency virus (HIV)-infected patients. This therapy, called highly active antiretroviral therapy (HAART), has demonstrated clinical, immunological, and survival benefits. However, unrecognized side effects of this therapy are becoming more evident as drug availability improves and treatment durations increase. A syndrome combining peripheral fat wasting (affecting facial, arm, and leg fat pads), insulin resistance, and hyperlipidemia, referred to as lipodystrophy syndrome, has been identified in HIV-infected patients receiving this treatment (4). The surprising feature of lipodystrophy syndrome is a difference in sensitivity to HAART between visceral and subcutaneous adipose tissues. In fact, under HAART, subcutaneous adipose tissue undergoes atrophy, whereas visceral adipose tissue does not (5, 34). Although the use of NRTIs was originally believed to be secondary to the use of the protease inhibitor class, it is now clear that NRTIs are also involved in this syndrome (3, 15). NRTIs are triphosphorylated by intracellular kinases. Even if these triphosphorylated molecules preferentially inhibit HIV reverse transcriptase, they are also able to inhibit mitochondrial enzymes such as adenylate kinase, ADP/ATP translocase, and mitochondrial DNA polymerase gamma; the inhibition of mitochondrial DNA polymerase gamma resulted in impaired synthesis of mitochondrial enzymes that are involved in oxidative phosphorylation (for a review, see reference 1). In a previous work, we demonstrated in vivo that 3'-azido-3'-deoxythymidine (AZT) significantly decreased the mitochondrial DNA content of rat inguinal adipose cells but did not modify that of epididymal adipose cells. The loss of mitochondrial DNA content per inguinal cell was associated with a parallel decrease in the cytochrome c oxidase activity but not in the activity of citrate synthase, a protein which, in contrast to the cytochrome oxidase complex, is exclusively encoded by the nuclear DNA (9). This mitochondrial enzymatic defect could in turn cause a modification of lipid metabolism in white adipocytes. Indeed, de novo lipogenesis requires cooperation between mitochondrial and cytoplasmic enzymes and involves fluxes of metabolites across mitochondrial membranes (for a review, see reference 17). Mitochondria are engaged in several pathways that are essential for fatty acid synthesis, namely, the generation of ATP, and they also support the formation of acetyl coenzyme A in cytoplasm via a mitochondrial efflux of citrate.

As we knew that de novo fatty acid synthesis in adipocytes is an important mechanism involved in the control of fat content not only in rodents but also in humans, in which adipose tissue may account for up to 40% of whole-body lipogenesis (6, 37), the aim of this study was to determine whether a decrease in oxidative capacity, evaluated through the measurement of cytochrome c oxidase, induced by AZT treatment, could be associated with an alteration of the lipogenic capacity of white adipose tissues. This study was carried out, in parallel, with subcutaneous and visceral adipose tissues to check for possible discrepancies. Moreover, this work was extended to the study of the influence of obesity, which has been shown to modify carbohydrate, lipid, and adipose metabolisms (2, 13). Indeed, standard and cafeteria diets were used to modulate the nutritional status of the animals. To distinguish the effects of NRTIs from those of PIs and nonnucleoside reverse transcriptase inhibitors, we chose to treat rats with AZT because thymidine analogues (i.e., stavudine and AZT) appear to play a major role in the development of lipoatrophy (23, 25). Inguinal and epididymal tissues were chosen, respectively, as subcutaneous and visceral adipose tissues because they represent the two main lipid storage sites of rat fat pads (10).


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animal care. Animal experimentation was performed in compliance with the French laws and regulations and in accordance with the Principles of Laboratory Animal Care (26). Male Wistar rats weighing 250 ± 10 g (Charles River Laboratories, l'Arbresle, France) were housed at 22 ± 2°C, on a 12-h/12-h photoperiod and were provided with food and water ad libitum. After 1 week of acclimatization, the rats were randomly divided in two groups. Control rats (n = 16) had free access to a standard nonpurified diet containing 51% carbohydrate, 4% fat, and 21% protein (SAFE, Augy, France) ad libitum. The second group, the cafeteria-fed group (n = 16), was given four foods palatable for human consumption in addition to the standard chow (22). The four supplementary items were changed each day on a weekly rotation. The list of palatable items included bacon, salami, potato chips, marshmallows, cheese, cookies, candies, and chocolate. The average composition of the diet was 12% protein, 57% fat, and 31% carbohydrate. One week later, each group of animals was randomly divided into two subgroups. The AZT-treated group (n = 8) received AZT in their drinking water for four weeks while the control group (n = 8) continued to receive plain tap water, both ad libitum. The dose of AZT used here in drinking water was 0.6 mg/ml, equivalent to 70 mg/kg body weight/day. This dose corresponds to human treatment regimens, i.e., 8 to 12 mg/kg/day, or to 50 to 75 mg/kg/day in rats after correction for the metabolic and drug disposal rate in rats being five- to sevenfold higher than that in humans (7).

Water consumption of all rats was recorded daily, thus allowing the calculation of the overall daily dose of AZT. The AZT solution was made up every day. All animals were weighed daily, and their overall appearances and activity levels were noted at that time. As reported in our previous work (9), the total area under the concentration-time curve for our rats was equal to 872 µg · min/ml. The total body clearance was 26 ml/min, and the apparent volume of distribution was 1.73 ml (with an elimination half-life of 15 min).

Before they were sacrificed, animals were anesthetized with 100 mg/kg ketamine and 10 mg/kg xylazine, mixed just before administration, and they were bled from the posterior vena cava by the drawing of 3 ml of blood on heparin sodium.

Determination of blood triglyceride, cholesterol, HDL cholesterol, and insulin. Serum triglyceride (TG) and total and high-density-lipoprotein (HDL) cholesterol concentrations were measured by routine enzymatic methods on an automated Hitachi 911 analyzer (Roche) with kits from Randox. The Friedewald equation was used to calculate low-density-lipoprotein (LDL) cholesterol (chol) concentrations ([LDL chol] = [total chol] – [HDL chol] – {[TG]/2.2}). Plasma insulin concentrations were determined in duplicate by using a commercial radioimmunoassay kit (INSIK 5; DiaSorin, Antony, France).

Tissue isolation. Two regions of adipose tissue were carefully dissected: the epididymal tissue, by a horizontal cut above the epididymus, and the inguinal subcutaneous tissue, by carefully dissecting all fat in the inguinal region up to a horizontal line parallel to the xyphoid cartilage. Tissues were dissected from visible blood vessels. Livers were also collected. Tissues were washed in phosphate-buffered saline (PBS) medium, blotted, weighed, and immediately frozen in liquid nitrogen. Aliquots were pulverized with a stainless steel mortar and pestle in liquid nitrogen. Powdered tissues were stored at –80°C until use.

Microscopic analysis. Freshly dissected inguinal and epididymal adipose tissues were fixed in 2.5% (vol/vol) glutaraldehyde in phosphate buffer (0.1 M, pH 7.2) and embedded in paraffin. Eight-micrometer sections were cut and stained with hematoxylin and eosin. Photomicrographs were captured with a Nikon Microphot-FXA camera at x4 and x10 magnifications. The cross-sectional measurement of the cell surface was determined using SigmaScan Pro 5 software, counting 200 to 300 cells in six different microscopic fields per tissue section.

DNA quantification. Powdered frozen tissues (200 to 300 mg) were homogenized in 1.5 ml of PBS by using a 3-ml glass-glass homogenizer (0.025-mm clearance; Kontes Glass Co., Vineland, NJ). Tissue homogenates were exposed to ultrasound energy (110 W) for 15 s on ice. Adipose samples were then centrifuged for 5 min at 600 x g and delipidated by aspirating the top of the supernatant.

Total DNA was quantified from delipidated tissue homogenates by using the method of Labarca and Paigen (19). This method uses the fluorescence enhancement of DNA-dye complex produced after the specific interaction of total cellular DNA with fluorescent dye (Hoescht H33258; Sigma). DNA content for each sample was extrapolated from a standard curve constructed using 0 to 1 µg/ml of calf thymus DNA (Sigma).

Mitochondrial enzymatic assays. Powdered frozen tissues (200 to 300 mg) were homogenized in 1.5 ml of PBS as described for DNA quantification assays.

Citrate synthase (EC 2.3.3.1, formerly EC 4.1.3.7) was measured according to the procedure of Srere (35), and one unit of citrate synthase was equal to the reduction of 1 µmol of 5-5'-dithiobis-2-nitrobenzoic acid per min. Cytochrome c oxidase (EC 1.9.3.1) activity was measured spectrophotometrically according to the method used by Rustin et al. (33) except that n-dodecylmaltoside (5% [vol/vol] final concentration) was added to homogenates prior to measuring cytochrome c oxidase activity. One unit of cytochrome c oxidase was equal to the oxidation of 1 µmol of ferricytochrome c per min. Tissue data were expressed as U/mg DNA.

Fatty acid synthase activity. Powdered frozen tissues (300 to 400 mg) were homogenized in 2 ml of buffer containing 0.25 M sucrose, 1 mM EDTA, and 0.1% (vol/vol) ß-mercaptoethanol with a 3-ml glass-glass homogenizer (0.025-mm clearance; Kontes Glass Co., Vineland, NJ). Tissue homogenates were then centrifuged for 15 min at 10,000 x g and delipidated. Supernatant was then centrifuged at 105,000 x g for 1 h. Enzymatic assays were carried out spectrophotometrically on this centrifuged supernatant as described by Hardie et al. (11) with minor modifications. Briefly, the reaction mixture contained 1 ml of a buffer containing 0.075% (wt/vol) delipidated bovine serum albumin, 1 mM EDTA, 3.75 mM glutathione, 0.2 M sodium phosphate (pH 6.6) supplemented with 0.125 mM NADPH, and 5 µM acetyl coenzyme A. The reaction, measured at 340 nm and at 37°C, was started by the addition of the sample and 10 µM malonyl coenzyme A. One unit of fatty acid synthase was taken to be equal to the oxidation of 1 µmol of NADPH per min. Tissue data were expressed as U/mg DNA.

AMPK phosphorylation. Protein extracts (200 mg) were prepared by complete homogenization of fat tissue powder in 1 ml homogenization buffer containing 50 mM Tris-HCl, 250 mM mannitol, 5 mM NaF, 1 mM sodium orthovanadate, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100 (vol/vol), 1 mM dithiothreitol, and antiprotease cocktail (Complete EDTA-free; Boehringer Mannheim). Dithiothreitol, NaF, sodium orthovanadate, and antiprotease cocktail were added just before use. The lysate was cleared by centrifugation at 14,000 x g for 10 min, and the supernatant was collected and used as protein extract. Protein concentration was determined using the bicinchoninic acid assay (Pierce). Equivalent protein amounts (50 µg) were diluted in sodium dodecyl sulfate (SDS) sample buffer (50 mM Tris-HCl, pH 6.8, 6% [wt/vol] SDS, 0.1% [wt/vol] bromophenol blue, 10% [vol/vol] glycerol) with 10% (vol/vol) ß-mercaptoethanol and were boiled for 5 min. They were separated on 12% SDS-polyacrylamide gels. Separated proteins were electroblotted onto polyvinylidene difluoride membranes (Millipore). Membranes were blocked for 4 h in TBS-Tween buffer (10 mM Tris-HCl, pH 7.4, 150 mM NaCl, 0.2% [vol/vol] Tween 20) containing 5% (wt/vol) skim milk and incubated overnight at 4°C in TBS-Tween buffer containing 5% (wt/vol) bovine serum albumin with antibody against AMP-activated protein kinase (AMPK) (which detects the {alpha}1 and {alpha}2 isoforms of the catalytic subunit) (1:1,000) and phospho-AMPK (which detects AMPK-{alpha} only when phosphorylated at threonine 172) (1/1,000) (Cell Signaling). Secondary antibodies conjugated to horseradish peroxidase were from Jackson ImmunoResearch Laboratories (Pennsylvania) ({alpha}-rabbit, 1:10,000). Antigen-antibody complexes were detected by chemiluminescence using an ECL kit (Amersham). The immunoblots were scanned and quantitatively analyzed by ImageJ software (NIH). Results are expressed as the ratios of phosphorylated proteins to total proteins (arbitrary units).

Statistical analysis. The results are expressed as means ± standard errors of the means (SEM). Statistical analysis was carried out using a nonparametric Mann-Whitney test (Statsdirect, United Kingdom). A probability value of less than 0.05 was considered significant.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Physiological characteristics of treated rats. By the end of the 5-week period, the cafeteria-fed group weighed significantly more than the lean group (447 ± 5 versus 395 ± 7 g, P < 0.01). AZT was administered in drinking water rather than by gavage or by injection. This means of administration limited animal stress which could have interfered with oxidative metabolism. Thus, rats were treated with 70 mg/kg/day for one month. Both the control and AZT-treated male rats gained weight steadily over the 4-week period of treatment and no significant difference in body weight gain existed between the two subgroups during the treatment (395 ± 7 and 401 ± 10 g for control and AZT-treated lean rats, respectively, and 447 ± 5 and 443 ± 8 g for control and AZT-treated cafeteria-fed rats). All animals appeared healthy and displayed comparable levels of alertness and activity when removed from their cages for weighing.

Effects of AZT treatment on blood levels of triglyceride, cholesterol, HDL cholesterol, and insulin. After triglyceride and total, HDL, and LDL cholesterol concentrations were measured, the ratio of total cholesterol to HDL cholesterol was calculated (Table 1) . This ratio is important because it is a good predictor of ischemic heart disease risk (21). The blood levels of triglycerides were significantly increased in AZT-treated lean rats, and their insulin levels were slightly increased (although not significantly different) compared to the lean controls. The levels of total cholesterol, HDL, and LDL cholesterol and the ratio of total cholesterol to HDL cholesterol ratio were not modified in AZT-treated lean rats compared to the lean control group. None of the parameters was significantly different in the AZT-treated cafeteria-fed group compared to the control cafeteria-fed group (Table 1).


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  		TABLE 1. Effects of AZT on blood triglyceride, cholesterol (total, HDL, and LDL), and insulina

 
It is worth noting that the total cholesterol-to-HDL cholesterol ratio was significantly decreased and the insulin level was increased (P = 0.097) for cafeteria-fed control rats in comparison to lean control rats. Finally, the total cholesterol level (1.79 ± 0.13 versus 1.50 ± 0.10 mM, P < 0.05), the ratio of total cholesterol to HDL cholesterol (8.35 ± 3.62 versus 3.71 ± 0.17 mM, P < 0.01) and insulin level (59.5 ± 19.9 versus 26.3 ± 2.5 mM, P < 0.05) were significantly higher in the AZT-treated cafeteria-fed group than in the control lean group (Table 1).

Effects of AZT treatment on inguinal and epididymal adipose tissues of lean and cafeteria-fed rats. (i) Tissue weight. As shown in Fig. 1, the adipose tissues of the cafeteria-fed group weighed significantly more than those of the lean group (10.16 ± 2.01 versus 6.99 ± 1.61 g for inguinal tissue and 12.27 ± 3.29 versus 6.55 ± 1.94 g for epididymal tissue). Moreover, the administration of AZT to rats for 4 weeks had no significant effect on the weights of inguinal or epididymal adipose pads (Fig. 1).


Figure 1
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  		FIG. 1. Effect of AZT on the weights of inguinal (ING) and epididymal (EPI) adipose tissues. The weights of inguinal and epididymal tissues for control (stippled column) and AZT-treated (hatched column) lean and cafeteria-fed rats were measured as described in Materials and Methods. Values are means ± SEM of three independent determinations performed for eight rats per group.

 
(ii) Cellularity, adipocyte size, and lipid accumulation. Cellularity, expressed as DNA content per tissue weight, represents the cell density of a given tissue. This tissular parameter may vary as the volume occupied by the lipid droplet in the adipocyte expands or decreases as a function of the steady-state lipid metabolism. Figure 2 shows the cellularity of the adipose tissues of all groups of rats. Even though AZT treatment did not significantly modify the cellularity of the inguinal adipose tissue in the lean group, it significantly increased that of the inguinal tissue for the cafeteria-fed group (625 ± 90 versus 401 ± 39 µg DNA/g wet weight, P < 0.05). No significant change of cellularity was shown in epididymal tissue by AZT treatment for any group of rats (Fig. 2).


Figure 2
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  		FIG. 2. Effect of AZT on the cellularities of inguinal (ING) and epididymal (EPI) adipose tissues. Cellularity was assessed by measuring DNA concentrations in triplicate on tissue homogenates as described in Materials and Methods for control (stippled column) and AZT-treated (hatched column) lean and cafeteria-fed rats. Values are means ± SEM of three independent determinations for eight rats per group. The asterisk indicates a significant difference between control and AZT-treated tissues (P < 0.05 [Mann-Whitney test]).

 
However, one should be aware that the cellularity of adipose tissues takes into account nonadipocyte cell population (i.e., resulting in the stromal vascular fraction) (8). To further analyze whether these changes in cellularity were correlated to changes in adipocyte size and, consequently, in cellular lipid accumulation, the inguinal and epididymal adipocyte sizes were measured under different conditions of diet and treatment. A representative photomicrograph of tissue sections is shown in Fig. 3A. It should be noted that adipocytes from cafeteria-fed rats are bigger than those of lean rats. After quantifications performed with photomicrographs (Fig. 3B), it can be concluded that AZT treatment significantly decreased the size of inguinal adipocytes from cafeteria-fed rats (2,082 ± 288 versus 2,866 ± 346 µm2, P < 0.05). Finally, there was no modification in adipocyte size in epididymal tissue upon AZT treatment, regardless of the nutritional status of the rats (Fig. 3B).


Figure 3
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  		FIG. 3. Effect of AZT on the sizes of inguinal (ING) and epididymal (EPI) adipocytes. (A) Representative micrograph of inguinal and epididymal cross-sectional adipose tissue from control and AZT-treated lean and cafeteria-fed rats. Tissue fixation and thin sections were prepared as described in Materials and Methods from inguinal (a) and epididymal (e) tissues from control lean rats, inguinal (b) and epididymal (f) tissues from AZT-treated lean rats, inguinal (c) and epididymal (g) tissues from control cafeteria-fed rats and inguinal (d) and epididymal (h) tissues from AZT-treated cafeteria-fed rats. Scale bar, 100 µm. (B) Inguinal and epididymal adipocyte sizes from control (stippled column) and AZT-treated (hatched column) lean and cafeteria-fed rats were assessed by measuring cross-sectional cell surfaces by using SigmaScan Pro 5 software, counting 200 to 300 cell surfaces in six different microscopic fields per tissue section as described in Materials and Methods. Values are means ± SEM of independent determinations for eight rats. The asterisk indicates a significant difference between control and AZT-treated tissues (P < 0.05 [Mann-Whitney test]).

 
(iii) Mitochondrial enzymatic equipment. As regards the enzymatic equipment, it is worth noting that it was expressed per mg of total DNA (cellular parameter) rather than per mg of total protein or per g of wet tissue (tissular parameters). This normalization made it possible to compare enzymatic activities per cell, which is particularly useful when comparing data on two types of adipose tissue cell. Figure 4 shows the specific activities of two mitochondrial enzyme markers measured on adipose tissue homogenates. AZT treatment significantly decreased the cytochrome c oxidase activities of inguinal adipose tissue of lean (6.61 ± 1.68 versus 9.60 ± 1.29 U/mg DNA, P < 0.05) and cafeteria-fed (6.43 ± 1.15 versus 10.75 ± 1.12 U/mg DNA, P < 0.01) rats (Fig. 4A). In contrast, regarding the activity of this complex, AZT had no significant effect on epididymal adipose tissue regardless of the nutritional status of the rats. Meanwhile, the specific activity of citrate synthase, a nuclear DNA-encoded enzyme which is commonly used as a probe of mitochondrial mass (14), was slightly decreased in inguinal tissue by AZT treatment for each subgroup of rats (lean or cafeteria fed), although it was not significantly different from the activity of the controls (Fig. 4B). The same observation can be made for epididymal tissue (Fig. 4).


Figure 4
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  		FIG. 4. Effect of AZT on the mitochondrial enzymatic equipments of inguinal (ING) and epididymal (EPI) adipose tissues. Specific activities of cytochrome c oxidase (A) and citrate synthase (B) were measured, as described in Materials and Methods, in triplicate on inguinal and epididymal tissue homogenates from control (stippled column) and AZT-treated (hatched column) lean and cafeteria-fed rats. Values are means ± SEM of three independent determinations for eight rats per group. Asterisks indicate significant differences between control and AZT-treated tissues (*, P < 0.05; **, P < 0.01 [Mann-Whitney test]).

 
(iv) Fatty acid synthase content. To gain further insight into the origin of the AZT-induced changes in cellular lipid accumulation, specific activity of the fatty acid synthase, one of the key enzymes for de novo lipogenesis (38), was measured in the cytosolic fraction of adipose tissue homogenates (Fig. 5). Fatty acid synthase activity was decreased in inguinal cells of AZT-treated lean rats (0.72 ± 0.2 versus 1.12 ± 0.2 U/mg DNA, P = 0.17) and was significantly altered in inguinal cells of AZT-treated cafeteria-fed rats (0.55 ± 0.39 versus 1.61 ± 1.00 U/mg DNA, P < 0.05). Moreover, there was no modification of fatty acid synthase activity in epididymal tissue upon AZT treatment for any group of rats (Fig. 5).


Figure 5
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  		FIG. 5. Effect of AZT on fatty acid synthase activity of inguinal (ING) and epididymal (EPI) adipose tissues. The specific activity of fatty acid synthase was measured as described in Materials and Methods in triplicate on inguinal and epididymal tissue homogenates from control (stippled column) and AZT-treated (hatched column) lean and cafeteria-fed rats. Values are means ± SEM of three independent determinations for eight rats per group. The asterisk indicates a significant difference between control and AZT-treated tissues (P < 0.05 [Mann-Whitney test]).

 
(v) AMPK phosphorylation. Inguinal adipose cells from AZT-treated rats presented with a reduction of oxidative and lipogenic capacities; it thus seemed interesting to measure AMPK activation in our different adipose tissues given the critical role of this kinase in regulating intracellular energy metabolism in response to energy crises. The activation of AMPK is mainly dependent on phosphorylation of the Thr172 residue of the {alpha} subunit by enzymes collectively referred to as AMPK kinase (12, 16, 36). As shown in Fig. 6, AZT tended to increase the steady-state AMPK phosphorylation level in inguinal adipose cells, regardless of the nutritional status of the rats. No difference in AMPK phosphorylation could be detected in epididymal tissue after AZT treatment among any of the groups of rats (Fig. 6).


Figure 6
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  		FIG. 6. Effect of AZT on AMPK activities of inguinal (ING) and epididymal (EPI) adipose tissues. Cytosolic proteins were extracted from inguinal and epididymal tissues from control (stippled column) and AZT-treated (hatched column) lean and cafeteria-fed rats. Equal amounts (50 µg) of proteins were subjected to 12% SDS-polyacrylamide gel electrophoresis and transferred onto a polyvinylidene difluoride membrane. AMPK and phospho-AMPK were detected by immunoblotting with specific anti-AMPK and anti-phospho-AMPK antibodies, respectively. Quantitative analysis of AMPK and phospho-AMPK was performed by densitometric analysis. Results are expressed as the ratios between phosphorylated and total proteins (arbitrary units [a.u.]) and are presented as means ± SEM (n = 8).

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Although treatments with NRTIs can modify fat metabolism and fat distribution in humans, the mechanisms underlying these modifications and the respective role of each NRTI are still unknown. Moreover, these alterations are probably the consequence of several nonexclusive factors, namely, HIV infection, PIs, and NRTIs. To distinguish the effects of NRTIs from these other factors, we decided to treat rats with AZT, rather than other available drugs, as lipoatrophy appears to be most closely associated with prolonged therapy with thymidine nucleoside analogues (23, 25). Adverse effects of AZT have been linked to mitochondrial toxicity. In a previous work with a rat model, we demonstrated that AZT treatment significantly decreased the mitochondrial DNA content of inguinal adipose cells, and this was associated with a decrease in the cytochrome c oxidase activity, thus leading to a lower oxidative capacity in this subcutaneous adipose tissue (9).

This study therefore set out to determine whether AZT-induced changes in oxidative capacity on inguinal and epididymal adipose tissues had effects on adipocyte lipogenic capacity and, if so, to check the site-specific effect of AZT. This work was also dedicated to the study of the influence of nutritional status on the goals mentioned above. Indeed, obesity and its metabolic complications already reflect, in themselves, an unbalanced lipid and energy metabolism (2). For this purpose, one group of rats underwent a cafeteria diet, which has been reported to induce increases in body weight and adipose fat pad mass in rats, even after a short period of time (28, 30). Cafeteria-fed rats are a useful model for human obesity because the cafeteria diet is a palatable hypercaloric and hyperlipidic diet that induces voluntary hyperphagia and fast body weight gain (30).

In our study, a cafeteria diet significantly increased the body weight of rats and the sizes of their inguinal and epididymal adipose pads. Furthermore, it decreased significantly the total cholesterol-to-HDL cholesterol ratio and increased almost significantly the insulin level. Moreover, the combination of AZT treatment with a cafeteria diet had synergistic effects on these biological parameters, namely, increased levels of total cholesterol and insulin as well as a higher total cholesterol-to-HDL cholesterol ratio than that of lean control rats.

Under our conditions of AZT treatment (i.e., 70 mg/kg/day for one month) and among any of the groups of rats, the weights of adipose pads were not significantly different, indicating that AZT did not induce lipodystrophy, even though a trend was noted for inguinal fat pad weight to be lower in the AZT-treated cafeteria-fed rats.

Before a lipogenic capacity analysis of adipose tissues was addressed, the oxidative capacity was assessed by measuring the cytochrome c oxidase activity. We confirmed that AZT significantly decreased the cytochrome c oxidase activity in inguinal tissue from lean rats (9) and demonstrated, for the first time, a higher sensitivity of the inguinal tissue to AZT treatment for the cafeteria-fed rats. No such pattern could be detected in the epididymal tissues of both groups of rats.

Cellular lipid accumulation, assessed in inguinal tissue by measuring the cellularity and the adipocyte size, was significantly decreased by AZT treatment for the cafeteria-fed group, whereas it had no significant effect on inguinal adipocytes of lean rats, even if a tendency for this level to decrease could be noted. In parallel, tissue cellularity increase was associated with adipocyte size decrease. These results obtained in vivo with subcutaneous tissue confirmed those obtained in vitro by Lagathu et al. (20), who found that NRTIs, used individually (i.e., AZT and stavudine), decreased lipid accumulation on differentiating (3T3-F442A) and fully differentiated (3T3-L1) adipocytes in culture. Kosmiski et al. (18) demonstrated that NRTI combinations do significantly decrease lipid accumulation in 3T3-L1 adipocytes in culture. Nolan et al. (27) observed an increased number of small subcutaneous adipocytes in NRTI-treated patients affected by fat wasting. However, our in vivo study clearly shows that AZT has differential effects depending on the localization of the fat pad, since AZT had no effect on a visceral tissue, i.e., the epididymal adipose tissue.

Lipogenic capacity was then assessed by measuring the fatty acid synthase activity. Fatty acid synthase is rate limiting in the long-term regulation of fatty acid synthesis (38). This enzyme activity in inguinal cells of AZT-treated lean rats decreased (compared with control lean rats), and that in inguinal cells of AZT-treated cafeteria-fed rats was significantly lower (compared with control cafeteria-fed rats). There was no modification in fatty acid synthase activity in epididymal tissue by AZT treatment for any group of rats. This report highlights the fact that a decrease in the activity of cytochrome c oxidase (i.e., a reduction of oxidative capacity), consecutive to AZT treatment, is associated with decreased activity of fatty acid synthase (i.e., a reduction of lipogenic capacity).

These data are consistent with those obtained by Rossmeisl et al. (32), who found that a decrease in ATP synthesis (i.e., by an acute uncoupling of oxidative phosphorylation by means of a protonophoric uncoupler addition on cultured cells) depressed fatty acid synthesis. They proposed that inhibition of fatty acid synthesis by mitochondrial uncoupling probably results from a short-term metabolic regulation, i.e., a limited availability of intramitochondrial ATP for the pyruvate carboxylase (31), whose inhibition probably slows down fatty acid synthesis as a result of a limited supply of acetyl units. On the other hand, by using transgenic mice expressing an ectopic UCP1 in white fat, the same authors showed a decrease in both acetyl coenzyme A carboxylase and fatty acid synthase expressions in white fat. In this case of an adaptative model of chronic mitochondrial uncoupling, they suggested that the expression of genes encoding lipogenic enzymes responds to changes in mitochondrial oxidative phosphorylation capacity (32).

Our animal model treated for one month with AZT appeared also to resemble an adaptative model of chronic decrease of mitochondrial oxidative phosphorylation capacity. Upon modulation of the mitochondrial energy metabolism, the changes in the lipogenic activity of adipose cells (and consequently, their steady-state lipid accumulation) may be the result of a short-term kinetic regulation of lipogenic enzymes, doubled by a long-term regulation of the expression of lipogenic enzymes. In contrast, whichever parameters are considered and regardless of the nutritional status of the rats, AZT had no significant effect on epididymal tissues.

To examine the causal link between these metabolic changes, we analyzed the steady-state AMPK phosphorylation level in our rat model. AMPK is a key regulatory enzyme in cellular energy homeostasis (12, 16, 36). This kinase is actually considered to be an energy status sensor, since it is activated allosterically by increases in the AMP/ATP ratio as well as by phosphorylation on Thr172 by upstream kinases. In this context, in response to energy depletion, AMPK activation may trigger the upregulation of genes involved in energy production and therefore regulate mitochondrial biogenesis (for a review, see reference 29). A twofold activation of this protein kinase upon expression of the uncoupling protein UCP1 in vivo on white adipose tissues of transgenic mice has already been described (24). Under our conditions of AZT treatment, AMPK phosphorylation tended to increase by 35% in inguinal adipose cells, regardless of the nutritional status of the rats. No difference in AMPK phosphorylation could be detected in epididymal tissue after AZT treatment among any of the groups of rats. The fact that the AMPK specifically responds to AZT treatment in inguinal tissue would argue for a lowered energy status of the inguinal adipocytes, the consequence of an AZT-induced decrease in the mitochondrial oxidative capacity. Nevertheless, the moderate response of AMPK in inguinal adipocytes could be explained by the fact that our rat model represents more an adaptative model of chronic limitation in ATP synthesis than a model of acute energy depletion.

The other main result of this study is that, under our treatment conditions, AZT produced greater effects on inguinal tissue of cafeteria-fed rats than on that of lean rats. Thus, in our rat model, it appeared that obesity and metabolic complications associated with this nutritional status, such as resistance to insulin and dyslipidemia, strongly increased the vulnerability of subcutaneous adipose tissue to AZT. Our results underline the widely accepted role of obesity as an aggravating factor in metabolic pathologies. However, it is the first time, to our knowledge, that the impact of obesity on adipose tissue AZT side effects has been investigated. It is therefore important to point out that the prevention of obesity could be a key factor in the control of lipodystrophy syndrome effects.


   ACKNOWLEDGMENTS
 
We thank Joanne Pageze for her contribution to the editing of the manuscript.

This work was supported by a grant from Sidaction (France). C.D. was a recipient of a grant from Sidaction (France).


   FOOTNOTES
 
* Corresponding author. Mailing address: INSERM U441, Avenue du Haut Lévêque, 33600 Pessac, France. Phone: (33) 5 57 89 19 79. Fax: (33) 5 56 36 89 79. E-mail: catherine.deveaud{at}wanadoo.fr. Back

{triangledown} Published ahead of print on 11 December 2006. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Barile, M., D. Valenti, E. Quagliariello, and S. Passarella. 1998. Mitochondria as cell targets of AZT (zidovudine). Gen. Pharmacol. 31:531-538.[Medline]
  2. Bjorntorp, P. 1993. Visceral obesity: a "civilization syndrome." Obes. Res. 1:206-222.[Medline]
  3. Brinkman, K., J. A. Smeitink, J. A. Romijn, and P. Reiss. 1999. Mitochondrial toxicity induced by nucleoside analogue reverse transcriptase inhibitors is a key factor in the pathogenesis of antiretroviral therapy related lipodystrophy. Lancet 354:1112-1115.[CrossRef][Medline]
  4. Carr, A., K. Samaras, S. Burton, M. Law, J. Freund, D. J. Chisholm, and D. A. Cooper. 1998. A syndrome of peripheral lipodystrophy, hyperlipidaemia and insulin resistance in patients receiving HIV protease inhibitors. AIDS 12:F51-F58.[CrossRef][Medline]
  5. Carr, A., K. Samaras, D. J. Chisholm, and D. A. Cooper. 1998. Abnormal fat distribution and use of protease inhibitors. Lancet 351:1736.[Medline]
  6. Chascione, C., D. H. Elwyn, M. Davila, K. M. Gil, J. Askanazi, and J. M. Kinney. 1987. Effect of carbohydrate intake on de novo lipogenesis in human adipose tissue. Am. J. Physiol. 253:E664-E669.
  7. Chiou, W. L., C. Ma, S. M. Chung, T. C. Wu, and H. Y. Jeong. 2000. Similarity in the linear and non-linear oral absorption of drugs between human and rat. Int. J. Clin. Pharmacol. Ther. 38:532-539.[Medline]
  8. Deveaud, C., B. Beauvoit, B. Salin, J. Schaeffer, and M. Rigoulet. 2004. Regional differences in oxidative capacity of rat white adipose tissue are linked to the mitochondrial content of mature adipocytes. Mol. Cell. Biochem. 267:157-166.[CrossRef][Medline]
  9. Deveaud, C., B. Beauvoit, S. Hagry, A. Galinier, A. Carriere, B. Salin, J. Schaeffer, S. Caspar-Bauguil, Y. Fernandez, J. B. Gordien, D. Breilh, L. Penicaud, L. Casteilla, and M. Rigoulet. 2005. Site specific alterations of adipose tissue mitochondria in 3'-azido-3'-deoxythymidine (AZT)-treated rats: an early stage in lipodystrophy? Biochem. Pharmacol. 70:90-101.[CrossRef][Medline]
  10. DiGirolamo, M., J. B. Fine, K. Tagra, and R. Rossmanith. 1998. Qualitative regional differences in adipose tissue growth and cellularity in male Wistar rats fed ad libitum. Am. J. Physiol. 274:R1460-R1467.
  11. Hardie, D. G., P. S. Guy, and P. Cohen. 1981. Acetyl-CoA carboxylase and fatty acid synthase from lactating rabbit and rat mammary gland. Methods Enzymol. 71:26-33.
  12. Hardie, D. G., and D. Carling. 1997. The AMP-activated protein kinase-fuel gauge of the mammalian cell? Eur. J. Biochem. 246:259-273.[Medline]
  13. Havel, P. J. 2002. Control of energy homeostasis and insulin action by adipocyte hormones: leptin, acylation stimulating protein, and adiponectin. Curr. Opin. Lipidol. 13:51-59.[CrossRef][Medline]
  14. James, A. M., Y. H. Wei, C. Y. Pang, and M. P. Murphy. 1996. Altered mitochondrial function in fibroblasts containing MELAS or MERRF mitochondrial DNA mutation. Biochem. J. 318:401-407.
  15. Kakuda, T. N., R. C. Brundage, P. L. Anderson, and C. V. Fletcher. 1999. Nucleoside reverse transcriptase inhibitor-induced mitochondrial toxicity as an etiology for lipodystrophy. AIDS 13:2311-2312.[CrossRef][Medline]
  16. Kemp, B. E., D. Stapleton, D. J. Campbell, Z. P. Chen, S. Murthy, M. Walter, A. Gupta, J. J. Adams, F. Katsis, B. van Denderen, I. G. Jennings, T. Iseli, B. J. Michell, and L. A. Witters. 2003. AMP-activated protein kinase, super metabolic regulator. Biochem. Soc. Trans. 31:162-168.[Medline]
  17. Kopecky, J., M. Rossmeisl, P. Flachs, K. Bardova, and P. Brauner. 2001. Mitochondrial uncoupling and lipid metabolism in adipocytes. Biochem. Soc. Trans. 29:791-797.[CrossRef][Medline]
  18. Kosmiski, L. A., H. L. Miller, and D. J. Klemm. 2006. In combination, nucleoside reverse transcriptase inhibitors have significant effects on 3T3-L1 adipocyte lipid accumulation and survival. Antivir. Ther. 11:187-195.[Medline]
  19. Labarca, C., and K. Paigen. 1980. A simple, rapid, and sensitive DNA assay procedure. Anal. Biochem. 102:344-352.[CrossRef][Medline]
  20. Lagathu, C., J. P. Bastard, M. Auclair, M. Maachi, M. Kornprobst, J. Capeau, and M. Caron. 2004. Antiretroviral drugs with adverse effects on adipocyte lipid metabolism and survival alter the expression and secretion of proinflammatory cytokines and adiponectin in vitro. Antivir. Ther. 9:911-920.[Medline]
  21. Lemieux, I., B. Lamarche, C. Couillard, A. Pascot, B. Cantin, J. Bergeron, G. R. Dagenais, and J. P. Despres. 2001. Total cholesterol/HDL cholesterol ratio vs. LDL cholesterol/HDL cholesterol ratio as indices of ischemic heart disease risk in men: the Quebec Cardiovascular Study. Arch. Intern. Med. 161:2685-2692.[Abstract/Free Full Text]
  22. Mandenoff, A., T. Lenoir, and M. Apfelbaum. 1982. Tardy occurrence of adipocyte hyperplasia in cafeteria-fed rat. Am. J. Physiol. 242:R349-R351.
  23. Martin, A., D. E. Smith, A. Carr, C. Ringland, J. Amin, S. Emery, J. Hoy, C. Workman, N. Doong, J. Freund, D. A. Cooper, and the Mitochondrial Toxicity Study Group. 2004. Reversibility of lipoatrophy in HIV-infected patients 2 years after switching from a thymidine analogue to abacavir: the MITOX Extension Study. AIDS 18:1029-1036.[CrossRef][Medline]
  24. Matejkova, O., K. J. Mustard, J. Sponarova, P. Flachs, M. Rossmeisl, I. Miksik, M. Thomason-Hughes, D. Grahame Hardie, and J. Kopecky. 2004. Possible involvement of AMP-activated protein kinase in obesity resistance induced by respiratory uncoupling in white fat. FEBS Lett. 569:245-248.[CrossRef][Medline]
  25. Moyle, G., C. Sabin, J. Cartledge, M. Johnson, E. Wilkins, D. Churchill, P. Hay, A. Fakoya, M. Murphy, G. Scullard, C. Leen, and G. Reilly. 2006. A randomized comparative trial of tenofovir DF or abacavir as replacement for a thymidine analogue in persons with lipoatrophy. AIDS 20:2043-2050.[Medline]
  26. National Institutes of Health. 1985. Principles of laboratory animal care. NIH publication 85-23, revised. National Institutes of Health, Bethesda, MD.
  27. Nolan, D., E. Hammond, A. Martin, L. Taylor, S. Herrmann, E. McKinnon, C. Metcalf, B. Latham, and S. Mallal. 2003. Mitochondrial DNA depletion and morphologic changes in adipocytes associated with nucleoside reverse transcriptase inhibitor therapy. AIDS 17:1329-1338.[CrossRef][Medline]
  28. Redonnet, A., R. Groubet, C. Noël-Suberville, S. Bonilla, A. Martinez, and P. Higueret. 2001. Exposure to an obesity-inducing diet early affects the pattern of expression of peroxisome proliferator, retinoic acid, and triiodothyronine nuclear receptors in the rat. Metabolism 50:1161-1167.[CrossRef][Medline]
  29. Reznick, R. M., and G. I. Shulman. 2006. The role of AMP-activated protein kinase in mitochondrial biogenesis. J. Physiol. 574:33-39.[Abstract/Free Full Text]
  30. Rodríguez, E., J. Ribot, A. M. Rodriguez, and A. Palou. 2004. PPAR-{gamma}2 expression in response to cafeteria diet: gender- and depot-specific effects. Obes. Res. 12:1455-1463.[Medline]
  31. Rognstad, R., and J. Katz. 1969. The effect of 2,4-dinitrophenol on adipose-tissue metabolism. Biochem. J. 111:431-444.[Medline]
  32. Rossmeisl, M., I. Syrovy, F. Baumruk, P. Flachs, P. Janovska, and J. Kopecky. 2000. Decreased fatty acid synthesis due to mitochondrial uncoupling in adipose tissue. FASEB J. 12:1793-1800.
  33. Rustin, P., D. Chretien, T. Bourgeron, B. Gérard, A. Rötig, J. M. Saudubray, and A. Munnich. 1994. Biochemical and molecular investigations in respiratory chain deficiencies. Clin. Chim. Acta 228:35-51.[CrossRef][Medline]
  34. Shaw, A. J., K. A. McLean, and B. A. Evans. 1998. Disorders of fat distribution in HIV infection. Int. J. STD AIDS 9:595-599.[Abstract/Free Full Text]
  35. Srere, P. A. 1969. Citrate synthase. Methods Enzymol. 13:3-11.
  36. Stein, S. C., A. Woods, N. A. Jones, M. D. Davison, and D. Carling. 2000. The regulation of AMP-activated protein kinase by phosphorylation. Biochem. J. 345:437-443.
  37. Swierczynski, J., E. Goyke, L. Wach, A. Pankiewicz, Z. Kochan, W. Adamonis, Z. Sledzinski, and Z. Aleksandrowicz. 2000. Comparative study of the lipogenic potential of human and rat adipose tissue. Metabolism 49:594-599.[CrossRef][Medline]
  38. Volpe, J. J., and P. R. Vagelos. 1976. Mechanisms and regulation of biosynthesis of saturated fatty acids. Physiol. Rev. 56:339-417.[Free Full Text]

Antimicrobial Agents and Chemotherapy, February 2007, p. 583-590, Vol. 51, No. 2
0066-4804/07/$08.00+0     doi:10.1128/AAC.01078-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.




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