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Klotho Deficiency Disrupts Hematopoietic Stem Cell Development and Erythropoiesis

2014; Elsevier BV; Volume: 184; Issue: 3 Linguagem: Inglês

10.1016/j.ajpath.2013.11.016

1525-2191

Sangeetha Vadakke‐Madathil, Lindsay M. Coe, Carla Casu, Despina Sitara,

Genetic Syndromes and Imprinting

Klotho deficiency is a characteristic feature of chronic kidney disease in which anemia and cardiovascular complications are prevalent. Disruption of the Klotho gene in mice results in hypervitaminosis D and a syndrome resembling accelerated aging that includes osteopenia and vascular calcifications. Given that the bone microenvironment and its cellular components considerably influence hematopoiesis, in the present study, we addressed the in vivo role of klotho in blood cell formation and differentiation. Herein, we report that genetic ablation of Klotho in mice results in a significant increase in erythropoiesis and a decrease in the hematopoietic stem cell pool size in the bone marrow, leading to impaired hematopoietic stem cell homing in vivo. Our data also suggest that high vitamin D levels are only partially responsible for these hematopoietic changes in Klotho−/− mice. Importantly, we found similar hematopoietic abnormalities in Klotho−/− fetal liver cells, suggesting that the effects of klotho in hematopoietic stem cell development are independent of the bone microenvironment. Finally, injection of klotho protein results in hematopoietic changes opposite to the ones observed in Klotho−/− mice. These observations unveil a novel role for the antiaging hormone klotho in the regulation of prenatal and postnatal hematopoiesis and provide new insights for the development of therapeutic strategies targeting klotho to treat hematopoietic disorders associated with aging. Klotho deficiency is a characteristic feature of chronic kidney disease in which anemia and cardiovascular complications are prevalent. Disruption of the Klotho gene in mice results in hypervitaminosis D and a syndrome resembling accelerated aging that includes osteopenia and vascular calcifications. Given that the bone microenvironment and its cellular components considerably influence hematopoiesis, in the present study, we addressed the in vivo role of klotho in blood cell formation and differentiation. Herein, we report that genetic ablation of Klotho in mice results in a significant increase in erythropoiesis and a decrease in the hematopoietic stem cell pool size in the bone marrow, leading to impaired hematopoietic stem cell homing in vivo. Our data also suggest that high vitamin D levels are only partially responsible for these hematopoietic changes in Klotho−/− mice. Importantly, we found similar hematopoietic abnormalities in Klotho−/− fetal liver cells, suggesting that the effects of klotho in hematopoietic stem cell development are independent of the bone microenvironment. Finally, injection of klotho protein results in hematopoietic changes opposite to the ones observed in Klotho−/− mice. These observations unveil a novel role for the antiaging hormone klotho in the regulation of prenatal and postnatal hematopoiesis and provide new insights for the development of therapeutic strategies targeting klotho to treat hematopoietic disorders associated with aging. Hematopoiesis is a complex and tightly regulated process of blood cell formation that is hierarchically coordinated. During normal hematopoiesis, diverse blood cell types are produced by the bone marrow (BM) in a manner related to physiologic requirement. Certain conditions may trigger additional production of blood cells. When the oxygen content of body tissues is low, the kidneys produce and release erythropoietin (Epo), a hormone that stimulates the BM to produce more red blood cells (RBCs). Aging is associated with disruption of normal hematopoiesis, resulting in an increase in the prevalence of anemia, the emergence of hematopoietic malignancies, and the development of leukemias.1Henry C.J. Marusyk A. 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Klotho exists in a membrane-bound form expressed at high levels in the kidney and, to a lesser extent, in other tissues, whereas a soluble form of klotho is secreted into blood, urine, and cerebrospinal fluid after cleavage of the extracellular domain.7Bloch L. Sineshchekova O. Reichenbach D. Reiss K. Saftig P. Kuro-o M. Kaether C. Klotho is a substrate for alpha-, beta- and gamma-secretase.FEBS Lett. 2009; 583: 3221-3224Crossref PubMed Scopus (200) Google Scholar, 8Chen C.D. Podvin S. Gillespie E. Leeman S.E. Abraham C.R. Insulin stimulates the cleavage and release of the extracellular domain of Klotho by ADAM10 and ADAM17.Proc Natl Acad Sci U S A. 2007; 104: 19796-19801Crossref PubMed Scopus (424) Google Scholar, 9Hu M.C. Shi M. Zhang J. Pastor J. Nakatani T. Lanske B. Razzaque M.S. Rosenblatt K.P. Baum M.G. Kuro-o M. Moe O.W. 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Klotho converts canonical FGF receptor into a specific receptor for FGF23.Nature. 2006; 444: 770-774Crossref PubMed Scopus (1474) Google Scholar Maintaining mineral ion homeostasis is critical and involves a delicate and concerted action between bone- and kidney-derived endocrine factors that operate through a complex feedback mechanism(s). Patients with CKD often present with bone diseases, such as osteopenia, osteoporosis, or osteomalacia, as a result of significant derangement of mineral metabolism.15Hruska K.A. Teitelbaum S.L. Renal osteodystrophy.N Engl J Med. 1995; 333: 166-174Crossref PubMed Scopus (195) Google Scholar, 16Mazzaferro S. Pasquali M. Pirro G. Rotondi S. Tartaglione L. The bone and the kidney.Arch Biochem Biophys. 2010; 503: 95-102Crossref PubMed Scopus (26) Google Scholar In patients with CKD, failure of appropriate fibroblast growth factor 23/Klotho signaling results in hyperphosphatemia and vascular calcifications.17Lim K. Lu T.S. Molostvov G. Lee C. Lam F.T. Zehnder D. Hsiao L.L. Vascular Klotho deficiency potentiates the development of human artery calcification and mediates resistance to fibroblast growth factor 23.Circulation. 2012; 125: 2243-2255Crossref PubMed Scopus (346) Google Scholar Klotho expression is decreased progressively with loss of renal function,18Hu M.C. Shi M. Zhang J. Quinones H. Griffith C. Kuro-o M. Moe O.W. Klotho deficiency causes vascular calcification in chronic kidney disease.J Am Soc Nephrol. 2011; 22: 124-136Crossref PubMed Scopus (714) Google Scholar whereas blood levels of fibroblast growth factor 23 are elevated and are associated with increased CVD and mortality in these patients and in patients undergoing dialysis.19Fliser D. Kollerits B. Neyer U. Ankerst D.P. Lhotta K. Lingenhel A. Ritz E. Kronenberg F. Kuen E. Konig P. Kraatz G. Mann J.F. Muller G.A. Kohler H. Riegler P. 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Suppression of aging in mice by the hormone Klotho.Science. 2005; 309: 1829-1833Crossref PubMed Scopus (1411) Google Scholar Loss of klotho is further known to cause endothelial dysfunction by promoting oxidative stress.29Kuro-o M. Klotho as a regulator of oxidative stress and senescence.Biol Chem. 2008; 389: 233-241Crossref PubMed Scopus (235) Google Scholar It has been well appreciated that aging and oxidative stress adversely affect hematopoiesis by altering the niche functions.30Hosokawa K. Arai F. Yoshihara H. Nakamura Y. Gomei Y. Iwasaki H. Miyamoto K. Shima H. Ito K. Suda T. Function of oxidative stress in the regulation of hematopoietic stem cell-niche interaction.Biochem Biophys Res Commun. 2007; 363: 578-583Crossref PubMed Scopus (98) Google Scholar, 31Lepperdinger G. Inflammation and mesenchymal stem cell aging.Curr Opin Immunol. 2011; 23: 518-524Crossref PubMed Scopus (106) Google Scholar An earlier report has also highlighted that klotho deficiency in mice results in reduced B lymphopoiesis, suggesting changes in immune regulatory functions by klotho.32Okada S. Yoshida T. Hong Z. Ishii G. Hatano M. Kuro-o M. Nabeshima Y. Nabeshima Y. Tokuhisa T. Impairment of B lymphopoiesis in precocious aging (klotho) mice.Int Immunol. 2000; 12: 861-871Crossref PubMed Scopus (51) Google Scholar In addition, klotho expression at the mRNA level has been found to be significantly decreased in resting human CD4+ lymphocytes proportionally to advancing age.33Witkowski J.M. Soroczynska-Cybula M. Bryl E. Smolenska Z. Jozwik A. Klotho: a common link in physiological and rheumatoid arthritis-related aging of human CD4(+) lymphocytes.J Immunol. 2007; 178: 771-777Crossref PubMed Scopus (59) Google Scholar Signals emanating from the BM microenvironment and extrinsic soluble factors associated with the bone and marrow milieu are known to modulate hematopoietic stem cell (HSC) proliferation and differentiation.34Mendez-Ferrer S. Michurina T.V. Ferraro F. Mazloom A.R. Macarthur B.D. Lira S.A. Scadden D.T. Ma'ayan A. Enikolopov G.N. Frenette P.S. Mesenchymal and haematopoietic stem cells form a unique bone marrow niche.Nature. 2010; 466: 829-834Crossref PubMed Scopus (2531) Google Scholar, 35Shen Y. Nilsson S.K. Bone, microenvironment and hematopoiesis.Curr Opin Hematol. 2012; 19: 250-255Crossref PubMed Scopus (58) Google Scholar Identifying the contributing factors involved in the regulation of hematopoiesis is an area of active research. 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Vitamin D receptor deletion leads to increased hematopoietic stem and progenitor cells residing in the spleen.Blood. 2010; 116: 4126-4129Crossref PubMed Scopus (25) Google Scholar Therefore, alterations in bone modeling and remodeling processes and/or mineralization seem to have a prominent effect on the modulation or formation of the hematopoietic niche. However, the regulation of mineral ion balance and hematopoiesis still remains largely a naive area. Because the bone environment and its components and the process of aging are closely linked to the regulation of hematopoiesis, and klotho deficiency is associated with a marked defect in skeletal mineralization and premature aging-like features, we hypothesized that klotho is involved in the regulation of RBC production and differentiation. In the present study, we demonstrate that loss of klotho severely affects erythropoiesis and HSC number and function. More important, we show that klotho affects hematopoiesis independently of changes in the BM environment and that the absence of klotho results in aberrant hematopoiesis prenatally, providing evidence for a novel and direct role for klotho in hematopoietic development. Although the kidney is the adult hematopoietic organ in zebra fish equivalent to mammalian BM,47Huang H.T. Zon L.I. Regulation of stem cells in the zebra fish hematopoietic system.Cold Spring Harb Symp Quant Biol. 2008; 73: 111-118Crossref PubMed Scopus (16) Google Scholar, 48Jin H. Xu J. Wen Z. Migratory path of definitive hematopoietic stem/progenitor cells during zebrafish development.Blood. 2007; 109: 5208-5214Crossref PubMed Scopus (130) Google Scholar, 49Murayama E. Kissa K. Zapata A. Mordelet E. Briolat V. Lin H.F. Handin R.I. Herbomel P. Tracing hematopoietic precursor migration to successive hematopoietic organs during zebrafish development.Immunity. 2006; 25: 963-975Abstract Full Text Full Text PDF PubMed Scopus (382) Google Scholar the present data demonstrate for the first time, to our knowledge, a link between the kidney-bone-hematopoiesis axes in the mammalian system and attest that klotho is a key factor in the process of hematopoiesis. Klotho heterozygous mice (Klotho+/−) were purchased from the Mutant Mouse Regional Resource Center (University of California, Davis, CA) and were interbred to obtain Klotho-null mice (Klotho−/−). 1α(OH)ase heterozygous mice were a gift from Dr. René St-Arnaud (Genetics Unit, Shriners Hospital, Montreal, QC, Canada). Klotho heterozygous and 1α(OH)ase heterozygous mice were bred to obtain Klotho−/−/1α(OH)ase−/− double mutants. B6.SJL-Ptprca/BoyAiTac (CD45.1; Ly5.1) mice were purchased from Taconic Farms Inc. (New York, NY). All mice were kept on a light/dark (12 hours/12 hours) cycle at 23°C and received standard laboratory chow and water ad libitum. Genomic DNA was obtained from tail snips, and routine PCR was performed to identify the genotypes. PCR conditions were as follows: klotho—initial denaturation at 94°C for 5 minutes, 35 cycles of denaturation at 94°C for 1 minute, annealing at 63°C for 1 minute and extension at 72°C for 30 seconds, and final extension at 72°C for 10 minutes; 1α(OH)ase—94°C for 5 minutes, annealing at 56°C for 1 minute and extension at 72°C for 1 minute, followed by final extension at 72°C for 10 minutes. The following primers were used: KL0787-12, 5′-GATGGGGTCGACGTCA-3′; KL0787-13, 5′-TAAAGGAGGAAAGCCATTGTC-3′; KL0787-20, 5′-ATGCTCCAGACATTCTCAGC-3′; Neo3a, 5′-GCAGCGCATCGCCTTCTATC-3′; and 1α(OH)ase (forward and reverse), 5′-GCACCTGGCTCAGGTAGCTCTTC-3′ and 5′-GTCCCAGACAGAGACATCCGT-3′. All the animals were maintained in the New York University (NYU) College of Dentistry Animal Facility in accordance with the general guidelines of the NYU School of Medicine Division of Laboratory Animal Resources. All the animal studies were approved by the NYU Institutional Animal Care and Use Committee. Peripheral blood was collected after euthanasia from 6-week-old mice by cardiac puncture into EDTA-coated tubes (BD Biosciences, San Jose, CA) to prevent clotting. Blood samples were then shipped overnight to Cornell University Animal Health Diagnostic Center (Ithica, NY) for automated complete blood cell count. BM was isolated from dissected tibiae and femora from 6-week-old mice by flushing in Iscove's modified Dulbecco's medium (IMDM) (Sigma-Aldrich, St. Louis, MO) supplemented with 20% fetal bovine serum (20% IMDM) (HyClone; Thermo Scientific, Wilmington, DE) through a 27-gauge needle (Becton Dickinson Co., Franklin Lakes, NJ). Marrow cells were dispersed by manual agitation and then were filtered to remove foreign particles. Spleens from 6-week-old mice were surgically removed and were homogenized into a cell suspension in 20% IMDM. Timed pregnant Klotho+/− female mice were sacrificed at 15.5 days postcoitum, and fetal livers were collected by caesarean section and were homogenized into a cell suspension in 20% IMDM. Genomic DNA was obtained from tail snips, and routine PCR as described previously herein was used to identify the genotypes of the embryos. Tissues were dissected from 6-week-old or E15.5 embryo mice, and flow cytometry analysis for peripheral blood, BM, spleen, and fetal liver cells was performed in a BD FACSort flow cytometer equipped with 488 argon lasers (BD Biosciences). For immunostaining, cells were washed and then resuspended in 1× PBS containing 0.1% bovine serum albumin. Mouse Fc receptor was blocked before staining using CD16/32 antibody to reduce nonspecific binding. After the addition of antibodies, cells were incubated for 40 minutes on ice; for peripheral blood, RBCs were further lysed using BD FACS lysing solution (BD Biosciences). Labeled cells were then washed with 1× PBS and were analyzed by flow cytometry. Appropriate isotype controls were kept for each set. Forward and side scatter patterns were gated to exclude debris. A total of 50,000 events were collected and analyzed using FlowJo software version 7.6.5 (Tree Star Inc., Ashland, OR). Erythroid lineage was assessed using Ter119 APC/CD71 phosphatidylethanolamine (PE) markers combined with the forward scatter (FSC) properties.50Koulnis M. Pop R. Porpiglia E. Shearstone J.R. Hidalgo D. Socolovsky M. Identification and analysis of mouse erythroid progenitors using the CD71/TER119 flow-cytometric assay.J Vis Exp. 2011; 54 (pii)Google Scholar CXCR4 expression was analyzed using PE-tagged CXCR4 antibody. Hematopoietic stem/progenitor cells were differentiated using SLAM markers (CD150 PE/CD48 APC), Sca1 fluorescein isothiocyanate (Ly6A-E), cKit Percp Cy5.5 (CD117), CD90 PE (Thy-1), and APC-tagged lineage cocktail composed of antibodies against CD3, B220 (CD45R), Ly6G and Ly6C (Gr1), CD11b (Mac1), and TER-119. CKit+Sca1+ cells were gated on the lineage-negative fraction to analyze LSK (lin−cKit+Sca1+). The LSK cells were then analyzed using a Thy-1low gate to obtain the KTLS population (LSK Thylow). CD45.1 PE and CD45.2 fluorescein isothiocyanate antibodies were used to differentiate donor and recipient populations after transplantation. All the antibodies except the SLAM markers were purchased from BD Pharmingen (San Jose, CA). SLAM markers CD150 and CD48 were purchased from eBioscience Inc. (San Diego, CA). Total RNA from bone, BM, spleen, kidney, adult liver, and fetal liver from 6-week-old or E15.5 wild-type (WT) and Klotho−/− mice was extracted using TRIzol reagent (Sigma-Aldrich) according to the manufacturer's protocol and was quantified using a NanoDrop 2000 spectrophotometer (Thermo Scientific). RNA (1 μg) from each sample was reverse transcribed to cDNA in 10 μL of final volume using a high-capacity cDNA reverse transcription kit with random hexamers (Applied Biosystems, Foster City, CA). Real-time PCR analysis was performed using an Eppendorf Mastercycler ep gradient S realplex2 machine (Eppendorf, Hamburg, Germany) in a final reaction volume of 25 μL containing 1 μL of the prepared cDNA of each gene, 12.5 μL of PerfeCta SYBR Green PCR SuperMix (Quanta BioSciences Inc., Gaithersburg, MD), and 1 μmol/L of primers amplifying the genes of interest. Thermal cycle conditions were as follows: 60°C for 2 minutes, 95°C for 10 minutes, and 40 cycles at 95°C for 15 seconds and 60°C for 1 minute. Analyses were performed in duplicate. Samples without reverse transcriptase were used as negative controls. The expression of klotho was measured in BM, spleen, kidney, and fetal liver; the expression of Epo, Hif-1α, and Hif-2α was analyzed in bone, BM, kidney, and liver; and the expression of transferrin, transferrin receptor, glucose transporter type 1, and phosphoglycerate kinase 1 was analyzed in BM and liver. All quantitative RT-PCR values were normalized to the housekeeping gene HPRT, and differences in gene expression between control (WT) mice and Klotho−/− mice were calculated based on the ΔCT method. The primer sequences are shown in Table 1.Table 1Mouse Primer Sequences Used for Real-Time PCR AnalysisGeneForward sequenceReverse sequenceKlotho5′-ACTTGGCCTTTATTAGCCGGGTCT-3′5′-AGATGGCCTCTTCCCTGTGTTCAA-3′Erythropoietin (Epo)5′-TCTACGTAGCCTCACTTCACT-3′5′-ACCCGGAAGAGCTTGCAGAAA-3′Hypoxia-inducible factor-1α (HIF-1α)5′-TCTCGGCGAAGCAAAGAGTCT-3′5′-TAGACCACCGGCATCCAGAAG-3′Hypoxia-inducible factor-2α (HIF-2α)5′-GGGAACACTACACCCAGTGC-3′5′-TCTTCAAGGGATTCTCCAAGG-3′Transferrin5′-CCCTCTGTGACCTGTGTATTG-3′5′-CTTTCTCAACGAGACACCTGAA-3′Transferrin receptor5′-TCCTGTCGCCCTATGTATCT-3′5′-CGAAGCTTCAAGTTCTCCACTA-3′Glucose transporter-1 (Glut-1)5′-CCCAGGTGTTTGGCTTAGA-3′5′-CAGAAGGGCAACAGGATACA-3′Phosphoglycerate kinase-1 (Pgk-1)5′-CACAGAAGGCTGGTGGATTT-3′5′-CTTTAGCGCCTCCCAAGATAG-3′HPRT5′-AAGCCTAAGATGAGCGCAAG-3′5′-TTACTAGGCAGATGGCCACA-3′ Open table in a new tab Serum Epo and stromal-derived factor-1α (SDF-1α) levels were determined using Quantikine enzyme-linked immunosorbent assay (ELISA) kits (R&D Systems, Minneapolis, MN) according to th