Early Senescence Is Not an Inevitable Fate of Human-Induced Pluripotent Stem-Derived Cells

Abstract

Human-induced pluripotent stem cells (hiPSCs) are expected to become a powerful tool for regenerative medicine. Their efficacy in the use of clinical purposes is currently under intensive verification. It was reported that hiPSC-derived hemangioblasts had severely limited expansion capability due to an induction of early senescence: hiPSC-derived vascular endothelial cells (VECs) senesced after one passage and hiPSC-derived hematopoietic progenitor cells (HPCs) showed substantially decreased colony-forming activities. Here we show that early senescence is not an inevitable fate of hiPSC-derived cells. Applying our unique feeder-free culture methods for the differentiations of human embryonic stem cells (hESCs), we successfully generated VECs and HPCs from three lines of hiPSCs that were established by using a retrovirus vector system. All hiPS-derived VECs could be subcultured by 2:1∼3:1 dilutions up to 10∼20 passages, after which the cells underwent senescence. Among the three lines of hiPSCs, two lines generated HPCs that bore comparable granulocyte colony-forming units to those of hESCs. Moreover, one line effectively reproduced HPCs within the sac-like structures, the fields of in vitro hematopoiesis, as in the case of hESCs. Surprisingly, release of neutrophils into culture supernatant persisted even longer (∼60 days) than the case of hESCs (∼40 days). Thus, the problem of early senescence can be overcome by selecting appropriate lines of hiPSCs and applying proper differentiation methods to them.

Introduction

Human induced pluripotent stem cells (hiPSCs) hold great promise for the development of regenerative medicine with their unlimited expansion capacities and pluripotency that were equivalent to human embryonic stem cells (hESCs). Before translating the achievements of basic researches to medical purposes, however, the efficacy and safety of hiPSCs must be carefully scrutinized from various points of view. Large quantities of useful findings have been accumulating in the case of murine iPSCs (miPSCs); however, performing the studies on hiPSCs is particularly important because hiPSCs may not be the real equivalent of miPSCs as the hESCs may not be the genuine counterpart of murine ESCs (mESCs) (Tesar et al., 2007; Vallier et al., 2009).

So far, two major issues have been raised concerning the disadvantageous outcomes of iPSCs after differentiation. One is a widely recognized matter of tumorigeneity, which was reported in the cases of miPSC chimeric mice (Okita et al., 2007) and the mice transplanted with miPSC-derived differentiated cells (Nelson et al., 2009). Already, several measures have been proposed to overcome this problem: an application of nonviral vectors (Okita et al., 2008; Yu et al., 2009), excisions of integrated transgenes (Kaji et al., 2009; Woltjen et al., 2009), a usage of L-myc instead of c-myc transgene (Nakagawa et al., 2008, 2010), and a selection of iPSC lines with least tumorigeneity after differentiation (Miura et al., 2009). Those strategies, along with the recent technological innovations to introduce the reprogramming factors via RNA-based (Fusaki et al., 2009; Seki et al., 2010; Warren et al., 2010) or protein-based systems (Kim et al., 2009; Zhou et al., 2009), are expected to make a great contribution toward the complete resolution of the problem.

The second issue is rather a newly proposed one: an induction of early senescence in hiPSC-derived differentiated cells (Feng et al., 2010). It was reported that every hiPSC established by using a retrovirus vector system suffered from expansion deficiency after differentiation into vascular endothelial cells (VECs), hematopoietic progenitor cells (HPCs), and retinal pigmented epithelium cells (Feng et al., 2010). Although the molecular mechanism of early senescence remains elusive, the effects of p53 inactivation, LIN28 activation and insertion of proviral transgenes into chromosomes were discussed (Feng et al., 2010). The finding of early senescence is worth reporting in that it has provoked a caution that we should o use during our researches on hiPSCs. Nevertheless, we thought that their conclusion must be further validated in other situations, where differentiation processes are performed by distinct methods, because early senescence is often caused by cellular stresses and the degrees of stresses substantially vary depending on culture conditions.

We previously established feeder-free methods for neutrophilic (Saeki et al., 2009) and vascular endothelial (Nakahara et al., 2009) differentiations of hESCs. By applying those methods to hiPSCs, we studied the incidence of early senescence during the differentiation of hiPSCs that were established by using a retroviral vector system. Our data indicate that early senescence is not an inevitable fate of the hiPSC-derived differentiated cells even in the presence of retroviral insertions of the reprogramming transgenes. We also show that hiPSC-derived VECs undergo senescence after 10∼20 passages, as in the cases of primary human VECs, without entering crisis. Our results clearly show that the problem of early cellular senescence can be overcome by selecting appropriate lines of hiPSCs and applying proper differentiation methods to them.

Materials and Methods

Cell culture

The use of hESCs was performed in accordance with the Guidelines for Derivation and Utilization of Human Embryonic Stem Cells of the Ministry of Education, Culture, Sports, Science, and Technology (MEXT) of Japan, after approval by the institutional review board of International Medical Center of Japan (IMCJ). The hESCs (KhES-1, KhES-3) (Suemori et al., 2006) were provided by Kyoto University (Kyoto, Japan). The hiPSC were provided by either CiRA at Kyoto University (253G1 and 253 G4 (Nakagawa et al., 2008), 201B7 and 201B2 (Takahashi et al., 2007) or by National Research Institute for Child Health and Development (#25). The cells were maintained on dishes coated with γ-irradiated murine embryonic fibroblasts (MEFs) in DMEM/F12 (Invitrogen Corp., Carlsbad, CA) supplemented with 20% Knockout™ Serum Replacement (KSR) (Invitrogen Corp.), 5 ng/mL fibroblast growth factor 2 (FGF-2; Pepro Tech Inc., Rocky Hill, NJ), 1% nonessential amino acids solution (Invitrogen Corp.), 1 mM sodium pyruvate solution (Invitrogen Corp.), 100 μM 2-mercaptethanol (Sigma Chemical Co., St. Louis, MO), 2 mM L-glutamine (Invitrogen Corp.), 20 U/mL penicillin (Invitrogen Corp.) and 20 μg/mL streptomycin (Invitrogen Corp.). The cells were passed twice a week (e.g., on Tuesday mornings and Friday evenings) by a treatment with dissociation liquid, which contains 0.25% trypsin (Invitrogen Corp.), 1 mg/mL collagenase IV (WAKO Pure Chemical Industries, Osaka, Japan), 20% KSR, 1 mM CaCl₂, at 37°C for 5 to 15 min and seeded at split ratios of 1:2 to 1:4 into new MEF-coated dishes. During the course of experiments, cells showed normal karyotypes.

Vascular endothelial differentiation of hESCs/hiPSCs

Differentiation was performed as previously described (Nakahara et al., 2009) with a slight modification. The hESCs/hiPSCs were detached from culture plates by using the dissociation liquid for 15 min at 37°C. The mildly dissociated hESC/iPSC clumps were cultured in a 6-cm diameter low-attachment dish (Nalge Nunc International K.K., Tokyo, Japan) to form spheres using the differentiation medium consisting Iscove's modified Dulbecco's medium (IMDM) (Sigma Chemical Co.) supplemented with 15% heat-inactivated fetal bovine serum (FBS) (PAA Laboratories GmbH, Linz, Austria), 0.1 mM 2-mercaptoethanol, 3 mM L-glutamine, 10 U/mL penicillin, 20 ng/mL vascular endothelial growth factor (VEGFA), 20 ng/mL bone morphogenetic protein 4 (BMP4), 20 ng/mL stem cell factor (SCF), 10 ng/mL FMS-related tyrosine kinase-3 ligand (Flt3-L), 20 ng/mL Interleukin 3 (IL3), and 10 ng/mL IL6. After incubation for 3 days at 37°C under a 100% humidified condition in a 5% CO₂ gas incubator, spheres were subjected to adherent culture using 100 mm × 20 mm 0.1% porcine type A gelatin (Sigma Chemical Co.)-coated dishes in the differentiation medium described above. Media were changed twice a week. For passage, cells were harvested by treatment with 0.25% trypsin and 1 mM EDTA and replated at split ratios of 1:2 on new gelatin-coated dishes.

Senescence-associated (SA)-β-galactosidase assays

The 1 × 10⁵ VECs were cultured in 6-cm culture plates. After an incubation at 37°C in 5% CO₂ incubator for 4∼5 days, cells were subjected to SA-β-galactosidase assays by using Senescence Detection Kit (BioVision Research Products Inc., Mountain View, CA) according to a manufacture's guidance.

Cord formation assays

Matrigel™ Basement Membrane Matrix, phenol-Red free (Cat 356237, BD Biosciences, San Jose, CA) was loaded into the 24 multiwell dishes (95 μL/well). After the dishes were incubated for 30 min at 37°C, 1 × 10⁴ cells per well were seeded in differentiation medium described above. Cell morphologies were observed after overnight culture under an inverted light microscope (Olympus Optical Co. Ltd, Japan).

Uptake of acetylated low-density lipoprotein (Ac-LDL)

Cells were transferred in four-well chamber slide system (Nalge Nunc International Corp., Naperville, IL). After overnight culture, cells were washed by Hank's balanced salt solution (HBSS) twice and incubated in serum-free medium containing 10 μg/mL of low-density lipoprotein from human plasma, acetylated, DiI complex (DiI Ac-LDL) (Invitrogen Corp.) for 4 h. After washing the cells by HBSS for three times, nuclei were counterstained using 10 nM of Hoechst 33342 (Sigma Chemical Co.). After washing the cells, samples were observed under the fluorescence microscope (Olympus Optical Co. Ltd).

Flow cytometry

Cells were collected by a treatment with 0.2% EDTA or Dispase (BD Biosciences). After a wash in phosphate-buffered saline (PBS), 1 × 10⁶ cells were reacted with first antibodies on ice for 30 min. The expression level of each protein was analyzed using a FACSCalibur^™ (BD Biosciences). The antibodies used were a mouse monoclonal antihuman Tie-2- allophycocyanin (APC) antibody (R&D Systems Inc., Minneapolis, MN), antihuman VEGF receptor 1 (VEGF R1)-PE antibody (R&D Systems Inc.), mouse antihuman CD45-PE (BD Biosciences), and mouse antihuman CD11b-PE (BD Biosciences). After antibody-staining procedures, cells were stained with propidium iodide (PI) (Sigma Chemical Co.), in the case of Tie-2 staining, or TO-PRO3 fluorescent dye (Invitrogen Corp.), in the cases of VEGF1, CD45, and CD11b staining, for 10 min. During analysis, dead cells were gated out as FL-2 higher fractions, in the case of PI staining, or FL4-higher fractions in the case of TO-PRO3 staining.

Immunostaining

The cells were fixed on slide glasses by using a cytospin apparatus (Cytospin 2) along with further fixation with acetone/methanol solution (1:3). The immunostaining procedure was performed as described elsewhere (Nakahara et al., 2009) with first antibody reactions using a rabbit polyclonal antihuman p16^INK4A antibody (SC-20) (Santa Cruz Biotechnology Inc., Santa Cruz, CA), a mouse monoclonal antihuman p21^CIP1 antibody (Santa Cruz Biotechnology Inc.), a rabbit polyclonal antihuman endothelial nitric oxide synthase (eNOS) antibody (H-159) (Santa Cruz Biotechnology Inc.), or a rabbit polyclonal antihuman von Willebrand factor (vWF) antibody (Sigma Chemical Co.), followed by second antibody reactions using Alexa Fluor^® 488 chicken antimouse IgG (H+L), Alexa Fluor^® 568 goat antirabbit IgG (H+L), or Alexa Fluor^® 594 chicken antigoat IgG (H+L) (Invitrogen Corp.). Nuclear counterstaining was performed using 300 nM of 4′,6-diamino-2-phenylindole (DAPI).

Hematopoietic differentiation of hESCs/hiPSCs

Differentiation was performed as previously described (Saeki et al., 2009). In brief, hESCs/hiPSCs were detached with 1 mg/mL collagenase IV (Invitrogen Corp.) and transferred to a 6 cm diameter low-attachment dish (Nalge Nunc International K.K.) coated with 2-methacryloyloxyethyl phosphorylcholine in 5 mL IMDM (Sigma Chemical Co.) supplemented with 15% FBS (PAA Laboratories GmbH), 2 mM L-glutamine, 100 μM 2-mercaptethanol, 20 U/mL penicillin, and 20 μg/mL streptomycin in the presence of 20 ng/mL insulin-like growth factor II (IGF-II; Pepro Tech Inc.), 20 ng/mL VEGFA (Pepro Tech Inc.), 100 ng/mL SCF (Pepro Tech Inc.), 100 ng/mL Flt3-L (Pepro Tech Inc.), 50 ng/mL thrombopoietin (TPO; Kirin Brewery Company, Ltd., Tokyo, Japan), and 100 ng/mL G-CSF (Kirin Brewery Company, Ltd.) (Differentiation medium) at a density of 4 × 10⁵ cells/mL. After primary differentiation for 3 days, the spheres were transferred to 10-cm diameter dish coated with 0.1% gelatin. Spheroid cells floating in the culture supernatant were collected over time and analyzed.

Colony assays

Colony assays were performed using Methocult TM GF⁺H4535 (Stemcell Technologies Inc., Vancouver, Canada) in accordance with the manufacturer's recommendations. In brief, 0.3 mL of cell suspension, which contained 10 cells, was mixed in 3 mL of methylcellulose solution consisting of 1% methylcellulose, 30% FBS, 1% bovine serum albumin, 100 μM 2-mercaptoethanol, 2 mM L-glutamine, 50 ng/mL SCF (Pepro Tech Inc.), 20 ng/mL interleukin 3 (IL-3; Pepro Tech Inc.), 20 ng/mL interleukin 6 (IL-6; Pepro Tech Inc.), 20 ng/mL granulocyte-macrophage colony-stimulating factor (GM-CSF; Pepro Tech Inc.), 20 ng/mL G-CSF and 3 U/mL erythropoietin (Kirin Brewery Company, Ltd.) in 3.5-cm culture dishes. After 2 weeks, the number of colonies was counted. The morphology of the colonies was observed using an inverted light microscope (Olympus Optical Co. Ltd).

Reverse transcription-polymerase chain reaction (RT-PCR)

RNA was extracted from 5 × 10⁶ cells using an RNeasy Mini Kit (Qiagen Inc., Valencia, CA) and cDNA was synthesized using a Superscript II Kit (Invitrogen Corp.) in accordance with the manufacture's protocol. The sequence of the primers used are as follows: vascular endothelial (VE)-cadherin; a forward primer 5′-TGGGCTCAGACATCCACATA-3′ and a reverse primer 5′-TCACAGTCTCCCATTGGGAAT-3′, platelet endothelial cell adhesion molecule-1 (PECAM1); a forward primer 5′-GCAAAATGGGAAGAACCTGA-3′ and a reverse primer 5′-CACTCCTTCCACCAACACCT-3′, interferon α1 (IFNA1); a forward primer 5′-GGAGTTTGATGGCAACCAGT-3′ and a revsese primer 5′-CTCTCCTCCTGCATCACACA-3′, interferon α2 (IFNA2); a forward primer 5′-GCAAGTCAAGCTGCTCTGTG-3′ and a reverse primer 5′-GATGGTTTCAGCCTTTTGGA-3′, interferon β1(IFNB1); a forward primer 5′-ATTGCCTCAAGGACAGGATG-3′ and a reverse primer 5′-AGCCAGGAGGTTCTCAACAA-3′. As a molecular maker, DNA MW Standard Marker 100 bp DNA Ladder (Takara Shuzo Co. Ltd., Shiga, Japan) was used.

Wright-Giemsa (WG) staining and special staining procedures

Viable cells in the dishes were observed directly using an inverted phase contrast light microscope (Olympus Optical Co. Ltd., Tokyo, Japan). Alternatively, the cells were fixed on glass slides using a cytospin centrifuge (Cytospin 2, SHANDON, Pittsburgh, PA), stained with WG solution (Muto Pure Chemical Co., Tokyo, Japan), and then observed using a light microscope (Olympus Optical Co. Ltd.). Double esterase staining was performed using the staining kit (Muto Pure Chemical Co.) according to the manufacturer's protocols.

Phagocytosis

hESC-derived neutrophils attracted to the lower chamber of Chemotaxel were suspended in HBSS containing 2.5% FBS and incubated at 37°C for 1 h with 5 μL zymosan (1 mg/mL) in the presence of 100 nM fMLP. Subsequently, the cells were collected using a cytospin apparatus and stained with WG solution. Phagocytosis was determined by a microscope observation.

Nitroblue tetrazolium reduction assay for respiratory burst activity

The floating cells were collected by mild centrifugation of the culture supernatant. After washing with PBS, the cells were resuspended in 1 mL RPMI 1640 (Sigma Chemical Co.) supplemented with 10% FBS containing 1 mg/mL nitroblue tetrazolium (NBT) (Nacalai Tesque Inc., Kyoto, Japan) and 100 nM fMLP for 30 min at 37°C. After washing with PBS, the cells were resuspended in 10 μL PBS and dropped onto Matsunami Adhesive Silane (MAS)-coated glass slides (Matsunami Glass Ind., Ltd. Osaka, Japan) and the formazan blue-black deposit-containing cells were observed using a light microscope (Olympus Optical Co. Ltd.).

Results

hiPSC can generate subculturable VECs

We previously established a feeder-free method for the vascular endothelial differentiation of hESCs (Nakahara et al., 2009). The unique points of our method are (1) it is a two-tired differentiation system with a sphere-forming floating culture and a subsequent attachment culture, (2) it uses multiple hematopoietic cytokines in addition to a conventionally used cytokine of VEGF, (3) it enables high-purity production of VECs without contamination by growth-competing pericytes, (4) it enables the production of VECs that can be subcultured up to 10∼20 passages (Nakahara et al., 2009). Applying this method, we performed VEC differentiation of the following lines of hiPSCs: 253G1 and 253G4, which were established from adult human dermal fibroblasts by introducing three retroviral transgenes of oct4, sox2, and klf-4 (OSK) (Nakagawa et al., 2008), 201B7 and 201B2, which were established from adult human dermal fibroblasts by introducing four retroviral transgenes of oct4, sox2, klf-4, and c-my (OSKM) (Takahashi et al., 2007) and #25, which were established from human embryonic lung fibroblasts of MRC-5 by introducing retroviral transgenes of OSKM.

Among the five lines, 253G1, 201B7, and #25 successfully generated VECs. Although cell morphologies slightly differed from one another (Fig. 1A), we could effectively expand the VECs generated from all the three lines: the hiPSC-derived VECs were subcultured by 2:1∼3:1 dilutions up to 10 passages, in the case of 201B7, or even higher, in the cases of 253G1 and #25 (Fig. 1B). Both 253G1-derived and #25-derived VECs expanded with comparable growth rates to hESCs-derived VECs, whereas the growth speed of 201B7-derived VECs was slightly lower. After 10∼20 passages, the hiPSC-derived VECs underwent senescence as demonstrated by SA-β-galactosidase assays as in the cases of hESC-derived VECs and HUVEC (Fig. 1C). In agreement with this, the expressions of senescence-associated gene products of p16^INK4A and/or p21^CIP1 were induced in the senesced cells (Fig. 1D). Using the cells at exponentially growing phases, we evaluated the functions and maker expressions. All the hiPSC-derived VECs showed high Ac-LDL-uptaking capacities (Fig. 2A) and cord-forming activities (Fig. 2B). Although the expressions of VE-cadherin and PECAM1 messages were hardly detectable in #25-derived VECs (Fig. 2C), these cells showed comparable protein expressions of eNOS (Fig. 2D), Tie-2 (Fig. 2E), and VEGFR1 (Fig. 2F) to the other hiPSCs-derived VECs and HUVEC.

FIG. 1.

The VEC differentiation of hiPSCs. (A) Phase contrast micrographs. Morphologies of hiPSC-derived and hESC-derived VECs were shown as indicated. Numbers on the photographs in white letters indicates the passage number of each VEC. Scale bars indicate 100 μm. (B) Growth curves of various VECs. Average data with standard deviations from three independent experiments are shown in the cases of hiPSC(253G1)-derived VECs (closed triangles), hiPSC(201B7)-derived VECs (open triangles), hiPSC (#25)-derived VECs (gray circles), and hESC(khES-1)-derived VECs (open circles), whereas typical results from one or two experiments were shown in the cases of HUVEC (gray diamonds), HAEC (closed squares), and hESC(khES-3)-derived VECs (gray triangles). (C) SA-β-galactosidase assays. The hiPSC (253G1, 201B7, and #25)-derived VECs, hESC (khES-3)-derived VECs, and HUEVC at indicated passage numbers were subjected to SA-β-galactosidase assays. Scale bars indicate 100 μm. (D) Expressions of senescence-associated genes. The hiPSC (253G1, 201B7, and #25)-derived VECs, hESC (khES-3)-derived VECs, and HUEVC at indicated passage numbers were subjected to immunostaining studies using an anti-p16 or anti-p21 antibody as indicated. Scale bars indicate 100 μm.

FIG. 2.

Evaluation of hiPSC-derived VECs. (A) Ac-LDL-uptaking assays. VECs derived from each hiPSC line (253G1, 201B7, and #25) and HUEVC were subjected to Ac-LDL-uptaking assays (red; dil-Ac-LDL, blue; nuclear staining by Hoechst dye). Scale bars indicate 100 μm. (B) Cord formation assays. VECs derived from each hiPSC line (253G1, 201B7, and #25) and HUEVC were subjected to cord formation assays. Scale bars indicate 20 μm. (C) RT-PCR analyses. The message expressions of VE-cadherin (vecad), PECAM1 (pecam1), and β-actin were determined by RT-PCR in undifferentiated hiPSCs (253G1, 201B7, and #25), as indicated “U,” or VEC-differentiated hiPSCs, as indicated “V.” As positive control, cDNA of HUVEC was used as indicated “H.” “M” indicates the molecular marker. (D) The expressions of the eNOS protein. The hiPSCs (253G1, 201B7, and #25 as indicated)-derived VECs and HUVEC were subjected to immunostaining studies using an antihuman eNOS antibody with nuclear counterstaining by DAPI. Scale bars indicate 50 μm. (E, F) The expressions of Tie-2 and VEGFR1 proteins. The VECs generated from hiPSCs (253G1, 201B7, and #25 as indicated) an HUVEC were subjected to flow cytometric analyses using an antihuman Tie-2 antibody (E) or an antihuman VEGFR1 antibody (F). A typical result from two to five experiments are shown.

Thus, hiPSCs can generate VECs with equivalent expansion potentials to hESCs, although maturation levels of hiPSCs-derived VECs vary depending on the lines of hiPSCs.

hiPSCs can generate reproducible HPCs with comparable colony-forming activities to hESCs

We previously established a feeder-free method for the neutrophil differentiation of hESCs (Saeki et al., 2009). By our system, HPCs are generated within a unique construction named the “sac-like structure.” Within this structure, HPCs are repeatedly generated: they are reproduced within a few days after manually puncturing the sac walls and releasing the inner HPCs into the culture supernatant (Saeki et al., 2009). The reproduction process can be repeated three or four times (Saeki et al., 2009).

By applying this method, we performed neutrophil differentiation of 251G3, 201B7, and #25. All three lines successfully generated sac-like structures that were filled with abundant spheroid cells (Fig. 3A–C). However, the walls of 201B7-derived sacs seemed rather fragile because the inner spheroid cells spontaneously permeated the sac-like structures (Fig. 3B). Furthermore, we failed in reproducing spheroid cells after manually puncturing the sac walls and releasing inner spheroid cells into the culture supernatant (data not shown). In the case of 253G1, the sac walls seemed solid; however, spheroid cells were scarcely reproduced after sac wall puncturing (Fig. 3D and E). In contrast, spheroid cells were actively reproduced in the case of #25 (Fig. 3F and G). Eventually, the reproduction process persisted up to 60 days after the start of differentiation (data not shown), which was longer than that of hESCs (up to 40 days) (Saeki et al., 2009). Thus, the #25 line bears an equivalent, or even higher, spheroid cell-reproducing potential to hESCs.

FIG. 3.

Phase contrast microscopic observation during hematopoietic differentiation of hiPSCs. (A–C) Micrographs of the sac-like structures generated from 253G1 (A), 201B7 (B), and #25 (C). (D, E) Micrographs of the sac-like structures of 253G1 several days after manually puncturing the sac walls and releasing the inner hematopoietic cells. Note that the sac remained empty (E). (F, G) Micrographs of the sac-like structures of #25 several days after manually puncturing the sac walls and releasing the inner hematopoietic cells. Note that the sacs were refilled with hematopoietic cells (G). Scale bars indicate 200 μm.

We further evaluated the qualities of hematopoietic differentiation of 253G1, 201B7, and #25. First, cytological examinations were performed around day 30 of differentiation, when abundant neutrophil production was observed in the case of hESCs (Saeki et al., 2009). However, vast majorities of the products of 253G1 and 201B7 were macrophages (Fig. 4A, left and middle). On the other hand, various stages of granulocyte-lineage cells, from azurophilc granule-positive myeloid progenitors to segmented granulocytes, were observed in the case of #25 (Fig. 4A, right). Flow cytometric analyses demonstrated that #25-derived spheroid cells were highly positive for CD45, a pan-hematopoietic cell marker, and the majority of cells were positive for CD11b, a granulo-monocytic marker (Fig. 4B). To confirm their hematopoietic activities, colony assays were performed (Fig, 4C). The #25-derived spheroid cells demonstrated equivalent, or even higher, colony-forming unit-granulocyte (CFU-G) (8.0 ± 5.3/10⁴ cells; n = 3), colony-forming unit-granulocyte/macrophage (CFU-GM) (12.3 ± 5.5/10⁴ cells; n = 3), colony-forming unit-macrophage (CFU-M) (21.0 ± 3.5/10⁴ cells; n = 3) to those of hESCs, whose average CFU-G, CFU-GM, and CFU-M per 10⁴ cells were 2.3, 7.9, and 3.1, respectively (n = 2). Thus, hiPSCs can produce HPCs with equivalent colony-forming activities to hESCs.

FIG. 4.

Evaluation of hematopoietic potentials of hiPSC-derived cells (A) Cytological observation. Hematopoietic cells generated from hiPSCc (253G1, 201B7, and #25 as indicated) were stained by Wright-Giemsa solution. In #25-derived cells (right), azure granule-positive myeloid precursor cells (arrows) and segmented neutrocytes (arrow heads) were observed, whereas only macrophages were detected in 253G1-derived (left) and 201B7-derived (middle) cells. Scale bars indicate 50 μm. (B) Flow cytometry. The #25-derived hematopoietic cells were subjected to flow cytometric analyses using antihuman CD34, CD45, and CD11b antibodies as indicated. Number on each figure indicates the percentage of the corresponding marker-positive cells. (C) Colony assays. The #25-derived hematopoietic cells were subjected to colony assays. CFU-M, CFU-GM, and CFU-G were determined by phase contrast microscopic observation (upper panels; scale bars indicate 500 μm), which were further confirmed by Wright-Giemsa staining studies (lower panels; scale bars indicate 50 μm).

We also confirmed the functional maturation of #25-derived neutrophils by performing a superoxide production study (Fig. 5A), a phagocytosis assay (Fig. 5B), and double esterase-staining test (Fig. 5C). Interestingly, the double-esterase staining test, in which neutrophil-specific esterase is stained blue while that of nonspecific monocyte/macrophage esterase is stained brown, demonstrated that #25-derived cells showed even clearer neutrophil-specific blue staining patterns than hESC-derived ones, the majority of which showed brownish blue or bluish brown staining (Fig. 5D). Thus, hiPSCs can generate neutrophil with equivalent, or even higher, maturation levels than hESCs.

FIG. 5.

Functional assessments and special staining. (A, B) The #25-derived hematopoietic cells were subjected to an NBT-reducing assay (A) or phagocytosis assay (B). Almost all the cells showed NBT-reducing activities (A) and zymosan-phagocytizing myeloid cells were often observed (B). Scale bars indicate 20 μm. (C, D) The #25-derived hematopoietic cells (C) or hESC (khES-3)-derived hematopoietic cells (D) were subjected to double esterase staining assay. Note that typical neutrophil-specific dark blue staining patterns were clearly observed in #25-derived cells, whereas hESC-derived hematopoietic cells showed rather mixed (brownish blue) staining patterns. Scale bars indicate 50 μm.

For further assessment of possible hematopoietic potentials of 253G1-derived and 201B7-derived spheroid cells, we checked the morphologies of spheroid cells over time. Surprisingly, small round cells with high nucleus/cytoplasm ratios were detected in 253G1-derived samples around day 20 of differentiation (Fig. 6A and B), whereas no such cells were detected in 201B7-derived samples (data not shown). Interestingly, hemophagocytosis-like scenes, where the small cells were phagocytized by macrophages, were often observed (Fig. 6C). Because the morphologies of the small cells resembled to hematopoietic stem/progenitor cells, we checked their colony-forming activities (Fig. 6D). Colony assays performed around day 20 indicated that 253G1-derived cells had comparable CFU-G (3.3 ± 2.3/10⁴ cells; n = 3), CFU-GM (3.3 ± 1.2/10⁴ cells; n = 3), CFU-M (15.0 ± 1.0/10⁴ cells; n = 3) to those of hESCs. On the other hand, few hematopoietic colonies were observed in the case of 201B7 at any time points (data not shown).

FIG. 6.

The hiPCS-derived differentiated cells at early phases. (A–C) The 253G1-derived nonadherent cells at day 21 of differentiation were collected and stained by Wright-Giemsa solution. Abundant small cells with particularly high nuclear cytoplasmic ratios were detected (A, B). Note that the macrophage contained condensed nucleus-like substances (closed arrow head) and/or chromatin-like substances (arrows) in addition to its own nucleus (open arrow). Small cells were occasionally encompassed by the cytoplasm of macrophages (open arrow head). Scale bars indicate 20 μm. (D) The 253G1-derived nonadherent cells at day 23 of differentiation were subjected to colony assays. CFU-M, CFU-GM, and CFU-G were determined by phase contrast microscopic observation. Scale bars indicate 500 μm.

Thus, hiPSCs can generate reproducible HPCs with equivalent colony-forming activities to hESC-derived HPCs, although some lines of hiPSCs suffer from defective hematopoietic differentiation.

Discussion

In this article, we have provided the counterexamples to a previously reported finding that hiPSC-derived hemangioblast, the common progenitor of hematopoietic and endothelial cells, suffered from early senescence. In that report, hiPSC-derived HPCs was shown to have substantially decreased colony-forming activities and the majority of hiPSC-derived endothelial cells senesced after one passage (Feng et al., 2010). However, our data have clearly shown that the issue of early senescence can be overcome by selecting appropriate lines of hiPSCs and applying proper differentiation methods to them. Moreover, our results proved that retroviral insertion of reprogramming transgenes was not the cause of early senescence contrary to the discussion by the authors (Feng et al., 2010). We have also shown that, after sequential passages, hiPSC-derived VECs enter senescence as in the cases of hESC-derived VECs and primary human VECs, guaranteeing that hiPSC-derived VECs bear very low tumorigeneity, if any.

The key to our success in producing hiPSC-derived VECs that bear as high growth potentials as hESC-derived counterparts may reside, at least in part, in our usage of multiple hematopoietic cytokines in addition to VEGF. As we have shown previously, the six cytokines, SCF, IL6, IL3, BMP4, Flt3-L, and VEGF, as a whole work for the stable and high-purity production of subculturable VECs (Saeki et al, 2008). Interestingly, we are also observing that, under serum-free conditions, the presence of hematopoietic cytokine cocktail is crucial for the formation of spheres and their subsequent growth on gelatin-coated plates (M.N., unpublished finding). Thus, the usage of hematopoietic cytokine cocktail is advantageous not only for an achievement of high-efficiency differentiation but also survival and proliferation of the differentiated cells. Alternatively, the differentiation process per se, which is often followed by apoptosis, might include antiapoptotic processes as far as the differentiated cells keep surviving. In any event, stressful conditions should be avoided as much as possible from the differentiation procedures of hESCs/hiPSCs as in the case of their maintenance culture, where chromosomal aberrations are reportedly induced via stressful handling of the cells (Draper et al., 2004, Mitalipova et al., 2005).

As we mentioned, two lines of hiPSCs, 253G4 and 201B2, failed in directed differentiation into VECs. The 253G4- and 201B2-derived cells showed poor cord-forming activities and lacked VEC marker expressions, although they possessed Ac-LDL-uptaking capacities and were subculturable over 10 passages (data not shown). Their disadvantageous natures concerning VEC differentiation may be resulted from the possible line dependency in differentiation propensity among hiPSCs as reported in the case of hESCs (Osafune et al., 2008). Indeed, 253G4 and 201B2 showed very poor or no hemaotopoietic differentiation (data not shown). The finding that hiPSC lines with poor VEC-differentiating potentials bear little hemaotocyte-producing capacities seems very reasonable, because hematopoietic cells are derived from a specific population of vascular endothelial cells (Eilken et al., 2009).

Our findings together indicate that, although hiPSCs may be imposed line-dependent limitations in their differentiation capacities, they are not put inevitable fates of differentiation-dependent early senescence.

Footnotes

Acknowledgments

The authors greatly thank Professor Yamanaka at Center for iPS Cell Research and Application, Kyoto University, Japan, for generously providing the hiPSC lines (201B2, 201B7, 253G1, and 253G4). This work was supported by a Grant-in-Aid Scientific Research from the Ministry of Health, Labour and Welfare (KHD1029).

Author Disclosure Statement

The authors declare that no conflicting financial interests exist.

References

Draper

J.S.

, Smith

, Gokhale

et al. 2004. Recurrent gain of chromosomes 17q and 12 in cultured human embryonic stem cells. Nat. Biotechnol., 22:53–54.

Eilken

H.M.

, Nishikawa

, Schroeder

2009. Continuous single-cell imaging of blood generation from haemogenic endothelium. Nature, 457:896–900.

Feng

, Lu

S.J.

, Klimanskaya

et al. 2010. Hemangioblastic derivatives from human induced pluripotent stem cells exhibit limited expansion and early senescence. Stem Cells, 28:704–712.

Fusaki

, Ban

, Nishiyama

et al. 2009. Efficient induction of transgene-free human pluripotent stem cells using a vector based on Sendai virus, an RNA virus that does not integrate into the host genome. Proc. Jpn. Acad. Ser. B Phys. Biol. Sci., 85:348–362.

Kaji

, Norrby

, Paca

et al. 2009. Virus-free induction of pluripotency and subsequent excision of reprogramming factors. Nature, 458:771–775.

Kim

, Kim

C.H.

, Moon

J.I.

et al. 2009. Generation of human induced pluripotent stem cells by direct delivery of reprogramming proteins. Cell Stem Cell, 4:472–476.

Mitalipova

M.M.

, Rao

R.R.

, Hoyer

D.M.

et al. 2005. Preserving the genetic integrity of human embryonic stem cells. Nat. Biotechnol., 23:19–20.

Miura

, Okada

, Aoi

et al. 2009. Variation in the safety of induced pluripotent stem cell lines. Nat. Biotechnol., 27:743–745.

Nakagawa

, Koyanagi

, Tanabe

et al. 2008. Generation of induced pluripotent stem cells without Myc from mouse and human fibroblasts. Nat. Biotechnol., 26:101–106.

10.

Nakagawa

, Takizawa

, Narita

et al. 2010. Promotion of direct reprogramming by transformation-deficient Myc. Proc. Natl. Acad. Sci. USA, 107:14152–14157.

11.

Nakahara

, Nakamura

, Matsuyama

et al. 2009. High-efficiency production of subculturable vascular endothelial cells from feeder-free human embryonic stem cells without cell-sorting technique. Cloning Stem Cells, 11:509–522.

12.

Nelson

T.J.

, Martinez-Fernandez

, Yamada

et al. 2009. Repair of acute myocardial infarction by human stemness factors induced pluripotent stem cells. Circulation, 120:408–416.

13.

Okita

, Ichisaka

, Yamanaka

2007. Generation of germline-competent induced pluripotent stem cells. Nature, 448:313–317.

14.

Okita

, Nakagawa

, Hyenjong

et al. 2008. Generation of mouse induced pluripotent stem cells without viral vectors. Science, 322:949–953.

15.

Osafune

, Caron

, Borowiak

et al. 2008. Marked differences in differentiation propensity among human embryonic stem cell lines. Nat. Biotechnol., 26:313–315.

16.

Saeki

, Yogiashi

, Nakahara

et al. 2008. Highly efficient and feeder-free production of subculturable vascular endothelial cells from primate embryonic stem cells. J. Cell Physiol., 217:261–280.

17.

Saeki

, Saeki

, Nakahara

et al. 2009. A feeder-free and efficient production of functional neutrophils from human embryonic stem cells. Stem Cells, 27:59–67.

18.

Seki

, Yuasa

, Oda

et al. 2010. Generation of induced pluripotent stem cells from human terminally differentiated circulating T cells. Cell Stem Cell, 7:11–14.

19.

Suemori

, Yasuchika

, Hasegawa

et al. 2006. Efficient establishment of human embryonic stem cell lines and long-term maintenance with stable karyotype by enzymatic bulk passage. Biochem. Biophys. Res. Commun., 345:926–932.

20.

Takahashi

, Tanabe

, Ohnuki

et al. 2007. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell, 131:861–872.

21.

Tesar

P.J.

, Chenoweth

J.G.

, Brook

F.A.

et al. 2007. New cell lines from mouse epiblast share defining features with human embryonic stem cells. Nature, 448:196–199.

22.

Vallier

, Touboul

, Chng

et al. 2009. Early cell fate decisions of human embryonic stem cells and mouse epiblast stem cells are controlled by the same signalling pathways. PLoS One, 4:e6082.

23.

Warren

, Manos

P.D.

, Ahfeldt

et al. 2010. Highly efficient reprogramming to pluripotency and directed differentiation of human cells with synthetic modified mRNA. Cell Stem Cell, 7:618–630.

24.

Woltjen

, Michael

I.P.

, Mohseni

et al. 2009. piggyBac transposition reprograms fibroblasts to induced pluripotent stem cells. Nature, 458:766–770.

25.

, Hu

, Smuga-Otto

et al. 2009. Human induced pluripotent stem cells free of vector and transgene sequences. Science, 324:797–801.

26.

Zhou

, Wu

, Joo

J.Y.

et al. 2009. Generation of induced pluripotent stem cells using recombinant proteins. Cell Stem Cell, 4:381–384.