Expression of nitric oxide synthases in subcutaneous adipose tissue of nonobese and obese humans
2000; Elsevier BV; Volume: 41; Issue: 8 Linguagem: Inglês
10.1016/s0022-2275(20)33432-5
ISSN1539-7262
AutoresMontserrat Elizalde, Mikael Rydén, Vanessa van Harmelen, P. Eneroth, Hans Gyllenhammar, Cecilia Holm, Stig Ramel, Anders Ölund, Peter Arner, Kurt Andersson,
Tópico(s)Adipokines, Inflammation, and Metabolic Diseases
ResumoStudies have shown evidence of production of nitric oxide (NO) in adipose tissue, as well as inhibition of lipolysis by NO. We have analyzed nitric oxide synthase (NOS) expression in subcutaneous adipose tissue from 13 nonobese and 18 obese male subjects. Using a competitive reverse transcription polymerase chain reaction method, endothelial (eNOS) and inducible (iNOS), but not neuronal (nNOS), nitric oxide synthase mRNA expression was detected in isolated fat cells and pieces of adipose tissue. Tissue mRNA levels for eNOS were 3,814 ± 825 and 5,956 ± 476 amol/mg RNA (P = 0.043), and for iNOS 306 ± 38 and 332 ± 48 amol/mg RNA, for nonobese and obese individuals, respectively. Western blotting revealed similar eNOS protein levels in isolated fat cells and adipose tissue pieces. Protein levels for eNOS in nonobese and obese individuals, respectively, were (in optical density [OD] units per mm2 per 100 μg of total protein) 0.11 ± 0.08 and 2.80 ± 1.30 (P = 0.043). iNOS protein was detectable, but not measurable, at low levels in a subset of obese patients (3 of 10). iNOS protein levels could not be detected in nonobese individuals. Hormone-sensitive lipase (HSL), the key regulating enzyme in lipolysis, is reduced in obesity. The expression of HSL protein in subcutaneous adipose tissue was studied in the same subset of patients; in agreement with previous results, HSL levels were reduced in obese subjects: 4.64 ± 1.10 and 1.27 ± 0.35 (P = 0.012) in nonobese and obese subjects, respectively. In conclusion, this study shows that eNOS and iNOS, but not nNOS, are present in human subcutaneous adipose tissue. Gene expression and protein levels of eNOS are increased, whereas HSL protein levels are decreased in obesity. It is speculated that increased NO production, preferably by eNOS, and decreased HSL levels may cause decreased subcutaneous adipose tissue lipolysis in obesity. —Elizalde, M., M. Rydén, V. van Harmelen, P. Eneroth, H. Gyllenhammar, C. Holm, S. Ramel, A. Ölund, P. Arner, and K. Andersson. Expression of nitric oxide synthases in subcutaneous adipose tissue of nonobese and obese humans. J. Lipid Res. 2000. 41: 1244–1251. Studies have shown evidence of production of nitric oxide (NO) in adipose tissue, as well as inhibition of lipolysis by NO. We have analyzed nitric oxide synthase (NOS) expression in subcutaneous adipose tissue from 13 nonobese and 18 obese male subjects. Using a competitive reverse transcription polymerase chain reaction method, endothelial (eNOS) and inducible (iNOS), but not neuronal (nNOS), nitric oxide synthase mRNA expression was detected in isolated fat cells and pieces of adipose tissue. Tissue mRNA levels for eNOS were 3,814 ± 825 and 5,956 ± 476 amol/mg RNA (P = 0.043), and for iNOS 306 ± 38 and 332 ± 48 amol/mg RNA, for nonobese and obese individuals, respectively. Western blotting revealed similar eNOS protein levels in isolated fat cells and adipose tissue pieces. Protein levels for eNOS in nonobese and obese individuals, respectively, were (in optical density [OD] units per mm2 per 100 μg of total protein) 0.11 ± 0.08 and 2.80 ± 1.30 (P = 0.043). iNOS protein was detectable, but not measurable, at low levels in a subset of obese patients (3 of 10). iNOS protein levels could not be detected in nonobese individuals. Hormone-sensitive lipase (HSL), the key regulating enzyme in lipolysis, is reduced in obesity. The expression of HSL protein in subcutaneous adipose tissue was studied in the same subset of patients; in agreement with previous results, HSL levels were reduced in obese subjects: 4.64 ± 1.10 and 1.27 ± 0.35 (P = 0.012) in nonobese and obese subjects, respectively. In conclusion, this study shows that eNOS and iNOS, but not nNOS, are present in human subcutaneous adipose tissue. Gene expression and protein levels of eNOS are increased, whereas HSL protein levels are decreased in obesity. It is speculated that increased NO production, preferably by eNOS, and decreased HSL levels may cause decreased subcutaneous adipose tissue lipolysis in obesity. —Elizalde, M., M. Rydén, V. van Harmelen, P. Eneroth, H. Gyllenhammar, C. Holm, S. Ramel, A. Ölund, P. Arner, and K. Andersson. Expression of nitric oxide synthases in subcutaneous adipose tissue of nonobese and obese humans. J. Lipid Res. 2000. 41: 1244–1251. Nitric oxide (NO) is produced in many different cells and is involved in the regulation of such physiologic events as inflammation, vascular tone, and metabolism (see refs. 1Christopherson K.S. Bredt D.S. Perspectives series: nitric oxide and nitric oxide synthases.J. Clin. Invest. 1997; 100: 2424-2429Google Scholar and 2Michel T. Feron O. Nitric oxide synthases: which, how, and why?.J. Clin. Invest. 1997; 100: 2146-2152Google Scholar). Depending on cell type, NO is formed in an enzymatic reaction catalyzed by one of the three isoforms of NO synthase (NOS) (see ref. 2Michel T. Feron O. Nitric oxide synthases: which, how, and why?.J. Clin. Invest. 1997; 100: 2146-2152Google Scholar). Neuronal NOS (nNOS) and endothelial NOS (eNOS) are constitutive, and produce small amounts of NO after stimuli that raise intracellular Ca2+ concentrations. The third isoform, inducible NOS (iNOS), is expressed after induction by agents such as cytokines and bacterial lipopolysaccharide. eNOS and iNOS have been shown to be present in white adipose tissue of the rat (3Ribiere C. Jaubert A.M. Gaudiot N. Sabourault D. Marcus M.L. Boucher J.L. Denis-Henriot D. Giudicelli Y. White adipose tissue nitric oxide synthase: a potential source for NO production.Biochem. Biophys. Res. Commun. 1996; 222: 706-712Google Scholar), suggesting that this tissue may be a potential source of NO production. We have demonstrated the presence of NOS activity and iNOS protein also in human subcutaneous adipose tissue (4Andersson K. Gaudiot N. Ribiere C. Elizalde M. Giudicelli Y. Arner P. A nitric oxide-mediated mechanism regulates lipolysis in human adipose tissue in vivo.Br. J. Pharmacol. 1999; 126: 1639-1645Google Scholar). Moreover, we showed that inhibition of NOS resulted in increased lipolysis in this tissue. Conversely, addition of NO caused an inhibition of lipolysis in intact adipose tissue in vivo, as well as in isolated fat cells in vitro in humans. Together, these results suggest an important role for NO in the regulation of lipolysis in humans (4Andersson K. Gaudiot N. Ribiere C. Elizalde M. Giudicelli Y. Arner P. A nitric oxide-mediated mechanism regulates lipolysis in human adipose tissue in vivo.Br. J. Pharmacol. 1999; 126: 1639-1645Google Scholar). It is still unclear which NOS isoforms are present in human adipose tissue. Work by Ribiere et al. (3Ribiere C. Jaubert A.M. Gaudiot N. Sabourault D. Marcus M.L. Boucher J.L. Denis-Henriot D. Giudicelli Y. White adipose tissue nitric oxide synthase: a potential source for NO production.Biochem. Biophys. Res. Commun. 1996; 222: 706-712Google Scholar), suggests that it is the stimulation of iNOS that substantially contributes to NO production in rodent adipose tissue. It is known from several studies that catecholamine-stimulated lipolysis is reduced in obesity (5Arner P. Regulation of lipolysis in fat cells.Diabetes Rev. 1996; 4: 450-463Google Scholar). However, the exact mechanisms underlying this phenomenon remain unknown, although a number of possible causes have been suggested. Our group demonstrated that expression of hormone-sensitive lipase (HSL), the rate-limiting enzyme in adipocyte lipolysis (6Langin D. Holm C. Lafontan M. Adipocyte hormone-sensitive lipase: a major regulator of lipid metabolism.Proc. Nutr. Soc. 1996; 55: 93-109Google Scholar), is reduced in adipocytes of obese subjects (7Large V. Reynisdottir S. Langin D. Fredby K. Klannemark M. Holm C. Arner P. Decreased expression and function of adipocyte hormone-sensitive lipase in subcutaneous fat cells of obese subjects.J. Lipid Res. 1999; 40: 2059-2066Google Scholar). Molecular mechanisms governing this finding remain to be resolved. Studies of rat isolated fat cells have shown that NO-mediated regulation of lipolysis is complex (8Gaudiot N. Jaubert A-M. Charbonnier E. Sabourault D. Lacasa D. Giudicelli Y. Ribière C. Modulation of white adipose tissue lipolysis by nitric oxide.J. Biol. Chem. 1998; 273: 13475-13481Google Scholar). Thus, the nitric oxide donor 1-propamine 3- (2-hydroxy-2-nitroso-1-propylhydrazine) (PAPA-NONOate) and NO gas have no effect on basal lipolysis but nitrosothiols such as S-nitroso-N-acetyl-penicillamine (SNAP) and S-nitrosglutathione stimulate basal lipolysis in rat adipose tissue. However, SNAP inhibits catecholamine-stimulated lipolysis, possibly by reducing the production of cyclic AMP (cAMP), and PAPA-NONOate and NO gas reduce lipolysis induced by catecholamines as well as other agents without altering cAMP production. Beside the inhibition of cAMP-dependent HSL activation it was suggested in the same study that NO can inactivate HSL by a direct mechanism that is not fully understood (8Gaudiot N. Jaubert A-M. Charbonnier E. Sabourault D. Lacasa D. Giudicelli Y. Ribière C. Modulation of white adipose tissue lipolysis by nitric oxide.J. Biol. Chem. 1998; 273: 13475-13481Google Scholar). It is thus feasible that NO exerts its antilipolytic effect by downregulating lipolytic pathways. A previous study showed an inhibitory mechanism of NO on both basal and stimulated lipolysis in human adipose tissue (4Andersson K. Gaudiot N. Ribiere C. Elizalde M. Giudicelli Y. Arner P. A nitric oxide-mediated mechanism regulates lipolysis in human adipose tissue in vivo.Br. J. Pharmacol. 1999; 126: 1639-1645Google Scholar), indicating important species differences in the action of NO on lipolysis in rats versus humans. In the earlier human study only the presence of iNOS protein was investigated. Here, we investigated more carefully which NOS isoforms are present in human subcutaneous adipose tissue. We also determined whether NOS gene expression and protein levels are changed in the obese state. In view of the previously demonstrated reduction in HSL expression in obese individuals, we also studied whether there was a parallel change in HSL and NOS in obesity. All subjects were male caucasians born in Sweden. The control group (ages 34–68 years) consisted of 13 nonobese men, and the study group (ages 24–57 years) consisted of 18 obese men. These groups took part in the studies of mRNA expression for eNOS, iNOS, and nNOS as well as in measurements of protein levels for eNOS and HSL. The control subjects were undergoing elective surgery for uncomplicated inguinal hernia or gallstone. The obese subjects were undergoing weight reduction surgery with adjustable gastric banding. All subjects were otherwise healthy. Some female subjects were also included for methodological studies, such as development of NOS gene expression and protein levels assays. These women underwent general surgery for nonmalignant disorders such as ovarian cysts or hysterectomy. The study was approved by the Ethics Committee of Karolinska Institutet (Stockholm, Sweden). All subjects gave informed consent to participate in the study. The subjects fasted overnight, and subcutaneous abdominal adipose tissue biopsies (1–2 g) were taken from the surgical wound within 30 min after the start of the operation. Only saline was administered until the tissue pieces were taken. Surgical anesthesia was given as previously described (9Large V. Reynisdottir S. Eleborg L. van Harmelen V. Strommer L. Arner P. Lipolysis in human fat cells obtained under local and general anesthesia.Int. J. Obes. 1997; 21: 78-82Google Scholar). Fat cells were isolated from 5 of the subjects according to the method of Rodbell (10Rodbell M. Metabolism of isolated fat cells.J. Biol. Chem. 1964; 239: 375-380Google Scholar). Peritumoral tissue from human brain was provided by S. Yakisich (Department of Neurology, Huddinge Hospital, Huddinge, Sweden). All tissue pieces were frozen in liquid nitrogen and stored at −70°C for subsequent mRNA analysis. The selected primers were derived from the eNOS and iNOS sequences described by Nadaud et al. (11Nadaud S. Bonnardeaux A. Lathrop M. Soubrier F. Gene structure, polymorphism and mapping of the human endothelial nitric oxide synthase gene.Biochem. Biophys. Res. Commun. 1994; 198: 1027-1033Google Scholar) and Xu et al. (12Xu W. Charles I.G. Liu L. Moncada S. Emson P. Molecular cloning and structural organization of the human inducible nitric oxide synthase gene (NOS2).Biochem. Biophys. Res. Commun. 1996; 219: 784-788Google Scholar), respectively. The sizes of the amplified fragments were 116 base pairs (bp) for eNOS and 114 bp for iNOS. The intervening genomic sequences included an intron. The amplified fragment for nNOS in human brain was 175 bp, and for 18S rRNA (used as a housekeeping gene) was 314 bp, as described by Torczynski et al. (13Torczynski R.M. Fuke M. Bollon A.P. Cloning and sequencing of a human 18S ribosomal RNA gene.DNA. 1985; 4: 283-291Google Scholar) and Hall et al. (14Hall A.V. Antoniou H. Wang Y. Cheung A.H. Arbus A.M. Olson S.L. Lu W.C. Kau C.L. Marsden P.A. Structural organization of the human neuronal nitric oxide synthase gene (NOS1).J. Biol. Chem. 1994; 269: 33082-33090Google Scholar), respectively. The forward primers were 5′-biotin labeled and the reverse primers were 5′-digoxigenin labeled, respectively (Cybergene, Stockholm, Sweden). The used primer sequences were: eNOS-primers Forward primer: 5′-TTG-GCG-GCG-GAA-GAG-GAA-GGA-GT-3′; 23-mer, found in exon 15 position 128–150. Sequence accession X76311 (11Nadaud S. Bonnardeaux A. Lathrop M. Soubrier F. Gene structure, polymorphism and mapping of the human endothelial nitric oxide synthase gene.Biochem. Biophys. Res. Commun. 1994; 198: 1027-1033Google Scholar)Reverse primer: 5′-CAA-AGG-CGC-AGA-AGT-GGG-GGT-ATG-3′; 24-mer, found in exon 16 position 339–316. Sequence accession X76311 (11Nadaud S. Bonnardeaux A. Lathrop M. Soubrier F. Gene structure, polymorphism and mapping of the human endothelial nitric oxide synthase gene.Biochem. Biophys. Res. Commun. 1994; 198: 1027-1033Google Scholar) iNOS-primers Forward primer: 5′-ACG-TGC-GTT-ACT-CCA-CCA-ACA-A-3′; 22-mer, found in exon 7 position 149–170. Sequence accession X85765 (12Xu W. Charles I.G. Liu L. Moncada S. Emson P. Molecular cloning and structural organization of the human inducible nitric oxide synthase gene (NOS2).Biochem. Biophys. Res. Commun. 1996; 219: 784-788Google Scholar)Reverse primer: 5′-CAT-AGC-GGA-TGA-GCT-GAG-CAT-T-3′; 22-mer, found in exon 8 position 216–195. Sequence accession X85766 (12Xu W. Charles I.G. Liu L. Moncada S. Emson P. Molecular cloning and structural organization of the human inducible nitric oxide synthase gene (NOS2).Biochem. Biophys. Res. Commun. 1996; 219: 784-788Google Scholar) nNOS from human brain Forward primer: 5′-CCT-CCC-GCC-CTG-CAC-CAT-CTT-3′; 21-mer, found in exon 22 position 78–98. Sequence accession U17319 (13Torczynski R.M. Fuke M. Bollon A.P. Cloning and sequencing of a human 18S ribosomal RNA gene.DNA. 1985; 4: 283-291Google Scholar)Reverse primer: 5′-CTT-GCC-CCA-TTT-CCA-TTC-CTC-GTA-3′; 24-mer, found in exon 23 position 109–87. Sequence accession U17320 (13Torczynski R.M. Fuke M. Bollon A.P. Cloning and sequencing of a human 18S ribosomal RNA gene.DNA. 1985; 4: 283-291Google Scholar) 18S rRNA Forward primer: 5′-TGC-ATG-TCT-AAG-TAC-GCA-CG-3′, found in position 152–171. Sequence accession M10098 (14Hall A.V. Antoniou H. Wang Y. Cheung A.H. Arbus A.M. Olson S.L. Lu W.C. Kau C.L. Marsden P.A. Structural organization of the human neuronal nitric oxide synthase gene (NOS1).J. Biol. Chem. 1994; 269: 33082-33090Google Scholar)Reverse primer: 5′-TTG-ATA-GGG-CAG-ACG-TTC-GA-3′, found in position 465–446. Sequence accession M10098 (14Hall A.V. Antoniou H. Wang Y. Cheung A.H. Arbus A.M. Olson S.L. Lu W.C. Kau C.L. Marsden P.A. Structural organization of the human neuronal nitric oxide synthase gene (NOS1).J. Biol. Chem. 1994; 269: 33082-33090Google Scholar) An oligonucleotide with the same sequence as the 5′ strand for eNOS (116 bases) and for iNOS (114 bases), containing a 9-base mutation in the middle of the sequence, was synthesized (Cybergene). The standard specific mutations for eNOS were generated at positions 182–190 (wild type, 5′-CAC-CCT-CAG-3′; mutation, 5′-TGG-AAC-AGT-3′ [sequence X76311]). The standard specific mutations for iNOS were generated at positions 155–163 (wild type, 5-CCC-CCA-GCG-3′; mutation, 5′-AAG-GAT-CGC-3′ [sequence X85766]). These synthetic oligonucleotides were used as templates in a first polymerase chain reaction (PCR). The mutated amplicon was amplified with the primers for eNOS and iNOS, respectively. The T7 RNA polymerase promotor was added to the mutated amplicon in a two-step process of ligation and PCR in the 5′ orientation using the Lig'nScribe− RNA polymerase promotor addition kit (Ambion, Austin, TX). This PCR product was used as DNA template in an in vitro transcription reaction to synthesize the RNA internal standard (IS). The 100-μL reaction consisted of 2 μl of DNA template, 40 mm Tris-HCl (pH 8.0, 20°C), 6 mm MgCl2, 10 mm dithiothreitol (DTT), 2 mm spermidine, 2 mm rNTPs, 80 U of RNase inhibitor, and 100 U of T7 RNA polymerase and was incubated overnight at 37°C. The reaction was stopped by heating to 65°C. The DNA template was degraded by incubating it for 5 h with 5 U of RQ1 RNase-free DNase and the RNA was isolated with the RNeasy mini kit from Qiagen (Hilden, Germany) and directly used as IS in the reverse transcriptase-PCR (RT-PCR). For the 18S rRNA, an IS was constructed that shared primer-binding sites with the target RNA, but that contained a fucosyl-transferase (FUC) internal sequence as well as an A20 tail. To construct the IS, a PCR was run with the CDM8 plasmid as template which contains a part of the FUC gene (provided by J. Holgersson, Clinical Immunology, Huddinge Hospital). The following primers were used: The forward primer contained a CGC-GGG clamp, a HindIII site, the 5′ specific primer sequence for 18S rRNA, and a 5′FUC primer sequence. The reverse primer contained a CGC-GGG clamp, an NOT I site, the 3′ specific primer sequence for 18S rRNA, and a 3′FUC primer sequence. The sequences of these primers were as follows (the 5′ and 3′ primer sequences for 18S rRNA are underlined): 18S rRNA -FUC 5′ primer: 5′-CGC-GGG-AAG-CTT-TGC-ATG-TCT-AAG-TAC-GCA-CGG-GAC-AGA-TAC-TTC-AAT-CTC-3′18S rRNA -FUC 3′ primer: 5′-CGC-GGG-GCG-GCC-GCT-TGA-TAG-GGC-AGA-CGT-TCG-AAC-ACG-TCC-ACC-TTG-AGA-T-3′ The PCR product was digested with HindIII and NotI and was ligated into a HindIII/NotI-digested CDM8/poly(A) vector (provided by J. Holgersson; and Lennart Råhlén, KFC, Novum, Huddinge Hospital, Sweden). The digested CDM8/poly(A) vector was treated with calf intestine alkaline phosphatase to prevent reannealing of incompletely digested vector. Competent bacteria (MC1061 with P3) were transformed with the ligated vector and spread on LB–agar plates containing ampicillin and tetracycline. A maxiprep was made and an aliquot of this DNA preparation was linearized with BamHI. The linearized DNA was in vitro transcribed and the DNA template was DNase treated. The RNA product (the 18S rRNA IS) was visualized on a 1% agarose gel and the concentration was determined spectrophotometrically. The size of the 18S IS rRNA PCR product was 250 bp. Total RNA was prepared from 300-mg subcutaneous fat tissue samples or 300-μL cell samples, using the RNeasy minikit (Qiagen). The integrity of the RNA was checked by electrophoresis in a 1% agarose gel containing ethidium bromide. The RNA concentration was measured spectrophotometrically. In the eNOS competitive RT-PCR, 250–500 ng of total sample RNA was reverse transcribed and amplified together with various amounts of IS (1 pg, 100 fg, 50 fg, 10 fg, and 5 fg); in the iNOS competitive RT-PCR, 750–1,000 ng of total RNA was reverse transcribed with 10 fg, 3 fg, 1 fg, and 300 ag of iNOS-IS; and in the 18S rRNA competitive RT-PCR, 2 pg of total RNA was reverse transcribed with 11, 3.8, 1.3, 0.4, 0.1, 0.05, and 0.01 pg of 18S rRNA-IS. RNA, IS, and 3′ primers were preincubated at 70°C for 10 min. The reaction mixture was then added and incubated for 5 min at 25°C, for 5 min at 37°C, and for 10 min at 70°C. The final 15-μL reaction mixture consisted of 50 mm Tris-HCl (pH 8.3), 75 mm KCl, 3 mm MgCl2, 10 mm DTT, 60 U of Moloney murine leukemia virus (Mo-MuLV) reverse transcriptase inhibitor, 8 U of RNasin® ribonuclease inhibitor, 3.4 mm dNTPs and the 3′ specific primer at 1.3 pmol/μL. The reverse transcription process was finished at 10°C. The RT products were used immediately in the PCR. The RT products were amplified by PCR. For iNOS the reaction contained 15 μL of the RT product, 20 mm Tris-HCl (pH 8.4, 20°C), 50 mm KCl, 2 mm MgCl2, 1 mm dNTPs, 2.25 U of Taq polymerase, and 0.8-pmol/μL concentration of each 5′ and 3′ primer in a final volume of 75 μL. The same conditions except another MgCl2 concentration (1.5 mm) were used for the eNOS PCR. For 18S rRNA, the reaction volume of 25 μL contained 5 μL of the RT product, 20 mm Tris-HCl (pH 8.3), 65 mm KCl, 2.1 mm MgCl2, 1.7 mm dNTPs, 1 pmol/μL concentration of each primer, and 0.75 U of Taq polymerase. The PCR program consisted of 2 min at 94°C, 30 cycles of amplification (each cycle consisting of 20 sec at 94°C, 20 sec at the corresponding annealing temperature and 30 sec at 72°C), and a final step of 10 min at 72°C. The annealing temperatures were as follows: eNOS, 60°C; iNOS and 18S rRNA, 55°C. The MJ Research (Watertown, MA) Peltier thermal cycler PTC-200 was used. The PCR products were stored at 4°C and examined on 3% MetaPhor® (FMC Bioproducts, Rockland, ME) agarose gels with ethidium bromide staining to verify the expected size of the amplicons. iNOS and eNOS. The quantification of PCR products was carried out according to Gross et al. (15Gross J. Müller I. Berndt C. Ungethüm U. Heldt J. Elizalde M. Peters L.E. Andersson K. Quantitation of dopamine D2 receptor mRNA in a mesencephalic cell culture using a nonradioactive competitive reverse transcription polymerase chain reaction method.J. Neurosci. Methods. 1998; 82: 187-194Google Scholar). The PCR products were absorbed to streptavidin-coated magnetic particles. The magnetic particles were then heated to 94°C for 5 min and cooled rapidly for strand separation of the PCR products. The supernatants containing digoxigenin-labeled single-stranded amplified DNA were hybridized with specific capture probes immobilized on microtiter plates. Capture probes were synthesized (33-mer oligonucleotides) and cartridge purified (Cybergene). The sequences of the capture probes for the amplicons were as follows (underlined sequence corresponds to the mutation): eNOS-RNA: 5′-AGG-GGC-CCT-GGG-CAC-CCT-CAG-GTT-CTG-TGT-GTT-3′eNOS-IS: 5′-AGG-GGC-CCT-GGG-TGG-AAC-AGT-GTT-CTG-TGT-GTT-3′iNOS-RNA: 5′-CAT-CAC-CGT-GTT-CCC-CCA-GCG-GAG-TGA-TGG-CAA-3′iNOS-IS: 5′-CAT-CAC-CGT-GTT-AAG-GAT-CGC-GAG-TGA-TGG-CAA-3′ The samples were incubated with anti-digoxigenin peroxidase enzyme and spectrophotometrically measured with 2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonate) (ABTS) reagent. The equivalence point was obtained by a two-step titration of the IS. The intersection calculated from the regression curves of log total IS versus log of the absorbance of RNA/IS represents the amount of RNA equivalent. At least 3 concentrations were used in the linear part of the respective competition curve to allow for calculation of the regression lines of the sample and the IS. Regression lines with correlation coefficients above 0.9 were accepted in calculating the equivalence points. A representative example is shown in Fig. 1. 18S rRNA. The PCR products were resolved on a 3% agarose gel and scanned with a charge-coupled device (CCD) camera (DIANA 1; Fuji Film AB, Stockholm, Sweden). The optical density (OD) of bands on the gel was determined with TINA 2.09Gsoftware. The log(OD IS/OD RNA) was plotted as a function of log IS, and the amount of RNA was calculated as described above. Fat tissue specimens from abdominal subcutaneous tissue of obese and nonobese subjects were snap-frozen in liquid nitrogen and stored at −70°C. From 5 subjects, isolated fat cells were prepared according to the method of Rodbell (10Rodbell M. Metabolism of isolated fat cells.J. Biol. Chem. 1964; 239: 375-380Google Scholar) before freezing. On the day of protein isolation from adipose tissue or cells, aliquots ranging from 200 to 600 mg were crushed and lysed in protein lysis buffer (1% Triton X-100, Tris-HCl [pH 7.6], 150 mm NaCl) supplemented with protease inhibitors (Complete; Boehringer Mannheim, Indianapolis, IN), and homogenized with a microtome. The homogenate was centrifuged at 14,000 rpm for 30 min and the infranatant removed and saved. All steps were performed at 4°C to minimize the risk of proteolysis. The protein content in each sample was determined with a kit of reagents from Pierce Biotech (Rockford, IL). One hundred (eNOS and HSL) or 200 (iNOS) μg of total protein was then loaded on polyacrylamide gels and separated by standard sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis. To control for differences in gel migration, exposure time, antibody incubation, and so on, samples from obese and nonobese subjects were run on the same gels and transferred to the same polyvinylidene difluoride (PVDF) membranes (Amersham, Arlington Heights, IL). Blots were blocked overnight in 2.5% nonfat dried milk and probed with antibodies directed against eNOS, iNOS (1:1,000; both from Transduction Laboratories, Lexington, KY) and HSL (1:1,000; provided by C.H.). Positive controls were included in all experiments as provided by the manufacturer (Transduction Laboratories) so as to confirm antibody specificity. Secondary antibodies conjugated to horseradish peroxidase were from Sigma (St. Louis, MO) (eNOS and iNOS, 1:5,000; HSL, 1:8,000). Antigen–antibody complexes were detected by chemiluminescence with a kit of ECL reagents (Amersham) and blots exposed to high-performance chemiluminescence film (Amersham). Films were scanned and the optical density of each specific band analyzed with the ImageMaster program and expressed as OD units per mm2 per 100 μg of total protein. The following chemicals were used: 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide methiodide (EDC methiodide; Sigma-Aldrich, St. Louis, MO); I-Block® (Tropix, Bedford, MA); recombinant RNasin® ribonuclease inhibitor, deoxynucleotide triphosphates (dNTPs), riboxynucleotide triphosphates (rNTPs), and RQ1 RNase-free DNase (Promega, Madison, WI); Mo-MuLV reverse transcriptase, Taq polymerase, and 10× TBE (GIBCO Life Technologies, Gaithersburg, MD); T7 RNA polymerase, RNase inhibitor, DNA molecular weight marker V, streptavidin magnetic particles, anti-digoxigenin-POD Fab fragments, ABTS tablets, enzyme test substrate 1, and blocking reagent (Boehringer Mannheim); primers (Cybergene); MetaPhor® agarose; Nunc-Immuno− module, MaxiSorp−, and F8 module (Nunc, Röskilde, Denmark); DNA sequencing (Cybergene); Lig'nScribe− RNA polymerase promotor addition kit (Ambion). RNA isolation was performed with the RNeasy minikit (Qiagen). Results are presented as means ± SEM. Statistical differences were analyzed by Student's unpaired t-test. P < 0.05 was regarded as a significant difference. mRNA for eNOS and iNOS were detected in adipose tissue, as well as in isolated fat cells (Fig. 2). Specific fragments were amplified for eNOS (116 bp) (Fig. 2A) and iNOS (114 bp) (Fig. 2B). The specificity of the amplified fragments was confirmed by DNA sequencing. In all subjects, an approximately 10-fold higher mRNA expression was observed for eNOS, as compared with iNOS (Table 1). Similar mRNA expression was found for eNOS and iNOS in adipose tissue and isolated fat cells, indicating that at least a substantial part of eNOS and iNOS mRNA derives from the adipocytes themselves; however, we cannot exclude a minor contribution of mRNA levels from stromal cells. Although it was possible to amplify both eNOS and iNOS from isolated fat cells, we were unable to quantify the gene expression in isolated fat cells (i.e., perform a reliablequantitative RT-PCR assay). This was a consistent finding for which at present we have no good explanation.TABLE 1Clinical characteristics and experimental dataNonobeseObeseP ValueAge, years44 ± 3.039 ± 2.4NSBMI, kg/m225.2 ± 0.340.7 ± 1.00.0001eNOS mRNA, amol/mg RNA3814 ± 8255956 ± 4760.024iNOS mRNA, amol/mg RNA306 ± 37332 ± 48NSnNOS mRNA, amol/mg RNANDND18S rRNA mRNA, amol/mg RNA0.11 ± 0.040.11 ± 0.09NSeNOS protein, OD/mm2/100 μg total protein0.11 ± 0.082.80 ± 1.300.043iNOS protein, OD/mm2/100 μg total proteinNDNDHSL protein, OD/mm2/100 μg total protein4.64 ± 1.101.27 ± 0.350.012The values (means ± SEM) were compared using Student's unpaired t-test. Adipose tissue mRNA was measured in 13 nonobese and 18 obese subjects. Adipose tissue protein levels were tested in 11 nonobese and 10 obese individuals. BMI, body mass index; NS, not significantly different; ND, not detectable. Open table in a new tab The values (means ± SEM) were compared using Student's unpaired t-test. Adipose tissue mRNA was measured in 13 nonobese and 18 obese subjects. Adipose tissue protein levels were tested in 11 nonobese and 10 obese individuals. BMI, body mass index; NS, not significantly different; ND, not detectable. Although a specific neuronal NOS fragment (175 bp) (Fig. 2C) could be amplified from human brain tissue, nNOS mRNA could not be amplified in adipose tissue. We were able to amplify a nonspecific band with a length from 167 bp, but sequence analysis showed that this product had no homology with nNOS. Studies (quantitative RT-PCR) performed on intact subcutaneous adipose tissue (Table 1) revealed that eNOS mRNA levels were significantly
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