human pluripotent stem cell derived erythropoietin ...erythropoietin-producing cells ameliorate...

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KIDNEY DISEASE Copyright © 2017 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works Human pluripotent stem cellderived erythropoietin-producing cells ameliorate renal anemia in mice Hirofumi Hitomi, 1,2 Tomoko Kasahara, 1 Naoko Katagiri, 1 Azusa Hoshina, 1 Shin-Ichi Mae, 1 Maki Kotaka, 1 Takafumi Toyohara, 1 Asadur Rahman, 2 Daisuke Nakano, 2 Akira Niwa, 3 Megumu K. Saito, 3 Tatsutoshi Nakahata, 3 Akira Nishiyama, 2 Kenji Osafune 1 * The production of erythropoietin (EPO) by the kidneys, a principal hormone for the hematopoietic system, is reduced in patients with chronic kidney disease (CKD), eventually resulting in severe anemia. Although recombinant human EPO treatment improves anemia in patients with CKD, returning to full red blood cell production without fluctuations does not always occur. We established a method to generate EPO-producing cells from human induced pluripotent stem cells (hiPSCs) by modifying previously reported hepatic differentiation protocols. These cells showed increased EPO expression and secretion in response to low oxygen conditions, prolyl hydroxylase domaincontaining enzyme inhibitors, and insulin-like growth factor 1. The EPO protein secreted from hiPSC-derived EPO-producing (hiPSC-EPO) cells induced the erythropoietic differentiation of human umbilical cord blood progenitor cells in vitro. Furthermore, transplantation of hiPSC-EPO cells into mice with CKD induced by adenine treatment improved renal anemia. Thus, hiPSC-EPO cells may be a useful tool for clarifying the mechanisms of EPO production and may be useful as a ther- apeutic strategy for treating renal anemia. INTRODUCTION Erythropoietin (EPO), a potent regulator of erythropoiesis, is secreted by the kidney and liver. The kidney is the major site of EPO production in adults, and the liver is the primary organ of EPO production in the fetal and early neonatal periods, but EPO is produced by the adult liver in states of severe anemia (1, 2). EPO production is reduced in patients with chronic kidney disease (CKD), eventually resulting in renal anemia. Treatment with recombinant human EPO (rhEPO) improves the control of renal anemia compared to conventional therapies, such as blood trans- fusions, which have been associated with viral infections and iatrogenic hemosiderosis. However, although rhEPO is relatively safe, the require- ment for infusions one to three times a week makes it difficult to return to physiological control of red blood cell production (3). In addition, the total cost of rhEPO is increasing due to the increasing number of CKD patients, and patients with anemia secondary to chronic diseases may not respond well to rhEPO ( 4). Finally, very rare cases of severe anemia caused by germline mutations in EPO (5) or by anti-rhEPO autoantibodies after the administration of rhEPO have also been reported (6, 7). There- fore, new physiological therapies for renal anemia are required. The differentiation of induced pluripotent stem cells (iPSCs) (8, 9) and embryonic stem cells (ESCs) (10), which have unlimited self-renewal capability and the potential to differentiate into any cell type in the body, provides promising cell sources for regenerative medicine. The somatic cell types differentiated from these stem cells have the potential for clin- ical applications and practical use, including cell therapy, drug screen- ing, toxicology, and disease modeling. Although vigorous efforts have been made to generate multiple somatic cell types from these stem cells, the directed differentiation of EPO-producing cells (EPO cells) from iPSCs or ESCs has not yet been achieved. EPO cells were originally iden- tified in the mouse kidney (11), but the development of culture methods for these cells has been proven difficult. For this reason, we considered cultured EPO cells derived from iPSCs or ESCs for both basic and clin- ical applications. One basic research application of iPSC/ESC-derived EPO cells is to clarify the mechanisms of EPO production and secretion, given that EPO cells isolated from the kidney or liver are not readily available. Clinical applications of iPSC- or ESC-derived EPO cells could include transplanting these cells as a cell therapy for treating renal anemia. Here, we aimed to establish a differentiation method for producing EPO cells from human iPSCs (hiPSCs) and human ESCs (hESCs). We modified previously reported differentiation protocols for hepatic lineages and generated cells expressing EPO from mouse iPSCs and ESCs (miPSCs/ESCs) and hiPSCs/ESCs. These cells could physiologi- cally regulate EPO production, as evidenced by increased EPO expres- sion in response to low oxygen conditions. EPO protein secreted by hiPSC-derived EPO (hiPSC-EPO) cells induced the differentiation of human umbilical cord blood progenitor cells into erythropoietic cells in vitro. Furthermore, we confirmed that the transplantation of hiPSC-EPO cells into a mouse model of CKD improved renal anemia. RESULTS EPO-producing cells can be generated from hiPSCs/ESCs Given that the liver is the primary organ of EPO production in the fetal stage, we examined EPO mRNA expression in the human fetal liver. In particular, we measured EPO mRNA expression in various human tis- sues including human fetal liver and adult kidney (Fig. 1A). To confirm EPO expression during normal development, we examined EPO ex- pression in the embryonic day 12.5 (E12.5) mouse fetal liver by immuno- staining (Fig. 1B). The results showed that almost all hepatoblasts were positively stained for both a-fetoprotein (AFP) and EPO protein, sug- gesting that most or all hepatocytes were derived from EPO-producing hepatoblasts. We thus aimed to generate EPO cells from the hiPSC 253G4 cell line (12) by modifying previously reported protocols for hepatic lineage dif- ferentiation (fig. S1) (13, 14). As a result, hepatic lineage cells derived 1 Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan. 2 Department of Pharmacology, Faculty of Medicine, Kagawa University, Kagawa, Japan. 3 Department of Clinical Application, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan. *Corresponding author. Email: [email protected] SCIENCE TRANSLATIONAL MEDICINE | RESEARCH ARTICLE Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017 1 of 15 by guest on June 26, 2020 http://stm.sciencemag.org/ Downloaded from

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Page 1: Human pluripotent stem cell derived erythropoietin ...erythropoietin-producing cells ameliorate renal anemia in mice Hirofumi Hitomi,1,2 Tomoko Kasahara,1 Naoko Katagiri,1 Azusa Hoshina,1

SC I ENCE TRANS LAT IONAL MED I C I N E | R E S EARCH ART I C L E

K IDNEY D I S EASE

1Department of Cell Growth and Differentiation, Center for iPS Cell Research andApplication, Kyoto University, Kyoto, Japan. 2Department of Pharmacology, Facultyof Medicine, Kagawa University, Kagawa, Japan. 3Department of Clinical Application,Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan.*Corresponding author. Email: [email protected]

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

Copyright © 2017

The Authors, some

rights reserved;

exclusive licensee

American Association

for the Advancement

of Science. No claim

to original U.S.

Government Works

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nloaded from

Human pluripotent stem cell–derivederythropoietin-producing cells ameliorate renalanemia in miceHirofumi Hitomi,1,2 Tomoko Kasahara,1 Naoko Katagiri,1 Azusa Hoshina,1 Shin-Ichi Mae,1

Maki Kotaka,1 Takafumi Toyohara,1 Asadur Rahman,2 Daisuke Nakano,2 Akira Niwa,3

Megumu K. Saito,3 Tatsutoshi Nakahata,3 Akira Nishiyama,2 Kenji Osafune1*

Theproduction of erythropoietin (EPO) by the kidneys, a principal hormone for the hematopoietic system, is reducedin patients with chronic kidney disease (CKD), eventually resulting in severe anemia. Although recombinant humanEPO treatment improves anemia inpatientswith CKD, returning to full redblood cell productionwithout fluctuationsdoes not always occur. We established a method to generate EPO-producing cells from human induced pluripotentstem cells (hiPSCs) bymodifying previously reported hepatic differentiation protocols. These cells showed increasedEPO expression and secretion in response to low oxygen conditions, prolyl hydroxylase domain–containing enzymeinhibitors, and insulin-like growth factor 1. The EPOprotein secreted fromhiPSC-derived EPO-producing (hiPSC-EPO)cells induced the erythropoietic differentiation of human umbilical cord blood progenitor cells in vitro. Furthermore,transplantation of hiPSC-EPO cells into mice with CKD induced by adenine treatment improved renal anemia. Thus,hiPSC-EPO cells may be a useful tool for clarifying the mechanisms of EPO production and may be useful as a ther-apeutic strategy for treating renal anemia.

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by guest on June 26, 2020tp://stm

.sciencemag.org/

INTRODUCTIONErythropoietin (EPO), a potent regulator of erythropoiesis, is secretedby the kidney and liver. The kidney is the major site of EPO productionin adults, and the liver is the primary organ of EPO production in thefetal and early neonatal periods, but EPO is produced by the adult liverin states of severe anemia (1, 2). EPO production is reduced in patientswith chronic kidney disease (CKD), eventually resulting in renal anemia.Treatmentwith recombinant humanEPO (rhEPO) improves the controlof renal anemia compared to conventional therapies, such as blood trans-fusions, which have been associated with viral infections and iatrogenichemosiderosis. However, although rhEPO is relatively safe, the require-ment for infusions one to three times a weekmakes it difficult to returnto physiological control of red blood cell production (3). In addition, thetotal cost of rhEPO is increasing due to the increasing number of CKDpatients, and patients with anemia secondary to chronic diseasesmay notrespondwell to rhEPO (4). Finally, very rare cases of severe anemia causedby germline mutations in EPO (5) or by anti-rhEPO autoantibodiesafter the administration of rhEPOhave also been reported (6, 7). There-fore, new physiological therapies for renal anemia are required.

The differentiation of induced pluripotent stem cells (iPSCs) (8, 9)and embryonic stemcells (ESCs) (10),whichhave unlimited self-renewalcapability and the potential to differentiate into any cell type in the body,provides promising cell sources for regenerative medicine. The somaticcell types differentiated from these stem cells have the potential for clin-ical applications and practical use, including cell therapy, drug screen-ing, toxicology, and disease modeling. Although vigorous efforts havebeenmade to generate multiple somatic cell types from these stem cells,the directed differentiation of EPO-producing cells (EPO cells) fromiPSCs or ESCs has not yet been achieved. EPO cells were originally iden-tified in themouse kidney (11), but the development of culturemethods

for these cells has been proven difficult. For this reason, we consideredcultured EPO cells derived from iPSCs or ESCs for both basic and clin-ical applications. One basic research application of iPSC/ESC-derivedEPOcells is to clarify themechanisms of EPOproduction and secretion,given that EPO cells isolated from the kidney or liver are not readilyavailable. Clinical applications of iPSC- or ESC-derived EPO cells couldinclude transplanting these cells as a cell therapy for treating renal anemia.

Here, we aimed to establish a differentiation method for producingEPO cells from human iPSCs (hiPSCs) and human ESCs (hESCs). Wemodified previously reported differentiation protocols for hepaticlineages and generated cells expressing EPO from mouse iPSCs andESCs (miPSCs/ESCs) and hiPSCs/ESCs. These cells could physiologi-cally regulate EPO production, as evidenced by increased EPO expres-sion in response to low oxygen conditions. EPO protein secreted byhiPSC-derived EPO (hiPSC-EPO) cells induced the differentiation ofhuman umbilical cord blood progenitor cells into erythropoietic cellsin vitro. Furthermore, we confirmed that the transplantation ofhiPSC-EPO cells into a mouse model of CKD improved renal anemia.

RESULTSEPO-producing cells can be generated from hiPSCs/ESCsGiven that the liver is the primary organ of EPO production in the fetalstage, we examined EPOmRNA expression in the human fetal liver. Inparticular, we measured EPOmRNA expression in various human tis-sues including human fetal liver and adult kidney (Fig. 1A). To confirmEPO expression during normal development, we examined EPO ex-pression in the embryonic day 12.5 (E12.5)mouse fetal liver by immuno-staining (Fig. 1B). The results showed that almost all hepatoblasts werepositively stained for both a-fetoprotein (AFP) and EPO protein, sug-gesting that most or all hepatocytes were derived from EPO-producinghepatoblasts.

We thus aimed to generate EPO cells from the hiPSC 253G4 cell line(12) by modifying previously reported protocols for hepatic lineage dif-ferentiation (fig. S1) (13, 14). As a result, hepatic lineage cells derived

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Fig. 1. Differentiation ofhiPSCs/ESCs into EPO-producing cells. (A) The ex-pression of EPO mRNA wasexamined in human fetaland adult liver tissue, hu-man fetal and adult kidney,and human adult brain tis-sue. Bacterial artificial chro-mosome (BAC) containingEPO complementary DNA(cDNA) was used as a posi-tive control. (B) EPO proteinexpression was evaluatedusingimmunocytochemistry.Fetalmouse liver (E12.5)waspositively stained with anti-EPO (green) and anti-AFP(red) antibodies. (C) Expres-sion of marker genes for en-doderm and liver lineagesand EPO in cultures of differ-entiatedhiPSCs.Human liverspecimens were used as apositive control. hiPSCsweredifferentiated into the he-patic lineage (stages 1 to 3),as shown in fig. S1. (D andE) Time-course analyses ofthe expressionof EPOmRNA(D) and protein (E). The ex-pression of EPO mRNA wasmeasured using qRT-PCR in(D), and Western blottingwas performed with anti-EPO antibodies in (E). Eachvalue was normalized tothat of the sample on day 0.(F) EPO protein expressionwasevaluatedusing immuno-cytochemistry.hiPSC-EPOcellswere positively stained withanti-EPO (green) and anti-AFP (red) antibodies. (G) Se-cretoryvesicles (blackarrows)were observed in the cyto-plasm of hiPSC-EPO cells bytransmissionelectronmicros-copy. The right lower panelis an image of E11.5 fetalmouse liver for comparison.(H) A temporal analysis ofthe EPO protein concentra-tions in the culture mediumofhiPSC-EPOcellsmeasuredusing ELISA. (I) EPO mRNAexpression was observed inthe differentiation culturesof multiple hiPSC/ESC lines.

Each value was normalized to that of the hiPSC 253G4 cell line on day 0. The data were obtained from four independent experiments and are means ± SEM in (D), (E), (H), and(I). *P < 0.05 versus day 0 in (D), (E), and (H); analysis of variance (ANOVA) with Bonferroni’s test. Scale bars, 40 mm (B and F) and 2 mm (G).

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from hiPSCs expressed EPOmRNA at stage 2, and the expression dis-appeared at stage 3 (Fig. 1C). The expression of both SOX17 and AFPwas also detected in stage 2 cells, confirming that the differentiated cellsobtained in the hepatic differentiation step (stage 2) but not in the mat-uration step (stage 3) mainly produced EPO. We next examined thetime course of EPO mRNA expression in stage 2 cells and found thatthe expression gradually increased with statistical significance onculture day 8 (55.49 ± 6.22–fold increase on day 8 versus day 0; P <0.05; Fig. 1D). Consistent with the results of the quantitative reversetranscription polymerase chain reaction (qRT-PCR) analysis, the in-crease in EPO protein expression became statistically significant onday 12 (3.42 ± 0.53–fold increase versus day 0; P < 0.05) according toWestern blot analysis (Fig. 1E). The EPO protein expression was alsoconfirmed by immunostaining, which showed signals in the cytoplasmof the induced cells at stage 2 and confirmed that the EPOcells were alsopositive for the hepatic lineage marker AFP (Fig. 1F). AFP and anotherhepatic lineage marker, albumin, along with the endodermmarkers he-patocyte nuclear factor-1b (HNF-1b),HNF-4, Sal-like protein 4 (SALL4),GATA4, and cytokeratin 19 (CK19), were expressed in almost all hiPSC-EPOcells (fig. S2). In addition, EPOexpression, as analyzedby immuno-staining, was similar across the hiPSC-EPO cell population (fig. S2).Ultrastructural observations using transmission electron microscopyrevealed that EPO cells contained numerous vesicles in the cytoplasm,but thesewere less dense than in themouse fetal liver (Fig. 1G).Next, weexaminedEPOprotein secretionbyhiPSC-EPOcells into the cultureme-dium. The concentration of EPO protein measured using the enzyme-linked immunosorbent assay (ELISA) showed a statistically significantincrease in the culturemediumof hiPSC-EPOcells at stage 2 after 8 daysof culture (0.20 ± 0.05 ng/ml on day 8 versus day 0; P < 0.05; Fig. 1H).

Different hiPSC/ESC lines vary in their differentiation potential(14, 15). We thus confirmed the generation of EPO cells from multiplehiPSC/ESC lines using our differentiation protocol, including eighthiPSC lines (201B6, 201B7, 253G1, 253G4, 585A1, 604A3, 606A1,and 606B1) (8, 12, 14) and two hESC lines (KhES3 and H9) (10, 16).The results suggested that EPO cells can be differentiated frommultiplehiPSC/ESC lines using our modified hepatic differentiation protocol,although the EPOmRNA expression was variable among the cell lines(Fig. 1I). We then investigated the correlation of EPO mRNA expres-sion and protein secretion and mRNA expression of the hepatoblastmarkers AFP and DLK1 (delta-like noncanonical Notch ligand 1) inthe 253G4, 585A1, and KhES3 cell lines. EPO mRNA expression andprotein secretion correlated with the expression of hepatoblast markersin the 253G4 and 585A1 cell lines but not when the 585A1 and KhES3cell lines were compared, suggesting that both the differentiation po-tential of hiPSC/ESC-EPO cells and EPOproduction and secretion ca-pacity were variable among the cell lines (fig. S3).

hiPSC-EPO cells show long-term proliferative andEPO-producing capacity in vitroWe evaluated the long-term proliferation and EPO-producing capacityof hiPSC-EPO cells in vitro. The number of hiPSC-EPO cells increasedin a time-dependent manner (Fig. 2A), and some hiPSC-EPO cells atstage 2 on day 12 of culture stained positively for the cell proliferationmarker Ki67 (Fig. 2B). By developing an in vitro maintenance culturemethod for hiPSC-EPO cells using a stage 2 culture medium and gentlecell passaging with Accutase treatment, we could increase the numberof cells about 1000 times after 10 cell passages in 70 days (Fig. 2C),during which time both EPOmRNA expression and protein secretionwere maintained (Fig. 2, D and E). These results suggest that hiPSC-

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

EPO cells have a long-term proliferative and EPO-producing capacityin vitro.

EPO-producing cells can be generated from miPSCs/ESCsWe next examined whether miPSCs/ESCs could also be induced to dif-ferentiate into EPO cells. By modifying our differentiation method fordriving hiPSCs to becomeEPOcells, we developed a differentiation pro-tocol for obtaining EPO cells from miPSCs/ESCs (fig. S4). qRT-PCRanalyses showed that the miPSC cell line 492B-4 treated with our differ-entiation protocol expressed EpomRNA during stage 2 of developmentand that the expression became statistically significant on culture day 10(5.88± 1.02–fold increase versus day 0;P<0.05; Fig. 3A).We subsequentlyconfirmed that our differentiation protocol for inducing EPO cells alsoworked with the mESC D3 cell line (Fig. 3B). EPO protein secretion intothe culturemediumwas also confirmed by ELISA, which showed that theprotein concentration increased in a time-dependent manner (Fig. 3C).These results indicate that our differentiation protocol could be used togenerate EPO cells from both miPSCs/ESCs and hiPSCs/ESCs.

IGF-1 treatment augments EPO production and secretion inhiPSC-EPO cellsTo increase EPO production and improve the differentiation efficiencyof EPO cells from hiPSCs/ESCs, we evaluated the effects of variousgrowth factors and chemicals on EPO expression during stage 2 culture.The screening results for 46 different factors showed that the additionof human insulin-like growth factor 1 (IGF-1) most potently aug-mented EPOmRNA expression (fig. S5). We next compared the effectsof IGF-1 onEPO expressionwith that induced by related factors, such asIGF-2 and insulin (Fig. 4A). IGF-1 treatment at 100 ng/ml increasedEPO gene expression about sixfold compared to that observed incontrols treated with no additional factors, whereas the same concen-tration of IGF-2 or insulin had little or no effect. Western blot analysesconfirmed that stage 2 hiPSC-EPO cells expressed the IGF-1 receptor(Fig. 4B). We then examined the dose response and time course ofEPO gene expression and protein secretion in the culture medium afterIGF-1 treatment. Both EPO gene expression and protein secretion wereaugmentedby IGF-1 treatment in adose-dependent and time-dependentmanner (Fig. 4, C to F). IGF-1 treatment at 100 ng/ml resulted in a sig-nificant increase in EPOmRNA expression after 4 days (82.14 ± 10.86–fold increase on day 4 versus day 0; P < 0.05) and EPOprotein secretionafter 8 days (0.19 ± 0.02 ng/ml on day 8; P < 0.05), which continued toincrease up to day 12. Neither AFP nor ALBUMINmRNA expressionwas significantly changed by IGF-1 treatment, excluding the possibilitythat IGF-1 altered the differentiation state of the hiPSC-EPO cells (Fig.4, G andH). These results indicate that IGF-1 treatment promoted EPOproduction by hiPSC-EPO cells.

Hypoxia augments EPO production by hiPSC-EPO cellsIt has been reported that EPO production is regulated by oxygen con-centrations through hypoxia-inducible factors (HIFs) and their regula-tors, prolyl hydroxylase domain–containing enzymes (PHDs) (17). Weexaminedwhether this was also the casewith hiPSC-EPOcells and eval-uated the effects of hypoxia on EPO gene expression and protein secre-tion in hiPSC-EPO cells (Fig. 5, A and B). The results showed that bothEPOmRNA expression and protein secretion gradually increased evenunder normoxic condition (21% oxygen) for the first 8 days of stage 2development in parallel with the AFP and ALBUMIN mRNA expres-sion (Fig. 5, C and D), indicating that the differentiation or maturationof hiPSC-EPO cells proceeded to some extent during these 8 days.

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Nevertheless, hypoxia (1% oxygen) increased EPO mRNA expressionandEPOprotein concentration in the culturemediummore thanundernormoxic conditions, with statistically significant differences in EPOmRNA expression on day 6 and in EPO protein secretion on day 10(P < 0.05; Fig. 5, A and B). In addition, the sensitivity of hiPSC-EPOcells to the hypoxic conditionswas evaluated.Compared to thenormoxiccondition (21%oxygen), twohypoxic conditions (1 and5%oxygen) aug-mentedEPOmRNAexpression andprotein secretiondose-dependently(fig. S6, A and B). In contrast, hypoxia did not significantly alterAFP orALBUMIN mRNA expression, excluding the possibility that hypoxiaaffected the differentiation state of the hiPSC-EPO cells (Fig. 5, C andD). These results suggest that hiPSC-EPO cells are functionally similarto EPO cells in vivo in that they produce and secrete EPO in response tothe oxygen concentration.

We then evaluated the effects of three PHD inhibitors: dimethyl-oxalylglycine (DMOG), deferoxamine (DFO), and FG4592 (18). First,we confirmed the expression of HIF subunits.HIF-1a,HIF-2a,HIF-3a,HIF-1b, andHIF-2bmRNAexpressionwas detected in both hiPSC-EPO

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

and HepG2 cells, an immortalized human cell line that secretes EPOprotein in an oxygen-dependent manner (fig. S6C). We also confirmedthe protein expression of HIF-1a and HIF-2a by immunostaining inhiPSC-EPOcells (fig. S6D). In addition, bothhiPSC-EPOandHepG2cellsexpressed PHD1, PHD2, and PHD3mRNAs, sharing the expression ofthe same HIF and PHD genes with the human fetal liver (fig. S6C). InhiPSC-EPO cells, DFO and FG4592 significantly augmented EPO pro-duction in a dose-dependentmanner, whereas DMOGdid not (Fig. 5, EandF, and fig. S6, E andF). Furthermore, an inhibitor ofHIF-1dimeriza-tion,Acriflavine, attenuatedEPOproduction inhiPSC-EPOcells (Fig. 5G).In contrast, inHepG2 cells, DFO andDMOGaugmented EPO produc-tion, but FG4592 did not (Fig. 5, E and F). We also confirmed that hy-poxic culture conditions stabilized HIF-1a and HIF-2a, as evidencedby the augmentation of nuclear signals revealed using anti–HIF-1a andanti–HIF-2a immunostaining of hiPSC-EPO and HepG2 cells (Fig.5H). IGF-1 treatment also stabilized HIF-1a andHIF-2a and increasedEPO expression (Fig. 4D and fig. S6, G and H). These data suggest thathiPSC-EPO cells produced EPO via the HIF-PHD pathway and that

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Fig. 2. Cell proliferationand EPO-producing ca-pacity of hiPSC-EPO cells.(A) Number of stage 2 hiPSC-EPOcellsduringculture.hiPSCswere differentiated into EPO-producing cells (stage 2), asshown in fig. S1. (B) The ex-pressionof a cell proliferationmarker, Ki67, was measuredby immunocytochemistry.(C)NumberofhiPSC-EPOcellsduring maintenance culture.(D and E) Time-course analy-ses of EPOmRNA expression(D) and EPO protein secre-tion (E) using RT-PCR andEPOELISA, respectively. Eachvaluewas normalized to thatof the stage 2 sample on day0 (D). The data from four in-dependent experiments aremeans ± SEM in (A), (C), (D),and (E). *P < 0.05 versus day0 in (A), (D), and (E); ANOVAwith Bonferroni’s test. Scalebars, 40 mm (B).

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hiPSC-EPO and HepG2 cells may use different regulatory machineriesto express EPO.

EPO protein produced by hiPSC-EPO cells showserythropoietic activity in vitroEPO is essential for erythropoiesis and is the primary regulator of thisprocess. To evaluate the erythropoietic potential of human EPOproteinproduced by hiPSC-EPO cells (hiPSC-EPO protein), we performedclonogenic hematopoietic progenitor cell assays inmethylcellulose withpurified and concentrated hiPSC-EPO protein (Fig. 6A). Similar to ob-servations for rhEPO, treatment with hiPSC-EPO protein significantlyincreased the number of erythroid burst-forming unit colonies [BFU-E;7.33 ± 1.20 colonies with hiPSC-EPO protein (2.8 ng/ml) (n = 3) versus2.00 ± 0.00 colonies without EPO treatment (n = 3); P < 0.05; Fig. 6, B andC] but not the other types of colonies, such as granulocyte-macrophagecolony-forming units [CFU-GM; 9.33 ± 1.45 colonies with hiPSC-EPOprotein (2.8 ng/ml) (n = 3) versus 6.00 ± 1.16 colonies without EPOtreatment (n = 3); P > 0.05], differentiated from cord blood CD34+

hematopoietic progenitor cells (Fig. 6C and table S1). The efficiencyof BFU-E formation induced by hiPSC-EPO protein was similar to thatachieved with rhEPO protein [6.67 ± 0.33 colonies with rhEPO protein(2.8 ng/ml); n = 3; P > 0.05]. However, the erythropoietic effects of thehiPSC-EPOprotein (2.8 ng/ml)weremarkedly suppressed by adding hu-

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

man EPO–specific antibodies (Fig. 6D andtable S1). These results suggest that hiPSC-EPO cells secrete functional EPO proteinthat can stimulate erythropoiesis to a simi-lar extent as conventional rhEPO protein.

EPO protein produced byhiPSC-EPO cells showserythropoietic activity in vivoA renal anemia mouse model was devel-oped using adenine treatment of C57BL/6mice (19, 20). An oral gavage of adenine(50 mg/kg body weight daily for 4 weeks)decreased hematocrit and hemoglobinconcentration. Themicewere then furthertreated after dividing them into the follow-ing five groups: group 1, no adenine treat-ment(control;n=6);group2, rhEPO(28ng),subcutaneous injection (n = 10); group 3,rhEPO (0.56 ng), subcutaneous injection(n = 10); group 4, hiPSC-EPO protein(0.56 ng), subcutaneous injection (n = 10);and group 5, saline, subcutaneous injection(n = 10). Notably, the subcutaneousinjection of rhEPO and hiPSC-EPO proteininto the mice for 4 weeks resulted in recov-ery of both the hematocrit [normal rangein C57BL/6 mice, 36.8 to 52.7%; 39.78 ±0.51% with hiPSC-EPO protein (n = 10)versus 33.85 ± 0.52% with saline (n = 10);P < 0.05; Fig. 7, A and B] and hemoglobinconcentration (normal range in C57BL/6mice, 11.0 to 15.2 g/dl; 11.40 ± 0.16 g/dlwith hiPSC-EPO protein versus 9.52 ±0.27 g/dl with saline; P < 0.05; Fig. 7C)(21) towithin the normal range.However,

this treatment did not increase the number of white blood cells or plate-lets (Fig. 7D), suggesting that hiPSC-EPO protein induces the differen-tiation of only erythrocyte progenitor cells. The adenine-treated micedid not show any significant changes in mean corpuscular volume ormean corpuscular hemoglobin concentration (Fig. 7E) or body weight(fig. S7A) after 4 weeks of treatment with rhEPO or hiPSC-EPO protein.Whereas themouseEPOconcentrationsmeasured usingELISAwerenotaffected by human EPO treatment in the adenine-treated mice (Fig. 7F),the human EPO concentrations increased after treatment with rhEPO(28 ng) or hiPSC-EPO protein (0.56 ng), suggesting that the mouse re-nal anemia was improved by treatment with human EPO protein (Fig.7G). We also confirmed that both hiPSC-EPO protein and rhEPO im-proved renal anemia in a dose-dependent manner (fig. S7B). The effi-ciency of renal anemia recovery was calculated on the basis of the increaseinhematocrit perweek.TreatmentwithhiPSC-EPOproteinmore efficientlyimproved the anemic state than did rhEPO treatment [3.46 ± 0.47%/weekwith hiPSC-EPO protein (0.56 ng) versus 1.00 ± 0.52%/week with rhEPO(0.56 ng); Fig. 7H]. To elucidate the molecular mechanisms responsible,we measured the glycosylation pattern of hiPSC-EPO protein by lectinmicroarray analysis (Fig. 7I). Although the glycosylation pattern resembledthat of two clinically available rhEPOs (epoetin-a and epoetin-b), theglycosylationof hiPSC-EPOproteinwasmore complicated, as evidencedby its reactivity with numerous lectins. Because glycosylation modulates

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Fig. 4. Effects of IGF-1 treatment on EPO expression and secretion in hiPSC-EPO cells. (A) Effects of IGF-1, IGF-2, and insulin treatment on the EPOmRNA expressionfor stage 2 hiPSC-EPO cells on culture day 8 using qRT-PCR analysis. Each value was normalized to that of control samples (no growth factor). A sample of human fetal liverwas used as a positive control. (B) IGF-1 receptor expression was measured using Western blot analysis. Representative data are shown for four independent experiments.(C and D) Concentration-dependent effects of IGF-1 treatment on EPO mRNA expression (C) and protein secretion (D) for stage 2 hiPSC-EPO cells on culture day 8. Eachvalue was normalized to that of control hiPSCs treated with no IGF-1. (E and F) Time-course analyses of the IGF-1–induced EPOmRNA expression (E) and protein secretion(F) for hiPSC-EPO cells. Each value was normalized to that of samples on day 0 in (E). (G andH) ThemRNA expression of AFP (G) and ALBUMIN (H) was analyzed using qRT-PCR.Each value was normalized to control stage 2 hiPSC-EPO cells on culture day 8 not treated with IGF-1. The data were obtained from four independent experiments and aremeans ± SEM in (A) and (C) to (H). *P < 0.05 versus control samples without IGF-1 treatment in (A) and (C) to (F); ANOVA with Bonferroni’s test.

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the efficacy and degradation of EPO (22), we also compared half-livesin vivo and found that hiPSC-EPO had a slightly longer half-life than didepoetin-b inmice, although thedifferencemaynotbe clinically significant(Fig. 7J). These data suggest that EPO protein produced by hiPSC-EPOcells was capable of inducing erythropoiesis in vivo and in vitro and that

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

different glycosylation patterns may help explain why hiPSC-EPO pro-tein induced a slightly greater hematocrit in mice with renal anemia thandid rhEPO. However, we should note that there may be no biologicaldifference between the two EPO proteins because increased doses ofrhEPO resulted in bioequivalence with hiPSC-EPO protein (fig. S7B).

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control samples (HepG2 cells) in (E) and (F); ANOVA with Bonferroni’s test (A, B, E, and F) and Student’s t test (G). Scale bars, 20 mm (H).

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Transplantation of hiPSC-EPO cells improves renal anemia inmiceImmunodeficient (NOD.CB17-Prkdcscid/J) mice were treated with ade-nine by oral gavage (50 mg/kg body weight daily for 5 weeks) to inducerenal anemia. To examine the feasibility of hiPSC-based cell therapy fortreating renal anemia, we transplanted 20 aggregates of hiPSC-EPO

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

cells (5.0 × 105 cells per aggregate) underthe kidney subcapsules of the anemic mice.Notably, transplantation of hiPSC-EPOcells significantly improved renal anemiaafter 4 weeks compared with saline treat-ment [hematocrit, normal range in NOD.CB17-Prkdcscid/Jmice, 42.3 to48.1%; 42.56±1.74% with hiPSC-EPO cells (n = 6) versus34.60 ± 5.23% with saline (n = 6); P < 0.05;Fig. 8A] (21). The human EPO concentra-tion in the host mouse serum at 4 weeksafter transplantation was detectable usingELISA and was significantly increased inthe mice that underwent transplantationwith hiPSC-EPO cells [1.42 ± 0.23 ng/mlwith hiPSC-EPO cells (n = 6) versus 0.07 ±0.02 ng/ml with saline (n= 6); P < 0.05; Fig.8B]. Mice with renal anemia transplantedwithhiPSC-EPOcells did not develop poly-cythemia during the observation period(for up to 28 weeks after transplantation),and their hematocrit remained within thenormal range. In contrast, renal anemiapersisted in adenine-treated mice that didnot undergo transplantation (46.40 ±3.49% with hiPSC-EPO cells (n = 4) versus37.23±2.86%with saline (n=4) at 28weeksafter transplantation;P<0.05; Fig. 8C). In ad-dition, humanEPOprotein inmouse serumwas also detectable for up to 28 weeks aftertransplantation (Fig. 8D). We performedhistological analyses of the hiPSC-EPOcell aggregates injected under the renal sub-capsule of immunodeficientmice. Hematox-ylin and eosin (H&E) staining revealedstructures consisting of homogeneous cellsin thegrafts (Fig. 8E). Immunohistochemicalanalysis revealed that the transplantedhiPSC-EPO cell aggregates maintainedEPO expression in the subcapsule of hostmouse kidneys for at least 28 weeks aftertransplantation (Fig. 8E). Although AFPexpression was decreased 8 weeks aftertransplantation, albumin expression wasdetectable for 28weeks after transplantation(Fig. 8E). Blood vessel–like tubular struc-tures were found inside the grafts and con-tained red blood cells (Fig. 8F). The newvasculature derived fromhostmicewas con-firmed by the expression of mouse CD31/platelet endothelial cell adhesionmolecule-1(PECAM-1), indicating that mouse hostblood vessels may have supplied nutritionandoxygen to thegrafts. EPOsecretion from

transplanted hiPSC-EPO cells after phlebotomy was evaluated (Fig. 8G).We found that moderate phlebotomy (with the hematocrit decreasedfrom 44% to 34%) in the group transplanted with hiPSC-EPO cell aggre-gates resulted in an increase in circulating human EPO, but this did notoccur in control untransplantedmice (Fig. 8G). These data indicate that

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Fig. 6. Effectsof hiPSC-EPOproteinonerythropoiesis invitro. (A) Schematic of clonogenic hematopoietic progenitorassays using methylcellulose-based semisolid medium. (B) Representative images of BFU-E induced by rhEPO (left) andhiPSC-EPO protein (right). (C) Number of clonal colonies on semisolidmedium containing ST3 supplementedwith rhEPO(0.28 and2.8 ng/ml) or hiPSC-EPOprotein (0.28 and2.8 ng/ml) (n=3). (D) Number of clonal colonies on semisolidmediumcontaining different concentrations of neutralizing antibodies against human EPO in addition to ST3 and hiPSC-EPO pro-tein (2.8 ng/ml) (n = 3). The datawere obtained from three independent experiments and aremeans ± SEM in (C) and (D).Scale bars, 200 mm (B).

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Fig. 7. Therapeutic effects of hiPSC-EPO protein on renal anemia in adenine-treatedmice. (A toH) Renal anemiawas induced by adenine treatment (50mg/kg bodyweightdaily for 4 weeks) in male C57BL/6 mice. The mice were then treated with rhEPO or hiPSC-EPO protein. (A) Time-course analyses of the hematocrit (Hct) measured using glass capillary

tubes. (B)Hematocrit after4weeksof treatmentwith rhEPOorhiPSC-EPOprotein. (C)Hemoglobinconcentrationsafter4weeksof treatmentwith rhEPOorhiPSC-EPOproteinanalyzedbyELISA. The gray shaded areas indicate the normal hematocrit in C57BL/6mice in (A) and (B) and the normal hemoglobin concentration in C57BL/6mice in (C). (D) Number of red bloodcells (RBC), white blood cells (WBC), and platelets after 4weeks of treatmentwith rhEPOor hiPSC-EPOprotein. (E)Mean corpuscular volume (MCV),mean corpuscular hemoglobin (MCH),andmean corpuscular hemoglobin concentration (MCHC) after 4 weeks of treatment with rhEPO or hiPSC-EPO protein. (F and G) The concentrations of mouse (F) and human (G) EPOprotein inmouse serumweremeasured after 4weeks of treatmentwith rhEPOor hiPSC-EPOprotein using ELISA. (H) The efficiency of renal anemia recovery after treatmentwith 0.56 ngof rhEPOor hiPSC-EPOproteinwas calculated on thebasis of the increase in hematocrit perweek (DHct/week). (I) Glycosylation patterns of hiPSC-EPOand rhEPOproteinweremeasuredusing lectin microarray assays. The data are means ± SEM (n = 4). Abbreviations are defined in table S3. (J) Half-lives of the hiPSC-EPO protein and rhEPO in vivo were evaluated bymeasuring EPOprotein concentrations inmouse serumafter subcutaneous injectionof hiPSC-EPOor rhEPOusing ELISA. Thedata from three independent experiments aremeans±SEMin (A) to (H) and (J).n=6 for control andn=10 for rhEPO (28ng), rhEPO (0.56ng), hiPSC-EPOprotein (0.56ng), andsaline. *P<0.05versus saline in (A) to (D), control in (G), and rhEPO in (H);ANOVA with Bonferroni’s test (A to G) and Student’s t test (H).

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hiPSC-EPOcells cansurvive inhostmicewithrenalanemia, cansecrete func-tionalhumanEPOprotein, andcan improve renal anemia in these animals.

DISCUSSIONAlthough rhEPO treatment is beneficial for CKD patients with renalanemia, several problems remain to be addressed. First, the increasing

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

number of CKD patients is expanding the demand for rhEPO treat-ment, which, in turn, increases the total cost of this therapy. Second,it is difficult to physiologically control renal anemia using rhEPO treat-ment. The intermittent administration of rhEPO causes fluctuations inhemoglobin concentrations (3), which is associated with an increased in-cidence of cardiovascular events (23). In addition, the target hemoglobinconcentration for rhEPOtreatment remains controversial, andhemoglobin

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Fig. 8. Therapeutic effects ofthe transplantation of hiPSC-EPO–producing cells on renalanemia in adenine-treatedmice. Renal anemia was in-duced using adenine treatment(50 mg/kg body weight dailyfor 5weeks) in immunodeficientmice (NOD.CB17-Prkdcscid/Jmice). Twenty aggregates ofhiPSC-EPO cells (5.0 × 105 cellsper aggregate) were trans-planted into the kidney sub-capsules of mice with renalanemia. (A) Hematocrit was ex-amined during the first 4 weeksafter transplantationusingglasscapillary tubes. (B) Human EPOconcentrations inmouse serumat 4 weeks after transplanta-tionweremeasuredusingELISA.(C) Hematocrit was examinedfor up to 28 weeks after trans-plantation. The gray shadedareas in (A) and (C) indicate thenormal hematocrit range inNOD.CB17-Prkdcscid/J mice.(D) Human EPO concentrationsin mouse serum after trans-plantation were measuredusing ELISA. (E) The hiPSC-EPO–producing cell grafts were eval-uated using immunohisto-chemistry for EPO (green), AFP(red) and ALBUMIN (red) andusing H&E staining. (F) Thenew vasculature in the graftsderived from host mice wasexamined by anti-mouse CD31/PECAM-1 immunostaining andH&E staining. (G) The humanEPO concentrations in hostmouse serum after phlebotomyweremeasured using ELISA. Thedata from three independentexperiments are means ± SEM;n = 6 for hiPSC-EPO–producingcells and saline in (A) and (B).The data from two independentexperiments are means ± SEM;n= 4 for hiPSC-EPO cells and sa-line in (C), (D), and (G). *P < 0.05versus control; ANOVA withBonferroni’s test (A, C, D, andG) and Student’s t test (B). Scalebars, 40 mm (E) and 20 mm (F).

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concentrations in most patients are lower than those observed inhealthy subjects (24). The physiological control of renal anemiabased on a stable, normal range of hemoglobin concentrations mayhelp in the treatment of CKD patients.

hiPSCs/ESCs are potential cell sources for regenerative medicine.Here, we generated EPO-producing cells from hiPSCs/ESCs andmiPSCs/ESCs. These cells expressed EPO mRNA and protein andincreased the production and secretion of EPO protein in response tolow oxygen conditions. The secreted EPO protein induced the erythro-poietic differentiation of human umbilical cord blood CD34+ hemato-poietic cells in vitro and improved renal anemia in adenine-treatedmicein vivo. Furthermore, the transplantation of hiPSC-EPO cells reversedrenal anemia, with a return to the normal range for the hematocrit inimmunodeficient mice. Therefore, EPO cells generated from hiPSCs/ESCs may be useful as a cell therapy for treating renal anemia.

rhEPO treatment is associated with a very rare side effect of severeanemia due to the generation of anti-rhEPO autoantibodies (6). Al-though further studies are needed to clarify the immunological effectsof transplanted hiPSC-EPO cells, the cell types generated from a pa-tient’s own iPSCs may provide a safer cell therapy in terms of the im-mune response.Moreover, we confirmed that hiPSC-EPO cells secretedfunctional EPO protein for at least 70 days using regular in vitro cultureconditions. In addition, hiPSC-EPOcells have the potential to survive ina long term (28 weeks) after transplantation in the body.

In addition to their application for clinical use, hiPSC/ESC-derivedEPO-producing cells will be valuable for basic research. Althoughmuchis known about the possible mechanisms of EPO production and secre-tion, such as the involvement of HIF-2a (17), the detailed molecularmechanisms have not yet been fully elucidated because of the limitedaccess to isolated and cultured EPO-producing cells, especially humancells. EPO cells have been identified in the peritubular interstitial spaceinmouse kidneys (11). Moreover, EPO cells were recently isolated fromprimary cultures of human (25) and mouse kidneys (26). These cellsexhibited gene expression and protein secretion of EPO in an oxygen-dependentmanner, suggesting that EPOcells removed from the kidneyscan be used to investigate the molecular mechanisms of human EPOproduction. However, it is difficult to obtain an adequate amount ofEPO cells because access to human kidney specimens is limited. In ad-dition, primary cultured cells isolated from normal human tissues areoften difficult to expand in vitro in a long term. HepG2 cells are widelyused to investigate EPO production (27, 28); however, because they arederived from hepatoma tissues, EPO production and secretion may dif-fer from that observed in healthy, nontumor cells. Here, hypoxia acti-vated EPO secretion from the hiPSC-EPO cells, and the secreted EPOprotein induced erythropoietic differentiation both in vitro and in vivo,suggesting that hiPSC-EPO cells may be functionally similar to theirin vivo counterparts. Given that these EPO cells can be readily derivedfrom hiPSCs/ESCs and miPSCs/ESCs and display robust in vitro ex-pansion potential, they could be reliable cellularmodels for investigatingthe intracellular signaling pathways involved in EPO production andsecretion.

Both clinical studies and in vivo research using experimental animalshave reported that IGF-1 is involved in EPO production and secretion.One clinical study revealed that IGF-1 stimulates EPO secretion fromthe kidneys of patients with insulin resistance (29). On the other hand,the acute injection of IGF-1 negatively controls EPO secretion from themouse kidney (30). In addition, IGF-1 suppresses EPO secretion fromthe kidney but increases its secretion from the liver (31). Here, we showthat IGF-1 augmented EPOproduction in hiPSC-EPOcells. Combined,

Hitomi et al., Sci. Transl. Med. 9, eaaj2300 (2017) 27 September 2017

these data suggest that IGF-1 is an important regulatory factor for EPOproduction and secretion.

Several limitations of our study deservemention. First, the EPO cellsin our study were generated from hiPSCs/ESCs through hepatic lineagedifferentiation. The liver is amajor organ of EPOproduction in the fetaland early neonatal periods. Although EPO production shifts to the kid-ney in late gestation and adulthood (32), previous studies have reportedthat EPO production by the adult liver can be induced under anemic orhypoxic conditions (1, 2, 33). Nevertheless, methods that inducehiPSCs/ESCs into becoming EPO-producing cells via differentiationthrough the kidney lineage would be helpful for elucidating the mech-anisms of EPO production, especially considering that gene regulationis different between the mouse kidney and the liver (34). It has recentlybeen reported that EPO-producing fibroblasts in the normal adultmouse kidney had originated from myelin protein zero–Cre lineage-labeled extrarenal cells, which entered the embryonic kidney on E13.5(35). Because myelin protein zero is expressed in migrating neural crestcells in early embryonic stages, the origin of EPO cells in the kidneymaybe the neural crest. Therefore, differentiationmethods that drive hiPSCsinto the neural crest lineage and then into EPO cells could provideinsights into the kidney-derived EPO production.

Second, themechanismsunderlying the variable responses seenwithPHD inhibitors and the different responses between hiPSC-EPO andHepG2 cells are still unclear. We evaluated the effects of three differentPHD inhibitors (DMOG, DFO, and FG4592) on EPO expression andsecretion. Previous studies reported that these PHD inhibitors induceddifferent responses in the expression or function of HIF-PHD pathwaymolecules and their biological effects (36, 37). Furthermore, PHD inhib-itors have variable selectivity and inhibitory effects on different celltypes, such as liver and retinal cells, and on PHD subtypes, such asPHD1, PHD2, and PHD3 (37, 38). Together, these differences amongPHD inhibitors may at least, in part, explain the variable responses ob-served between hiPSC-EPO and HepG2 cells. Nevertheless, a recent re-port indicated that FG4592 has more efficient and broader inhibitoryeffects on various cell types compared to DMOG (37); several clinicaltrials are using FG4592 (39, 40). The present findings that FG4592augmentedEPOproductiononly in hiPSC-EPOcells, but not inHepG2cells, suggest that hiPSC-EPO cells may provide a good model forscreening PHD inhibitors for their effects on renal anemia.

One final limitation of our study is that a large number of hiPSC-EPO cells need to be used. At the beginning of this study, we decided onthe cell number for transplantation based on EPO secretion in vitro.The results of EPO secretion into the cell culture medium detected byELISA indicated that 1 × 107 hiPSC-EPO cells secreted about 12 ng/dayof EPO protein. The findings of in vivo dose-response experiments (fig.S7B) indicated that this amount of EPO protein was enough to reverserenal anemia. Therefore, we transplanted 1 × 107 hiPSC-EPO cells un-der the left renal subcapsule of each immunodeficient mouse. However,future studies should optimize EPO production capacity and the num-ber of hiPSC-EPOcells for transplantation. In addition, our data suggestthat there was variability in both the differentiation potential and EPOsecretion capacity among the different hiPSC/ESC lines. Further opti-mization and improvement of the differentiation protocol for EPO cellswill be required to achieve broad application.

In summary, we successfully generated EPO cells fromhiPSCs/ESCsand miPSCs/ESCs. These cells produced and secreted functional EPOprotein in a hypoxia-dependent manner, and IGF-1 treatment aug-mented EPO production. From the point of view of basic research,iPSC/ESC-derived EPO cells may be a powerful tool for investigating

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the molecular mechanisms of EPO production and secretion. hiPSC/ESC-derived EPO-producing cells may potentially provide a useful cellpopulation for cell therapy for the treatment of renal anemia in CKDpatients.

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MATERIALS AND METHODSStudy designThe aim of this study was to establish a differentiation method thatproduces EPO cells from hiPSCs/ESCs and miPSCs/ESCs and to eval-uate the transplantation of the resulting hiPSC-EPO cells in a CKDmouse model. EPO production and secretion were evaluated by RT-PCR, immunostaining, and ELISA. All mice analyzed in this study werehandled and the experimental procedures were performed under theguidelines for the care and use of animals established by Kagawa Uni-versity andKyotoUniversity. All animal experiments were approved bythe Animal Care and Use Committee for Kagawa University and theCenter for iPSCell Research andApplicationAnimal ExperimentCom-mittee. The sample size (n = 6 to 10) for the animal experiments wasbased on the results of preliminary experiments. Exact numbers for eachexperiment are described in the figure legends. The investigators werenot blinded when evaluating the experiments. Themice were randomlyassigned to the treatment and control groups.

Cell culturehiPSCs (201B6, 201B7, 253G1, 253G4, 585A1, 604A3, 606A1, and606B1) (8, 14) and hESCs (KhES-3 and H9) (10, 16) were grown onfeeder layers of mitomycin C–treated SNL cells in Primate ES medium(ReproCELL) supplemented with penicillin/streptomycin (500 U/ml)(Thermo Fisher Scientific) and recombinant human basic fibroblastgrowth factor (4 ng/ml) (Wako), as previously described (41). The col-onies of hiPSCs were dissociated using an enzymatic method with CTKsolution consisting of 0.25% trypsin (Thermo Fisher Scientific), 0.1%collagenase IV (Thermo Fisher Scientific), 20% knockout serum re-placement (KSR; Thermo Fisher Scientific), and 1 mM CaCl2 inphosphate-buffered saline (PBS).

miPSCs (492B-4) andmESCs (D3)weremaintained on feeder layersof mitomycin C–treated SNL cells in Dulbecco’s modified Eagle’s me-dium (DMEM; Nacalai Tesque) supplemented with 15% fetal bovineserum (FBS; Hyclone Laboratory), penicillin/streptomycin (500 U/ml),0.1 mM of nonessential amino acids (Thermo Fisher Scientific), 2 mMglutamine (Thermo Fisher Scientific), 0.55 mM 2-mercaptoethanol(Thermo Fisher Scientific), and leukemia inhibitory factor. miPSCs/ESCs were passaged using enzymatic dissociation with 0.25% trypsin/EDTA. HepG2 cells were cultured in DMEM with 10% FBS andpenicillin/streptomycin (500 U/ml).

Differentiation protocols for EPO cellsThe differentiation of EPO cells from hiPSCs/ESCs (fig. S1) was per-formed by modifying previously reported protocols into hepatic line-ages (13, 14, 41). Colonies of hiPSCs/ESCs grown on an SNL feederlayer were first dissociated, according to an enzymatic method, withCTK dissociation solution to remove SNL cells. Then, the cells weredissociated to single cells via gentlepipetting after treatmentwithAccutase(Innovative Cell Technologies) for 20min and seeded onMatrigel-coatedplates (BD Biosciences) at a density of 4.5 × 105 cells/cm2 with stage1 medium containing RPMI 1640 (Nacalai Tesque) supplemented withpenicillin/streptomycin (500 U/ml), B27 supplement (Thermo FisherScientific), recombinant human/mouse/rat activin A (100 ng/ml)

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(R&D Systems), and 1 mM CHIR99021 (Wako). Y-27632 (10 mM;Wako) was added to the stage 1 medium for the first 24 hours. After24 hours, the stage 1 medium was supplemented with 0.5 mM sodiumbutyrate (NaB; Sigma-Aldrich) until culture day 3. On day 7, the me-dium was changed to stage 2 medium containing KnockOut DMEM(Thermo Fisher Scientific) supplemented with penicillin/streptomycin(500 U/ml), 20% KSR, 1% dimethyl sulfoxide, 1 mM L-glutamine, 1%nonessential amino acid, and 0.1 mM b-mercaptoethanol. For hepato-cyte differentiation, the cells were shifted to the hepatocyte maturationstep after 7 days of stage 2 treatment using incubation with stage 3 me-dium containing hepatocyte culture medium (Lonza) supplementedwithhepatocyte growth factor (10ng/ml) (R&DSystems) andoncostatinM (20 ng/ml) (R&D Systems). For cell passaging, stage 2 cells weredissociated to single cells via gentle pipetting after treatment withAccutase for 8 min and split on Matrigel-coated plates at a ratio of 1:4with stage 2 medium containing Y-27632 (10 mM).

The differentiation of EPO cells from miPSCs/ESCs was performedusing a similar protocol to that for hiPSCs/ESCs (fig. S4). Briefly,miPSCs/ESCs dissociated to single cells with Accutase treatmentwere cultured with stage 1 medium containing RPMI 1640, penicillin/streptomycin (500 U/ml), B27 supplement, recombinant activin A(100 ng/ml), and 1 mMCHIR99021. After 2 days, the mediumwas sup-plemented with 0.5 mM NaB until day 4. On day 5, the medium waschanged to stage 2 medium.

RT-PCR and real-time qRT-PCRTotal RNAwas isolated using the RNeasy kit (Qiagen) according to themanufacturer’s recommendations, followed by cDNA synthesis usingstandard protocols. Briefly, first-strand cDNA was synthesized from1 mg of total RNA using ReverTra Ace (Toyobo). The cDNA sampleswere subjected to PCR amplification using a thermal cycler (Veriti96-Well Thermal Cycler; Thermo Fisher Scientific), and PCR was per-formed using the Ex-Taq PCR kit (Takara Bio) according to the man-ufacturer’s protocol. The PCR cycles were as follows: For b-ACTIN,initial denaturation at 94°C for 2.5 min, followed by 25 cycles of 94°Cfor 30 s, 60°C for 1 min, 72°C for 30 s, and final extension at 72°C for10 min. For the other genes, the cycles consisted of initial denaturationat 94°C for 2.5 min, followed by 30 to 40 cycles of 94°C for 30 s, 58° to62°C for 30 s, 72°C for 30 s, and final extension at 72°C for 7 min. qRT-PCR was performed using the StepOnePlus Real-Time PCR System(Thermo Fisher Scientific) and SYBR Green PCR Master Mix (TakaraBio).Denaturationwas performed at 95°C for 30 s, followed by 45 cyclesat 95°C for 5 s and at 60°C for 30 s. The threshold cycle method wasused to analyze the data for the gene expression levels and calibrated tothose of either housekeeping gene b-ACTIN or GAPDH. Primer se-quences are described in table S2.

ImmunostainingImmunostaining of the cultured cells was carried out as previously de-scribed (41). Briefly, the cells were fixedwith 4%paraformaldehyde/PBSfor 20 min at 4°C. After washing with PBS, the cells were blocked with1% normal donkey serum and 3% bovine serum albumin (NacalaiTesque)/PBST (PBS/0.25% Triton X-100) for 30min at room tempera-ture. A fetal mouse liver (E12.5) and transplanted aggregates were alsofixed with 4% paraformaldehyde/PBS for 20 min at 4°C, embedded inparaffin, and sectioned into 4-mm slices. The sections were stained withH&E. The samples were incubated overnight at 4°C with the followingprimary antibodies: anti-EPO (Santa Cruz Biotechnology), AFP(Sigma-Aldrich), albumin (Bethyl Laboratories), HNF-1b (Santa Cruz

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Biotechnology), HNF-4 (Santa Cruz Biotechnology), SALL4 (Abcam),GATA4 (Santa Cruz Biotechnology), CK19 (Dako), Ki67 (BD Bio-sciences), CD31 (Dianova), HIF-1a (Sigma-Aldrich), and HIF-2a(Novus Biologicals). Secondary antibodies (Alexa Fluor antibodies;Thermo Fisher Scientific) were incubated for 30 min at room tempera-ture. The stained cells were evaluated using fluorescence microscopy(BZ-9000; Keyence).

Measurement of EPO protein expressionThe concentrations of EPO protein in the culture medium weremeasured using ELISA according to the manufacturer’s protocol(ALPCO Diagnostics for human EPO and Abnova for mouse EPO).Briefly, the supernatant of the EPO cell cultures was added to a 96-wellplate with enzyme-labeled anti-human or anti-mouse EPO antibodies.The plates were incubated for 2 hours at room temperature. Then, thesubstrate was added, and the reactionwas stoppedwith 0.5mM sulfuricacid. The plates were read at 450 nm with a microplate reader (2104EnVision; PerkinElmer), and the concentration of EPO was calculatedon the basis of the standard curve of lyophilized synthetic EPO protein.

For theWestern blot analysis, the EPO protein expression in the celllysate was measured as previously described (42). Briefly, the cells werelysed, and solubilized proteins were isolated via centrifugation andquantified using a Bradford assay. The proteins were separated usingSDS–polyacrylamide gel electrophoresis and transferred to nitrocellulosemembranes (GE Healthcare). After blocking with Odyssey blockingbuffer (LI-CORBiosciences), blots were incubated with anti-EPO (SantaCruz Biotechnology) or anti–IGF-1 receptor (Cell Signaling Technol-ogy) antibodies overnight at 4°C. Blots with embedded infrared dyewerevisualized with the Odyssey Infrared Imaging System (LI-COR Bio-sciences). To confirm the loadingof equal amounts of protein, eachmem-brane was reprobed with anti–b-ACTIN antibodies (Sigma-Aldrich).The band intensity was quantified according to immunoblot densitom-etry using the ImageJ software (National Institutes of Health).

Electron microscopyFor observation with transmission electronmicroscopy, the cells werefixed with 4% formaldehyde and 2% glutaraldehyde in 0.1 Mphosphate buffer at 4°C overnight and washed in 0.1 M phosphatebuffer. Then, the cells were refixed with phosphate-buffered 1%OsO4, dehydrated in a graded series of ethanol, and embedded inLUVEAK-812 (Nacalai Tesque). Thin sections were cut with an EMUC6 ultramicrotome (Leica Microsystems), stained with uranyl acetateand lead citrate, and observed using a Hitachi H-7650 electron micro-scope (Hitachi).

Lectin microarrayThe glycosylation patterns of hiPSC-EPO and rhEPO proteins weremeasured by lectin microarray assays. hiPSC-EPO proteins were con-centrated using a centrifugal filter device (Amicon Ultra-15; MerckMillipore) and then purified and isolated with biotin-labeled anti-EPO antibody (Fitzgerald Industries) using Dynabeads MyOneStreptavidin T1 (Thermo Fisher Scientific). Lectin microarray assayswere performed by GlycoTechnica, as previously reported (43).

Clonogenic hematopoietic progenitor assayHuman CD34+ cells were isolated from cord blood via immuno-magnetic bead separation (Miltenyi Biotec) according to the manufac-turer’s instructions. The purity of the isolated CD34+ cells was >95%, asanalyzed using flow cytometry. A total of 1 × 105 CD34+ cells were in-

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cubated with rhEPO or hiPSC-EPO protein from hiPSC-EPO cells at0.28 and 2.8 ng/ml for 2 hours. hiPSC-EPO protein was purified andconcentrated usingAmiconUltra-30 (MerckMillipore).Hematopoieticcells were cultured at a low cell density, with a concentration of 1 ×104 cells/ml in 35-mmpetri dishes (BectonDickinson) usingMethoCultGF+ semisolid medium (1 ml per dish) (STEMCELL Technologies) in-cluding stem cell factor, thrombopoietin, and interleukin-3 (ST3), aspreviously described (44). The number of colonies was counted after14 to 21 days of culture, and the colony types [CFU-Mix (mixed colony-forming units), BFU-E, and CFU-GM] were determined with in situobservation using an inverted microscope, according to the criteria de-scribed previously (44).

Animal experimentsAll experimental procedures were performed under the guidelines forthe care and use of animals established byKagawaUniversity andKyotoUniversity. Six-week-oldmale C57BL/6mice (CLEA Japan) were usedfor the adenine-induced renal anemia model. Renal injury was inducedbyoral gavagewithadenine (50mg/kgbodyweight in0.5%methylcellulose)daily for 4 weeks. The presence of renal anemia was confirmed bymeasuring hematocrit and hemoglobin levels and red blood cell count.

One week after the end of adenine treatment, the following fivemouse groups were prepared: group 1, no adenine treatment (control;n = 6); group 2, rhEPO (28 ng), subcutaneously injected after adeninetreatment [rhEPO (28 ng); n = 10]; group 3, rhEPO (0.56 ng), sub-cutaneously injected after adenine treatment [rhEPO (0.56 ng); n =10]; group 4, hiPSC-EPO protein (0.56 ng), subcutaneously injected af-ter adenine treatment [hiPSC-EPOprotein (0.56 ng);n=10]; and group5, saline injection after adenine treatment (saline; n = 10). The rhEPOand hiPSC-EPO protein treatments were performed via subcutaneousinjection three times aweek for 4weeks. For EPOprotein dose-responseexperiments in vivo, the following nine mouse groups were prepared:group 1, no adenine treatment (control; n = 6); group 2, hiPSC-EPOprotein (0.56 ng), subcutaneously injected after adenine treatment(n = 6); group 3, hiPSC-EPO protein (5.6 ng), subcutaneously injectedafter adenine treatment (n = 6); group 4, hiPSC-EPO protein (14 ng),subcutaneously injected after adenine treatment (n = 6); group 5,hiPSC-EPO protein (28 ng), subcutaneously injected after adeninetreatment (n = 6); group 6, rhEPO (0.56 ng), subcutaneously injectedafter adenine treatment (n=6); group 7, rhEPO (5.6 ng), subcutaneouslyinjected after adenine treatment (n = 6); group 8, rhEPO (14 ng), sub-cutaneously injected after adenine treatment (n = 6); and group 9,rhEPO (28 ng), subcutaneously injected after adenine treatment (n =6). To measure the half-life of EPO protein, rhEPO and hiPSC-EPOproteins were subcutaneously injected (28 ng) into an adenine-inducedrenal anemia model. Serum EPO concentrations were measured usingELISA at 1, 6, 24, and 48 hours after injection. The half-life of EPO wasevaluated by decreases in serum EPO protein concentration after in-jection. Blood samples were collected to measure the hematocrit andhemoglobin levels and EPO protein concentrations. The hematocritlevels were determined in blood samples withdrawn into glass capillarytubes. The hemoglobin levels were measured using ELISA (Abcam),and the numbers of red blood cells, white blood cells, and platelets werecounted using a Coulter Counter Multisizer 3 (Beckman Coulter).

For the transplantation experiments, renal anemia was inducedusing adenine treatment (50 mg/kg body weight daily for 5 weeks) inimmunodeficient mice (NOD.CB17-Prkdcscid/J). To prepare the EPOcell aggregates, EPO cells were dissociated to single cells via pipettingafter incubation with Accutase for 10 min. Then, the EPO cells were

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split into spindle-bottom low-adhesion 96-well plates at a density of5.0 × 105 cells per well and incubated in stage 2 medium supplementedwith 10 mMY-27632 for 48 hours. The formation of cellular aggregateswas confirmed with microscopic observation. After washing the aggre-gates with saline, 20 EPO cell aggregates were transplanted into therenal subcapsules of immunodeficient mice with renal anemia. Thehematocrit levels were measured for up to 28 weeks after transplanta-tion. The human EPO concentrations were alsomeasured using ELISA.For immunohistochemistry analysis,micewere sacrificed 2, 4, 8, 16, and28 weeks after transplantation, and the serial sections of transplantedaggregates were examined by immunostaining. To elucidate the effectsof phlebotomy, serum EPO concentrations of control (no transplanta-tion) and hiPSC-EPO cell–transplanted mice were measured by aGoldenrod animal lancet (MEDIpoint).

Statistical analysisResults were expressed as the mean ± SEM. Multiple group com-parisons were made using one-way or two-way ANOVA, followed byBonferroni’s test. Student’s t tests were performed to compare themeanvalues when the experimental design was composed of two individualgroups. P < 0.05 was considered statistically significant.

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SUPPLEMENTARY MATERIALSwww.sciencetranslationalmedicine.org/cgi/content/full/9/409/eaaj2300/DC1Fig. S1. Differentiation method for generating EPO-producing cells from hiPSCs/ESCs.Fig. S2. Expression of hepatic lineage and endoderm markers in hiPSC-EPO cells.Fig. S3. Variable expression and secretion of EPO and expression of hepatoblast markersamong three hiPSC/ESC lines.Fig. S4. Differentiation method for generating EPO-producing cells from miPSCs/ESCs.Fig. S5. Effects of 46 different factors on EPO mRNA expression in the hiPSC-EPO cells.Fig. S6. The HIF-PHD pathway regulates EPO production induced by hypoxia orIGF-1 treatment.Fig. S7. Effects of hiPSC-EPO protein on body weight and renal anemia in adenine-treatedmice.Table S1. Effects of hiPSC-EPO protein on in vitro erythropoiesis (related to Fig. 6).Table S2. The sequences of sense and antisense primers used for RT-PCR in this study.Table S3. A list of lectins and their specificity for microarray analysis.

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Acknowledgments: We thank T. Toyoda, A. Watanabe, and M. Nagao for the helpfulsuggestions; T. Sudo, H. Tanaka, Y. Fujita, and M. A. Sufiun for their excellent technicalassistance; and P. Karagiannis for reading the manuscript. Pathological analysis was done byCenter of Anatomical, Pathological and Forensic Medical Research, Graduate School ofMedicine, Kyoto University. Funding: This work was supported by the Japan Agency forMedical Research and Development through its research grant “Core Center for iPSCell Research, Research Center Network for Realization of Regenerative Medicine,” by a grant-in-aid for scientific research from the Ministry of Education, Culture, Sports, Science andTechnology of Japan (#22790786, #22790792, #24591204, and #15K09266), by the iPS CellResearch Fund, by a Sanju Alumni Research Grant, by the Kanae Foundation for the Promotionof Medical Science, and by the Daiichi Sankyo Foundation of Life Science. Authorcontributions: H.H., A.N., and K.O. conceived the project and designed the experiments. H.H.,T.K., N.K., A.H., S.M., M.K., T.T., A.R., D.N., A.N., M.K.S., T.N., A.N., and K.O. performed theexperiments. H.H., S.M., A.N., M.K.S., T.N., A.N., and K.O. analyzed the data. H.H., A.N.,M.K.S., T.N., A.N., and K.O. wrote the manuscript. Competing interests: H.H. and K.O. areco-inventors on patent numbers US61/621,256, PCT/JP2013/060878, JP2014-509231, US14/390853, US9334475, and EP13773017.2 “Method for inducing erythropoietin-producing cells.”All other authors declare that they have no competing interests.

Submitted 16 September 2016Accepted 27 April 2017Published 27 September 201710.1126/scitranslmed.aaj2300

Citation: H. Hitomi, T. Kasahara, N. Katagiri, A. Hoshina, S.-I. Mae, M. Kotaka, T. Toyohara,A. Rahman, D. Nakano, A. Niwa, M. K. Saito, T. Nakahata, A. Nishiyama, K. Osafune, Humanpluripotent stem cell–derived erythropoietin-producing cells ameliorate renal anemia inmice. Sci. Transl. Med. 9, eaaj2300 (2017).

ce

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anemia in micederived erythropoietin-producing cells ameliorate renal−Human pluripotent stem cell

OsafuneAsadur Rahman, Daisuke Nakano, Akira Niwa, Megumu K. Saito, Tatsutoshi Nakahata, Akira Nishiyama and Kenji Hirofumi Hitomi, Tomoko Kasahara, Naoko Katagiri, Azusa Hoshina, Shin-Ichi Mae, Maki Kotaka, Takafumi Toyohara,

DOI: 10.1126/scitranslmed.aaj2300, eaaj2300.9Sci Transl Med

anemia.the perspective of clinical research, these results may provide a physiological therapeutic agent for treating renal may be a useful tool for investigating the molecular mechanisms of erythropoietin production and secretion. Frommouse model ameliorated renal anemia. From the perspective of basic research, erythropoietin-producing cells secreted functional erythropoietin protein in a hypoxia-dependent manner. Transplantation of these cells into aprotocol for erythropoietin-producing cells from human and mouse iPSCs and ESCs. These cells produced and

. have developed a differentiationet alis beneficial and safe, more physiological therapies are required. Hitomi Erythropoietin dysregulation is a hallmark of renal anemia. Although recombinant erythropoietin treatment

Cell therapy for renal anemia

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