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News Article | February 15, 2017

No statistical methods were used to predetermine sample size. Affinity-purified antibodies against total and Ser886 and Ser999 phospho-sites of EPRS were generated as described7, 31. Antibody against phospho-Ser was from Meridian Life Science. Antibodies specific for the C terminus of S6K1 and N terminus of S6K2 were purchased from Abcam and LifeSpan, respectively. Antibodies against PKA, DMPK, PKN, GAPDH, caveolin1, CD36/FAT, GLUT4, His-tag, β-actin and FATP1, FATP3, and FATP4 were from Santa Cruz. Antibody specific for FABP4 and FABP5 were from R&D and for FABPpm/GOT2 was from GeneTex. All other antibodies and rapamycin were from Cell Signaling. SignalSilence siRNAs targeting RSK1, AKT and S6K1 were from Cell Signaling, and those targeting raptor and rictor were from Santa Cruz. The 3′-UTR-specific duplex siRNAs, 5′-UGAUACGAAGAUCUUCUCAG-3′ and 5′-GCCUAAAUUAACAGUGGAA-3′, targeting mouse EPRS were from Origene. Smart pool siRNA targeting the coding sequence of mouse FATP1 (SLC27a1) was from Dharmacon and 3′-UTR-specific trilencer siRNA targeting human S6K1 was from Origene. Recombinant wild-type and Ser-to-Ala (S886A and S999A) mutant His-tagged linker proteins spanning Pro683 to Asn1023 of human EPRS were expressed and purified as described7, 8. Recombinant active S6K1 (ref. 32) and RSK1–3 were from Cell Signaling; Akt1 and Akt2 were from EMD Millipore. Mouse EPRS domains ERS (Met1 to Gln682), linker (Pro683 to Asn1023), and PRS (Leu1024 to Tyr1512) were cloned into pcDNA3 vector with an N terminus Flag tag using full-length mouse EPRS cDNA (Origene) as template. Flag-tagged mouse wild-type linker and linker with Ser999-to-Ala (S999A) and Ser999-to-Asp (S999D) mutations were generated as described33. Full-length human S6K1 cDNA in pCMV6-entry vector was purchased from Origene and recloned, deleting the 23-amino acid N terminus nuclear localization signal, and adding an in-frame upstream 6-His tag and a downstream Myc tag in pcDNA3. Specific Thr389-to-Ala (T389A) and Thr389-to-Glu (T389E) mutations were introduced using primers with the desired mutation and GENEART Site-Directed Mutagenesis System (Invitrogen). Human U937 monocytic cells (CRL 1593.2; ATCC authenticated by STR DNA profiling) were cultured in RPMI 1640 medium and 10% fetal bovine serum (FBS) with penicillin and streptomycin at 37 °C in 5% CO . Bone-marrow-derived macrophages (BMDM) were flushed from femur and tibia marrows of S6K1−/−S6K2−/−, and double-knockout S6K1−/−S6K2−/− mice (from G. Thomas and S. Kozma), and then cultured for one week in RPMI 1640 medium containing 10% FBS and 20% L929 cell-conditioned medium at 37 °C in 5% CO . 1 × 107 cells were treated with 500 U ml−1 IFNγ (R&D) for up to 24 h, as described previously34, 35. 3T3-L1 fibroblasts (CL-173; ATCC-certified) were cultured in high glucose containing Dulbecco’s modified Eagle’s medium (DMEM), 10% FBS and antibiotics/antimycotic at 37 °C in 10% CO to near 75% confluence. Confluent fibroblasts were induced to differentiate in medium containing DMEM and 10% FBS supplemented with 1× solutions of insulin:dexamethasone:3-isobutyl-1-methylxanthine (Cayman). After 72 h, the medium was replaced with 10% FBS and DMEM containing only insulin and maintained for a week with 3 changes in the same medium. Adipocytes were maintained in DMEM medium with 10% calf serum and antibiotics/antimycotic for at least 3 d before utilization. Differentiated adipocytes were serum-deprived for 4 h followed by treatment with 100 nM insulin (Sigma-Aldrich) for 4 h, or as indicated. Cell lysates were prepared using Phosphosafe Extraction buffer (Novagen) supplemented with protease inhibitors. As certified U937 monocytes and 3T3-L1 fibroblasts were directly procured from ATCC, they were not subjected to any further testing for contamination. Primary adipocytes from white adipose tissue (WAT) were prepared as described14, 36. Briefly, after mouse sacrifice, fat pads were removed and minced in Krebs-Ringer-bicarbonate-HEPES (KRBH) buffer (pH 7.4) containing 10 mM sodium bicarbonate, 30 mM HEPES, 200 nM adenosine, and 1% fatty acid-free bovine serum albumin (BSA, Sigma). WATs were digested with collagenase (2 mg g−1) in KRBH buffer at 37 °C for 1 h. Digested WATs were suspended in DMEM supplemented with 10% FBS, and filtered through 100-μm mesh cell strainer (BD Falcon) to remove undigested material. The cell suspension was incubated for 10 min at room temperature, and adipocytes collected from the floating layer after centrifugation. Adipocytes were incubated for 1 h at room temperature with gentle shaking and washed three times with DMEM. Differentiated human adipocytes in adipocyte maintenance medium were obtained from Cell Applications. Adipocytes were maintained in DMEM medium with 10% calf serum and antibiotics/antimycotic for 2 d before utilization, and 5 × 106 cells were serum-deprived for 4 h followed by treatment with 100 nM insulin for 4 h. Hepatocytes were isolated by collagenase perfusion of mouse livers and cells seeded for 4 h on collagen-coated 6-well plates (1 × 106 cells per well) in Williams’ medium E with 10% FBS, 25 mM HEPES, 100 nM insulin, and 100 nM triiodothyronine37, 38, 39. Cells were cultured for 48 h in serum-free Williams medium E with two medium changes. Before experiments, hepatocytes were pre-incubated overnight in serum-, insulin-, and triiodothyronine-free DMEM, and then with 100 nM insulin. Adult mouse cardiac cells were isolated by sequential plating using non-perfusion adult cardiomyocyte isolation kit (Cellutron)40. After isolation, 1 × 106 cardiac cells were incubated for 24 h in serum containing AS medium, and then with serum-free AW medium for another 24 h. Before experiments, cells were incubated for 4 h in serum-free DMEM, and then with 100 nM insulin. All studies using cultured cells were repeated at least three times. The number of replicates was estimated from comparable published studies that gave statistically significant results. U937 cells (1 × 107), PBMs and differentiated 3T3-L1 adipocytes (5 × 106 cells for both) were transfected with endotoxin-free plasmid DNAs or siRNAs (target-specific and scrambled control) using nucleofector (100 μl solution V for U937 cells and PBMs and 100 μl solution L for 3T3-L1 adipocytes) from Amaxa nucleofection kit (Lonza) following the manufacturer’s protocol. Transfected cells were immediately transferred to pre-warmed Opti-MEM media for 6 h and then to RPMI 1640 (for U937 cells and PBMs) and DMEM (for 3T3-L1 adipocytes) containing 10% FBS supplemented with penicillin, streptomycin, and geneticin (G418; 20 μg ml−1) for 18 to 24 h before treatment with insulin and inhibitors. Cell lysates or purified active kinases were pre-incubated with recombinant EPRS linker (wild-type and mutant) for 5 min in kinase assay buffer (50 mM Tris-HCl (pH 7.6), 1 mM dithiotheitol, 10 mM MgCl , 1 mM CaCl , and phosphatase inhibitor cocktail)7, 8, 33. Phosphorylation was initiated by addition of 5 μCi [γ-32P]ATP (Perkin-Elmer) for 15 min, and terminated using SDS gel-loading buffer and heat denaturation. Phosphorylated proteins were detected after resolution on Tris-glycine SDS–PAGE, fixation in 40% methanol and 10% acetic acid, and autoradiography. Immunoblot with anti-His tag antibody to detect EPRS linker served as control. To assay kinase activity using peptide substrates, 50 μM of synthetic peptides were phosphorylated with 1 μCi [γ-32P]ATP in kinase assay buffer. Equal volumes were spotted onto P81-phosphocellulose squares, washed in 0.5% H PO , and 32P incorporation determined by scintillation counting. U937 cell lysates were pre-cleared using protein A-sepharose, and target AGC kinase members and a non-member, MK2 were immunoprecipitated by incubation with specific antibodies for 4 h. The immunocomplex was captured by incubating with protein A-sepharose beads for 4 h, and washed three times with kinase assay buffer supplemented with 0.1% Triton X-100. The immunocomplex was resuspended in kinase assay buffer and used to phosphorylate EPRS linker as above, and 32P incorporation into peptide substrates was determined by scintillation counting41. Target peptides for S6K1, RSK1, MSK1, SGK494, NDR1, MRCKα, CRIK, RSKL1, ROCK1 and 2 (RRRLSSLRA), GRK2 (CKKLGEDQAEEISDDLLEDSLSDEDE), LATS1 (CKKRNRRLSVA), MAST1 (KKSRGDYMTMQIG), PRKX (RRRLSFAEPG), DMPK (KKSRGDYMTMQIG), and PDK1 (KTFCGTPEYLAPEVRREPRILSEEEQEMFRDFDYIADWC) were from SignalChem; for MK2 (KKLNRTLSVA) from Enzo Life Sciences; for PKA (RRKASGP), SGK1/AKT (RPRAATF), PKC/PKN (HPLSRTLSVAAKK), PKG, (RKISASEFDRPLR), and Cdk5 (PKTPKKAKKL) were from Santa Cruz. All mice were housed in microisolator cages (maximum 5 per cage of same-sex littermates) and maintained in climate/temperature- and photoperiod-controlled barrier rooms (22 ± 0.5 °C, 12–12 h dark–light cycle) with unrestricted access to water and standard rodent diet (Harlan Teklad 2918) deriving 24, 18 and 58 kcal% from protein, fat and carbohydrate, respectively. Mice were fed standard rodent diet unless otherwise indicated. The number of animals used in each experiment was estimated from examination of comparable published studies that gave statistically significant results. All mouse studies were performed in compliance with procedures approved by the Cleveland Clinic Lerner Research Institute Institutional Animal Care and Use Committee. Genetically-modified EPRS phospho-deficient S999A and phospho-mimetic S999D knock-in mice were generated (Xenogen Biosciences, Taconic). The RP23-86H18 BAC clone from mouse chromosome 1 containing full-length mouse Eprs gene was used to generate 5′ and 3′ homology arms, the knock-in region for the gene targeting vector, and Southern blot probes for screening targeted events. The homology arms and the knock-in region were generated by high-fidelity PCR, and cloned into the pCR4.0 vector. The S999A and S999D mutations (TCA to GCA or GAT, respectively) in exon 20 were introduced by PCR-based site-directed mutagenesis. The final vector also contained Frt sequences flanking the Neo expression cassette for positive embryonic stem cell selection, and a DTA expression cassette for negative selection. The targeting vector was electroporated into C57BL/6 embryonic stem cells and screened with G418. Positive expanded clones with confirmed mutation were selected. Neo was deleted by Flp electroporation, and blastocysts injected. Male chimaeras were bred with C57BL/6 wild-type females, and resulting F1 heterozygotes interbred to generate homozygotes in C57BL/6 background. Genotyping was done using forward primer 5′-CAGCATAAGAACAGTTGCCAAATAAAGG-3′ and reverse primer 5′-TTCTTGAACACACACATGCACAGACTC-3′. For all experiments the wild-type (EprsS/S), EprsA/A and EprsD/D were generated exclusively by breeding heterozygotes (EprsS/A and EprsS/D), and most experiments shown use male mice unless otherwise indicated. Mice were not randomized and studies were performed unblinded with respect to mouse genotype. S6K1−/− mice in C57BL/6 background were generated at the National Jewish Medical and Research Centre (Denver, Colorado) by blastocyst injection of embryonic stem cells with targeted disruption of the S6K1 gene as described previously19, 42. Briefly, neomycin (Neo) selection cassette was inserted to disrupt the exon corresponding to amino acids 207–237 in the catalytic domain of S6K1, thereby frame-shifting the downstream coding region. S6K1−/− mice exhibited phenotypes consistent with the previously reported mice that were generated by similar approach that is, replacing the catalytic domains of S6K1 with a Neo selection cassette9, 20. EprsD/DS6K1−/− and EprsS/SS6K1−/− were generated by EprsS/DS6K1−/− × EprsS/DS6K1−/− crosses. Mice wild-type for both Eprs and S6K1 genes (EprsS/SS6K1+/+) were generated from crosses of S6K1+/− heterozygotes. Male and female mice of EprsS/S and EprsA/A genotypes were recruited (n = 212 total mice) exclusively from crosses of heterozygotes (EprsS/A). All mice were housed in microisolator cages (maximum 5 per cage of same-sex littermates) with routine cage maintenance as above. Weaned mice (>21 days), born between June 2010 and December 2012 from 40 heterozygous parents, were monitored daily and weighed biweekly for the entire duration of their life. Mice that spontaneously developed conditions common in the C57BL/6 strain, such as malocclusion and hydrocephalus, were sacrificed and excluded from the study43. Assessments of deterioration in general health and quality of individual life were made in consultation with veterinary services of the Biological Resources Unit (BRU) of the Cleveland Clinic Lerner Research Institute. Severely sick and moribund mice that were judged to not survive another 48 h were euthanized with this date considered date of death, and included in the longevity analysis. Mice euthanized owing to imminent death include 11.5% (6 out of 52) male and 11.1% (6 out of 54) female of EprsS/S genotype, and 7.7% (4 out of 52) male and 9.3% (5 out of 54) female of EprsA/A genotype. Longevity was analysed by Kaplan–Meier survival curves from 212 mice (52 male and 54 female of each genotype, EprsS/S and EprsA/A) using known birth and death dates. Statistical differences were evaluated by log-rank Mantel–Cox and Gehan–Breslow–Wilcoxon tests using GraphPad Prism 5. Male and female mice of EprsS/S and EprsD/D genotypes were recruited (n = 89 total mice) exclusively from crosses of heterozygotes (EprsS/D). All weaned mice (>21 days born between February, 2011 and September, 2014 from 23 EprsS/D parents) were housed in microisolator cages (maximum 5 per cage of same-sex littermates) with routine cage maintenance and health monitoring as above. Mice killed owing to imminent death (as described above) include 8.7% (2 out of 23) male and 9.5% (2 out of 21) female of EprsS/S genotype, and 8.3% (2 out of 24) male and 4.8% (1 out of 21) female of EprsA/A genotype. Longevity was analysed by Kaplan–Meier survival curves from 89 mice (23, 21 male and 24, 21 male of genotype, EprsS/S and EprsA/A, respectively) using known birth and death dates and statistical analysis, as above. Male and female mice of S6K1+/+ and S6K1−/− genotypes were recruited (n = 112 total mice) exclusively from crosses of heterozygotes (S6K1+/−). All weaned mice (>21 days born between February 2011 and December 2013 from 23 S6K1+/− parents) were housed in microisolator cages (maximum 5 per cage of same-sex littermates) with routine cage maintenance and health monitoring as above. Mice killed owing to imminent death (as described above) include 13.8% (4 out of 29) male and 10.3% (3 out of 29) female of S6K1+/+ genotype, and 14.3% (4 out of 28) male and 14.3% (3 out of 21) female of S6K1−/− genotype. Longevity estimation was analysed by Kaplan–Meier survival curves from 112 mice (29, 29 male and 28, 26 female of genotype, S6K1+/+ and S6K1−/−, respectively) using known birth and death dates and statistical analysis as above. Univariate and multivariate CPH regression models were performed to analyse the effects of 4 variables; genotype, date of birth (DOB), gender, and parental identity (PID), on longevity of mice recruited for the study. The independent variables were fitted as categorical variables in the model. Genotype and gender were coded as binary variables. DOB and PID were coded as multiple categories. For CPH regression analysis of EprsS/S and EprsA/A mice (n = 212), the data were coded as follows: genotype, EprsS/S (1) and EprsA/A (0); gender, male (0) and female (1). On the basis of unique occurrences, DOB and PID were categorized into 79 (0–78, 0 being the DOB for oldest mice in the study) and 40 (1–40) categories, respectively. Oldest DOB category represents the reference for DOB. PID-1 was considered reference for PID variable. Models were fit using Cox proportional hazards regression in R package ‘survival’ using coxph function. Univariate model was built fitting each of the four variables individually and multivariate model was built fitting all four variables simultaneously. For CPH regression analysis of EprsS/S and EprsD/D mice (n = 89), the data were coded as follows: genotype, EprsS/S (1) and EprsD/D (0); gender, male (0) and female (1). On the basis of unique occurrences, DOB and PID were categorized into 38 (0–37, 0 being the DOB for oldest mice in the study) and 23 (1–23) categories, respectively. For CPH regression analysis of S6K1+/+ and S6K1−/− mice (n = 112), the data were coded as follows: genotype, S6K1+/+ (1) and S6K1−/− (0); gender, male (0) and female (1). On the basis of unique occurrences, DOB and PID were categorized into 36 (0–35, 0 being the DOB for oldest mice in the study) and 23 (1–23) categories, respectively. Scanning electron microscopy was performed by the Cleveland Clinic Imaging Core. WAT from 20-week-old male mice was fixed using 2.5% glutaraldehyde and 4% paraformaldehyde in phosphate-buffered saline (PBS) overnight at 4 °C. Tissues were washed three times in PBS followed by post-fixation with 1% osmium tetroxide in PBS for 1 h at 4 °C. Finally, the tissues were dehydrated through graded alcohol (50, 70, 90, and 100%), twice in ethanol:hexamethyldisilizane (HMDS; 1:1), and three times in 100% HMDS for 10 min each, and dried at room temperature. Samples were mounted on aluminium stubs and coated with palladium-gold using a sputter-coater, and viewed at X500 magnification with a Jeol JSM 5310 Electron Microscope (EOL). Adipose tissues from 20-week-old male mice were fixed in formalin, dehydrated in ethanol, embedded in paraffin, and cut at 5-μm thickness. Sections were deparaffinized, rehydrated, and stained with haematoxylin and eosin by the Cleveland Clinic Histology Core. Stained tissues were visualized with Leica DM2500 microscope, captured with Micropublisher 5.0 RTV digital camera (QImaging) using a 5X objective lens for magnification, and QCapture Pro 6.0 (QImaging) software for image acquisition. Adipocytes from 100 mg EWAT of 20-week male mice were isolated as described above and suspended in DMEM. Cells were counted in a haemocytometer. Basal lipolysis in primary adipocytes from EprsS/S, EprsA/A, and EprsD/D EWAT was measured by glycerol release using adipolysis assay kit (Cayman). Fatty acid oxidation in EWAT of 20-week-old male mice was performed as described13, 44. Explants were placed in an Erlenmeyer flask (Kimble-chase Kontes) containing the reaction mixture (DMEM with 0.1 μCi of [14C]oleic acid, 100 mM l-carnitine, and 0.2% fat-free BSA), and conditioned for 5 min in a 37 °C CO incubator. The flask was sealed with a rubber stopper containing a centre-well (Kimble-chase Kontes) fitted with a loosely folded filter paper moistened with 0.2 ml of 1 N NaOH, and incubated for 5 h at 37 °C. 14CO in the filter paper was trapped by addition of 200 μl of perchloric acid to the reaction mixture followed by incubation at 55 °C for 1 h. Radioactivity in the filter paper was determined by scintillation counting. At 16 weeks, mice were individually housed and given standard rodent diet and water ad libitum. Cumulative food intake was measured by weighing the mouse and food every second day for 30 consecutive days. Intraperitoneal glucose tolerance test (GTT) and insulin tolerance test (ITT) in EprsS/S, EprsA/A, and EprsD/D mice were determined as described22, 39. Briefly, GTT was done after an overnight (12 h) fast followed by peritoneal injection of glucose (2 mg g−1 body weight, Sigma). ITT was performed in 6-h fasted mice by injection of 0.75 U kg−1 body weight of insulin (Sigma). Blood glucose was determined using a commercial glucometer (Contour, Bayer). Serum triglycerides, free fatty acids, glucose, and insulin in 12-h fasted and in 1-h post-prandial (fed) mice were determined using commercially available kits. Serum triglycerides, free fatty acids, and glucose kits were from Wako. Insulin was determined using enzyme-linked immunoassay-based, ultra-sensitive mouse insulin kit (Crystal). Determination of serum β-hydroxybutyrate (for ketone body analysis) from 6-h fasted mice was done using colorimetric assay kit from Cayman. White blood cell counts in blood freshly collected by cardiac puncture in the presence of 10 mM EDTA were determined using Advia hematology system. Lipid content in mouse faeces was determined after extraction with chloroform:methanol (2:1)45, 46. GAIT system activity in insulin-treated adipocytes was determined by in vitro translation of capped poly(A)-tailed Luc-Cp GAIT and T7 gene 10 reporter RNAs as described35, 47. Gel-purified RNAs were incubated with lysates from U937 monocytes and differentiated 3T3-L1 adipocytes in the presence of rabbit reticulocyte lysate and [35S]methionine. Translation of the two transcripts was determined following resolution on 10% SDS–PAGE and autoradiography. Cytokine levels in mouse serum (100 μg protein) were determined using mouse cytokine antibody array C3 kit (RayBiotech). Mouse liver triglyceride content was determined by measurement of glycerol following saponification in ethanolic KOH (2:1, ethanol: 30% KOH)48. For assessment of total neutral lipid, freshly isolated liver slices were frozen in OCT, 5-μm sections stained with Oil Red O, and analysed by densitometry using NIH image J as described49. Mouse energy metabolism was determined by indirect calorimetry using the Oxymax CLAMS system (Columbus Instruments) in the Rodent Behavioural Core of the Cleveland Clinic Lerner Research Institute. Mice were housed individually in CLAMS cages and allowed to acclimate for 48 h with unrestricted excess to food and water. Thereafter, O consumption (VO ), CO release, RER and heat generation were recorded for 24 h spanning a single light–dark cycle. Adipocytes from 500 mg WAT from wild-type and EprsA/A mice were labelled with 150 μCi of 32P-orthophosphate (MP Biomedicals) in phosphate-free DMEM medium in absence or presence of insulin (100 nM) for 4 h. EPRS was immunoprecipitated with antibodies cross-linked to protein A-sepharose beads (Sigma) in 50 mM Tris-HCl (pH 7.6), 150 mM NaCl, 1% Triton X-100, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, and protease/phosphatase inhibitor cocktail. Immunoprecipitated beads were washed with 50 mM Tris-HCl (pH 7.6), 150 mM NaCl, and 0.1% Triton X-100, and then in 50 mM Tris-HCl (pH 7.6) and 150 mM NaCl. 32P incorporation in immunoprecipitated proteins was determined by Tris-glycine SDS–PAGE, fixation and autoradiography. Adipocytes (0.25 × 106 cells) were pre-incubated in serum-free DMEM for 4 h. Subsequently, the medium was supplemented with 2.5 μCi [14C]Glu or [14C]Pro (Perkin-Elmer), and cells incubated for additional 6 h. Adipocytes were lysed and 14C incorporation determined by trichloroacetic acid-precipitation and scintillation counting. Mouse adipocytes (0.25 × 106 cells) were pre-incubated in methionine-free RPMI medium (Invitrogen) with 10% FBS for 30 min. [35S]Met/Cys (250 μCi, Perkin-Elmer) was added and incubated at 37 °C with 5% CO for 15 min. Labelled cells were lysed in RIPA buffer (Thermo Fisher) and analysed by Tris-glycine SDS–PAGE, fixation and autoradiography. Cell lysates or immunoprecipitates were denatured in Laemmli sample buffer (Bio-Rad) and resolved on Tris-glycine SDS–PAGE (10, 12, or 15% polyacrylamide) prepared using 37.5:1 acrylamide:bis-acrylamide stock solution (National Diagnostics). After transfer to polyvinyl difluoride membrane, the membranes were probed with target-specific antibody, followed by incubation with horseradish peroxidase conjugated secondary antibody and detection with Amersham ECL prime western blotting detection reagent (GE Healthcare). Immunoblots shown are typical of experiments independently done at least three times. Pre-cleared cell lysates (1 mg) were incubated with antibody cross-linked to protein A-sepharose beads in detergent-free buffer containing 50 mM Tris-HCl (pH 7.6), 150 mM NaCl, and EDTA-free protease/phosphatase inhibitor cocktail. Immunoprecipitates were analysed by Tris-glycine SDS–PAGE and immunoblotting either after washing the beads three times in the same buffer or after elution, followed by neutralization with 0.2 M glycine-HCl (pH 2.6) or 50 mM Tris-HCl (pH 8.5), respectively. Fatty acid uptake assay kit (QBT, Molecular Devices) that utilizes fluorescent bodipy-C , a LCFA analogue, was used to determine fatty acid uptake50. Differentiated 3T3-L1 adipocytes were plated at 5 × 104 cells per well in a 96-well plate. Adipocytes were first incubated in serum-free Hank’s balanced salt (HBS) solution for 4 h, and then with 100 nM insulin and bodipy-C for an additional 4 h. After 30 min, relative fluorescence was read at 485 nm excitation and 515 nm emission wavelength in bottom-read mode (SpectraMax GeminiEM, Molecular Devices). LCFA uptake was also determined in differentiated 3T3-L1 adipocytes as cellular accumulation of [14C]oleate (Perkin-Elmer). Adipocytes (10,000 cells) were seeded in a 24-well plate in DMEM with 10% calf serum overnight. Cells were serum-deprived for 4 h, treated with 100 nM insulin for 3.5 h, and then with 50 μM of [14C]oleate in HBS containing 0.1% fatty acid-free BSA for 30 min51, 52. Cells were washed extensively in cold HBS with 0.1% fatty acid-free BSA to remove unincorporated [14C]oleate, lysed in RIPA buffer (Thermo Fisher), and centrifuged at 2000 rpm for 5 min. Supernatant radioactivity was determined by scintillation counting and normalized to protein. LCFA uptake by mouse WAT, hepatocytes, cardiac cells, BMDM, and soleus muscle strips were measured using essentially the same method13. Adipocytes from wild-type and mutant mice were pre-incubated for 4 h in serum- and glucose-free DMEM and then rinsed with Krebs-Ringer buffer containing 20 mM HEPES (pH 7.4), 5 mM sodium phosphate, 1 mM MgSO , 1 mM CaCl , 136 mM NaCl, and 4.7 mM KCl53, 54. Adipocytes were incubated for 4 h in the presence of 1 μCi of [14C]2-deoxy-d-glucose (DG; Perkin-Elmer) and 100 nM insulin in the same buffer supplemented with 100 mM unlabelled 2-DG (Sigma). Uptake was stopped using ice-cold PBS containing 50 μM cytochalasin, followed by four washes with PBS. Lysate radioactivity was determined by scintillation counting. Membrane fraction from differentiated 3T3-L1 adipocytes was isolated by phase partitioning using Mem-PER Eukaryotic Membrane Protein Extraction Reagent Kit (Thermo-Scientific). Plasma membrane fractions from 3T3-L1 adipocytes were prepared as described14. Differentiated 3T3-L1 adipocytes were washed in buffer containing 250 mM sucrose, 10 mM Tris (pH 7.4), and 0.5 mM EDTA. Lysates were prepared by homogenization in the same buffer supplemented with protease and phosphatase inhibitor cocktail, and centrifuged at 16,000g for 20 min at 4 °C. The re-suspended pellet was layered onto a solution containing 1.12 M sucrose, 10 mM Tris (pH 7.4), and 0.5 mM EDTA, and centrifuged at 150,000g for 20 min at 4 °C. The resulting pellet was suspended in RIPA buffer (Sigma) and plasma membrane was obtained by centrifugation at 74,000g for 20 min at 4 °C. All data generated are included in the published article and in the supplementary information files. Additional statistical data sets generated are available from the corresponding author upon request.

Chistiakov D.A.,Moscow State University | Chistiakov P.A.,National Diagnostics
Hepatology Research | Year: 2012

Human embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs) are a potent source for unlimited production of hepatocytes and hepatocyte-like cells that may replace primary human hepatocytes in a variety of fields including liver cell therapy, liver tissue engineering, manufacturing bioartificial liver, modeling inherited and chronic liver diseases, drug screening and toxicity testing. Human ESCs are able to spontaneously form embryoid bodies, which then spontaneously differentiate to various tissue-specific cell lineages containing a total of 10-30% albumin-producing hepatocytes and hepatocyte-like cells. Enrichment of embryoid bodies with the definitive endoderm, from which hepatocytes arise, yields increasing the final ratio of hepatocyte population up by 50-65%. Current strategies of the directed differentiation of human ESCs (and iPSCs) to hepatocytes that reproduce liver embryogenesis by sequential stimulation of culturing ESCs with tissue-specific growth factors result in achieving the differentiation rate up to 60-80%. In the future, directed differentiation of human ESCs and iPSCs to hepatocytes should be further optimized towards generating homogeneous cultures of hepatocytes in order to avoid expensive procedures of separation and isolation of hepatocytes and hepatocyte-like cells. © 2011 The Japan Society of Hepatology.

Byron S.K.,National Institute for Health and Care Excellence (NICE) | Crabb N.,National Institute for Health and Care Excellence (NICE) | George E.,National Institute for Health and Care Excellence (NICE) | Marlow M.,National Institute for Health and Care Excellence (NICE) | Newland A.,National Diagnostics
Clinical Cancer Research | Year: 2014

Companion diagnostics are used to aid clinical decision making to identify patients who aremost likely to respond to treatment. They are becoming increasingly important as more new pharmaceuticals receive licensed indications that require the use of a companion diagnostic to identify the appropriate patient subgroupfor treatment.Thesepharmaceuticalshave proven benefit inthe treatment of some cancers andother diseases, and also have potential to precisely tailor treatments to the individual in the future. However, the increasing use of companion diagnostics could place a substantial burden on health system resources to provide potentially high volumes of testing. This situation, in part, has led policy makers and Health Technology Assessment (HTA) bodies to review the policies and methods used to make reimbursement decisions for pharmaceuticals requiring companion diagnostics. The assessment of a pharmaceutical alongside the companion diagnostic used in the clinical trialsmay be relatively straightforward, although there are a number of challenges associated with assessing pharmaceuticals where a range of alternative companion diagnostics are available for use inroutine clinical practice.TheUKHTAbody, theNational Institute forHealth and Care Excellence (NICE), has developed policy for considering companion diagnostics using its Technology Appraisal and Diagnostics Assessment Programs. Some HTA bodies in other countries have also adapted their policies and methods to accommodate the assessment of companion diagnostics. Here, we provide insight into the HTA of companion diagnostics for reimbursement decisions and how the associated challenges are being addressed, in particular by NICE.© 2014 American Association for Cancer Research.

Chistiakov D.A.,National Diagnostics
Journal of Biomedical Science | Year: 2010

Rapid repair of the denuded alveolar surface after injury is a key to survival. The respiratory tract contains several sources of endogenous adult stem cells residing within the basal layer of the upper airways, within or near pulmonary neuroendocrine cell rests, at the bronchoalveolar junction, and within the alveolar epithelial surface, which contribute to the repair of the airway wall. Bone marrow-derived adult mesenchymal stem cells circulating in blood are also involved in tracheal regeneration. However, an organism is frequently incapable of repairing serious damage and defects of the respiratory tract resulting from acute trauma, lung cancers, and chronic pulmonary and airway diseases. Therefore, replacement of the tracheal tissue should be urgently considered. The shortage of donor trachea remains a major obstacle in tracheal transplantation. However, implementation of tissue engineering and stem cell therapy-based approaches helps to successfully solve this problem. To date, huge progress has been achieved in tracheal bioengineering. Several sources of stem cells have been used for transplantation and airway reconstitution in animal models with experimentally induced tracheal defects. Most tracheal tissue engineering approaches use biodegradable three-dimensional scaffolds, which are important for neotracheal formation by promoting cell attachment, cell redifferentiation, and production of the extracellular matrix. The advances in tracheal bioengineering recently resulted in successful transplantation of the world's first bioengineered trachea. Current trends in tracheal transplantation include the use of autologous cells, development of bioactive cell-free scaffolds capable of supporting activation and differentiation of host stem cells on the site of injury, with a future perspective of using human native sites as micro-niche for potentiation of the human body's site-specific response by sequential adding, boosting, permissive, and recruitment impulses. © 2010 Chistiakov; licensee BioMed Central Ltd.

Apostolou P.,National Diagnostics | Fostira F.,National Diagnostics
BioMed Research International | Year: 2013

Breast cancer is the most common malignancy among females. 5%-10% of breast cancer cases are hereditary and are caused by pathogenic mutations in the considered reference BRCA1 and BRCA2 genes. As sequencing technologies evolve, more susceptible genes have been discovered and BRCA1 and BRCA2 predisposition seems to be only a part of the story. These new findings include rare germline mutations in other high penetrant genes, the most important of which include TP53 mutations in Li-Fraumeni syndrome, STK11 mutations in Peutz-Jeghers syndrome, and PTEN mutations in Cowden syndrome. Furthermore, more frequent, but less penetrant, mutations have been identified in families with breast cancer clustering, in moderate or low penetrant genes, such as CHEK2, ATM, PALB2, and BRIP1. This paper will summarize all current data on new findings in breast cancer susceptibility genes. © 2013 Paraskevi Apostolou and Florentia Fostira.

Chistiakov D.A.,National Diagnostics
Viral Immunology | Year: 2010

In addition to genetic factors, environmental triggers, including viruses and other pathogens, are thought to play a major role in the development of autoimmune disease. Recent findings have shown that viral-induced autoimmunity is likely to be genetically determined. In large-scale genetic analyses, an association of interferon induced with helicase C domain 1 (IFIH1) gene variants encoding a viral RNA-sensing helicase with susceptibility to several autoimmune diseases was found. To date, the precise role of IFIH1 in pathogenic mechanisms of viral-induced autoimmunity has yet to be fully elucidated. However, recent reports suggest that IFIH1 may play a role in the etiology of type 1 diabetes. Rare IFIH1 alleles have been shown to be protective against diabetes, and their carriage correlates with lower production of this helicase and its functional disruption. In contrast, upregulation of IFIH1 expression by viruses is associated with more severe disease, and could exacerbate the autoimmune process in susceptible individuals. © 2010 Mary Ann Liebert, Inc.

Phinney D.G.,Scripps Research Institute | Isakova I.A.,National Diagnostics
Brain Research | Year: 2014

Lysosomal storage diseases are a heterogeneous group of hereditary disorders characterized by a deficiency in lysosomal function. Although these disorders differ in their etiology and phenotype those that affect the nervous system generally manifest as a profound deterioration in neurologic function with age. Over the past several decades implementation of various treatment regimens including bone marrow and cord blood cell transplantation, enzyme replacement, and substrate reduction therapy have proved effective for managing some clinical manifestations of these diseases but their ability to ameliorate neurologic complications remains unclear. Consequently, there exists a need to develop alternative therapies that more effectively target the central nervous system. Recently, direct intracranial transplantation of tissue-specific stem and progenitor cells has been explored as a means to reconstitute metabolic deficiencies in the CNS. In this chapter we discuss the merits of bone marrow-derived mesenchymal stem cells (MSCs) for this purpose. Originally identified as progenitors of connective tissue cell lineages, recent findings have revealed several novel aspects of MSC biology that make them attractive as therapeutic agents in the CNS. We relate these advances in MSC biology to their utility as cellular vectors for treating neurologic sequelae associated with pediatric neurologic disorders. © 2014 Elsevier B.V. All rights reserved.

Chistiakov D.A.,National Diagnostics
Advances in Experimental Medicine and Biology | Year: 2010

Abstract Ligase IV (LIG4) syndrome belongs to the group of hereditary disorders associated with impaired DNA damage response mechanisms. Clinically and morphologically, patients affected with this syndrome are characterized by microcephaly, unusual facial features, growth retardation, developmental delay, skin anomalies and are typically pancytopenic. The disease leads to acute radiosensitivity, immunodeficiency and bone marrow abnormalities. LIG4 syndrome arises from hypomorphic mutations in the LIG4 gene encoding DNA ligase IV; a component of the nonhomologous end-joining machinery, which represents a major mechanism of repair of double strand DNA breaks in mammals. The hypomorphic mutations do not completely abolish but significantly reduce enzyme function. This results in impaired V(D)J recombination, the essential rejoining process in T- and B-cell development, in whose ligase IV plays the key role. As a consequence, patients with LIG4 syndrome frequently develop multiple immune abnormalities, clinically overlapping with severe combined immunodeficiency syndrome. © 2010 Landes Bioscience and Springer Science+Business Media.

Chistiakov D.A.,National Diagnostics
Diabetes and Metabolic Syndrome: Clinical Research and Reviews | Year: 2011

Since diabetes is now a global epidemic, the incidence of retinopathy, a leading cause of blindness in patients aged 20-74 years, is also expected to rise to alarming levels. The risk of development and progression of diabetic retinopathy is closely associated with the type and duration of diabetes, blood glucose, blood pressure and possibly lipids. It is an unmet medical need that can lead to severe and irreversible loss of vision in people of working age worldwide. The aim of this review is to give an overview of the clinical and anatomical changes during the progression of retinopathy, the underlying pathogenic mechanisms that link hyperglycemia with retinal tissue damage, current treatments, and the emerging pharmacological therapies for this sight-threatening complication of diabetes. © 2012 Diabetes India.

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