Weill Cornell Medicine

New York City, NY, United States

Weill Cornell Medicine

New York City, NY, United States
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NORWALK, Conn.--(BUSINESS WIRE)--The Multiple Myeloma Research Foundation (MMRF) today announced that Ronald O. Perelman and Dr. Anna Chapman, through the Perelman Family Foundation, have committed more than $4 million in funding to launch the first ever research program solely dedicated to the early detection and prevention of multiple myeloma. This generous donation will seed the launch of the groundbreaking Perelman Family Foundation Early Disease Translational Research Program, part of the MMRF Prevention Project, to speed efforts toward early detection, delayed disease progression, and eventually, ultimately, prevention of this incurable disease. “The goal of this initiative is to develop a completely new paradigm for research in multiple myeloma, focusing on early detection and ultimately, prevention. Right now, detection of this terrible disease often comes too late. Unlike most cancers, early detection of multiple myeloma doesn't increase a person’s chance of survival under current treatment options. The Perelman Family Foundation Early Disease Translational Research Program will support research focused on improving outcomes after early detection. With the MMRF and our university partners, we are confident that we will be able to make breakthroughs for multiple myeloma patients, and that the program will serve as a model for future initiatives,” said Dr. Anna Chapman. The gift from the Perelman Family Foundation provides a catalyst for essential research focused on: better understanding genomic determinants of early disease progression; how microenvironment factors influence early disease progression; and enhancing patient tumor immunity. Perelman Family Foundation Early Disease Translational Research Program brings together six leading cancer research centers: Dana-Farber Cancer Institute, Memorial Sloan Kettering Cancer Center, MD Anderson Cancer Center, Rockefeller University, University of Arkansas for Medical Science, Yale University, as well as the MMRF. The studies conducted by these teams will identify novel targets and biomarkers of disease progression and enable the development of therapeutic approaches to delay or even stop progression to myeloma. “We are so thankful to Ronald and Anna for supporting our vision for a bold program that will take us one step closer to a future where our children and grandchildren will never need to worry about incurable cancers,” said the Multiple Myeloma Research Foundation Founder Kathy Giusti. “Not only does the MMRF answer the questions of patients today and urgently deliver them the precise information and treatment they need to fight their multiple myeloma, but, with this generous donation, we will now also be able to focus on the patients of tomorrow.” Inspired by the dedication and vision of its Chairman and CEO Ronald O. Perelman and his family, the Perelman Family Foundation is firmly committed to philanthropy, focusing on women’s health, education and the arts. Ranked among the top philanthropists in the United States, Mr. Perelman is the founder of the Revlon/UCLA Women’s Cancer Research Program, which analyzes the causes of and develops groundbreaking treatment for breast and ovarian cancer. Launched in 1994, the program was responsible for the development of Herceptin, the first genetically-based treatment for a major cancer to be approved by the FDA, which currently cures more than thirty percent of breast cancer cases in women. In 2014, he co-founded, along with Barbra Streisand, the Women’s Heart Alliance to raise awareness, encourage action and drive new research to fight heart disease in women. Through the Perelman Family Foundation, Mr. Perelman supports the Ronald O. Perelman Center for Emergency Services and the Ronald O. Perelman Department of Dermatology at NYU Langone Medical Center; the Ronald O. Perelman Heart Institute at New York Presbyterian Hospital, an internationally-recognized center offering comprehensive, innovative, and world-class cardiovascular care and heart health education; and the Ronald O. Perelman and Claudia Cohen Center for Reproductive Medicine at Weill Cornell Medicine. Multiple myeloma is a cancer of the plasma cell. It is the second most common blood cancer. An estimated 30,280 adults will be diagnosed this year and 12,590 people are predicted to die from the disease. The mission of the Multiple Myeloma Research Foundation (MMRF) is to find a cure for multiple myeloma by relentless pursuing innovation that accelerates the development of next-generation treatments to extend the lives of patients. Founded in 1998 by Kathy Giusti, a multiple myeloma patient, and her twin sister Karen Andrews as a 501 (c) (3) nonprofit organization, the MMRF is a world-recognized leader in cancer research. Together with its partners, the MMRF has created the only end-to-end solution in precision medicine and the single largest genomic dataset in all cancers. The MMRF continues to disrupt the industry today, as a pioneer and leader at the helm of new research efforts. Since its inception, the organization has raised over $350 million and directs nearly 90% of the total funds to research and related programs. As a result, the MMRF has been awarded by Charity Navigator’s coveted four-star rating for 12 years, the highest designation for outstanding fiscal responsibility and exceptional efficiency.


News Article | May 23, 2017
Site: www.eurekalert.org

ATS 2017, WASHINGTON, DC--A lung cancer diagnosis appears to put patients at the greatest risk of suicide when compared to the most common types of non-skin cancers, according to new research presented at the ATS 2017 International Conference. Researchers analyzed 3,640,229 patients in the Surveillance, Epidemiology, and End Results (SEER) database and looked at suicide deaths for all cancers and for lung, prostate, breast and colorectal cancers individually. Over a 40 year period, cancer diagnoses were associated with 6,661 suicides. "We wanted to see what the impact of one of life's most stressful events is on patients," said Mohamed Rahouma, MD, a post-doctoral cardiothoracic research fellow at Weill Cornell Medical College/New York Presbyterian Hospital. "I think it's fair to say that most clinicians don't think about suicide risk in cancer patients. This study, I hope, will change that by making us more aware of those at greatest risk of suicide so that this catastrophe in the care of our patients doesn't happen." Among lung cancer patients, Asians have a more than 13-fold and men a nearly 9-fold increase in suicides. Other factors that increased suicide risk were being older, being widowed, refusing surgical treatment and having a difficult-to-treat (metastatic) type of lung cancer. The authors noted that over the 40-year study period, suicide rates decreased, most notably for lung cancer when compared to the other three most common cancers. "While cancer diagnosis counselling is an established practice, especially if a patient seems depressed, referral for ongoing psychological support and counseling typically does not happen," Dr. Rahouma said. "This represents a lost opportunity to help patients with a devastating diagnosis." Lung cancer patients have the highest malignancy-associated suicide rate in USA: a population based analysis. Authors: M. Rahouma, M. Kamel, A. Nasar, S. Harrison, B. Lee, B. Stiles, N. Altorki, J.L. Port; Weill Cornell Medicine, New York Presbyterian Hospital, New York, NY - New York, NY/US Previous studies have reported that the psychological and social distresses associated with cancer diagnosis have led to an increase in suicides compared to the general population. In the current study we sought to explore lung cancer associated suicide rates in a large national database compared to the general population as well as to the three most prevalent non-skin cancers [breast, prostate, colorectal cancer (CRC)]. The Surveillance, Epidemiology, and End Results (SEER) database (1973-2013) was retrospectively queried to identify cancer associated suicide deaths in all cancers combined, as well as, for each of lung, prostate, breast, or colorectal cancers. Suicide incidence and standardized mortality ratio (SMR) were estimated by using SEER*Stat 8.3.2 program. Suicidal rate for the general US population was obtained from National Vital Statistics Reports, Vol.64, No. 2. Furthermore, suicidal trends over time and timing from cancer diagnosis to suicide were estimated for each cancer type. Among lung cancer patients, suicide SMR of different demographic, social and tumor related factors were identified. Among 3,640,229 patients diagnosed with cancer in the study period, 6661 patients committed suicide. The cancer associated suicide rate was 27.5/100,000 person-years (SMR=1.6) compared to 13 /100,000 person-years for the general US population. The highest suicide risk was observed in lung cancer patients (SMR= 4.2) followed by CRC (SMR=1.4), breast cancer (SMR=1.4) and prostate cancer (SMR=1.2). Median time to suicide was 7 months from diagnosis in lung cancer, 56 months in prostate ca, 52 months in breast cancer and 37 months in CRC (p There was a trend towards the decrease in suicide SMR over time, which is most notable for lung cancer compared to the other three cancers (Figure). Among lung cancer patients; suicide SMR was higher in males (SMR=8.8), Asians (SMR=13.7), widowed patients (SMR=11.6), older patients (70-75 years; SMR=12), patients with undifferentiated tumors (SMR 8.6) or small cell lung carcinoma (SCLC) histology (SMR=11.2), patients presenting with metastatic disease (SMR=13.9) and in patients who refused to receive surgical treatment (SMR=13). The cancer associated suicide rate is nearly twice that of US-general population. Suicide risk is highest among lung cancer patients, particularly older patients, widowed, males, and patients with unfavorable tumor characteristics. It is important to identify these high risk patients in order to provide the proper psychological assessment, support, and counselling to reduce these rates.


News Article | May 24, 2017
Site: www.nature.com

No statistical methods were used to predetermine sample size. The experiments were not randomized and the investigators were not blinded to allocation during experiments and outcome assessment. All animal experiments were performed under the approval of Weill Cornell Medicine Institutional Animal Care and Use Committee. We used Runx1-IRES-GFP (Runx1tm4Dow) mice, provided by J. Downing (St Jude Hospital). These were crossed with Rosa26 rtTa mice (B6.Cg-Gt(ROSA)26Sortm1(rtTA*M2)Jae/J, Jackson laboratory, strain 006965) to produce Runx1-IRES-GFP;Rosa26-rtTa mice (referred to as Runx1), and maintained as heterozygous for both Runx1-IRES-GFP and rtTa. Floxed Cxcr4 mice were obtained from Y.-R. Zou (Feinstein Institute for Medical Research). Floxed Cxcr4 mice were crossed with Runx1-IRES-GFP;Rosa26-rtTa mice to produce Cxcr4fl/fl;Runx1-IRES-GFP+/−;Rosa26-rtTa mice. Rag1−/− mice with the genotype of B6.129S7-Rag1tm1Mom/J were obtained from the Jackson Laboratory (strain 002216). For rEC-HSPC transplantation, we used CD45.1+ congenic 10–12-week-old male recipients (B6.SJL-Ptprca Pepcb/BoyJ). These mice are referred here as CD45.1+ recipients, Jackson Laboratory, strain 002014. All cell preparations were routinely tested for mycoplasma contamination. To distinguish between vascular mECs and lymphatic endothelial cells, we performed intravital staining by injecting 25 μg of anti-VE-cadherin-AF647 antibody (clone BV13, Biolegend) retro-orbitally in 8–10-week-old male C57BL/6J (CD45.2+) mice under anaesthesia 8 min before they were killed and the organs harvested. For flow cytometry and cell sorting, organs, including lungs, brain, kidney and liver were minced and incubated with collagenase A (25 mg ml−1), dispase II (25 mg ml−1), and DNase (250 μg ml−1) (Roche) at 37 °C for 20–30 min to create a single-cell suspension. Cells were filtered through a 40-μm filter immediately before counter staining. The single-cell suspension was first blocked with an antibody against CD16/32 (2.4G2) before antibody staining with anti-mouse CD31-PE-Cy7 (390, Biolegend), anti-mouse TER119 (TER119) and anti-mouse CD45 (30-F11). Haematopoietic and erythroid cells were removed via CD45 and TER119 gate exclusion, and adult mouse endothelial cells (mECs) were defined and sorted as VEcad+CD31+CD45−TER119− cells. Resulting adult mEC cultures were cultured on fibronectin-coated (Sigma-Aldrich) plates in mEC media. Purity of mECs and absence of contaminating haematopoietic cells were confirmed by flow cytometry and confocal microscopy. Human umbilical vein endothelial cells (HUVECs) were isolated as described29, 40 and cultured in endothelial cell growth medium. Once endothelial cells were transduced with lentiviral vectors expressing E4ORF1 gene they were then cultured in human endothelial cell media (M199 (Sigma, M4530), 10% FBS (Omega Scientific, FB07), 50 μg ml−1 endothelial mitogen (Alfa Aesar J65416), and 100 μg ml−1 heparin (Sigma, H3393)11, 13, 28, 29. E4ORF1 confers endothelial cells with the capacity to survive in the absence of serum and exogenous growth factors, while sustaining the pre-determined endothelial cell signatures and repertoire of the pro-haematopoietic angiocrine factors. The aforementioned conversion experiments with FGRS transduction of endothelial cells would not be possible without use of HUVEC-E4ORF1 vascular-niche cells (VN-ECs) as reprogramming in the presence of serum there is no conversion of endothelial cells into haematopoietic cells9. Thus, because E4ORF1 allows endothelial cells to survive and perform as inductive vascular-niche cells, this platform enables FGRS-driven conversion of adult endothelial cells without the supplementation with xenobiotic factors, serum or exogenous angiogenic growth factors. Both serum and angiogenic growth factors could interfere with and complicate the conversion of the endothelial cells into haematopoietic cells, by aberrantly interfering with the survival, self-renewal and expansion of the converted endothelial cells into haematopoietic cells (rEC-HSPCs and rEC-HSCs). Indeed, in the presence of serum the conversion of endothelial cells to HSPCs or HSCs is completely blocked9. All niche-dependent experiments described herein were performed by using HUVEC-derived E4ORF1+ endothelial cells (VN-ECs, HUVEC-E4ORF1, Angiocrine Bioscience)11, 13, 19, 28, 29 as a vascular-niche feeder monolayer, except as specifically noted. Open reading frames (Fosb: NM_008036.2, Gfi1: NM_010278.1, Runx1 (Runx1b isoform): NM_001111022.2, Sfpi1: BC003815.1) were cloned into pLVx TET-3G lentiviral plasmid. Lentiviruses (LV) were produced were cloned into doxycycline-inducible pLVx TET-3G lentiviral plasmids. Lentiviral vectors expressing Cxcl12 (NM_001012477.2) were obtained from Cyagen (pLV(Exp)-Neo-EF1A>mCxcl12:IRES:EBFP). Viral particles were produced in the HEK 293T Lenti-X cell line (Clontech, 632180) using Lenti-X packaging single shot (Clontech, 631275) following the manufacturer’s instructions, and titred using Lenti-X p24 Rapid Titer Kit (Clontech, 632200). Transduction was carried out in 6-well plates. Doxycycline (dox) (Sigma-Aldrich) was added at a concentration of 1 μg ml−1 for FGRS transgene induction. Adult mECs were directly converted into haematopoietic cells by conditional enforced expression of transcription factors followed by replating onto a serum-free inductive vascular niche (VN-ECs, HUVEC-E4ORF1). Purified populations of VEcad+CD31+CD45−TER119− mECs were cultured in the mEC growth medium, consisting of DMEM:Ham’s F-12 (Sigma, D6421) supplemented with 20% FBS (Omega Scientific, FB07), 20 mM HEPES (Invitrogen, 15630080), 100 μg ml−1 heparin (Sigma, H3393), 50 μg ml−1 endothelial mitogen (Alfa Aesar J65416) and 5 μM SB431542 (R&D, 1614), in a humidified incubator at 37 °C, 5% CO and normoxia 5% O . Then, adult mECs were transduced with doxycycline-inducible TET-3G lentiviral vectors for a combination of transcription factors and the reverse tetracycline-controlled trans-activator—FosB, Gfi1, Runx1, Spi1 (FGRS), and rtTA—and maintained in mEC media for 3 days. FGRS-ECs were then selected for rtTA expression by puromycin resistance. 72 h after puromycin selection, FGRS expression was induced by adding 1 μg ml−1 dox in mEC media for 48 h. Dox was added every 24 h. FGRS-ECs were then reseeded onto confluent VN-EC monolayers supplemented with conversion media (StemSpan SFEM, STEMCELL Technologies, 09650), 10% KnockOut Serum Replacement (Invitrogen, 10828028), 10 ng ml−1 hFGF-2 (bFGF, Peprotech, 100-18), 50 ng ml−1 mouse c-Kit ligand (SCF, Peprotech, 250-03). Conversion media was then replaced every 48 h. Adult organs, including lung, brain, liver and kidney VEcad+CD31+CD45− (C57BL/6J CD45.2 background) mECs were purified to remove contaminating lymphoid endothelial cells and haematopoietic cells. After cultivation for 30 days, pure populations of mECs were then used for direct conversion into HSPCs (rEC-HSPCs) and HSCs (rEC-HSCs), as described above. Adult lung mECs, devoid of any haematopoietic cells, were primarily used to assess the capacity of the rEC-HSPC and rEC-HSCs to reconstitute primary and secondary, clonal and serial haematopoietic transplantations. On day 28, FGRS-ECs and residual VN-ECs were transplanted at 8.0 × 105 cells per recipient into lethally irradiated (950 cGy) congenic recipients. Dox was not administered in the drinking water to ensure no exogenous FGRS expression during the transplant. Serial transplantation from reprogrammed adult lung mECs to HSPCs experiments were performed by transplanting 1.0 × 107 unfractionated bone marrow cells into secondary recipients (CD45.1+ or Rag1−/−). Serial transplantation of unfractionated whole bone marrow cells (WBM) from C57BL/6J (CD45.2+) were used as controls. In all transplant experiments, peripheral blood and bone marrow analysis was performed at 4-week intervals with antibodies against c-Kit/CD117 (2B8), Sca-1/Ly-6A (D7), CD48 (HM48-1), and CD150 (mShad150). Lineage antibody cocktail included: CD41 (MWReg30) TER119 (TER119), B220 (RA3-6B2), CD11b (M1/70), Gr1 (RB6-8C5), CD3 (17A2), CD4 (GK1.5), CD8 (YTS156.7.7), CD44 (IM7), CD62l (MEL-14), CD25 (3C7), FoxP3 (150D), TCRγδ (GL3), CD45.1 (A20), CD45.2 (104), and DAPI to discriminate and eliminate dead cells from analysis. All antibodies were obtained from Biolegend unless mentioned. For calculation of competitive repopulating units (CRU), recipient mice were transplanted with limiting dilutions of donor-LKS cells (1 to 4,500) together with 500,000 recipient-derived bone marrow cells. Mice were bled every 4 weeks and killed after 16 weeks. Multi-lineage myelo-lymphoid donor-derived contribution in the peripheral blood was assessed using flow cytometry analysis. HSC-CRU frequency and statistical significance was determined using ELDA software (http://bioinf.wehi.edu.au/software/elda/). Samples were permeabilized in PBST and blocked in 5% donkey serum. Samples were incubated for 2 h in primary antibodies blocking solution, washed 3 times in PBS and incubated in secondary antibodies (Jackson Laboratories) for 1 h. Following washing, some sections were counterstained for nucleic acids by DAPI (Invitrogen) before mounting and imaging by confocal microscopy. The primary antibodies used for immunostaining are listed in the previous section. All imaging was performed using a Zeiss 710 META confocal microscope. At least 100 ng of total RNA from both freshly harvested and cultured cells was isolated (phenol-chloroform separation of TRIzol LS) and purified using Qiagen RNeasy Mini Kit. RNA quality was verified using an Agilent Technologies 2100 Bioanalyzer. RNA library preps were prepared and multiplexed using Illumina TruSeq RNA Library Preparation Kit v2 (non-stranded and poly-A selection) and 10 nM of cDNA was used as input for high-throughput sequencing via Illumina’s HiSeq 2500 producing 51 bp paired-end reads. Sequencing reads were de-multiplexed (bcl2fastq v2.17), checked for quality (FastQC v0.11.5), and trimmed/filtered when appropriate (Trimmomatic v0.36). The resultant high-quality reads were mapped (TopHat2 v2.1.0; Bowtie2 v2.2.6) to the transcriptome sequence reference of the UCSC mm10 genome build. Unique and multi-mapped reads were then assembled into transcripts, and abundance measures (FPKM values) quantified (Cufflinks v2.2.1). All subsequent transcriptome data analysis utilized the estimated measurements of transcripts abundance (that is, FPKMs). Genes with FPKM < 1 were filtered out, and log -transformed FPKM values were used for principal component analysis and hierarchical clustering. Transcriptome-wide and gene-set-specific analysis of the RNA-seq expression dataset were summarized and represented in the forms of scatter plots, dendrograms, and heatmaps. TCR repertoire analysis on RNA-seq reads were performed by custom BLAST-mapping. The reads were submitted for nucleotide BLAST-mapping against custom databases comprising TCR Vα genes, Vβ genes, Cα genes, and Cβ genes downloaded from IMGT (http://www.imgt.org). A table of the counts of reads meeting BLAST-expected value cutoffs for each α and β variable and constant gene was formulated for each sample and normalized to counts per million sequenced reads. The Relative Shannon Index was calculated using the Shannon entropy of the counts of TCR Vβ genes normalized by the logarithm of the number of different Vβ genes occurring in each sample and P values showing differences in the Relative Shannon Index were calculated using the Wilcoxon rank-sum test. Heatmaps and clustering were then performed in R using ‘heatmap.2’ function from gplots package. RNA was isolated from CD45.2+CD3+CD8+ and CD45.2+CD3+CD4+ sorted T lymphocytes from peripheral blood (1 × 105 cells) with TRIzol (Invitrogen; 15596-026). The diversity of CDR3 regions for 24 TCR Vβ regions was assessed with the TCRExpress Quantitative Analysis Kit (Biomed Immunotech; H0533) per the manufacturer’s instructions. BALB/c peripheral blood cells were labelled with the fluorescent membrane dye PKH26 (Sigma, PKH26GL) to distinguish them from effector cells (FGRS-CD45.2+CD3+CD8+ T cells) upon FACS analysis according to the manufacturer’s instructions. Wells of 24-well culture plates were seeded with 1 × 104 dye-labelled BALB/c peripheral blood cells; 2–3 h later, 500 μl of FGRS-CD45.2+CD3+CD8+ T cells pre-activated for 8 h with anti-CD3/CD28 beads according to the manufacturer’s instructions (Miltenyi, 130-097-627), as described in ref. 41. Reactions were performed in presence of 10 U ml−1 of IL-2 with effector:target ratios of 1:2, 1:5, 1:10, 1:25, 1:50, 1 :100. Samples were analysed using a SORP-LSR2 flow cytometer (BD Biosciences). All cytotoxicity assays were performed in triplicate. Dead target cells were defined as PKH26−DAPI+CD45.2− cells. Mice transplanted with either rEC-HSPCs or WBM mononuclear haematopoietic cells were injected with full-length chicken OVA emulsified in complete Freund’s adjuvant (CFA) subcutaneously at two sites on the back, injecting 0.1 mg at each site. A booster injection of OVA emulsified in incomplete Freund’s adjuvant (IFA) was administered 14 days after immunization with ovalbumin/CFA emulsion. The booster is given as a single subcutaneous injection with 0.1 ml of IFA emulsion, at one site on the back. (1) Adult mECs were purified by multicolour flow cytometry to eliminate contaminating lymphatic endothelial cells, pericytes, mesenchymal, and especially haematopoietic cells. To rule out the possibility of contamination with host-derived haematopoietic cells, freshly sorted mECs (8 × 105 cells from the CD45.2 strain) were transplanted into lethally irradiated (950 cGy) 10 to 12 weeks CD45.1+ male recipients with 500,000 radio-protective CD45.1+ bone marrow haematopoietic cells. CD45.1+ recipient mice were assessed for CD45.2+ engraftment with contaminating HSPCs cells, as described above in section transplantation assays. (2) To demonstrate that the freshly isolated mEC expansion culture conditions, not supplemented with haematopoietic cytokines, will prevent proliferation or survival of any contaminating haematopoietic cells, limiting dilutions of CD45.2+ wild-type LKS-SLAM cells were introduced into mEC expansion cultures. In these experiments LKS-SLAM cells were transduced with FGRS transgenes to rule out the possibility that these factors will enhance LKS-SLAM cell survival and expansion and seeded in limiting dilution on top of mEC–VN-EC co-culture. As per conversion protocol, these cultures were grown in mEC media (absent haematopoietic cytokines) for 4 weeks. ‘Contaminated’ mEC–VN-EC co-cultures were then grown in conversion media for 28 days. The resulting cultures were transplanted into three lethally irradiated (950 cGy) CD45.1+ male recipient mice with 500,000 CD45.1+ bone marrow haematopoietic cells to demonstrate that LKS-SLAM cells could not survive such haematopoietic intolerant culture conditions. (3) Purified mECs were expanded for conversion experiments routinely for 30 days in mEC media in complete absence of haematopoietic cytokines. Then, mECs were transduced with dox-inducible FGRS transgenes but were never exposed to dox (no-dox) and therefore never expressed FGRS transgenes. 8 × 105 cultured no-dox FGRS-transgene-transduced 30-day-expanded mECs were transplanted into lethally irradiated (950 cGy) recipient mice in a rescue dose setting to rule out existence of any contaminating haematopoietic cells. Recipient CD45.1+ transplanted mice were assessed for survival. (4) It is possible that if any host HSPCs could survive the intolerant 30 days of mEC culturing, upon FGRS transduction these contaminating HSPCs might revert to functional HSPCs or HSCs. To disprove this possibility, and mimic the standard procedure used in isolation of adult mouse lung mECs, we re-expressed FGRS in terminally differentiated CD45.2+ haematopoietic cells (TER119+, Gr1+CD11b+, B220+, CD3+) isolated from lungs of CD45.2+ rEC-HSPC week 16 secondary engrafted recipients. To this end, 150,000 terminally differentiated CD45.2+ (TER119+, Gr1+CD11b+, B220+, CD3+) were cultured for 28 days in conversion media in presence of doxycycline in co-culture with VN-ECs. Resulting cultures were transplanted into three lethally irradiated (950 cGy) CD45.1+ male recipient mice with 500,000 radio-protective bone marrow haematopoietic cells. Then, the recipient CD45.1+ transplanted mice were assessed for CD45.2+ engraftment as described above in the section on transplantation assays. (5) If host HSPC contamination contributes to rEC-HSPCs and rEC-HSCs, then there should be no hierarchical FGRS-dependence induction, specification or expansion phases during rEC-HSPC generation. In addition, within the first 8 days of culture (induction phase), the contaminating HSPCs should give rise to engraftable HSPCs. To assess this, dox-on FGRS-transduced mECs were passaged through stepwise shutting-off FGRS expression either during the induction, specification or expansion phases. The number of rEC-HSCs and rEC-HSPCs at each stage were then quantified, as described above. Adult lung mECs (VEcad+CD31+CD45−) were isolated from Runx1-IRES-GFP mice. VN-ECs were discriminated from Runx1-IRES-GFP-FGRS-ECs by anti-human CD31 (hCD31). Expression of VEcad and CD45 in Runx1-IRES-GFP-FGRS-ECs and derivatives were monitored during endothelial cell to haematopoietic cell reprogramming. Adult mECs were treated with different small molecules at their known IC on mouse endothelial cells (CXCR4 antagonist, AMD3100 = 44 μmol l−1; CXCR7 agonist, TC14012 = 350 nmol l−1; BMP antagonist, Noggin = 0.5 μg ml−1; TGFβ antagonist, SB431542 = 10 μmol l−1). Toxicity of each small molecule was assessed by annexin V/DAPI staining 48 h after the first treatment, as well as population-doubling time. Relative CD45+ percentages were then acquired by flow cytometry at day 28. Adult lung mECs were generated from Runx1-IRES-GFP-Cxcr4fl/fl. Cxcr4−/− endothelial cells were generated by transduction with LV-CMV-Cre-Puro lentivirus (SignaGen, SL100272) followed by 7 days of puromycin selection. Cxcr4fl/fl and Cxcr4−/− endothelial cells were transduced with FGRS along with VN-EC induction and converted. Then, the frequency and functionality of the emerging CD45+ rEC-HSPCs and rEC-HSCs were assessed, as described above, by performing phenotypic flow cytometry. We analysed recipient organ architecture and histological profile after 20 weeks of primary or secondary transplantation for any evidence of malignant alterations. For each organ, including bone marrow, lung, kidney, spleen, liver, intestine and brain, Wright–Giemsa, Masson and PicroSirius Red staining were performed on 10-μm paraffin-embedded sections (Histoserv). All images were acquired using a colour CCD camera. The primer (Integrated DNA) sets used to detect each sequence were as follows. B1 repeated sequence: 5′-GTGGTGGCGCACGCCT-3′ and 5′-TAGCCCTGGCTGTCCTGGAA-3′; LTR: 5′-TCCACAGATCAAGGATATCTTGTC-3′. Reactions contained 1× Taqman universal master mix (Perkin-Elmer), 300 nM forward primer, 300 nM reverse primer, 100 nM probe primer and 100−500 ng of template DNA in a 30-μl volume. After initial incubations at 50 °C for 2 min and 95 °C for 10 min, 40 cycles of amplification were carried out at 15 s at 95 °C followed by 1 min 30 s at 60 °C. PCR products were then analysed on a 4% TBE–EtBR gel. All data are presented as either median ± s.e.m., mean ± s.d., or mean ± s.e.m. (as indicated in figure legends). The data presented in the figures reflect multiple independent experiments performed on different days using different mice. Unless otherwise mentioned, most of the data presented in figure panels are based on at least three independent experiments. The significance of differences was determined using a two-tailed Student’s t-test, unless otherwise stated. P > 0.05 was considered not significant; *P < 0.05; **P < 0.01; ***P < 0.001. In all the figures, n refers to the number of mice when applicable. All statistical analyses were performed using Graphpad Prism software. No animals were excluded from analyses. Sample sizes were selected on the basis of previous experiments. Unless otherwise indicated, results are based on three independent experiments to guarantee reproducibility of findings. The RNA sequencing dataset was submitted to the Gene Expression Omnibus database with accession number GSE88840. Source Data for this study are included in the online version of the paper. Single-cell RNA-seq datasets for embryonic day 11 (E11) aorta–gonad–mesonephros endothelium, E11 CD201− pre-HSC type 1, E11 CD201+ pre-HSC type 1, E11 CD201+ pre-HSC type 2, E12.5 fetal liver HSCs (lin−Sca-1+Mac1loCD201+), E14.5 fetal liver HSCs (lin−CD45+CD150+CD48−CD201−), adult bone marrow HSCs (LKS-SLAM) were obtained from ref. 31 (GSE67120); the LKS-SLAM RNA-seq datasets were obtained from ref. 32 (GSE60808); embryonic-stem-cell-derived endothelial cells, embryonic stem cell RNA-seq datasets were obtained from ref. 42. A step-by-step protocol describing in vitro conversion of endothelial cells into HSCs can be found at Protocol Exchange43.


"Nurses are at the center of delivering a quality, compassionate patient care experience. AMN Healthcare is honored to recognize the exceptional service of nurses throughout the world this week," said Susan Salka, AMN Healthcare President and CEO.  "We are also very proud that some of the finest nurses in the country have or are currently working with the AMN team providing outstanding care for our clients. Their commitment to their profession and their advocacy for patients and their families is inspiring to us all." Per diem and travel nurses have always been an essential part of any effective healthcare system and are becoming increasingly important as the healthcare industry faces a growing nurse shortage and increased need for workforce flexibility. Research confirms that per diem and travel nurses provide quality of care that is equivalent to their permanent counterparts. The following four outstanding AMN travel nurses were chosen by a committee of AMN clients, clinicians and recruiters. These nurses showed an unwavering commitment to excellence in the nursing profession that goes far beyond their job requirements. The nominations were a testament to the high quality of nurses that AMN Healthcare has on assignment. The Innovation Award went to Jennifer Ordonez, RN BSN, an ICU Nurse on assignment in Palm Springs, CA, for her ongoing pursuit of personal and professional excellence through innovation and the advancement of patient care. As her nomination stated, "Jennifer goes beyond the care and comfort of the patient to provide life-changing care. This is the definition of excellence." The Passion Award for exemplifying the highest standards of professional excellence through leadership and extraordinary commitment to service throughout their healthcare community went to Allison Griffin, RN, BLS, CHEMO, PALS, PMD, a Pediatric Nurse on assignment in Boston, MA. Allison was selected because she "is able to impact the lives of the children and their families with her care and compassion in a meaningful way each and every day!" The Customer Focus Award for demonstrating an unwavering dedication to the improvement of patient care across all specialties and embodying the core values of the nursing profession in actions and words went to Sandra Shrago, RN, ACLS, BSN, an ICU Nurse on assignment at NewYork-Presbyterian/ Columbia University Medical Center in New York, NY. Sandra was nominated for this award because she "encompasses all that nursing should represent, going above and beyond what is standard, setting a bar for herself, for her care of others. Her amiable demeanor, warm smile and linguistic adeptness proves to be a pleasure and an asset." The Overall Commitment to Excellence Award for all of these qualities, and continually striving to improve patient care through education and innovation, displaying an unmatched passion for the profession was given to Sonia Washington, RN, ACLS, BLS, PALS, an ICU Nurse on assignment at Hilo Medical Center in Hilo, HI. "Sonia's respect of patients, families, physicians, and quick adaption to local culture make her shine," said Sonia's supervisors at Hilo Medical Center. "Sonia demonstrated the professional conduct and compassion our ICU Beacon Status department is known for and expects. Sonia has an outgoing personality, participates in our teamwork philosophy… and works above and beyond as a patient advocate." NewYork-Presbyterian NewYork-Presbyterian is one of the nation's most comprehensive, integrated academic healthcare delivery systems, whose organizations are dedicated to providing the highest quality, most compassionate care and service to patients in the New York metropolitan area, nationally, and throughout the globe. In collaboration with two renowned medical schools, Weill Cornell Medicine and Columbia University Medical Center, NewYork-Presbyterian is consistently recognized as a leader in medical education, groundbreaking research and innovative, patient-centered clinical care. For more information, visit www.nyp.org and on Facebook, Twitter and YouTube. Hilo Medical Center As the Big Island's leading provider of nationally recognized 4-star care, Hilo Medical Center (HMC) delivers a full range of services and programs. Our 20-acre campus consists of 276 beds located throughout the 137-bed acute hospital, 20-bed behavioral health unit and a 119-bed long-term care facility. We have over 1,000 employees and a medical staff comprised of 250 physicians, physician assistants and Advanced Practice Registered Nurses, representing 33 specialties. As a medical center, we have a network of nine outpatient clinics offering primary and specialty care. The hospital is a Level III Trauma Center which includes the second busiest emergency room in the state that provides 24-hour care to nearly 48,000 patients annually. In 2016, the Centers for Medicare & Medicaid Services (CMS) ranked HMC 4 stars for Overall Hospital Quality, putting our hospital in the top 20% in the nation, among the top 5 hospitals in the state, and named HMC the only 4-star hospital on Hawaii Island. Also in 2016, our Intensive Care Unit was designated as a bronze level for Beacon Award of Excellence – only the second ICU in the state to receive this designation. HMC received the 2016 American Heart Association Gold Plus Award for heart failure. HMC has also been recognized for quality long term care by Providigm for Quality Assurance & Performance Improvement Accredited Facility and Embracing Quality Award for the Prevention of Hospital Readmissions. Our long term care met the requirements for the American Health Care Association's Three Tier Level Quality Initiative Recognition Program. HMC ranks in the top 2% of hospitals in country and best in the state of Hawaii for preventing Hospital Acquired Conditions, according to CMS in 2015. The hospital is also a recipient of the 2015 Healthgrades Patient Safety Excellence Award™ and a past recipient of the Mountain Pacific Quality Health's Quality Achievement Award. HMC received the HIMSS Nicholas E. Davies Award for Excellence in 2015 for demonstrating EMR utilization to improve quality of care and financial management. We are part of the Hawaii Health Systems Corporation, a public entity established in 1996 by the State of Hawaii to fulfill the promise to provide quality, hometown healthcare. For more information, go to: www.hilomedicalcenter.org. About AMN Healthcare AMN Healthcare is the leader and innovator in healthcare workforce solutions and staffing services to healthcare facilities across the nation. The Company provides unparalleled access to the most comprehensive network of quality healthcare professionals through its innovative recruitment strategies and breadth of career opportunities. With insights and expertise, AMN Healthcare helps providers optimize their workforce to successfully reduce complexity, increase efficiency, and improve patient outcomes. AMN delivers managed services programs, healthcare executive search solutions, vendor management systems, recruitment process outsourcing, predictive modeling, medical coding and consulting, and other services. Clients include acute-care hospitals, community health centers and clinics, physician practice groups, retail and urgent care centers, home health facilities, and many other healthcare settings. For more information about AMN Healthcare, visit www.amnhealthcare.com. To view the original version on PR Newswire, visit:http://www.prnewswire.com/news-releases/national-nurses-week-amn-healthcare-commitment-to-excellence-awards-recognize-importance-of-nurses-in-patient-care-300456603.html


NEW YORK, NY--(Marketwired - May 10, 2017) - Dr. Peter M. Fleischut has been named senior vice president and chief transformation officer at NewYork-Presbyterian, effective May 1. In this new role, Dr. Fleischut will lead enterprise integration, transformation for clinical operations, innovation initiatives, logistics and pharmacy. He will also oversee the implementation of clinical technology operations for the new David H. Koch Center at NewYork-Presbyterian/Weill Cornell Medical Center, which is slated to open in 2018. "Dr. Fleischut has already led NewYork-Presbyterian to national prominence in telemedicine and patient-focused innovation," said Daniel Barchi, chief information officer at NewYork-Presbyterian. "As chief transformation officer, he will help drive integration and optimization of clinical processes across our entire enterprise." Most recently, Dr. Fleischut served as NewYork-Presbyterian's chief innovation officer, where he led the development of NYP OnDemand, NewYork-Presbyterian's comprehensive digital health program that offers a suite of services including Second Opinion, Urgent Care and Express Care. As chief transformation officer, he will maintain these responsibilities with the added focus of developing and instituting innovative solutions that are scaled across NewYork-Presbyterian's enterprise, developing governance structure for innovation, establishing cross-departmental collaborations and recruiting and mentoring innovative talent. "It's an honor to continue to revolutionize how we define and offer patient care," said Dr. Fleischut, an associate professor of anesthesiology at Weill Cornell Medicine. "I am excited to continue this work at such a wonderful institution." Dr. Fleischut joined NewYork-Presbyterian/Weill Cornell Medical Center in 2006 and has held many roles since then, including: medical director of the operating rooms, deputy quality patient safety officer, founding director of the Center for Perioperative Outcomes, chief medical information officer, chief medical operating officer and vice chairman of the Department of Anesthesiology at Weill Cornell Medicine. Dr. Fleischut retains his appointment as associate professor of anesthesiology at Weill Cornell Medicine. A graduate of Jefferson Medical College and the Wharton School at the University of Pennsylvania, Dr. Fleischut completed his residency training in anesthesiology at NewYork-Presbyterian/Weill Cornell Medical Center. He distinguished himself by becoming one of the founding members of the Housestaff Quality Council©, an organization formed to improve patient care and safety by creating a culture that promotes greater housestaff participation, and served as resident quality and patient safety officer for NewYork-Presbyterian. Dr. Fleischut has received numerous awards and honors, including NewYork-Presbyterian's Physician of the Year, the David A. Leach Award for Innovation in Quality from the Accreditation Council on Graduate Medical Education and the Weill Cornell Medical College (now Weill Cornell Medicine) Healthcare Leadership Award. In 2016 and 2017, he led NewYork-Presbyterian to CIO100 recognition, the InformationWeek Elite 100 and the President's Award from the American Telehealth Association for healthcare redesign for the release of NYP OnDemand. NewYork-Presbyterian is one of the nation's most comprehensive, integrated academic healthcare delivery systems, whose organizations are dedicated to providing the highest quality, most compassionate care and service to patients in the New York metropolitan area, nationally, and throughout the globe. In collaboration with two renowned medical schools, Weill Cornell Medicine and Columbia University Medical Center, NewYork-Presbyterian is consistently recognized as a leader in medical education, groundbreaking research and innovative, patient-centered clinical care. For more information, visit www.nyp.org and find us on Facebook, Twitter and YouTube.

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