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• W1992830797 abstract "Treatment of cancer using radiation can be significantly compromised by the development of severe acute and late damage to normal tissue. Treatments that either reduce the risk and severity of damage or that facilitate the healing of radiation injuries are being developed, including autologous adipose tissue grafts to repair tissue defects or involutional disorders that result from tumor resection. Adipose tissue is specialized in energy storage and contains different cell types, including preadipocytes, which could be used for autologous transplantation. It has long been considered a poorly proliferative connective tissue; however, the acute effects of ionizing radiation on adipose tissue have not been investigated. Therefore, the aim of this study was to characterize the alterations induced in adipose tissue by total body irradiation. A severe decrease in proliferating cells, as well as a significant increase in apoptotic cells, was observed in vivo in inguinal fat pads following irradiation. Additionally, irradiation altered the hematopoietic population. Decreases in the proliferation and differentiation capacities of non-hematopoietic progenitors were also observed following irradiation. Together, these data demonstrate that subcutaneous adipose tissue is very sensitive to irradiation, leading to a profound alteration of its developmental potential. This damage could also alter the reconstructive properties of adipose tissue and, therefore, calls into question its use in autologous fat transfer following radiotherapy. Treatment of cancer using radiation can be significantly compromised by the development of severe acute and late damage to normal tissue. Treatments that either reduce the risk and severity of damage or that facilitate the healing of radiation injuries are being developed, including autologous adipose tissue grafts to repair tissue defects or involutional disorders that result from tumor resection. Adipose tissue is specialized in energy storage and contains different cell types, including preadipocytes, which could be used for autologous transplantation. It has long been considered a poorly proliferative connective tissue; however, the acute effects of ionizing radiation on adipose tissue have not been investigated. Therefore, the aim of this study was to characterize the alterations induced in adipose tissue by total body irradiation. A severe decrease in proliferating cells, as well as a significant increase in apoptotic cells, was observed in vivo in inguinal fat pads following irradiation. Additionally, irradiation altered the hematopoietic population. Decreases in the proliferation and differentiation capacities of non-hematopoietic progenitors were also observed following irradiation. Together, these data demonstrate that subcutaneous adipose tissue is very sensitive to irradiation, leading to a profound alteration of its developmental potential. This damage could also alter the reconstructive properties of adipose tissue and, therefore, calls into question its use in autologous fat transfer following radiotherapy. Radiation therapy remains the cornerstone of modern cancer management, with an estimated half of all newly diagnosed cancer patients receiving radiotherapy at some point during the course of their disease. Compared with surgery, radiation therapy has the advantage of being potentially organ-preserving, although the functional outcome might be negatively impacted by the side effects. Indeed, irradiation perturbs the homeostatic network linking parenchymal, mesenchymal, and vascular cells within tissues. Normal communication between cells through soluble, matrix- and cell-associated ligands and receptors is altered, as is an inexorable series of events leading to tissue regeneration and healing.1Barcellos-Hoff MH Park C Wright EG Radiation and the microenvironment- tumorigenesis and therapy.Nat Rev Cancer. 2005; 5: 867-875Crossref PubMed Scopus (392) Google Scholar, 2Bentzen SM Preventing or reducing late side effects of radiation therapy: radiobiology meets molecular pathology.Nat Rev Cancer. 2006; 6: 702-713Crossref PubMed Scopus (727) Google Scholar The use of radiation therapy to treat cancer inevitably involves the exposure of normal tissues that could develop complications. The damage in normal tissues differs depending on the target organ and cell type. Radiation injury is commonly classified into acute, consequential, or late effects, depending on the time before the appearance of symptoms. Acute (early) effects are those that are observed during the course of treatment or within a few weeks following the treatment. Acute radiation damage is most prominent in tissues with rapidly proliferating cells such as the epithelial surfaces of the skin or alimentary tract.3Bergonié J Tribondeau L Interpretation of some results from radiotherapy and an attempt to determine a rational treatment technique. 1906.Yale J Biol Med. 2003; 76: 181-182PubMed Google Scholar, 4Stone HB Coleman CN Anscher MS McBride WH Effects of radiation on normal tissue: consequences and mechanisms.Lancet Oncol. 2003; 4: 529-536Abstract Full Text Full Text PDF PubMed Scopus (682) Google Scholar Ionization events cause damage to vital cellular components, leading to cell death within the first few divisions following irradiation. Radiation also activates various cellular signaling pathways that lead to expression and activation of pro-inflammatory and pro-fibrotic cytokines, vascular injury, and activation of the coagulation cascade.4Stone HB Coleman CN Anscher MS McBride WH Effects of radiation on normal tissue: consequences and mechanisms.Lancet Oncol. 2003; 4: 529-536Abstract Full Text Full Text PDF PubMed Scopus (682) Google Scholar Late reactions occur months to years following radiation exposure and are primarily the result of radiation-dependent depletion of tissue-specific stem cells or progenitors leading to fibrosis, organ dysfunction, and necrosis. In late-responding normal tissues, where cell death is not compensated for by rapid regeneration, this process unfortunately often culminates in the symptomatic complications of radiation exposure.5Brush J Lipnick SL Phillips T Sitko J McDonald JT McBride WH Molecular mechanisms of late normal tissue injury.Semin Radiat Oncol. 2007; 17: 121-130Abstract Full Text Full Text PDF PubMed Scopus (112) Google Scholar, 6Rodemann HP Blaese MA Responses of normal cells to ionizing radiation.Semin Radiat Oncol. 2007; 17: 81-88Abstract Full Text Full Text PDF PubMed Scopus (153) Google Scholar Treatments that reduce the risk or the severity of damage to normal tissue, or that facilitate the healing of radiation injuries, are being developed. These treatments could greatly improve the quality of life of patients treated for cancer. Plastic and reconstructive surgical procedures are thus performed to repair tissue defects or involutional disorders resulting from tumor resection. Different strategies have been used, including the use of autologous tissue transfer of tissues such as fat tissue.7Locke MB de Chalain TM Current practice in autologous fat transplantation: suggested clinical guidelines based on a review of recent literature.Ann Plast Surg. 2008; 60: 98-102Crossref PubMed Scopus (77) Google Scholar Adipose tissue is a highly specialized connective tissue whose primary function is to provide the body with an energy source. The primary cellular component for adipose tissue is a large collection of lipid-filled cells known as adipocytes. Other cellular components contained in adipose tissue are stroma-vascular cells, including endothelial and hematopoietic cells, and preadipocytes.8Caspar-Bauguil S Cousin B Galinier A Segafredo C Nibbelink M André M Casteilla L Pénicaud L Adipose tissues as an ancestral immune organ: site-specific change in obesity.FEBS Lett. 2005; 579: 3487-3492Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar, 9Prunet-Marcassus B Cousin B Caton D André M Pénicaud L Casteilla L From heterogeneity to plasticity in adipose tissues: site-specific differences.Exp Cell Res. 2006; 312: 727-736Crossref PubMed Scopus (229) Google Scholar, 10Mitchell JB McIntosh K Zvonic S Garrett S Floyd ZE Kloster A Di Halvorsen Y Storms RW Goh B Kilroy G Wu X Gimble JM Immunophenotype of human adipose-derived cells: temporal changes in stromal-associated and stem cell-associated markers.Stem Cells. 2006; 24: 376-385Crossref PubMed Scopus (934) Google Scholar, 11Yoshimura K Shigeura T Matsumoto D Sato T Takaki Y Aiba-Kojima E Sato K Inoue K Nagase T Koshima I Gonda K Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates.J Cell Physiol. 2006; 208: 64-76Crossref PubMed Scopus (654) Google Scholar Either preadipocytes or whole subcutaneous pads have been transplanted in patients to restore the volume of tissue lost at defect sites12Gomillion CT Burg KJ Stem cells and adipose tissue engineering.Biomaterials. 2006; 27: 6052-6063Crossref PubMed Scopus (285) Google Scholar or for the treatment of degenerative chronic lesions induced by oncologic radiation.13Moseley TA Zhu M Hedrick MH Adipose-derived stem and progenitor cells as fillers in plastic and reconstructive surgery.Plast Reconstr Surg. 2006; 118: 121S-128SCrossref PubMed Scopus (1) Google Scholar, 14Rigotti G Marchi A Galiè M Baroni G Benati D Krampera M Pasini A Sbarbati A Clinical treatment of radiotherapy tissue damage by lipoaspirate transplant: a healing process mediated by adipose-derived adult stem cells.Plast Reconstr Surg. 2007; 119: 1409-1422Crossref PubMed Scopus (900) Google Scholar The sensitivity of healthy subcutaneous adipose tissue to radiation exposure has, however, never been studied. In other words, it is not known whether irradiated adipose tissue presents healing or reconstructive properties in autologous transplantation therapy, as healthy stromal cells do,15Casteilla L Dani C Adipose tissue-derived cells: from physiology to regenerative medicine.Diabetes Metab. 2006; 32: 393-401Abstract Full Text PDF PubMed Scopus (71) Google Scholar or if irradiation of adipose tissue may be an issue for the patients who undergo total body radiotherapy. Therefore, the aim of this study was to determine the characteristics of subcutaneous adipose tissue isolated from mice after total body irradiation (TBI). Proliferation and apoptosis were quantified in vivo. Phenotypic analysis of the stroma-vascular fraction was performed, and proliferation and differentiation potentials of progenitor cells were evaluated in vitro. For adipose tissue digestion, bovine serum albumin and collagenase were purchased from Sigma Aldrich (St. Quentin Fallavier, France). Culture medium and newborn calf serum were provided by Invitrogen (Cergy-Pontoise, France). The methylcellulose used was Methocult M3534 (StemCell Technologies, Vancouver). For fluorescence-activated cell sorter (FACS) analysis, we used directly conjugated primary murine monoclonal antibodies (all BD Biosciences, Heidelberg, Germany, unless indicated) against mouse CD34-fluorescein isothiocyanate (FITC) or -phycoerythrin (clone RAM34), mouse CD45-peridinin chlorophyll-a protein (PerCP) (clone 30-F11), mouse CD90-allophycocyanin (clone 53–2.1), mouse Ly-6A/E (Sca-I)-phycoerythrin (clone D7), and mouse CD31-allophycocyanin (clone MEC13.3) We used directly conjugated rat immunoglobulin (BD Biosciences) for isotype controls. Lethal (10Gy) or sublethal (7Gy) irradiation, given in one dose (BIOBEAM 8000, Cs137, 4,4 Gy/min), was performed on 5- to 6-week-old C57Bl/6 mice (Harlan, France). Animals were housed in a controlled environment (12 hours light/dark cycle at 21°C), and maintained on acidified water and autoclaved food for 7 days. At the end of the experiments, the mice were euthanized by cervical dislocation under CO2 anesthesia. Blood samples were immediately drawn and tissues were quickly removed and processed for analyses as described. Before and 7 days after irradiation, 200 μl of peripheral blood was collected from the retro-orbital plexus and immediately transferred into vials containing heparin. Blood cell counts were performed automatically using a hematology analyzer. The differential of the nucleated cells was determined automatically by the analyzer (Micros OT 60 ABX, No. 21CFR864.5220 Montpellier, France). Subcutaneous adipose tissue was removed, fixed on 95% ethanol, and embedded in paraffin. Sections were deparaffinized in xylene, and rehydrated in descending grades (100% to 50%) of ethanol with a final wash in tris-buffered saline (TBS). Slides were then either stained with May-Grünwald Giemsa or processed for Ki-67 staining or terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) analysis. Slides stained with May-Grünwald Giemsa were then used for determination of mature adipocyte size, by using image J software (NIH, USA). For Ki-67 staining, sections were incubated in 10 mmol/L citrate buffer (pH 6.0), for antigen retrieval. Endogenous peroxidase activity was removed by incubating with H2O2, 3% in TBS for 15 minutes. Ki-67 protein was detected using a 1:25 dilution of rat monoclonal antibody (Clone TEC3. Dakocytomation, Denmark). Histofine simple stain mouse “MAX PO” anti-rat (Microm, Francheville, France) was used as a secondary antibody. Finally, sections were stained using 3-Amino-9-ethylcarbazole (AEC), and counterstained using hematoxylin. Control experiments were performed using purified rat IgG. Resulting immunostaining was observed using an optical microscope (DMRB Leica) and quantified using Visilog 6.3 image analyzer software (Noesis, France). For each group of three mice, 15 fields were analyzed. Apoptotic cells were detected with a commercial in situ cell-death detection kit, POD (Roche DIAGNOTICS, Mannheim, Germany) according to the manufacturer's protocol. Slides were washed in TBS and incubated in 20 mmol/L citrate buffer (pH 6.0) under 750W microwave irradiation for 1 minute. Non-specific sites were blocked with Tris-HCl buffer (100 mmol/L Tris-HCl, 3% bovine serum albumin, 10% newborn calf serum) for 30 minutes. The tissue section was covered with 50 μl of TUNEL reaction mixture containing dUTP-fluorescein (2′-deoxyuridine 5′-triphosphate) supplemented with TdT (terminal deoxynucleotidyl transferase) for 60 minutes at 37°C. After three washes with TBS, endogenous peroxidase activity was removed by incubating with H2O2, 3% in TBS for 10 minutes. Non-specific sites were blocked again and 50 μl of Convert-peroxidase (Roche Diagnostics, Mannheim, Germany) diluted at 1:2 in Tris-HCl buffer was applied to each slide for 30 minutes at 37°C. Finally sections were stained using 3,3′Diaminobenzidine (DAB) and counterstained using hematoxylin. Control experiments were performed using dUTP-fluorescein without TdT. Staining was observed under an optical microscope (DMRB Leica) and quantified using Visilog 6.3 image analyzer software (Noesis, France). For each group of three mice, 15 fields were analyzed. For PCR analysis, subcutaneous adipose tissue was excised and stored at −80°C. Total RNA from tissue was prepared using RNAeasy Lipid Tissue Mini Kit (Qiagen, France) according to the manufacturer's recommendations, and 1 μg was reversed transcribed with the high-capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA, USA) in the presence of random primers. Quantitative PCR was performed by monitoring in real time the increase in fluorescence of the SYBR Green dye on an ABI PRISM 7000 Sequence Detector System (Applied Biosystems) according to the manufacturer's instructions. The nucleotide sequences of the PCR primers used were as follows: 36B4: Forward (5′–AGTCGGAGGAATCAGATGAGGAT-3′), Reverse (5′- GGCTGACTTGGTTGCTTTGG-3′); superoxide dismutase (MnSOD): Forward (5′-CTTACAGATTGCTGCCTGCTCTAA-3′), Reverse (5′-AATCCCCAGCAGCGGAAT-3′); nicotinamide-adenine dinucleotide phosphate oxidase (NADPH)ox: Forward (5′-ACCAAATGTTGGGCCTAGGAT-3′), Reverse (5′- AAAA GGATGAGGCTGCAGTTGA-3′). Each PCR was performed in duplicate in a 25 μl volume using SYBR Green I Master Mix Plus (Applied Biosystems), 0.3 μmol/L of each primer, for 15 minutes at 95°C for initial denaturing step, followed by 40 cycles of 95°C for 30 seconds, and 60°C for 30 seconds in the ABI Prism 7000 sequence Detector System. To exclude contamination by nonspecific PCR products such as primer dimers, melting curve analysis was applied to all final PCR products after the cycling protocols. Values for each gene were normalized to the expression levels of the 36B4 mRNA. Relative quantification of studied genes was calculated by using δCt formula, as recommended by the manufacturer (Applied Biosystems). Results were expressed relative to the control condition, which was arbitrary assigned a value of 1. Aconitase enzymatic activity was measured using a Bioxytech aconitase-340 assay kit (OxisResearch, Foster City, CA, USA) on tissue homogenates. The assay is based on measurement of concomitant formation of NADPH from NADP+ when isocitrate (produced by aconitase) is decarboxylated by isocitrate dehydrogenase. Briefly, around 50 mg inguinal adipose tissue removed from control, sublethally, or lethally irradiated mice was minced and homogenized in 500 μl assay buffer provided by the kit with tissue lyser beads (Qiagen, France). After elimination of lipids by 10 minutes centrifugation at 380g at 4°C, the enzymatic reaction was started by mixing 200 μl of homogenate (about 2 mg proteins/ml) with 200 μl citrate, 200 μl isocitrate dehydrogenase, and 200 μl NADP+. Absorbance was recorded during 40 minutes at 340 nm at 37°C. Then, the slope was estimated in the linear part of the curve, and aconitase activity was calculated according to the supplier's instructions with normalization to the protein concentration of the sample measured by Biorad DC protein assay (BioRad, Marne la Coquette, France). Cells were isolated according to Björntorp et al with minor modifications.16Björntorp P Karlsson M Pertoft H Pettersson P Sjöström L Smith U Isolation and characterization of cells from rat adipose tissue developing into adipocytes.J Lipid Res. 1978; 19: 316-324Abstract Full Text PDF PubMed Google Scholar Inguinal subcutaneous adipose tissue was dissected from visible blood vessels and ganglions and was digested at 37°C in phosphate buffer PBS containing 0.2% bovine serum albumin and 2 mg/ml collagenase for 30 minutes (collagenase A, Roche Diagnostics, Meylan, France). After elimination of undigested fragments by filtration through 25 μm filters, mature adipocytes were separated from the pellets of the stroma-vascular fraction (SVF) by centrifugation (600 × g, 10 minutes). SVF cells were incubated for 5 minutes in hemolysis buffer (140 mmol/L NH4Cl and 20 mmol/L Tris, pH 7.6) to eliminate red blood cells and washed by centrifugation in PBS. The number of isolated SVF cells was then counted and their viability assessed by trypan blue exclusion. SVF cells were either used for flow cytometry analyses or plated in vitro. Mature adipocytes were directly counted on thoma grid. Isolated cells were analyzed by flow cytometry (FACS) using the following protocol. Freshly-isolated SVF cells were stained in staining buffer consisting of phosphate-buffered saline containing 0.5% new calf serum and FcR Block reagent (StemCell Technologies, Vancouver, Canada). Cells were incubated with anti-mouse monoclonal antibodies (mAb) or rat immunoglobulins (isotypes) conjugated with FITC, phycoerythrin, PerCP, or allophycocyanin for 30 minutes at 4°C. Triple or quadruple staining was performed by incubating cells with FITC, phycoerythrin, PerCP, and allophycocyanin-conjugated primary antibodies in one step, to precisely characterize the different cell populations. Cells were washed in staining buffer and then analyzed on a FACS (FACS Calibur, Becton Dickinson, Mountain View, CA). Data acquisition and analysis were then performed (Cell Quest Pro software, Becton Dickinson). To evaluate the frequency of mesenchymal-like progenitors in the inguinal SVF, cells were cultured in T-25 flasks at a final concentration of 16 or 32 cells/cm2 in Dulbeco's Modified Eagles Medium (DMEM) F12 10% newborn calf serum and incubated at 37°C, 5% CO2. The medium was renewed every 2 days. After 14 days, the cells were washed with PBS and fixed with methanol for 15 minutes. For scoring the colony forming unit-fibroblasts (CFU-f), flasks were stained with Giemsa 6% for 30 minutes. Plates were scored under an optical microscope, and colonies were considered aggregates of more than 50 cells. For adipogenic differentiation, cells were plated at a density of 8000 cells/cm2 in DMEM:F12 supplemented with 10% newborn calf serum, biotin (16 μmol/L), panthotenic acid (18 μmol/L), ascorbic acid (100 μmol/L), and amphotericin (25 μg/ml), streptomycin (10 mg/ml), and penicillin (10000 U/ml). At confluence, adipogenic differentiation was induced by adding dexamethasone (33 mmol/L), insulin (2 nmol/L), 3, 3′, 5-tri-iodo-l-thyronin (T3; 2 nmol/L) and transferrin (10 μg/ml) for 10 days. Medium was changed every 2 days. Adipocytes were characterized by oil red O staining. Differentiation was quantified by cellular triglyceride (TG) content measurement by using a commercially available test combination (Triglycerides enzymatiques PAP 150, Biomerieux) after cell lysis in 0.1N NaOH. The TG content was calculated per μg of proteins. The protein content was determined by using the DC Protein Assay Kit (BioRad, Marne la Coquette, France). SVF cells from each fat pad were isolated as described above and plated at 7 × 103 cells/ml in 1.5 ml of methylcellulose. Colony number was assessed at day 21 after plating. Adipocyte colonies were identified as lipid droplet-containing cells. All statistical analyses were done by unpaired t-test using Prism software (GraphPad software, San Diego, CA). Results are expressed as mean ± SEM. Total body irradiation induces changes in blood and tissue cell-composition, depending on the dose of ionizing agents and on the sensitivity of the tissue considered. The sensitivity of adipose tissue to irradiation was investigated 7 days after sublethal (7Gy) or lethal (10Gy) total body irradiation. Blood numeration was used as a control of the irradiation efficiency. As expected, leukocyte number was severely decreased by day 7 in lethally irradiated mice (5.62 ± 0.86% of control values), although sublethal irradiation induced a slight and non-significant decrease in leukocyte blood cell count (75.61 ± 14% of control values; Table 1).Table 1Physiological Parameters and Characteristics of Inguinal Adipose Tissue 7 days after Sublethal or Lethal Total Body IrradiationControlSublethal IrradiationLethal IrradiationMice weight (g)22.14 ± 0.3920.44 ± 0.18**P < 0.01;18.57 ± 0.18***P < 0.001 in irradiated versus control mice.Leucocyte blood count (103/μl)3.6 ± 0.293.42 ± 0.510.125 ± 0.01***P < 0.001 in irradiated versus control mice.Fat pad weight (g)0.231 ± 0.0140.165 ± 0.003**P < 0.01;0.15 ± 0.016 **P < 0.01;Mature adipocyte number (×106)/mice0.577 ± 0.0780.32 ± 0.03**P < 0.01;0.360 ± 0.007*P < 0.05;Mature adipocyte mean diameter (μm)65.63 ± 3.2239.38 ± 1.93***P < 0.001 in irradiated versus control mice.22.26 ± 1.57***P < 0.001 in irradiated versus control mice.* P < 0.05;** P < 0.01;*** P < 0.001 in irradiated versus control mice. Open table in a new tab Seven days after irradiation, mice weight was significantly decreased, by 8 and 16%, in sublethally and lethally irradiated mice, respectively, as compared with controls (Table 1). The subcutaneous (inguinal) fat pad weight was significantly decreased in irradiated mice compared with controls, regardless the dose of irradiation (Table 1). Additionally, inguinal fat pads removed from irradiated mice presented morphological changes as revealed by May-Grünwald Giemsa coloration (Figure 1). Indeed, 7 days after irradiation, the size of mature adipocytes was reduced (Table 1, Figure 1D), and the presence of non-adipose cells was more apparent (Figure 1). These modifications were emphasized after lethal irradiation compared to sublethal irradiation. Indeed, the number of small adipocytes (diameter <50 μm) was increased by up to five in inguinal fat pads removed from irradiated mice as compared with controls (Figure 1D). In contrast, no large mature adipocytes (diameter >100 μm) were observed in irradiated samples, although they represented 11.02 ± 3.62% of total adipocytes in control mice (Figure 1D). In addition to decreased adipocyte size, the total number of mature adipocytes per fat pad was also decreased in irradiated mice compared with controls, and this decrease was not irradiation dose-dependent (Table 1). The presence of numerous small mature adipocytes in inguinal fat pads after irradiation compared to controls induced an apparent increase of adipocyte number, as appears in Figure 1. However, the total adipocyte number has to take into account the fat pad weight, which is severely decreased in irradiated animals. The total number of adipocytes in fat pads after irradiation is thus significantly lower than in control animals. We next investigated the cellular mechanisms involved in the effects of radiation in adipose tissue. Inguinal fat pads isolated from either control, sublethally, or lethally irradiated mice were processed for immunohistochemistry 7 days after irradiation and were stained with an antibody against the nuclear antigen Ki-67 to detect proliferating cells (Figure 2, A–C) or processed for TUNEL analysis to quantify apoptotic cells (Figure 2, D–F). Quantification of immunostaining demonstrated the presence of 3.9 ± 0.7% of proliferating cells in control adipose tissue (Figure 2A). Irradiation induced a total loss of these proliferating cells at sublethal and lethal doses (Figure 2, B,C). Control adipose tissue also harbored numerous apoptotic cells (5.5 ± 1.3 × 103 nuclei/cm2; Figure 2D). This percentage was significantly increased after sublethal (11.18 ± 1.1 × 103 nuclei/cm2; factor 2.01 ± 0.20; Figure 2E) or lethal (25.79 ± 5.4 × 103 nuclei/cm2; factor 4.65 ± 0.98; Figure 2F) irradiation. Most of the apoptotic cells were SVF cells, localized between mature adipocytes (Figure 2, D–F, black arrows). However, some mature adipocytes were also positive for TUNEL (Figure 2, D–F, red arrow). We thus showed that control adipose tissue harbored both proliferating and apoptotic cells, suggesting a high cell dynamic. In addition, ionizing radiation significantly altered this parameter in subcutaneous adipose tissue. To investigate whether oxidative stress and especially radical oxygen species (ROS) production may lead to irradiation damage in adipose tissue, we measured by quantitative reverse transcription PCR the expression level of genes encoding ROS metabolism enzymes and quantified aconitase activity, considered to be a marker of oxidative stress.17Bota DA Davies KJA Lon protease preferentially degrades oxidized mitochondrial aconitase by an ATP-stimulated mechanism.Nat Cell Biol. 2002; 4: 674-680Crossref PubMed Scopus (460) Google Scholar NADPH oxidase (NADPHox) is a major source of ROS in various cells including adipose derived cells,18Furukawa S Fujita T Shimabukuro M Iwaki M Yamada Y Nakajima Y Nakayama O Makishima M Matsuda M Shimomura I Increased oxidative stress in obesity and its impact on metabolic syndrome.J Clin Invest. 2004; 114: 1752-1761Crossref PubMed Scopus (4024) Google Scholar and its expression was significantly increased after lethal irradiation, although a sublethal dose gives rise to a slight but insignificant rise in its expression level (Figure 3A). In parallel, we measured the expression of an antioxidant enzyme, MnSOD, and showed that its expression level was reduced in adipose tissue of irradiated mice (Figure 3A). These results suggest that irradiation leads to an increase in ROS production via the activated NADPHox pathway and impaired antioxidant defense system. To confirm the presence of oxidative stress in adipose tissue following ionizing radiation, we quantified aconitase activity. Aconitase is an essential mitochondrial enzyme known to be particularly susceptible to oxidative damage.17Bota DA Davies KJA Lon protease preferentially degrades oxidized mitochondrial aconitase by an ATP-stimulated mechanism.Nat Cell Biol. 2002; 4: 674-680Crossref PubMed Scopus (460) Google Scholar Similarly to MnSOD expression, aconitase activity was reduced in adipose tissue after irradiation, and the magnitude of the decrease was dependent on the dose of radiation (Figure 3B). This result thus confirms the presence of oxidative stress in adipose tissue after irradiation, particularly after lethal irradiation. To analyze the potential changes at a cellular level in adipose tissue following irradiation, we focused on lethal irradiation and analyzed the proportion and phenotype of the different cell subsets present in SVF. We next investigated the proliferation and differentiation potentials of progenitors. Flow cytometry was performed by using different cell surface markers previously shown to be expressed in SVF and allowed the identification of cell subsets. CD45 was used to discriminate between hematopoietic and non-hematopoietic cells present in SVF. As shown in representative dot plots, the percentage of CD45-positive hematopoietic cells in the inguinal SVF (region R1) was significantly reduced (two fold) 7 days after lethal irradiation (Figure 4). As already described,9Prunet-Marcassus B Cousin B Caton D André M Pénicaud L Casteilla L From heterogeneity to plasticity in adipose tissues: site-specific differences.Exp Cell Res. 2006; 312: 727-736Crossref PubMed Scopus (229) Google Scholar a large majority of CD45-negative (non-hematopoietic) cells present in control adipose tissue expressed CD90, CD34, and ScaI (Fig" @default.
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• W1992830797 date "2009-01-01" @default.
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• W1992830797 title "Adipose Tissue Sensitivity to Radiation Exposure" @default.
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