Yale Stem Cell Center

New Haven, CT, United States

Yale Stem Cell Center

New Haven, CT, United States

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Ge X.,Yale University | Ren Y.,Yale University | Bartulos O.,Yale University | Lee M.Y.,Yale University | And 15 more authors.
Circulation | Year: 2012

BACKGROUND-: Supravalvular aortic stenosis (SVAS) is caused by mutations in the elastin (ELN) gene and is characterized by abnormal proliferation of vascular smooth muscle cells (SMCs) that can lead to narrowing or blockage of the ascending aorta and other arterial vessels. Having patient-specific SMCs available may facilitate the study of disease mechanisms and development of novel therapeutic interventions. METHODS AND RESULTS-: Here, we report the development of a human induced pluripotent stem cell (iPSC) line from a patient with SVAS caused by the premature termination in exon 10 of the ELN gene resulting from an exon 9 four-nucleotide insertion. We showed that SVAS iPSC-derived SMCs (iPSC-SMCs) had significantly fewer organized networks of smooth muscle α-actin filament bundles, a hallmark of mature contractile SMCs, compared with control iPSC-SMCs. The addition of elastin recombinant protein or enhancement of small GTPase RhoA signaling was able to rescue the formation of smooth muscle α-actin filament bundles in SVAS iPSC-SMCs. Cell counts and BrdU analysis revealed a significantly higher proliferation rate in SVAS iPSC-SMCs than control iPSC-SMCs. Furthermore, SVAS iPSC-SMCs migrated at a markedly higher rate to the chemotactic agent platelet-derived growth factor compared with the control iPSC-SMCs. We also provided evidence that elevated activity of extracellular signal-regulated kinase 1/2 is required for hyperproliferation of SVAS iPSC-SMCs. The phenotype was confirmed in iPSC-SMCs generated from a patient with deletion of elastin owing to Williams-Beuren syndrome. CONCLUSIONS-: SVAS iPSC-SMCs recapitulate key pathological features of patients with SVAS and may provide a promising strategy to study disease mechanisms and to develop novel therapies. © 2012 American Heart Association, Inc.


PubMed | Yale Stem Cell Center and Yale University
Type: Journal Article | Journal: Blood | Year: 2016

Bipotent megakaryocyte/erythroid progenitors (MEPs) give rise to progeny limited to the megakaryocyte (Mk) and erythroid (E) lineages. We developed a novel dual-detection functional in vitro colony-forming unit (CFU) assay for single cells that differentiates down both the Mk and E lineages (CFU-Mk/E), which allowed development and validation of a novel purification strategy for the identification and quantitation of primary functional human MEPs from granulocyte colony-stimulating factor-mobilized peripheral blood and bone marrow. Applying this assay to fluorescence-activated cell sorter-sorted cell populations, we found that the Lin(-)CD34(+)CD38(mid)CD45RA(-)FLT3(-)MPL(+)CD36(-)CD41(-) population is much more highly enriched for bipotent MEPs than any previously reported subpopulations. We also developed purification strategies for primary human lineage-committed Mk and E progenitors identified as CFU-Mk and burst forming unit-E. Comparative expression analyses in MEP, MkP, and ErP populations revealed differential expression of MYB We tested whether alterations in MYB concentration affect the Mk-E fate decision at the single cell level in MEPs and found that short hairpin RNA-mediated MYB knockdown promoted commitment of MEPs to the Mk lineage, further defining its role in MEP lineage fate. There are numerous applications for these novel enrichment strategies, including facilitating mechanistic studies of MEP lineage commitment, improving approaches for in vitro expansion of Mk and E cells, and developing improved therapies for benign and malignant hematologic disease.


News Article | April 20, 2016
Site: www.nature.com

Mouse TT2 ES cells were cultured on gelatin coating plates with recombinant LIF. ES cells were grown in DMEM supplemented with 15% fetal bovine serum, 1% non-essential amino acids, 2 mM l-glutamine, 1,000 units of mLIF (EMD Millipore), 0.1 mM β-mercaptoethanol (Sigma) and antibiotics. A doxycycline (Dox)-inducible Cas9–eGFP ES cell line was established with TT2 ESC. Guide RNA oligos (5′-accgAGTGCCTCTGGCATCCCGGG-3′, 5′-aaacCCCGGGATGCCAGAGGCACT-3′) were annealed and cloned into a pLKO.1-based construct (Addgene: 52628). Guide RNA virus was made in 293FT cells and infected inducible Cas9 ES cells. ES cells were first selected with Puromycin (1 μg ml−1) for two days, and Dox (0.5 μg ml−1) was added to induce Cas9–eGFP expression for 24 h. ES cells were then seeded at low density to obtain single-derived colonies. Then, 72 ES cell colonies were randomly picked up and screened by PCR-enzyme digestion that is illustrated in Extended Data Fig. 3a. PCR screening primers flanking guide RNA sequence were designed as following: 5′-AGGCAGATTTCTGAGTTCAAGG-3′ and 5′-TTTAGTCATGTGCTTGTCCAGG-3′. PCR products were digested by XmaI overnight at 37 degrees and separated on 2% agarose gel. A total of 8 mutants from which PCR products show resistance to XmaI digestion were subjected to DNA sequencing. Clones that harbour deletion and coding frame shift (premature termination mutation) were expanded and used in this study. Human Alkbh–Flag DNA sequence was inserted into pCW lenti-virus based vector (puromycin or hygromycin resistance). The amino acid of D233 was mutated to A by QuickChange Site-Directed Mutagenesis (QuikChange II XL Site-Directed Mutagenesis Kit, number 200521, Agilent) according to the manual. For Alkbh1 rescue experiment, wild-type and D233A mutated Alkbh1 constructs were introduced to Alkbh1 knockout ES cells, pCW-Hygromycin was chosen as control. After infections, the cells were selected with hygromycin at 200 μg ml−1 for 4 days, and then the cells were expanded to isolate genomic DNA for N6-mA dot blotting or other tests. The 293FT cells were transfected with pCW-hAlkbh1 and pCW-hAlkbh1-D233A mutant plasmids along with package plasmids of pMD2.G and pSPAX2. Culture medium was changed 10 h after transfection. The viruses were collected and concentrated 24 and 48 h after transfecction according to manufacturer’s instructions (Lenti-X Concentrator, Clontech). To establish stable expression of hAlkbh1 and hAlkbh1-D233A cell lines, 293FT cells were infected the corresponding virus, and then select with puromycin at 1 μg ml−1 for 4 days. The stable cell lines of hAlkbh1-293FT and D233A-293FT were expanded to purify the proteins according to the previous reported method with some modifications34. Briefly, M2 Flag antibody was added to the nuclear extract and incubated overnight, and then Dynabeads M-280 (sheep anti-mouse IgG, from Life Technology) was added to the above solution and incubated for 3–4 h. Subsequently, the beads were separated from the solution and washed clean with washing buffer34. Finally, the beads were eluted with 3× Flag peptides, followed by standard chromatography purification to 95% purity. Proteins were analysed by mass spectrometry. Demethylation assays were performed in 50 μl volume, which contained 50 pmol of DNA oligos and 500 ng recombinant ALKBH1 (or D233A mutant) protein. The reaction mixture also consisted of 50  μM KCl, 1mM MgCl , 50 μM HEPES (pH = 7.0), 2 mM ascorbic acid, 1 mM-KG, and 1 mM (NH ) Fe(SO ) .6H O. Reactions were performed at 37 degrees for 1 h and then stopped with EDTA followed by heating at 95 degrees for 5 min. Then the reaction product was subjected to dot blotting. Substrate sequences are listed in Supplementary Table 2. First, DNA samples were denatured at 95 degrees for 5 min, cooled down on ice, neutralized with 10% vol of 6.6 M ammonium acetate. Samples were spotted on the membrane (Amersham Hybond-N+, GE) and air dry for 5 min, then UV-crosslink (2× auto-crosslink, 1800 UV Stratalinker, STRATAGENE). Membranes were blocked in blocking buffer (5% milk, 1% BSA, PBST) for 2 h at room temperature, incubated with 6mA antibodies (202-003, Synaptic Systems, 1:1000) overnight at 4 degrees. After 5 washes, membranes were incubated with HRP linked secondary anti-rabbit IgG antibody (1:5,000, Cell Signaling 7074S) for 30 min at room temperature. Signals were detected with ECL Plus Western Blotting Reagent Pack (GE Healthcare). DNA samples were purified by standard N-ChIP protocol. 5 μg anti-H2A.X antibodies were used per 10 million cells. DNA (250 ng) from ChIP pull-down were converted to SMRTbell templates using the PacBio RS DNA Template Preparation Kit 1.0 (PacBio catalogue number 100-259-100) following manufacturer’s instructions. Control samples were amplified by PCR (18 cycles). In brief, samples were end-repaired and ligated to blunt adaptors. Exonuclease incubation was carried out in order to remove all unligated adapters. Samples were extracted twice (0.6× AMPure beads) and the final ‘SMRTbells’ were eluted in 10 μl embryoid bodies. Final quantification was carried out on an Agilent 2100 Bioanalyzer with 1 μl of library. The amount of primer and polymerase required for the binding reaction was determined using the SMRTbell concentration (ng μl−1) and insert size previously determined using the manufacturer-provided calculator. Primers were annealed and polymerase was bound using the DNA/Polymerase Binding Kit P4 (PacBio catalogue number 100-236-500) and sequenced using DNA sequencing reagent 2.0 (PacBio catalogue number 100-216-400). Sequencing was performed on PacBio RS II sequencer using SMRT Cell 8Pac V3 (PacBio catalogue number 100-171-800). In all sequencing runs, a 240 min movie was captured for each SMRT Cell loaded with a single binding complex. Base modification was detected using SMRT Analysis 2.3.0 (Pacific Biosciences), which uses previously published methods for identifying modified bases based on inter-pulse duration ratios in the sequencing data35. All calculations used the Mus musculus mm10 genome as a reference. For the detection of modified bases in individual samples, the RS_Modification_Detection.1 protocol was used with the default parameters. Modifications were only called if the computed modification QV was better than 20, corresponding to P < 0.01 (versus in silico model, Welch’s t-test). The in silico model considers the IPDs from the eight nucleotides 5′ through the three nucleotides 3′ of the site in question. Only the sites with a sequencing coverage higher than 25 fold were used for subsequent analyses. To assess the significance of the overlap between N6-mA sites by SMRT-ChIP and peaks from DIP-seq, intersection with DIP-seq peaks was analysed for each of the N6-mA site called by SMRT-ChIP. To assess if the overlap is higher than expected by random chance, a permutation based approach was used, in which we randomly shuffle the original mapping between “As” that meet coverage cutoff and their corresponding QV scores, and estimated the expected overlap by random chance. As preparation for PacBio RS II sequencing, these relatively short DNA fragments (200–1,000 base pairs on average) were made topologically circular, allowing each base to be read many times by a single sequencing polymerase. Thus, the coverage requirement for modification detection was achieved both by sequencing different fragments pulled down from the same genomic regions and by sequencing the same fragment with many passes. Of note, the SMRT-ChIP approach did not identify more N6-mA sites in Alkbh1 knockout cells than wild-type cells. Although the exact reason remain to be identified, our analysis showed that much fewer adenines are sequenced at a comparable coverage in Alkbh1 knockout cells than wild-type cells (Extended Data Fig. 5c and Extended Data Fig. 1b), presumably due to the difficulty of using native ChIP approach to isolate H2A.X-deposition regions from Alkbh1 knockout cells because of heterochromatinization. Genomic DNA from wild-type or knockout ES cells was purified with DNeasy kit (QIAGEN, 69504). For each sample, 5 μg DNA was sonicated to 200–500 bp with Bioruptor. Then, adaptors were ligated to genomic DNA fragments following the Illumina protocol. The ligated DNA fragments were denatured at 95 degree for 5 min. Then, the single-stranded DNA fragments were immunoprecipitated with 6 mA antibodies (5 μg for each reaction, 202-003, Synaptic Systems) overnight at 4 degrees. N6-Me-dA enriched DNA fragments were purified according to the Active Motif hMeDIP protocol. IP DNA and input DNA were PCR amplified with Illumina indexing primers. The same volume WT and KO DNA samples were subjected to multiplexed library construction and sequencing with Illumina HiSeq2000. After sequencing and filter, high quality raw reads were aligned to the mouse genome (UCSC, mm10) with bowtie (2.2.4, default)36. By default, bowtie searches for multiple alignments and only reports the best match; for repeat sequences, such as transposons, bowtie reports the best matched locus or random one from the best-matched loci. After alignment, N6-mA enriched regions were called with SICER (version 1.1, FDR <1.0 × 10−15, input DNA as control)37. Higher FDR cut-off could not further reduce N6-mA peak number. MACS2 was also used for peak calling, which generated similar results as SICER. Part of the data analysis was done by in-house customized scripts in R, Python or Perl. Genomic DNA samples from mouse fibroblast cells (where the endogenous N6-mA level is undetectable) were spiked with increasing amount of N6-mA-containing, or unmodified (control), oligonucleotides, and the N6-mA levels were determined by qPCR approach after DIP and library construction. Followed manufacture’s protocol (Active Motif 5mC MeDIP kit). The 5 mC data processed with MEDIPS in Bioconductor, and in-house scripts in R, Python or Perl. Native chromatin immunoprecipitation (N-ChIP) assay was performed as previously described. 10 million ES cells were used for each ChIP and massive parallel sequencing (ChIP-seq) experiment. Cell fractionation and chromatin pellet isolation were performed as described. Chromatin pellets were briefly digested with micrococcal nuclease (New England BioLabs) and the mononucleosomes were monitored by electrophoresis. Co-purified DNA molecules were isolated and quantified (100–200 ng for sequencing). Co-purified DNA and whole cell extraction (WCE) input genomic DNA were subject to library construction, cluster generation and next-generation sequencing (Illumina HiSeq 2000). The output sequencing reads were filtered and pre-analyzed with Illumina standard workflow. After filtration, the qualified tags (in fastq format) were aligned to the mouse genome (UCSC, mm10) with bowtie (2.2.4, default)36. Then, these aligned reads were used for peak calling with the SICER algorithm (input control was used as control in peak calling). H3K4Me1 and H3K27Ac ChIP-seq data were aligned to mouse genome (mm10) and peaks were called with SICER. H3K4Me1 and H3K27Ac enriched regions were defined as enhancers. Then, RSEG38 (mode 3) was to call the H3K27Ac differentiated regions. Decommissioned enhancers in KO cells are determined by H3K27Ac downregulation (compared to wild-type cells). Native ChIP-qPCR assay was used to validate H4K4Me3 at levels on gene promoters (Extended Data Fig. 8). All procedures were similar to what has been described in ChIP-seq experiments, except that the co-purified DNA molecules were diluted and subject to qPCR (histone H3K4Me3 antibodies: Abcam Ab8580). Real-time PCR was performed with SybrGreen Reagent (Qiagen, QuantiTect SYBR Green PCR Kit, Cat: 204143) and quantified by a CFX96 system (BioRAD, Inc.). RNA was extracted with miRNeasy kit (QIAGEN, 217004) and standard RNA protocol. The quality of RNA samples was measured using the Agilent Bioanalyzer. Then, RNA was prepared for sequencing using standard Illumina ‘TruSeq’ single-end stranded or ‘Pair-End’ mRNA-seq library preparation protocols. 50 bp of single-end and 100 bp of pair-end sequencing were performed on an Illumina HiSeq 2000 instrument at Yale Stem Cell Center Genomics Core. RNA-seq reads were aligned to mm9 with splicing sites library with Tophat39 (2.0.4, default parameters). The gene model and FPKM were obtained from Cufflink2. The differentially expressed genes were identified by Cuffdiff40 (2.0.0, default parameters). To make sure the normalization is appropriate, the data were also analysed with DESeq2 (default parameters), which generated similar results (Extended Data Fig. 4b). For transposons analysis, unique best alignment reads were used (alignment with bowtie (0.12.9), -m 1; or BWA) and calculated RPKM for each subfamily. For qPCR, the cDNA libraries were generated with First-strand synthesis kit (Invitrogen). Real-time PCR was performed with SybrGreen Reagent (Qiagen, QuantiTect SYBR Green PCR Kit, Cat: 204143) and quantified by a CFX96 system (BioRAD, Inc.). For Fig. 3d, the specific loci L1Md elements primers were designed and optimized based on ref. 27. For embryoid body differentiation experiment, feeder-free cultured ES cells were treated with 0.5% trypsin-EDTA free solution and resuspended with culture medium and counted. Then, cells were seeded at 200,000 cells per ml to Petri dishes with embryoid body differentiation medium (ESC medium without LIF and beta-ME). Medium was changed every 2 days. Histones were isolated in biological triplicate from wild-type and Alkbh1 knockout cells by acid-extraction and resolved/visualized by SDS–PAGE/Coomassie staining. The low molecular weight region of the gel corresponding to core histones was excised and de-stained. The excised gel region containing the histones was treated with d6-acetic anhydride to convert unmodified lysine resides to heavy acetylated lysines (45 Da mass addition) as reported in ref. 41. Following d6-acetic anhydride treatment, the gel region was subjected to in-gel trypsin digestion. Histone peptides were analysed with a Thermo Velos Orbitrap mass spectrometer coupled to a Waters nanoACQUITY LC system as detailed in ref. 42. Tandem mass spectrometric data was searched with Mascot for the following possible modifications: heavy lysine acetylation, lysine acetylation, lysine monomethylation, lysine dimethylation and lysine trimethylation. For each biological replicate, histone H2A was identified with 100% sequence coverage across K118/119 that revealed predominately no detectable lysine methylation DNA was digested with DNA Degradase Plus (Zymo Research) by following the manufacturer’s instructions with small modification. Briefly, the digestion reaction was carried out at 37 °C for 70 min in a 25 μl final volume containing 5 units of DNA Degradase Plus and 5 fMol of internal standard. Following digestion, reaction mixture was diluted to 110 μl and the digested DNA solution was filtered with a Pall NanoSep 3kDa filter (Port Washington, NY) at 8,000 r.p.m. for 15 min. After centrifugal filtration, the digested DNA solution was injected onto an Agilent 1200 HPLC fraction collection system equipped with a diode-array detector (Agilent Technologies, Santa Clara, CA). Analytes were separated by reversed-phase liquid chromatography using an Atlantis C T3 (150 × 4.6 mm, 3 μm) column. The column temperature was kept at 30 °C. For the purification of N6-mA, the mobile phases were water with 0.1% acetic acid (A) and acetonitrile with 0.1% acetic acid (B). The flow rate was 1.0 ml min−1 with a starting condition of 2% B, which was held for 5 min, followed by a linear gradient of 4% B at 20 min, 10% B at 30 min, followed by 6 min at 80% B, then re-equilibration at the starting conditions for 20 min. dA and 6-Me-dA eluted with retention times of 14.7 and 27.0 min, respectively. The amount of dA in samples was quantitated by the UV peak area (λ = 254 nm) at the corresponding retention time using a calibration curve ranging from 0.2 to 5 nMol dA on column. For the simultaneous purification of N3-Me-dC, N1-Me-dA, N3-Me-dA, N6-Me-dA and dA, the mobile phases were water with 5 mM ammonium acetate (A) and acetonitrile (B). The flow rate was 0.45 ml min−1 and the gradient elution program was set at following conditions: 0 min, 1% B; 2 min, 1% B; 40 min, 4% B; 60 min, 30% B; 65 min, 30% B; 65.5 min, 1% B, and 75 min, 1% B. N3-Me-dC, N1-Me-dA, N3-Me-dA, N6-Me-dA and dA eluted with retention times of 24.8, 25.0, 22.0, 60.2 and 54.2 min, respectively. The amount of dA in samples was quantitated by the UV peak area (λ = 254 nm) at the corresponding retention time using a calibration curve ranging from 0.9 to 7.2 nMol dA on the column. HPLC fractions containing target analyte were dried in a SpeedVac and reconstituted in 22 μl of D.I. water before LC-MS/MS analysis. LC-MS-MS analysis of N3-Me-dC, N1-Me-dA, N3-Me-dA and N6-Me-dA was performed on Ultra Performance Liquid Chromatography system from Waters Corporation (Milford, MA) coupled to TSQ Quantum Ultra triple-stage quadrupole mass spectrometer (Thermo Scientific, San Jose, CA). 20 μl of sample was introduced into mass spectrometry through a 100 mm × 2.1 mm HSS T3 column (Waters) at flow rate of 0.15 ml/min. Mobile phases were comprised of water with 0.1% formic acid (A) or acetonitrile (B). Elution gradient condition was set as following: 0 min, 1%B; 3 min, 1%B; 15 min, 7.5%B; 15.5 min, 1%B; 20 min, 1%B. Ionization was operated in positive mode and analytes were detected in selected reaction monitoring (SRM) mode. Specifically, 6-Me-dA and its internal standard were detected by monitoring transition ions of m/z = 266.1 to m/z = 150.1 and m/z = 271.1 to m/z = 155.1, respectively. Similarly, N3-Me-dC, N1-Me-dA and N3-Me-dA was detected by monitoring transition ions of m/z = 242.1 to m/z = 126.1, m/z = 266.1 to m/z = 150.1 and m/z = 266.1 to m/z = 150.1, respectively. Mass spectrometry conditions were set as following: source voltage, 3,000 V; temperature of ion transfer tube, 280 °C; skimmer offset, 0; scan speed, 75 ms; scan width, 0.7 m/z; Q1 and Q3 peak width, 0.7 m/z; collision energy, 17 eV; collision gas (argon), 1.5 arbitrary units. For quantification of N6-Me-dA, the linear calibration curves ranging from 1.5 to 750 fMol, were obtained using the ratio of integrated peak area of the analytical standard over that of the internal standard. The linear calibration curves for analysis of N3-Me-dC, N1-Me-dA and N3-Me-dA were obtained using integrated peak area of the analytical standard. N3-Me-dA is not commercial available and was prepared from the reaction between 3-methyladenine and deoxythymidine in the presence of nucleoside deoxyribosyltransferase II. The chemical identity of purified N3-Me-dA was confirmed by using an Agilent 1200 series Diode Array Detector (DAD) HPLC system coupled with Agilent quadrupole-time-of-flight (QTOF)-MS (Agilent Technologies, Santa Clara, CA). Electrospray ionization (ESI)-MS-MS spectrum of N3-Me-dA was obtained by in source fragmentation. One product ion was observed from MS/MS spectra of the protonated precursor ion of N3-Me-dA, resulting from the loss of the deoxyribosyl group. The accurate masses for parent and fragment ion are m/z = 266.1253 and m/z = 150.0774, with mass error 0.4 p.p.m. and 3.8 p.p.m., respectively. The method sensitivity for N3-Me-dC, N1-Me-dA, N3-Me-dA and N6-Me-dA was detected at 1.0 fmol, 1.6 fmol, 1.0 fmol and 1.6 fmol, respectively. In order to confirm the chemical identity of the N6-Me-dA isolated from HLPC purification, HPLC fractions containing N6-Me-dA was analysed by HPLC-QTOF-MS/MS. The chemical identity of N6-Me-dA in HPLC fractions was characterized on an Agilent 1200 series Diode Array Detector (DAD) HPLC system coupled with Agilent quadrupole-time-of-flight (QTOF)-MS (Agilent Technologies, Santa Clara, CA). HPLC separation was carried out on a C18 reverse phase column (Waters Atlantis T3, 3  μM, 150 mm × 2.1 mm) with a flow rate at 0.15 ml min−1 and mobile phase A (0.05% acetic acid in water) and B (acetonitrile). The gradient elution program was set at following conditions: 0 min, 1% B; 2 min, 1% B; 15 min, 30% B; 15.5 min, 1% B; and 25 min, 1% B. N6-Me-dA was eluted with retention times of 12.7 min. The electrospray ion source in positive mode with the following conditions were used: gas temperature, 200 °C; drying gas flow, 12 litres per min; nebulizer, 35 psi; Vcap, 4000 V; fragmentor, 175 V; skimmer, 67 V. Electrospray ionization (ESI)-MS-MS spectrum of N6-Me-dA isolated from genomic DNA was obtained by in source fragmentation. One product ion was observed from MS/MS spectra of the protonated precursor ion of N6-Me-dA, resulting from the loss of the deoxyribosyl group. The accurate masses for parent and fragment ion are m/z = 266.1245 and m/z = 150.0775, with mass error 3.0 p.p.m. and 3.1 p.p.m., respectively. The same MS/MS fragmentation spectra was obtained from analytical standard of N6-Me-dA. For in vitro demethylation assay, sample was treated with EDTA to remove Fe2+. The mixture was transferred to Amicon Ultra Centrifugal Filter (EMD Millipore Corporation, 10K MWCO), followed by spin at 11,000 r.p.m. and 4 °C for 14 min. The concentrated sample was wash three times by adding 500 μl DI-H2O, followed spin at 11,000 r.p.m. and 4 °C for 14 min. The washed sample was digested with DNA Degradase Plus (Zymo Research) by following manufacturer’s instruction with small modification. Briefly, the digestion reaction was carried out at 37 °C for 60 min in 60 μl final volume containing 0.17 units per μl of DNA Degradase Plus and 50 fmol of Internal Standard of N6-Me-dA. Following digestion, reaction mixture was filtered with a Pall NanoSep 3kDa filter (Port Washington, NY) at 10000g and room temperature for 10 min to remove enzyme. The LC-MS/MS conditions for the quantification of dA and N6-Me-dA were set the same as those for quantification of N6-Me-dA in in vivo samples. The linear calibration curves for quantification of dA and N6-Me-dA was obtained using the ratio of integrated peak area of the analytical standard over that of the internal standard of N6-Me-dA.


Xiang Y.,Yale Cardiovascular Research Center | Xiang Y.,Yale University | Cheng J.,Yale Stem Cell Center | Wang D.,Yale Cardiovascular Research Center | And 16 more authors.
Blood | Year: 2015

An elevated level of von Willebrand factor (VWF) in diabetic patients is associated with increased risk of thrombotic cardiovascular events. The underlying mechanism of how VWF expression is upregulated in diabetes mellitus is poorly understood. We now report that hyperglycemia-induced repression of microRNA-24 (miR-24) increases VWF expression and secretion in diabetes mellitus. In diabetic patients and diabetic mouse models (streptozotocin/high-fat diet-induced and db/db mice), miR-24 is reduced in both tissues and plasma. Knockdown of miR-24 in mice leads to increased VWF mRNA and protein levels and enhanced platelet tethering (spontaneous thrombosis). miR-24 tightly controls VWF levels through pleiotropic effects, including direct binding to the 3′ untranslated region of VWF and targeting FURIN and the histamine H1 receptor, known regulators of VWF processing and secretion in endothelial cells. We present a novel mechanism for miR-24 downregulation through hyperglycemia-induced activation of aldose reductase, reactive oxygen species, and c-Myc. These findings support a critical role for hyperglycemic repression of miR-24 in VWF-induced pathology. miR-24 represents a novel therapeutic target to prevent adverse thrombotic events in patients with diabetes mellitus. © 2015 by The American Society of Hematology.


Guo Y.,Yale University | Guo Y.,Yale Stem Cell Center | Liu J.,Yale University | Liu J.,Yale Stem Cell Center | And 9 more authors.
Nucleic Acids Research | Year: 2015

Steady state cellular microRNA (miRNA) levels represent the balance between miRNA biogenesis and turnover. The kinetics and sequence determinants of mammalian miRNA turnover during and after miRNA maturation are not fully understood. Through a large-scale study on mammalian miRNA turnover, we report the co-existence of multiple cellular miRNA pools with distinct turnover kinetics and biogenesis properties and reveal previously unrecognized sequence features for fast turnover miRNAs. We measured miRNA turnover rates in eight mammalian cell types with a combination of expression profiling and deep sequencing. While most miRNAs are stable, a subset of miRNAs, mostly miRNA∗s, turnovers quickly, many of which display a two-step turnover kinetics. Moreover, different sequence isoforms of the same miRNA can possess vastly different turnover rates. Fast turnover miRNA isoforms are enriched for 5′ nucleotide bias against Argonaute-(AGO)-loading, but also additional 3′ and central sequence features. Modeling based on two fast turnover miRNA∗s miR-222-5p and miR-125b-1-3p, we unexpectedly found that while both miRNA∗s are associated with AGO, they strongly differ in HSP90 association and sensitivity to HSP90 inhibition. Our data characterize the landscape of genome-wide miRNA turnover in cultured mammalian cells and reveal differential HSP90 requirements for different miRNA∗s. Our findings also implicate rules for designing stable small RNAs, such as siRNAs. © 2015 The Author(s).


Ma Y.,Yale Stem Cell Center | Jin J.,Yale University | Jin J.,Wenzhou University | Dong C.,Yale Stem Cell Center | And 5 more authors.
RNA | Year: 2010

Loss-of-function studies in human embryonic stem cells (hESCs) and induced pluripotent stem cells (iPSCs) via nonviral approaches have been largely unsuccessful. Here we report a simple and cost-effective method for high-efficiency delivery of plasmids and siRNAs into hESCs and iPSCs. Using this method for siRNA delivery, we achieve >90% reduction in the expression of the stem cell factors Oct4 and Lin28, and observe cell morphological and staining pattern changes, characteristics of hESC differentiation, as a result of Oct4 knockdown. Published by Cold Spring Harbor Laboratory Press. Copyright © 2010 RNA Society.


PubMed | University of Minnesota, Yale Center for Science and Medicine, Yale Stem Cell Center, Yale University and 2 more.
Type: | Journal: Blood | Year: 2017

The hematopoietic-stem-cell-enriched miR-125-family miRNAs are critical regulators of hematopoiesis. Overexpression of miR-125a or miR-125b are frequent in human acute myeloid leukemia (AML), and their overexpression in mice leads to expansion of hematopoietic stem cells accompanied by perturbed hematopoiesis with mostly myeloproliferative phenotypes. However, whether and how miR-125 family miRNAs cooperate with known AML oncogenes in vivo, and how the resultant leukemia is dependent on miR-125 overexpression is not well understood. We modeled the frequent co-occurrence of miR-125b overexpression and MLL-translocations by examining functional cooperation between miR-125b and MLL-AF9 By generating a knock-in mouse model in which miR-125b overexpression is controlled by doxycycline (Dox) induction, we demonstrate that miR-125b significantly enhances MLL-AF9-driven AML in vivo and the resultant leukemia is partially dependent on continued overexpression of miR-125b Surprisingly, miR-125b promotes AML cell expansion and suppresses apoptosis involving a non-cell-intrinsic mechanism. MiR-125b expression enhances VEGFA expression and production from leukemia cells, in part by suppressing TET2 Recombinant VEGFA recapitulates miR-125bs leukemia-promoting effects, whereas knockdown of VEGFA or inhibition of VEGFR2 abolishes miR-125bs effects. In addition, significant correlation between miR-125b and VEGFA expression is observed in human AMLs. Our data reveal cooperative and dependent relationships between miR-125b and MLL oncogene in AML leukemogenesis, and demonstrate a miR-125b-TET2-VEGFA pathway in mediating non-cell-intrinsic leukemia promoting effects by an oncogenic miRNA.


News Article | March 31, 2016
Site: phys.org

"Basically, these viruses appear to allow the mammalian genome to continuously evolve, but they can also bring instability," said Andrew Xiao of the Department of Genetics and Yale Stem Cell Center, senior author of the paper published online March 30 in the journal Nature. "Aside from the embryo, the only other places people have found this virus active is in tumors and neurons." Xiao and the Yale team discovered a novel mechanism by which the early embryo turns off this virus on the X chromosome, which ultimately determines the sex of an organism. If the level of this molecular marker is normal, X chromosomes remain active, and females and males will be born at an equal ratio. If this marker is overrepresented, X chromosomes will be silenced, and males will be born twice as often as females. "Why mammalian sex ratios are determined by a remnant of ancient virus is a fascinating question," Xiao said. Tens of millions of years ago viruses invaded genomes and duplicated themselves within the DNA of their hosts. Xiao estimated that more than 40% of the human genome is made up of such remnants of viral duplications. In most cases, these remnants remain inactive, but recently scientists have discovered they sometimes take on surprising roles in developing embryos and may even push mammalian evolution. Researchers found that the virus active in the mouse genome that influences sex ratios is relatively recent—in evolutionary terms—and is enriched on the X chromosome. The Yale-led team found the mechanism that disables the virus. The newly discovered modification in mammals is a surprising expansion of the epigenetic toolbox, say the researchers. Epigenetics modulates gene expression during development without actually altering the sequences of genes. In the new marker, a methyl bond is added to adenine—one of the four nucleotides that comprise base pairs in DNA—allowing it to silence genes. For decades, most researchers assumed that a modification of the nucleotide cytosine was the only form of gene silencing in mammals. Xiao said it is possible that this mechanism might be used to suppress cancer, which has been known to hijack the same virus to spread. He also noted in other organisms, such as C elegans and the fruit fly Drosophila, this mechanism plays an entirely opposite role and activates genes, not suppresses them. "Evolution often uses the same piece but for different purposes and that appears to be the case here," Xiao said. More information: DNA Methylation on N6-adenine in mammalian embryonic stem cells, Nature, DOI: 10.1038/nature17640

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