Mediators of Inflammation

Mediators of Inflammation / 2010 / Article
Special Issue

Mediators of Inflammation in Obesity and Its Co-Morbidities

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Research Article | Open Access

Volume 2010 |Article ID 784343 |

Neira Sáinz, Amaia Rodríguez, Victoria Catalán, Sara Becerril, Beatriz Ramírez, Javier Gómez-Ambrosi, Gema Frühbeck, "Leptin Administration Downregulates the Increased Expression Levels of Genes Related to Oxidative Stress and Inflammation in the Skeletal Muscle of ob/ob Mice", Mediators of Inflammation, vol. 2010, Article ID 784343, 15 pages, 2010.

Leptin Administration Downregulates the Increased Expression Levels of Genes Related to Oxidative Stress and Inflammation in the Skeletal Muscle of ob/ob Mice

Academic Editor: Giamila Fantuzzi
Received21 Jan 2010
Revised31 Mar 2010
Accepted24 Apr 2010
Published30 Jun 2010


Obese leptin-deficient ob/ob mice exhibit a low-grade chronic inflammation together with a low muscle mass. Our aim was to analyze the changes in muscle expression levels of genes related to oxidative stress and inflammatory responses in leptin deficiency and to identify the effect of in vivo leptin administration. Ob/ob mice were divided in three groups as follows: control ob/ob, leptin-treated ob/ob (1 mg/kg/d) and leptin pair-fed ob/ob mice. Gastrocnemius weight was lower in control ob/ob than in wild type mice ( ) exhibiting an increase after leptin treatment compared to control and pair-fed ( ) ob/ob animals. Thiobarbituric acid reactive substances, markers of oxidative stress, were higher in serum ( ) and gastrocnemius ( ) of control ob/ob than in wild type mice and were significantly decreased ( ) by leptin treatment. Leptin deficiency altered the expression of 1,546 genes, while leptin treatment modified the regulation of 1,127 genes with 86 of them being involved in oxidative stress, immune defense and inflammatory response. Leptin administration decreased the high expression of Crybb1, Hspb3, Hspb7, Mt4, Cat, Rbm9, Serpinc1 and Serpinb1a observed in control ob/ob mice, indicating that it improves inflammation and muscle loss.

1. Introduction

Obesity is associated with a low-grade proinflammatory state resulting in an increase of circulating cytokines and inflammatory markers [1]. Inflammatory cytokines have been involved in the impairment of insulin signaling, thus providing molecular links between inflammation and insulin resistance [2]. Inflammation reportedly produces metabolic alterations in skeletal muscle with both inflammatory response and insulin resistance being associated with loss of muscle mass by decreased protein synthesis and increased proteolysis [35]. Recently, our group has shown that leptin reverses muscle loss of ob/ob mice by inhibiting the activity of the transcriptional factor forkhead box class O3a (FoxO3a) [6].

Leptin is an adipocyte-derived peptidic hormone [7] that inhibits food intake and increases thermogenesis by acting through its hypothalamic receptors [8, 9]. Leptin-deficient ob/ob mice are obese, hyperphagic, exhibit type 2 diabetes, decreased body temperature and hypogonadotropic hypogonadism [10]. Leptin is a member of the long-chain helical cytokine family and its receptors, which belong to the class I cytokine receptors, are present in bone marrow and spleen as well as on peripheral monocytes and lymphocytes [1]. Leptin increases in response to acute infection and sepsis and it has been reported to exert a profound influence on the function and proliferation of T lymphocytes and natural killer cells [11], on the phagocytosis of macrophages/monocytes [12], and to have a direct effect on the secretion of anti- and proinflammatory cytokines [13]. In this regard, impaired cellular and humoral immunity have been shown in leptin-deficient ob/ob mice as well as in leptin receptor-deficient db/db mice [14, 15]. These studies reflect the molecular nature of leptin as a cytokine and are consistent with leptin signaling playing a pivotal role in the pathogenesis of obesity-associated inflammation and muscle loss.

In the present paper, gastrocnemius muscle samples from wild type and ob/ob mice were analyzed for mRNA presence of over 41,000 transcripts by microarray analysis to identify genes involved in inflammation and oxidative stress that are affected by leptin deficiency and leptin administration in ob/ob mice. It was shown that leptin increases the gastrocnemius weight and reduces the high expression levels of genes related to the obesity-associated low-grade inflammation in skeletal muscle of ob/ob mice.

2. Material and Methods

2.1. Animals and Treatments

Ten-week-old male genetically obese ob/ob mice (C57BL/6J) ( ) and their lean control littermates wild type ( ) supplied by Harlan (Barcelona, Spain) were housed in a room with controlled temperature ( ) and a 12:12 light-dark cycle (lights on at 08:00 am). Body weight of ob/ob mice was measured before randomization into control, leptin-treated (1 mg/kg/d) and pair-fed groups ( per group). The control and pair-fed groups received vehicle (PBS), while leptin-treated mice were intraperitoneally administered with leptin (Bachem, Bubendorf, Switzerland) twice daily at 08:00 am and 08:00 pm for 28 days. Control and leptin-treated groups were provided with water and food ad libitum with a standard rodent chow (2014S Teklad, Harlan), while daily food intake of the pair-fed group was matched to the amount consumed by the leptin-treated group the day before in order to discriminate the inhibitory effect of leptin on appetite. Animals were sacrificed on the 28th day of treatment by inhalation 20 hours after the last PBS or leptin administration (in order to avoid picking up effects reflecting an acute response) and after 8 hours of fasting. Serum samples and gastrocnemius muscles were obtained and stored at . All experimental procedures conformed to the European Guidelines for the Care and Use of Laboratory Animals (directive 86/609) and were approved by the Ethical Committee for Animal Experimentation of the University of Navarra (080/05).

2.2. Blood Analysis

Serum glucose was analyzed using a sensitive-automatic glucose sensor (Ascensia Elite, Bayer, Barcelona, Spain). Free fatty acid (FFA) concentrations were measured by a colorimetric determination using the NEFA C kit (WAKO Chemicals, Neuss, Germany). Serum glycerol concentrations were evaluated by enzymatic methods as previously described [6]. Serum triglycerides (TG) concentrations were spectrophotometrically determined using a commercial kit (Infinity, Thermo Electron, Melbourne, Australia). Insulin and leptin were determined using specific mouse ELISA kits (Crystal Chem Inc., Chicago, IL, USA). Intra- and interassay coefficients of variation for measurements of insulin and leptin were 3.5% and 6.3%, respectively, for the former, and 2.8% and 5.8%, for the latter. Adiponectin concentrations were also assessed using a mouse ELISA kit (BioVendor Laboratory Medicine, Inc., Modrice, Czech Republic). Intra- and interassay coefficients of variation for adiponectin were 2.6% and 5.3%, respectively. Insulin resistance was calculated using the homeostasis model assessment score (HOMA; fasting insulin ( U/mL) fasting glucose (mmol/L)/22.5) [16]. An indirect measure of insulin sensitivity was calculated by using the quantitative insulin sensitivity check index (QUICKI; 1/[log(fasting insulin mU/mL) log(fasting glucose mg/dL)] [17].

Lipid peroxidation was analyzed by the measurement of thiobarbituric acid reactive substances (TBARS) in serum and gastrocnemius as previously described by Conti et al. [18] with some modifications. Since the best-known specific TBARS is malondialdehyde (MDA), we used serum MDA levels, a secondary product of lipid peroxidation, as an indicator of lipid peroxidation and oxidative stress. Gastrocnemius samples (20–30 mg) were homogenized in 20 volumes of phosphate buffer pH 7.4. Serum, muscle homogenates (5  L) or standard (MDA) were mixed with 120  L of diethyl thiobarbituric acid (DETBA) 10 mM and vortexed for 5 seconds. The reaction mixture was then incubated at for 60 minutes. After cooling to room temperature DETBA-MDA adducts were extracted in 360  L n-butanol vortexing for 1 minute and centrifuged at 1,600 g for 10 minutes at room temperature. Then, the chromophore of the DETBA-MDA adduct was quantified in 200  L of the upper butanol phase by fluorescence emission at 535 nm with an excitation at 590 nm. MDA equivalents (TBARS) were quantified using a calibration curve prepared using MDA standard working solutions and expressed as serum MDA M and gastrocnemius MDA M/mg protein. Protein concentrations were determined using a Bradford protein assay kit (BioRad, Hercules, CA, USA).

2.3. Microarray Experiments and Analysis

Total RNA was extracted from 20–30 mg of gastrocnemius muscle samples by homogenization with an ULTRA-TURRAX T 25 basic (IKA Werke GmbH, Staufen, Germany) using TRIzol reagent (Invitrogen, Barcelona, Spain). RNA was purified using the RNeasy Mini kit (Qiagen, Barcelona, Spain) and treated with DNase I (RNase-free DNase Set, Qiagen) in order to remove any trace of genomic DNA.

Gene expression analyses were conducted using the Agilent Whole Mouse Genome array (G4121B, Agilent Technologies, Santa Clara, CA, USA) containing ~41 mouse genes and transcripts. Fluorescence-labeled cDNA probes were prepared from 1  g of total RNA from each sample (5 animals per group) to be subsequently amino-allyl labeled and amplified using the Amino Allyl MessageAmp II aRNA Amplification Kit (Ambion, Austin, TX, USA). Aliquots (1.2  g) of amplified aRNA were fluorescently labeled using Cy3/Cy5 (Amersham Biosciences, Buckinghamshire, UK) and then appropriately combined and hybridized to Agilent microarrays. Hybridizations were performed following a reference design, where control samples were pools of RNA from all individual samples. Two hybridizations with fluor reversal (Dye-swap) were performed for each sample. After washing, microarray slides were scanned using a Gene Pix 4100A scanner (Axon Instruments, Union City, CA, USA) and image quantization was performed using the software GenePiX Pro 6.0. Gene expression data for all replicate experiments were analyzed using the GeneSpring GX software version 7.3.1 (Agilent Technologies). Clustering was accomplished with the Gene and Condition Tree algorithms. In addition, Gene Ontology database ( and the KEGG website ( were used in conjunction with GeneSpring ( to identify pathways and functional groups of genes. All microarray data reported are described in accordance with MIAME guidelines ( More information regarding the microarray experiments can be found at the EMBL-European Bioinformatics Institute ( ArrayExpress accession number: E-MEXP-1831). To validate the microarray data, a number of representative differentially expressed genes were selected to be individually studied by Real-Time PCR (7300 Real Time PCR System, Applied Biosystems, Foster City, CA, USA) ( per group) as previously described [19]. Primers and probes were designed using the software Primer Express 2.0 (Applied Biosystems) and purchased from Genosys (Sigma, Madrid, Spain) (Table 1).

GeneGene SymbolGenBank
accesión number
Oligonucleotide sequence (5-3)

Peroxisome proliferator-activated receptor-γ coactivator-1αPgc1aNM_008904Forward: GTCTGAAAGGGCCAAACAGAGA
Forkhead box O1Foxo1NM_019739Forward: GCGGGCTGGAAGAATTCAAT
Muscle atrophy F boxMAFbxNM_026346Forward: CCATCCTGGATTCCAGAAGATTC
Muscle RING finger 1MuRF1NM_001039048Forward: CGCCATGAAGTGATCATGGA

2.4. Statistical Analysis

Data are expressed as error of the mean (SEM). Differences between groups were assessed by Kruskal-Wallis followed by Mann Whitney’s U test. As previously outlined, Gene Ontology groupings were used to identify pathways significantly affected by leptin deficiency as opposed to its administration. Furthermore, statistical comparisons for microarray data to identify differentially expressed genes across different groups were performed using one-way ANOVA and Student’s t-tests as appropriate. Spearman’s correlations were used to evaluate the relations among different variables. All statistical analyses were performed by using the SPSS statistical program version 15.0 for Windows (SPSS, Chicago, IL, USA) and statistical significance was defined as .

3. Results

3.1. Leptin Treatment Improves the Metabolic Profile of ob/ob Mice

The morphological and biochemical characteristics of wild type and ob/ob mice are reported in Table 2. As expected, leptin treatment corrected the obese and diabetic phenotype of ob/ob mice. Body weight was significantly higher ( ) in the control ob/ob group as compared to wild type mice. Leptin-treated mice exhibited a decreased body weight ( ) as compared to control and pair-fed ob/ob animals. Importantly, leptin treatment normalized body weight of ob/ob mice as compared to wild type ( ). In addition, the gastrocnemius of control ob/ob mice exhibited a lower ( ) muscle weight than that of wild type mice and it was increased ( ) by leptin administration in comparison with that of control and pair-fed ob/ob rodents. As depicted in Table 2, higher fasting glucose ( ) and insulin ( ) concentrations were observed in the control ob/ob mice compared to wild types. Although no differences in glucose concentrations were observed in pair-fed as compared to leptin-treated ob/ob mice, higher serum insulin concentrations ( ) were detected in the pair-fed animals than in the leptin-treated ob/ob group. Furthermore, leptin administration normalized both the glucose and insulin levels in ob/ob mice compared to wild types. These data suggest that leptin increases the insulin sensitivity in peripheral tissues, as evidenced by the lower HOMA and higher QUICKI indices ( ) in the leptin-treated in comparison with the control ob/ob animals. Serum glycerol was markedly increased ( ) in the control ob/ob mice, while FFA and TG levels remained unchanged as compared to wild type mice. Interestingly, leptin not only decreased circulating concentrations of FFA ( ) and glycerol ( ) levels as compared to control ob/ob mice, but also FFA ( ), glycerol ( ) and TG ( ) concentrations as compared to pair-fed mice. Leptin administration to ob/ob mice reduced serum glycerol concentrations ( ) and tended to decrease FFA ( ) as compared to wild types. Furthermore, leptin treatment increased the low concentrations of adiponectin of ob/ob mice, but the differences fell out of statistical significance ( ).

wild typecontrol ob/ob pair-fed ob/ob leptin-treated ob/ob

Body weight (g)25.6±0.347.8±4.9b35.7±0.724.7±1.2d,f
Gastrocnemius (mg)142.9±3.490.7±10.0b68.5±1.6104.9±2.6b,f
Gastrocnemius (mg/g)5.59±0.121.91±0.11b1.92±0.074.28±0.15b,d,f
Glucose (mg/dL)149±42430±59a160±24d178±29d
FFA (mmol/L)1.62±0.491.61±0.301.65±0.120.78±0.13c,f
Glycerol (mmol/L)42.8±6.781.6±19.6a39.6±4.9c12.3±4.7a,d,f
TG (mg/dL)122±18169±32151±1086±17e
Insulin (ng/mL)0.42±0.098.60±1.51b2.40±0.68c0.47±0.09d,e
Adiponectin (μg/mL)30.2±3.028.3±5.439.1±1.840.2±3.0
Leptin (ng/mL)1.36±0.42UDUD3.48±1.02

Data are mean±SEM (n=5 per group). Differences between groups were analyzed by Kruskal-Wallis followed by Mann Whitney’s U test. aP<.05 and bP<.01 versus wild type. cP<.05 and dP<.01   versus ob/ob. eP<.05 and fP<.01 versus pair-fed ob/ob. FFA: free fatty acids. TG: triglycerides. UD: undetectable. HOMA: homeostasis model assessment. QUICKI: quantitative insulin sensitivity check index.

Control ob/ob mice exhibited significantly higher serum TBARS than wild type littermates ( ), which were significantly reduced after leptin administration as compared to the control ( ) and pair-fed ( ) ob/ob groups (Figure 1(a)). In addition, leptin decreased ( ) the high concentrations of MDA measured in the gastrocnemius muscle of control ob/ob mice, while this effect was not observed in the pair-fed group (Figure 1(b)). Serum and gastrocnemius TBARS levels were positively associated with body weight, FFA, insulin, and the HOMA index. Oppositely, TBARS levels were negatively associated with adiponectin and the QUICKI index both in serum and muscle. Importantly, a high positive relation were found between serum and gastrocnemius concentrations of TBARS ( , ) (Table 3).


Body weight0.57.0090.46.040

Values are Spearman's correlation coefficients (ρ) and associated P values. TBARS: thiobarbituric acid reactive substances. FFA: free fatty acids. TG: triglycerides. HOMA: homeostasis model assessment. QUICKI: quantitative insulin sensitivity check index.
3.2. Leptin Induces Changes in Gene Expression—Effect of Leptin on Genes Invoved in Oxidative Stress and Inflammation

Differential gene expression profiles in gastrocnemius muscle of wild type and ob/ob groups were compared by microarray analysis. Only genes whose mRNA levels were changed 1.5-fold or higher and identified as significantly changed by statistical analysis were designated as differentially expressed genes. Applying these criteria, microarray data showed that 7,582 genes were differentially expressed by leptin deficiency and leptin administration in ob/ob mice. In particular, leptin deficiency altered the expression of 1,127 genes between wild type and control ob/ob mice. Of these, 580 were upregulated and 547 were downregulated in ob/ob mice. Leptin treatment modified the expression of 1,546 genes in ob/ob mice, upregulating 512 and repressing 1,034. In addition, leptin repressed 736 genes that were upregulated in gastrocnemius muscle of control ob/ob and increased the transcript levels of 846 downregulated genes. Functional enrichment analysis using GeneOntology and KEGG databases revealed that the set of genes with altered expression levels induced by leptin deficiency and administration represents a broad spectrum of biological processes. However, for the purpose of the present paper we focused on the effects of leptin on the set of genes encoding proteins involved in oxidative stress and inflammation. Table 4 shows that leptin deficiency and leptin administration altered the expression of a large number of genes involved in oxidative stress and inflammation. The biological processes mainly affected between control ob/ob mice and wild types included “response to oxidative stress” ( ), “response to stress” ( ) and “acute-phase response” ( ). Furthermore, several processes regulating proliferation, differentiation, and activity of lymphocytes were also significantly affected by leptin deficiency. Importantly, comparison of leptin-treated and control ob/ob groups showed that leptin administration altered the expression of genes implicated in the “positive regulation of lymphocyte activation” ( ), “positive regulation of immune response” ( ) and “response to stress” ( ), as well as genes involved in the “chaperone cofactor dependent protein folding” ( ).

CategoryGenes in Categorywild type vs ob/ob ob/ob vs leptinleptin vs pair-fed
Altered genesP valueAltered genesP valueAltered genesP value

GO:6950: response to stress115661.0031469.018722.0757
GO:6952: defense response101043.18247.510333.83 e-6
GO:6955: immune response83536.18645.165335.53 e-8
GO:45321: immune cell activation2309.47513.2706.0974
GO:46649: lymphocyte activation2089.35913.1706.0673
GO:6954: inflammatory response1994.9384.9842.7590
GO:50776: regulation of immune response1489.09712.04268.00102
GO:6959: humoral immune response1237.1698.2114.0891
GO:42110: T cell activation1125.3967.2635.0191
GO:30098: lymphocyte differentiation1078.04418.1234.0597
GO:42113: B cell activation1013.7247.1883.1610
GO:6800: oxygen and reactive oxygen species metabolism9211.000567.1357.00027
GO:50778: positive regulation of immune response917.050811.003283.6 e-5
GO:51249: regulation of lymphocyte activation897.04610.008085.0076
GO:19882: antigen presentation819.00299.012581.53 e-5
GO:31098: stress-activated protein kinase signaling pathway808.009215.3131.6690
GO:30333: antigen processing7811.00013135.65 e-581.16 e-5
GO:7254: JNK cascade758.006294.4611.6450
GO:46651: lymphocyte proliferation672.7125.1992.2340
GO:6979: response to oxidative stress659.00067.030372.99 e-5
GO:50863: regulation of T cell activation625.07796.06675.0016
GO:7249: I-kappaB kinase/NF-kappaB cascade612.6633.5423.0512
GO:51251: positive regulation of lymphocyte activation586.019610.00035.00118
GO:30217: T cell differentiation545.04816.03804.00638
GO:9266: response to temperature stimulus54124.78 e-7137.96 e-71.5260
GO:30183: B cell differentiation502.5543.4102.1500
GO:50670: regulation of lymphocyte proliferation462.5093.3601.4700
GO:50864: regulation of B cell activation462.5095.06062.1310
GO:42087: cell-mediated immune response441.8091.8762.1220
GO:50777: negative regulation of immune response433.2102.5991.4480
GO:50870: positive regulation of T cell activation435.02036.01375.000294
GO:42088: T-helper 1 type immune response411.7861.8572.1080
GO:9408: response to heat4091.17 e-5121.54 e-71.4240
GO:45619: regulation of lymphocyte differentiation366.001865.02424.00144
GO:42100: B cell proliferation321.6995.01502.0709
GO:19884: antigen presentation, exogenous antigen3191.17 e-697.62 e-686.81 e-9
GO:50851: antigen receptor-mediated signaling pathway301.6763.1601.3390
GO:50871: positive regulation of B cell activation301.6765.01152.0633
GO:51250: negative regulation of lymphocyte activation302.3041.7591.3390
GO:50671: positive regulation of lymphocyte proliferation292.2903.1491.3300
GO:1909: immune cell mediated cytotoxicity272.2622.3583.00584
GO:45580: regulation of T cell differentiation265.002325.006174.00041
GO:30888: regulation of B cell proliferation241.5943.09751.2820
GO:45621: positive regulation of lymphocyte differentiation224.007885.002883.00323
GO:19886: antigen processing, exogenous antigen via MHC class II2192.37 e-882.45 e-681.98 e-10
GO:45058: T cell selection202.1671.6133.00244
GO:50868: negative regulation of T cell activation201.5281.6131.2410
G O:42591: antigen presentation, exogenous antigen via MHC class II1964.42 e-56.00015761.47 e-7
GO:45582: positive regulation of T cell differentiation194.004565.001433.0021
GO:1910: regulation of immune cell mediated cytotoxicity182.1412.2023.00178
GO:19724: B cell mediated immunity181.4911.5741.2200
GO:45577: regulation of B cell differentiation161.4521.5322.0198
GO:46328: regulation of JNK cascade161.4522.1681.1980
GO:30890: positive regulation of B cell proliferation141.4093.02461.1760
GO:45060: negative thymic T cell selection141.4091.4851.1760
GO:51085: chaperone cofactor dependent protein folding132.08094.002343.00066
GO:1912: positive regulation of immune cell mediated cytotoxicity111.3381.4073.00039
GO:48002: antigen presentation, peptide antigen1051.45 e-554.39 e-546.8 e-6
GO:48005: antigen presentation, exogenous peptide antigen751.33 e-654.11 e-641.17 e-6
GO:45620: negative regulation of lymphocyte differentiation62.01841.2481.0794
GO:46330: positive regulation of JNK cascade41.1391.1731.0537
GO:45581: negative regulation of T cell differentiation21.07231.09051.0272

P values reflect the significance of change in prevalence of genes in each category under the leptin deficiency (ob/ob), leptin administration (leptin) and pair-feeding (pair-fed) conditions in ob/ob mice to the expected prevalence of genes in each category. Statistically significant P values are highlighted in bold.

Noteworthy, leptin reduced the expression of several genes related to inflammatory conditions. DNA microarray analysis showed that 86 genes encoding proteins related to defense, stress, and inflammatory responses were altered in the gastrocnemius muscle of control ob/ob mice and modified by leptin administration. Leptin reduced the mRNA levels of various isoforms of the family of heat shock proteins (HSPs) (Dnajc16, Dnaja4, Dnajb4, Hspa2, Hspa4, and Hspb7), metallothioneins (Mt2, Mt4), crystallins (Cryab, Crybb1) and RNA binding proteins (RBMs) (Rbm9, Rbm22) in ob/ob mice (Table 5). In addition, histocompatibility 2, complement component factor B H2-Bf and several genes of the acute-phase response or inflammatory processes, such as kallikrein 5 (Klk5), and serine (or cysteine) proteinase inhibitor clade C member 1 (Serpinc1) and clade B member 1a (Serpinb1a), displayed an increased expression in ob/ob mice that was reduced by leptin administration. On the contrary, gene expression of Cryl1, Hsp105, Rbm5, and H2-Aa were enhanced in ob/ob mice after treated with leptin. Pair-feeding, which accounts for the decrease in food intake that is independent of the direct action of leptin, altered the expression of 1,960 genes, upregulating 984 while downregulating 976 genes. In the context of a food intake reduction as compared to the simple effect due to the caloric restriction, leptin administration further significantly altered the expression of genes involved in processes encompassing “immune response” ( ) “defense response” ( ), “response to oxidative stress” ( ), “positive regulation of T cell activation” ( ) and “positive regulation of immune cell mediated cytotoxicity” ( ) (Table 4). In particular, the gene array analysis provided evidence for elevated Hspa4, Mt4, Crybb1, and Serpinb8 mRNA levels in the pair-fed group as compared to the leptin-treated ob/ob mice (Table 6). On the contrary, leptin increased the gene expression of H2-Ab1 and H2-Eb1 in ob/ob mice. To confirm the microarray data, the mRNA expression of several representative transcripts was analyzed by Real-Time PCR (Figure 2). In this sense, leptin administration reduced the mRNA levels of the muscle atrophy-related transcription factor forkhead box O1 (Foxo1) and of the E3 ubiquitin-ligases muscle atrophy F-box (MAFbx) and muscle RING finger 1 (MuRF1) in leptin-treated ob/ob mice, while no effect of leptin was evidenced on the mRNA levels of the transcriptional coactivator peroxisome proliferator-activated receptor- coactivator-1   (Pgc1 ). The expression of the selected genes was concordant with that of the microarray.

GeneBank NumberGene SymbolGene NameFold changeRatio

Genes downregulated by leptin

NM_007705CirbpCold inducible RNA binding protein1.681.140.68
NM_009964CryabCrystallin, α B1.321.150.87
NM_023695Crybb1Crystallin, β B12.211.390.63
NM_023646Dnaja3DnaJ (Hsp40) homolog, subfamily A, member 30.950.640.67
NM_021422Dnaja4Heat shock protein, DNAJ-like 40.880.300.34
NM_018808Dnajb1DnaJ (Hsp40) homolog, subfamily B, member 10.440.330.74
NM_026400Dnajb11DnaJ (Hsp40) homolog, subfamily B, member 111.110.930.84
NM_027287Dnajb4DnaJ (Hsp40) homolog, subfamily B, member 41.090.600.55
NM_019874Dnajb5DnaJ (Hsp40) homolog, subfamily B, member 51.030.730.72
NM_011847Dnajb6DnaJ (Hsp40) homolog, subfamily B, member 6 isoform c0.700.470.67
NM_013760Dnajb9DnaJ (Hsp40) homolog, subfamily B, member 90.620.390.63
NM_007869Dnajc1DnaJ (Hsp40) homolog, subfamily C, member 10.820.520.63
NM_028873Dnajc14DnaJ (Hsp40) homolog, subfamily C, member 141.120.870.77
NM_172338Dnajc16DnaJ (Hsp40) homolog, subfamily C, member 161.150.660.57
NM_009584Dnajc2DnaJ (Hsp40) homolog, subfamily C, member 21.010.820.81
NM_008929Dnajc3DnaJ (Hsp40) homolog, subfamily C, member 3B1.020.830.82
NM_016775Dnajc5DnaJ (Hsp40) homolog, subfamily C, member 50.740.500.67
NM_010344GsrGlutathione reductase 11.170.710.61
NM_008180GssGlutathione synthetase1.130.880.78
NM_010357Gsta4Glutathione S-transferase, α 41.501.460.97
NM_010362Gsto1Glutathione S-transferase ο 11.421.150.81
NM_008198H2-BfHistocompatibility 2, complement component factor B2.001.440.72
NM_013558Hspa1lHeat shock 70kDa protein 1-like1.601.040.65
NM_008301Hspa2Heat shock protein 21.490.980.65
NM_008300Hspa4Heat shock protein 40.920.300.32
NM_031165Hspa8Heat shock protein 80.910.570.62
NM_010481Hspa9aHeat shock protein 91.030.880.86
NM_024441Hspb2Heat shock protein 21.451.210.83
NM_019960Hspb3Heat shock protein 31.661.270.77
NM_013868Hspb7Heat shock protein family, member 71.830.350.19
NM_008302HspcbHeat shock protein 1, β0.860.690.80
NM_008416JunbJun-B oncogene0.590.360.61
NM_010592Jund1Jun D proto-oncogene1.490.940.63
NM_008456Klk5Kallikrein 52.231.430.64
NM_026346MAFbxMuscle atrophy F box0.650.430.67
NM_008209Mr1Histocompatibility-2 complex class 1-like1.190.980.82
NM_008630Mt2Metallothionein 21.110.500.46
NM_008872PlatPlasminogen activator, tissue1.561.120.72
NM_029397Rbm12RNA binding motif protein 121.401.030.74
NM_026453Rbm13RNA binding motif protein 131.010.870.86
NM_026434Rbm18RNA binding motif protein 180.940.590.63
BC080205Rbm22RNA binding motif protein 221.140.750.66
BC040811Rbm28Rbm28 protein0.690.490.71
NM_172762Rbm34RNA binding motif protein 341.010.670.66
NM_009032Rbm4RNA binding motif protein 41.040.810.78
NM_148930Rbm5RNA binding motif protein 50.690.630.91
NM_144948Rbm7RNA binding motif protein 70.810.740.91
NM_025875Rbm8aRNA binding motif protein 8a0.910.690.76
NM_175387Rbm9RNA binding motif protein 9 isoform 21.960.460.23
NM_025429Serpinb1aSerine (or cysteine) proteinase inhibitor, clade B, member 1a2.732.090.77
NM_080844Serpinc1Serine (or cysteine) proteinase inhibitor, clade C (antithrombin), member 14.981.930.39
NM_008871Serpine1Serine (or cysteine) proteinase inhibitor, clade E, member 12.120.970.46
NM_011340Serpinf1Serine (or cysteine) proteinase inhibitor, clade F, member 12.431.500.62
NM_009776Serping1Serine (or cysteine) proteinase inhibitor, clade G, member 11.411.150.81
NM_009776Serping1Serine (or cysteine) proteinase inhibitor, clade G, member 11.411.150.81
NM_013749Tnfrsf12aTumor necrosis factor receptor superfamily, member 12a0.780.290.37

Genes upregulated by leptin

NM_030004Cryl1Crystallin λ 11.251.721.38
NM_016669CrymCrystallin μ1.371.641.19
NM_133679Cryzl1Crystallin, ζ (quinone reductase)-like
NM_008161Gpx3Glutathione peroxidase 3 isoform 20.470.541.15
NM_024198Gpx7Glutathione peroxidase 71.001.341.33
NM_010359Gstm3Glutathione S-transferase, μ
NM_010360Gstm5Glutathione S-transferase, μ 51.091.391.27
NM_013541Gstp1Glutathione S-transferase, π 10.871.041.20
NM_010361Gstt2Glutathione S-transferase, θ 21.211.701.40
NM_133994Gstt3Glutathione S-transferase, θ 31.531.691.11
NM_010363Gstz1Glutathione transferase zeta 1 (maleylacetoacetate isomerase)
NM_010378H2-AaHistocompatibility 2, class II antigen A, α0.461.262.76
NM_010379H2-Ab1Histocompatibility 2, class II antigen A, β 10.371.042.84
NM_010382H2-Eb1Histocompatibility 2, class II antigen E β0.431.032.40
NM_010395H2-T10Histocompatibility 2, T region locus 101.111.411.27
NM_013559Hsp105Heat shock protein 1050.410.731.79
NM_008303Hspe1Heat shock protein 1 (chaperonin 10)0.670.981.48
AK_052911MuRF1M muscle RING finger
XM_131139Rbm15RNA binding motif protein 150.811.341.66
NM_197993Rbm21RNA binding motif protein 210.670.731.08
BC029079Rbm26Rbm26 protein0.751.191.59
AK087759Rbm27RNA binding motif protein 270.881.191.36
NM_148930Rbm5RNA binding motif protein 50.771.181.55
NM_011251Rbm6RNA binding motif protein 6 isoform a0.800.971.21
NM_207105Rmcs1histocompatibility 2, class II antigen A, β 10.380.892.37
NM_011454Serpinb6bSerine (or cysteine) proteinase inhibitor, clade B, member 6b1.061.231.16
NM_009825Serpinh1Serine (or cysteine) proteinase inhibitor, clade H, member 10.650.991.53
NM_145533SmoxSpermine oxidase0.411.233.00
AK080908Sod1Superoxide dismutase0.580.621.07
NM_011723XdhXanthine dehydrogenase0.681.011.47

Differential expression of genes is indicated as fold changes with respect to the wild type group presenting only the genes which were significantly different (P<.05) between the leptin-treated and the ob/ob groups. Ratio: fold change value for leptin-treated between the ob/ob groups.

GeneBank NumberGene symbolGene nameFold change

Genes downregulated by leptin

NM_023695Crybb1Crystallin, β B10.51
NM_021422Dnaja4Heat shock protein, DNAJ-like 40.63
NM_019739Foxo1Forkhead box O10.34
NM_008300Hspa4Heat shock protein 40.64
NM_013868Hspb7Heat shock protein family, member 70.34
NM_010592Jund1Jun D proto-oncogene0.50
NM_008456Klk5Kallikrein 50.46
NM_008491Lcn2Lipocalin 20.34
NM_008631Mt4Metallothionein 40.63
NM_026346MAFbxMuscle atrophy F box0.37
AK_052911MuRF1M muscle RING finger 10.29
NM_011459Serpinb8Serine (or cysteine) proteinase inhibitor, clade B, member 80.38
NM_011459Serpinb8Serine (or cysteine) proteinase inhibitor, clade B, member 80.59
NM_008871Serpine1Serine (or cysteine) proteinase inhibitor, clade E, member 10.42

Genes upregulated by leptin

NM_009735B2mβ -2-microglobulin1.92
NM_010361Gstt2Glutathione S-transferase, θ 21.94
NM_010379H2-Ab1Histocompatibility 2, class II antigen A, β 14.72
NM_010379H2-Ab1Histocompatibility 2, class II antigen A, β 13.66
NM_010386H2-DMaHistocompatibility 2, class II, locus Dma2.35
NM_010387H2-DMb1Histocompatibility 2, class II, locus Mb13.31
NM_010382H2-Eb1Histocompatibility 2, class II antigen E β4.65
NM_013559Hsp105Heat shock protein 1051,79
AK220167Hspa4MKIAA4025 protein1,59
NM_207105Rmcs1Histocompatibility 2, class II antigen A, β 14.24
NM_207105Rmcs1Histocompatibility 2, class II antigen A, β 14.17
NM_009255Serpine2Serine (or cysteine) proteinase inhibitor, clade E, member 21.53
NM_009825Serpinh1Serine (or cysteine) proteinase inhibitor, clade H, member 12.21
NM_145533SmoxSpermine oxidase4.67

Differential expression of genes is indicated as fold changes presenting only the genes which were significantly different (P<.05) between the leptin-treated and the pair-fed ob/ob groups.

4. Discussion

Obesity is accompanied by a chronic proinflammatory state associated not only with insulin resistance, but also with muscular atrophy [4, 5]. Our study provides evidence that leptin constitutes a negative regulator of oxidative stress and inflammation in the gastrocnemius, which is a representative skeletal muscle of the whole skeletal musculature. This statement is supported by findings reported herein: (a) leptin deficiency is accompanied by systemic and skeletal muscle oxidative stress, muscle inflammation, and reduced muscle mass; (b) systemic and skeletal muscle oxidative stress, muscle atrophy and inflammation of ob/ob mice are reversed by leptin administration independently of the effects of food intake inhibition. Therefore, leptin is able to prevent the muscle atrophy associated with obese and inflammatory states.

Skeletal muscle constitutes an important target for leptin playing a key role on the regulation of lipid and glucose metabolism [20]. Since obese ob/ob mice exhibit an increased oxidative stress and impaired immune response [14, 15] and a reduced skeletal muscle mass [21] compared with their lean littermates, we aimed to identify the genes related to inflammatory processes differentially altered by leptin in the gastrocnemius muscle of obese ob/ob mice. In particular, 86 transcripts encoding inflammation-related proteins were shown to be modified by exogenous leptin administration. However, it has to be taken into account that many of these genes are multifunctional and may have important functions in other biological processes. Among them, leptin repressed the high expression levels of acute-phase reactants and several members of the HSP and RBM families. In addition, confirming a previous study of our group [6], leptin treatment increased the reduced muscle weight of gastrocnemius muscle of ob/ob mice. Taken together, these data suggest that leptin may prevent the obesity-associated inflammatory state and the muscle mass loss related to inflammatory states in leptin-deficient ob/ob mice.

Leptin-deficient ob/ob and leptin receptor-deficient db/db mice display many abnormalities in the immune response similar to those observed in starved animals and malnourished humans [14, 15, 22]. In this respect, exogenous leptin replacement to ob/ob mice modulates T cell responses in mice and prevents starvation-induced immunosuppression, suggesting that lack of leptin is directly involved in these immune system abnormalities [23, 24]. In agreement with these studies, our findings show that leptin deficiency and administration differentially regulate biological processes related to the immune response as well as the T and B cell differentiation and activation in gastrocnemius muscle of ob/ob mice.

Oxidative stress is defined as the imbalanced redox state in which prooxidants overwhelm the antioxidant capacity, resulting in an increased production of reactive oxygen species (ROS), ultimately leading to oxidative damage of cellular macromolecules. The major ROS is the superoxide anion (• ). Dismutation of • by superoxide dismutase (SOD) produces hydrogen peroxide ( ), a more stable ROS, which, in turn, is converted to water by catalase and glutathione peroxidase (GPx) [25]. Oxidative stress is increased in diabetes [26, 27] with leptin administration reportedly improving insulin sensitivity in normal and diabetic rodents [2830]. However, the relationship between leptin and oxidative stress has not been clearly exhibited. Leptin stimulates in vitro ROS production by inflammatory cells [31] and endothelial cells [32] and the level of systemic oxidative stress in nonobese animals [33, 34], suggesting a “prooxidative” role of leptin. However, administration of recombinant leptin reduces the oxidative stress induced by a high-fat diet in mice [35]. In this sense, findings of our study show a high oxidative stress in diabetic ob/ob mice, as reflected by increased TBARS concentrations in serum and the gastrocemius muscle. These observations are in agreement with a large number of studies related to increased plasma TBARS or MDA in diabetic rats [36] and humans [37]. Lipid peroxidation is a common index of free radical mediated injury and induction of antioxidant enzyme is a common cellular response [38]. More importantly, leptin administration decreased serum and gastrocnemius TBARS concentrations as compared to control ob/ob mice, with TBARS levels in gastrocnemius muscle from pair-fed ob/ob animals remaining very similar to those of control ob/ob mice. In this sense, from a molecular perspective, our results further show that transcript levels of Sod1, Gpx3 and glutathione S-transferase 1 Gstp1 are downregulated in control ob/ob mice as compared to wild type controls being upregulated after leptin treatment. Furthermore, leptin administration also upregulated Gpx7, glutathione S-transferase 5 (Gstm5) and glutathione S-transferase 2 (Gstt2). On the contrary, the high expression of catalase (Cat) was repressed by the exogenous injection of leptin to ob/ob mice. These findings are in line with previous observations showing the restoration of the defective antioxidant enzyme activity in plasma of ob/ob mice [39] and humans with a leptin gene mutation [40].

Acute-phase reactants have been suggested to contribute to the maintenance of the chronic low-grade inflammation state involved in the progression of obesity and related diseases [41]. Interestingly, our study provides evidence that genes of the acute-phase response were altered in gastrocnemius muscle of ob/ob mice, which were counteracted by exogenous leptin administration. Leptin reduced the elevated gene expression of tissue-type plasminogen activator (Plat) and lipocalin-2 (Lcn2), which are upregulated in many inflammatory conditions [42, 43], including human obesity [44]. In addition, a pivotal role for oxidative stress in the pathogenesis of muscle wasting in disuse and in a variety of pathological conditions is now being widely recognized [45]. A potential link between oxidative stress and muscle atrophy involves the redox regulation of the proteolytic system [46]. Moreover, various inflammatory cytokines induce oxidative stress [47] and muscle atrophy through the activation of the lysosomal [48, 49] and the ubiquitin-proteolysis system [50]. In this context, biological processes related to oxidative stress and inflammatory responses were altered in the gastrocnemius muscle of ob/ob mice and improved following leptin treatment. In spite of the usual upregulation of the E3 ubiquitin-ligases MAFbx and MuRF1 in most conditions associated with atrophy, their gene expression levels in ob/ob were lower as compared to wild type animals, although no statistically significant differences were observed. Contrarily to what would be expected, leptin administration prevented the increase of both MAFbx and MuRF1 mRNA expression levels induced by pair-feeding in ob/ob mice. A plausible explanation for this surprising finding may relate to the fact that in extreme conditions the energy homeostasis system is overriden whereby leptin is able to inhibit muscular protein degradation associated to food intake reduction. These data are in accordance with a previous study of our group evidencing that leptin replacement inhibits the ubiquitin proteolysis system activity in leptin-deficient mice [6]. Muscle atrophy is associated with increased expression of genes coding for RBM proteins which facilitate the translation, protection, and restoration of native RNA conformations during oxidative stress. It has been suggested that the gene expression of RBM proteins may increase as a compensatory mechanism in response to loss of muscle proteins [51, 52]. Other proteins involved in oxidative stress are metallothioneins, endogenous antioxidants [53] that have been shown to be overexpressed in muscle atrophy in rodents [5456]. In the present work, we have observed that administration of leptin inhibits the gene expression of several members of the RBM (Rbm9, Rbm22) and metallothioneins (Mt2, Mt4) families in the gastrocnemius of ob/ob mice, suggesting that leptin may modulate the inflammatory and oxidative stress responses and consequently, the muscle loss related to inflammatory states.

Genes involved in the chaperone system were also differentially expressed in ob/ob mice as compared to wild types and modified by leptin treatment. HSPs represent a family of molecular chaperones induced in response to cellular stress, responsible for maintaining the structure of proteins and contributing to the repair of damaged or malformed proteins in highly oxidative and lipotoxic conditions. As a result, HSPs are considered antiproteolytic proteins [57]. Muscle atrophy is also associated with an increased gene expression of HSPs [58]. In fact, HSPs are repressed in many rat models of skeletal muscle atrophy [54, 59, 60]. HSP70 is constitutively expressed in skeletal muscle, but its levels are increased in response to oxidative stress [61] with the induction of HSP70 expression by hyperthermia and during inactivity attenuating muscle atrophy [62, 63]. In this regard, a recent study has shown that HSP70 prevents muscle atrophy induced by physical inactivity through inhibition of the muscle atrophy-related transcription factor FoxO3a and the expression of MAFbx and MuRF1 [64]. Among the HSPs, HSP70 and αB-crystallin in particular, are considered negative regulators of muscle cell apoptosis [65, 66] and may inhibit the loss of nuclei taking place during muscle atrophy. In addition, ROS induce the activity of FoxO [67] and gene expression of members of the ubiquitin-proteolysis system in myotubes [68]. In this sense, our results provide evidence that leptin inhibits the increased gene expression of different members of the HSPs (Hspb7, Dnajc16, Hspa4, Cryab, and Crybb1) in the gastrocnemius muscle of ob/ob mice. Taken together, the elevated expression of HSPs in the control and pair-fed ob/ob groups suggests a high defense and stress response in these mice. Moreover, induction of HSPs may confer broader health benefits to patients who are insulin resistant or diabetic [69]. In mammals, caloric restriction has been shown to upregulate HSP induction [70, 71], while expression of HSP72 has been found to be low in skeletal muscle of patients with insulin resistance or type 2 diabetes [72, 73]. Figueiredo et al. [74] have recently shown that leptin downregulates HSP70 gene expression in chicken liver and hypothalamus but not in muscle, which was independent of food intake restriction. On the contrary, Bonior et al. [75] reported an increase in HSP60 gene expression in pancreatic cells by leptin.

Obesity is accompanied by a chronic proinflammatory state resulting in an increase in circulating cytokines and inflammatory markers. In this regard, inflammation produces metabolic alterations in skeletal muscle with both inflammatory response and insulin resistance being associated with muscle mass loss. Findings of our study provide evidence that systemic and skeletal muscle oxidative stress, muscle atrophy and the elevated expression of genes involved in oxidative stress and inflammation of ob/ob mice are reversed by leptin administration. Taken together, these data thereby support that leptin is able to prevent the muscle atrophy associated with obese and inflammatory states in ob/ob mice. Most obese people develop muscle atrophy in spite of exhibiting high leptin circulating concentrations, which may be explained by the leptin resistance present in these patients. Our paper sheds light on the relation between obesity and the loss of muscle mass associated to inflammatory states suggesting that leptin treatment may be an attractive therapeutic approach to prevent muscle loss associated with inflammatory diseases.


The authors would like to thank all the staff of the animal housing facilities for their technical expertise in animal care and handling and, in particular, to Javier Guillén and Juan Percaz. This paper was supported by grants from the Fundación Mutua Madrileña to GF; from the Instituto de Salud Carlos III, Fondo de Investigación Sanitaria (FIS) del Ministerio de Sanidad y Consumo to GF and JG-A; and from the Department of Health of the Gobierno de Navarra of Spain to GF and JG-A. CIBER de Fisiopatología de la Obesidad y Nutrición (CIBEROBN) is an initiative of the Instituto de Salud Carlos III, Spain. The funding bodies had no role in study design, data collection and analysis, decision to publish, or preparation of the paper.


  1. G. Fantuzzi and R. Faggioni, “Leptin in the regulation of immunity, inflammation, and hematopoiesis,” Journal of Leukocyte Biology, vol. 68, no. 4, pp. 437–446, 2000. View at: Google Scholar
  2. C. X. Andersson, B. Gustafson, A. Hammarstedt, S. Hedjazifar, and U. Smith, “Inflamed adipose tissue, insulin resistance and vascular injury,” Diabetes/Metabolism Research and Reviews, vol. 24, no. 8, pp. 595–603, 2008. View at: Publisher Site | Google Scholar
  3. X. Wang, Z. Hu, J. Hu, J. Du, and W. E. Mitch, “Insulin resistance accelerates muscle protein degradation: activation of the ubiquitin-proteasome pathway by defects in muscle cell signaling,” Endocrinology, vol. 147, no. 9, pp. 4160–4168, 2006. View at: Publisher Site | Google Scholar
  4. L. A. Schaap, S. M. F. Pluijm, D. J. H. Deeg, and M. Visser, “Inflammatory markers and loss of muscle mass (Sarcopenia) and strength,” American Journal of Medicine, vol. 119, no. 6, pp. 526–529, 2006. View at: Publisher Site | Google Scholar
  5. S. K. Powers, A. N. Kavazis, and J. M. McClung, “Oxidative stress and disuse muscle atrophy,” Journal of Applied Physiology, vol. 102, no. 6, pp. 2389–2397, 2007. View at: Publisher Site | Google Scholar
  6. N. Sáinz, A. Rodríguez, V. Catalán et al., “Leptin administration favors muscle mass accretion by decreasing FoxO3a and increasing PGC-1α in ob/ob mice,” PLoS ONE, vol. 4, no. 9, article e6808, 2009. View at: Publisher Site | Google Scholar
  7. Y. Zhang, R. Proenca, M. Maffei, M. Barone, L. Leopold, and J. M. Friedman, “Positional cloning of the mouse obese gene and its human homologue,” Nature, vol. 372, no. 6505, pp. 425–432, 1994. View at: Publisher Site | Google Scholar
  8. J. M. Friedman and J. L. Halaas, “Leptin and the regulation of body weight in mammals,” Nature, vol. 395, no. 6704, pp. 763–770, 1998. View at: Publisher Site | Google Scholar
  9. G. Frühbeck and J. Gómez-Ambrosi, “Rationale for the existence of additional adipostatic hormones,” FASEB Journal, vol. 15, no. 11, pp. 1996–2006, 2001. View at: Publisher Site | Google Scholar
  10. M. A. Pelleymounter, M. J. Cullen, M. B. Baker et al., “Effects of the obese gene product on body weight regulation in ob/ob mice,” Science, vol. 269, no. 5223, pp. 540–543, 1995. View at: Google Scholar
  11. M. Otero, R. Lago, F. Lago et al., “Leptin, from fat to inflammation: old questions and new insights,” FEBS Letters, vol. 579, no. 2, pp. 295–301, 2005. View at: Publisher Site | Google Scholar
  12. P. Mancuso, A. Gottschalk, S. M. Phare, M. Peters-Golden, N. W. Lukacs, and G. B. Huffnagle, “Leptin-deficient mice exhibit impaired host defense in Gram-negative pneumonia,” Journal of Immunology, vol. 168, no. 8, pp. 4018–4024, 2002. View at: Google Scholar
  13. S. Loffreda, S. Q. Yang, H. Z. Lin et al., “Leptin regulates proinflammatory immune responses,” FASEB Journal, vol. 12, no. 1, pp. 57–65, 1998. View at: Google Scholar
  14. M. A. Mandel and A. A. F. Mahmoud, “Impairment of cell-mediated immunity in mutation diabetic mice (db/db),” Journal of Immunology, vol. 120, no. 4, pp. 1375–1377, 1978. View at: Google Scholar
  15. R. K. Chandra, “Cell-mediated immunity in genetically obese (C57BL/6J ob/ob) mice,” American Journal of Clinical Nutrition, vol. 33, no. 1, pp. 13–16, 1980. View at: Google Scholar
  16. D. R. Matthews, J. P. Hosker, and A. S. Rudenski, “Homeostasis model assessment: insulin resistance and β-cell function from fasting plasma glucose and insulin concentrations in man,” Diabetologia, vol. 28, no. 7, pp. 412–419, 1985. View at: Google Scholar
  17. A. Katz, S. S. Nambi, K. Mather et al., “Quantitative insulin sensitivity check index: a simple, accurate method for assessing insulin sensitivity in humans,” Journal of Clinical Endocrinology and Metabolism, vol. 85, no. 7, pp. 2402–2410, 2000. View at: Publisher Site | Google Scholar
  18. M. Conti, P. C. Morand, P. Levillain, and A. Lemonnier, “Improved fluorometric determination of malonaldehyde,” Clinical Chemistry, vol. 37, no. 7, pp. 1273–1275, 1991. View at: Google Scholar
  19. V. Catalán, J. Gómez-Ambrosi, F. Rotellar et al., “Validation of endogenous control genes in human adipose tissue: relevance to obesity and obesity-associated type 2 diabetes mellitus,” Hormone and Metabolic Research, vol. 39, no. 7, pp. 495–500, 2007. View at: Publisher Site | Google Scholar
  20. R. B. Ceddia, “Direct metabolic regulation in skeletal muscle and fat tissue by leptin: implications for glucose and fatty acids homeostasis,” International Journal of Obesity, vol. 29, no. 10, pp. 1175–1183, 2005. View at: Publisher Site | Google Scholar
  21. N. Trostler, D. R. Romsos, W. G. Bergen, and G. A. Leveille, “Skeletal muscle accretion and turnover in lean and obese (ob/ob) mice,” Metabolism, vol. 28, no. 9, pp. 928–933, 1979. View at: Google Scholar
  22. G. Matarese, “Leptin and the immune system: how nutritional status influences the immune response,” European Cytokine Network, vol. 11, no. 1, pp. 7–14, 2000. View at: Google Scholar
  23. G. M. Lord, G. Matarese, J. K. Howard, R. J. Baker, S. R. Bloom, and R. I. Lechler, “Leptin modulates the T-cell immune response and reverses starvation—induced immunosuppression,” Nature, vol. 394, no. 6696, pp. 897–901, 1998. View at: Publisher Site | Google Scholar
  24. J. K. Howard, G. M. Lord, G. Matarese et al., “Leptin protects mice from starvation-induced lymphoid atrophy and increases thymic cellularity in ob/ob mice,” Journal of Clinical Investigation, vol. 104, no. 8, pp. 1051–1059, 1999. View at: Google Scholar
  25. A. Fortuño, G. San José, M. U. Moreno, J. Díez, and G. Zalba, “Oxidative stress and vascular remodelling,” Experimental Physiology, vol. 90, no. 4, pp. 457–462, 2005. View at: Publisher Site | Google Scholar
  26. J. V. Hunt, C. C. T. Smith, and S. P. Wolff, “Autoxidative glycosylation and possible involvement of peroxides and free radicals in LDL modification by glucose,” Diabetes, vol. 39, no. 11, pp. 1420–1424, 1990. View at: Google Scholar
  27. C. Feillet-Coudray, E. Rock, C. Coudray et al., “Lipid peroxidation and antioxidant status in experimental diabetes,” Clinica Chimica Acta, vol. 284, no. 1, pp. 31–43, 1999. View at: Publisher Site | Google Scholar
  28. P. Muzzin, R. C. Eisensmith, K. C. Copeland, and S. L. C. Woo, “Correction of obesity and diabetes in genetically obese mice by leptin gene therapy,” Proceedings of the National Academy of Sciences of the United States of America, vol. 93, no. 25, pp. 14804–14808, 1996. View at: Publisher Site | Google Scholar
  29. W. I. Sivitz, S. A. Walsh, D. A. Morgan, M. J. Thomas, and W. G. Haynes, “Effects of leptin on insulin sensitivity in normal rats,” Endocrinology, vol. 138, no. 8, pp. 3395–3401, 1997. View at: Publisher Site | Google Scholar
  30. N. Chinookoswong, J.-L. Wang, and Z.-Q. Shi, “Leptin restores euglycemia and normalizes glucose turnover in insulin- deficient diabetes in the rat,” Diabetes, vol. 48, no. 7, pp. 1487–1492, 1999. View at: Publisher Site | Google Scholar
  31. F. Maingrette and G. Renier, “Leptin increases lipoprotein lipase secretion by macrophages: involvement of oxidative stress and protein kinase C,” Diabetes, vol. 52, no. 8, pp. 2121–2128, 2003. View at: Publisher Site | Google Scholar
  32. A. Bouloumié, T. Marumo, M. Lafontan, and R. Busse, “Leptin induces oxidative stress in human endothelial cells,” FASEB Journal, vol. 13, no. 10, pp. 1231–1238, 1999. View at: Google Scholar
  33. J. Beltowski, G. Wójcicka, A. Marciniak, and A. Jamroz, “Oxidative stress, nitric oxide production, and renal sodium handling in leptin-induced hypertension,” Life Sciences, vol. 74, no. 24, pp. 2987–3000, 2004. View at: Publisher Site | Google Scholar
  34. V. Balasubramaniyan and N. Nalini, “Effect of leptin on peroxidation and antioxidant defense in ethanol-supplemented Mus musculus heart,” Fundamental and Clinical Pharmacology, vol. 21, no. 3, pp. 245–253, 2007. View at: Publisher Site | Google Scholar
  35. J. B. K. Sailaja, V. Balasubramaniyan, and N. Nalini, “Effect of exogenous leptin administration on high fat diet induced oxidative stress,” Pharmazie, vol. 59, no. 6, pp. 475–479, 2004. View at: Google Scholar
  36. S. Gülen and S. Dinçer, “Effects of leptin on oxidative stress in healthy and Streptozotocin-induced diabetic rats,” Molecular and Cellular Biochemistry, vol. 302, no. 1-2, pp. 59–65, 2007. View at: Publisher Site | Google Scholar
  37. R. D. Hoeldtke, K. D. Bryner, D. R. McNeill, S. S. Warehime, K. Van Dyke, and G. Hobbs, “Oxidative stress and insulin requirements in patients with recent-onset type I diabetes,” Journal of Clinical Endocrinology and Metabolism, vol. 88, no. 4, pp. 1624–1628, 2003. View at: Publisher Site | Google Scholar
  38. E. D. Harris, “Regulation of antioxidant enzymes,” FASEB Journal, vol. 6, no. 9, pp. 2675–2683, 1992. View at: Google Scholar
  39. A. M. Watson, S. M. Poloyac, G. Howard, and R. A. Blouin, “Effect of leptin on cytochrome P-450, conjugation, and antioxidant enzymes in the ob/ob mouse,” Drug Metabolism and Disposition, vol. 27, no. 6, pp. 695–700, 1999. View at: Google Scholar
  40. M. Ozata, G. Uckaya, A. Aydin, A. Isimer, and I. C. Ozdemir, “Defective antioxidant defense system in patients with a human leptin gene mutation,” Hormone and Metabolic Research, vol. 32, no. 7, pp. 269–272, 2000. View at: Google Scholar
  41. J. C. Pickup and M. B. Mattock, “Activation of the innate immune system as a predictor of cardiovascular mortality in Type 2 diabetes mellitus,” Diabetic Medicine, vol. 20, no. 9, pp. 723–726, 2003. View at: Publisher Site | Google Scholar
  42. L. Kjeldsen, J. B. Cowland, and N. Borregaard, “Human neutrophil gelatinase-associated lipocalin and homologous proteins in rat and mouse,” Biochimica et Biophysica Acta, vol. 1482, no. 1-2, pp. 272–283, 2000. View at: Publisher Site | Google Scholar
  43. C. Gabay and I. Kushner, “Acute-phase proteins and other systemic responses to inflammation,” The New England Journal of Medicine, vol. 340, no. 6, pp. 448–454, 1999. View at: Publisher Site | Google Scholar
  44. V. Catalán, J. Gómez-Ambrosi, A. Rodríguez et al., “Increased adipose tissue expression of lipocalin-2 in obesity is related to inflammation and matrix metalloproteinase-2 and metalloproteinase-9 activities in humans,” Journal of Molecular Medicine, vol. 87, no. 8, pp. 803–813, 2009. View at: Publisher Site | Google Scholar
  45. J. S. Moylan and M. B. Reid, “Oxidative stress, chronic disease, and muscle wasting,” Muscle and Nerve, vol. 35, no. 4, pp. 411–429, 2007. View at: Publisher Site | Google Scholar
  46. Y.-P. Li, Y. Chen, A. S. Li, and M. B. Reid, “Hydrogen peroxide stimulates ubiquitin-conjugating activity and expression of genes for specific E2 and E3 proteins in skeletal muscle myotubes,” American Journal of Physiology, vol. 285, no. 4, pp. C806–C812, 2003. View at: Google Scholar
  47. P. Matthys and A. Billiau, “Cytokines and cachexia,” Nutrition, vol. 13, no. 9, pp. 763–770, 1997. View at: Publisher Site | Google Scholar
  48. C. Ebisui, T. Tsujinaka, T. Morimoto et al., “Interleukin-6 induces proteolysis by activating intracellular proteases (cathepsins B and L, proteasome) in C2C12 myotubes,” Clinical Science, vol. 89, no. 4, pp. 431–439, 1995. View at: Google Scholar
  49. C. Deval, S. Mordier, C. Obled et al., “Identification of cathepsin L as a differentially expressed message associated with skeletal muscle wasting,” Biochemical Journal, vol. 360, no. 1, pp. 143–150, 2001. View at: Publisher Site | Google Scholar
  50. Y.-P. Li, Y. Chen, J. John et al., “TNF-α acts via p38 MAPK to stimulate expression of the ubiquitin ligase atrogin1/MAFbx in skeletal muscle,” FASEB Journal, vol. 19, no. 3, pp. 362–370, 2005. View at: Publisher Site | Google Scholar
  51. J. St-Amand, K. Okamura, K. Matsumoto, S. Shimizu, and Y. Sogawa, “Characterization of control and immobilized skeletal muscle: an overview from genetic engineering,” FASEB Journal, vol. 15, no. 3, pp. 684–692, 2001. View at: Publisher Site | Google Scholar
  52. M. Wittwer, M. Flück, H. Hoppeler, S. Müller, D. Desplanches, and R. Billeter, “Prolonged unloading of rat soleus muscle causes distinct adaptations of the gene profile,” FASEB Journal, vol. 16, no. 8, pp. 884–886, 2002. View at: Google Scholar
  53. R. Nath, D. Kumar, T. Li, and P. K. Singal, “Metallothioneins, oxidative stress and the cardiovascular system,” Toxicology, vol. 155, no. 1–3, pp. 17–26, 2000. View at: Publisher Site | Google Scholar
  54. E. J. Stevenson, P. G. Giresi, A. Koncarevic, and S. C. Kandarian, “Global analysis of gene expression patterns during disuse atrophy in rat skeletal muscle,” Journal of Physiology, vol. 551, no. 1, pp. 33–48, 2003. View at: Publisher Site | Google Scholar
  55. S. H. Lecker, R. T. Jagoe, A. Gilbert et al., “Multiple types of skeletal muscle atrophy involve a common program of changes in gene expression,” FASEB Journal, vol. 18, no. 1, pp. 39–51, 2004. View at: Publisher Site | Google Scholar
  56. M. L. Urso, P. M. Clarkson, and T. B. Price, “Immobilization effects in young and older adults,” European Journal of Applied Physiology, vol. 96, no. 5, pp. 564–571, 2006. View at: Publisher Site | Google Scholar
  57. R. I. Morimoto, “Cells in stress: transcriptional activation of heat shock genes,” Science, vol. 259, no. 5100, pp. 1409–1410, 1993. View at: Google Scholar
  58. C.-K. Lee, R. G. Klopp, R. Weindruch, and T. A. Prolla, “Gene expression profile of aging and its retardation by caloric restriction,” Science, vol. 285, no. 5432, pp. 1390–1393, 1999. View at: Publisher Site | Google Scholar
  59. J. M. Lawler, W. Song, and H.-B. Kwak, “Differential response of heat shock proteins to hindlimb unloading and reloading in the soleus,” Muscle and Nerve, vol. 33, no. 2, pp. 200–207, 2006. View at: Publisher Site | Google Scholar
  60. J. T. Selsby, S. Rother, S. Tsuda, O. Pracash, J. Quindry, and S. L. Dodd, “Intermittent hyperthermia enhances skeletal muscle regrowth and attenuates oxidative damage following reloading,” Journal of Applied Physiology, vol. 102, no. 4, pp. 1702–1707, 2007. View at: Publisher Site | Google Scholar
  61. Y. Liu, L. Gampert, K. Nething, and J. M. Steinacker, “Response and function of skeletal muscle heat shock protein 70,” Frontiers in Bioscience, vol. 11, no. 3, pp. 2802–2827, 2006. View at: Publisher Site | Google Scholar
  62. H. Naito, S. K. Powers, H. A. Demirel, T. Sugiura, S. L. Dodd, and J. Aoki, “Heat stress attenuates skeletal muscle atrophy in hindlimb-unweighted rats,” Journal of Applied Physiology, vol. 88, no. 1, pp. 359–363, 2000. View at: Google Scholar
  63. J. T. Selsby and S. L. Dodd, “Heat treatment reduces oxidative stress and protects muscle mass during immobilization,” American Journal of Physiology, vol. 289, no. 1, pp. R134–R139, 2005. View at: Publisher Site | Google Scholar
  64. S. M. Senf, S. L. Dodd, J. M. McClung, and A. R. Judge, “Hsp70 overexpression inhibits NF-κB and Foxo3a transcriptional activities and prevents skeletal muscle atrophy,” FASEB Journal, vol. 22, no. 11, pp. 3836–3845, 2008. View at: Publisher Site | Google Scholar
  65. C. Garrido, S. Gurbuxani, L. Ravagnan, and G. Kroemer, “Heat shock proteins: endogenous modulators of apoptotic cell death,” Biochemical and Biophysical Research Communications, vol. 286, no. 3, pp. 433–442, 2001. View at: Publisher Site | Google Scholar
  66. M. C. Kamradt, F. Chen, S. Sam, and V. L. Cryns, “The small heat shock protein αB-crystallin negatively regulates apoptosis during myogenic differentiation by inhibiting caspase-3 activation,” The Journal of Biological Chemistry, vol. 277, no. 41, pp. 38731–38736, 2002. View at: Publisher Site | Google Scholar
  67. T. Nakamura and K. Sakamoto, “Forkhead transcription factor FOXO subfamily is essential for reactive oxygen species-induced apoptosis,” Molecular and Cellular Endocrinology, vol. 281, no. 1-2, pp. 47–55, 2008. View at: Publisher Site | Google Scholar
  68. M. C. C. Gomes-Marcondes and M. J. Tisdale, “Induction of protein catabolism and the ubiquitin-proteasome pathway by mild oxidative stress,” Cancer Letters, vol. 180, no. 1, pp. 69–74, 2002. View at: Publisher Site | Google Scholar
  69. M. F. McCarty, “Induction of heat shock proteins may combat insulin resistance,” Medical Hypotheses, vol. 66, no. 3, pp. 527–534, 2006. View at: Publisher Site | Google Scholar
  70. K. B. Aly, J. L. Pipkin, W. G. Hinson et al., “Chronic caloric restriction induces stress proteins in the hypothalamus of rats,” Mechanisms of Ageing and Development, vol. 76, no. 1, pp. 11–23, 1994. View at: Publisher Site | Google Scholar
  71. A. R. Heydari, S. You, R. Takahashi, A. Gutsmann, K. D. Sarge, and A. Richardson, “Effect of caloric restriction on the expression of heat shock protein 70 and the activation of heat shock transcription factor,” Developmental Genetics, vol. 18, no. 2, pp. 114–124, 1996. View at: Google Scholar
  72. I. Kurucz, A. Morva, A. Vaag et al., “Decreased expression of heat shock protein 72 in skeletal muscle of patients with type 2 diabetes correlates with insulin resistance,” Diabetes, vol. 51, no. 4, pp. 1102–1109, 2002. View at: Google Scholar
  73. C. R. Bruce, A. L. Carey, J. A. Hawley, and M. A. Febbraio, “Intramuscular heat shock protein 72 and heme oxygenase-1 mRNA are reduced in patients with type 2 diabetes: evidence that insulin resistance is associated with a disturbed antioxidant defense mechanism,” Diabetes, vol. 52, no. 9, pp. 2338–2345, 2003. View at: Publisher Site | Google Scholar
  74. D. Figueiredo, A. Gertler, G. Cabello, E. Decuypere, J. Buyse, and S. Dridi, “Leptin downregulates heat shock protein-70 (HSP-70) gene expression in chicken liver and hypothalamus,” Cell and Tissue Research, vol. 329, no. 1, pp. 91–101, 2007. View at: Publisher Site | Google Scholar
  75. J. Bonior, J. Jaworek, S. J. Konturek, and W. W. Pawlik, “Leptin is the modulator of HSP60 gene expression in AR42J cells,” Journal of Physiology and Pharmacology, vol. 57, no. 7, pp. 135–143, 2006. View at: Google Scholar

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