Hum. Reprod. Advance Access published online on October 9, 2008
Human Reproduction, doi:10.1093/humrep/den356
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Engraftment potential of human placenta-derived mesenchymal stem cells after in utero transplantation in rats
1 Division of High Risk Pregnancy, Mackay Memorial Hospital, 92 Sec. 2 Chung-San North Road, Taipei 104, Taiwan 2 Department of Medical Research, Mackay Memorial Hospital, Taipei 104, Taiwan 3 Mackay Medicine, Nursing and Management College, Taipei 112, Taiwan 4 Maternal and Fetal Health Research Group, University of Manchester, St Mary’s Hospital, Manchester M13 0JH, UK
5Correspondence address.Tel: +886-2-2543-3535; Fax: +886-2-2754-3769; E-mail: cpchen{at}ms2.mmh.org.tw
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Abstract |
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BACKGROUND: Human placental mesenchymal stem cells (hPMCs) are thought to be multipotent, but their fate after in utero transplantation is not known.
METHODS: hPMCs isolated from term placenta were assessed for their phenotype markers, mutilineage capacity, and immunomodulatory properties. Their engraftment potential was analyzed in a pregnant rat model after in utero transplantation at embryonic day 17. Immunohistochemistry, tracing of labeled cells, fluorescence in situ hybridization and real-time PCR were used to assess post-transplant chimerism.
RESULTS: In vitro, lineage-negative, CD34-negative hPMCs differentiated into osteocytes, adipocytes, hepatocytes and endothelial cells with tube formation, and actively suppressed the rat lymphocyte proliferative response to allogeneic lymphocyte stimulation (P < 0.0001). After in utero transplantation into pregnant rats, a low level of engraftment was achieved in various fetal tissues. Engraftment occurred in more than 60% of the fetal rats. Cells persisted for at least 12 weeks after delivery and evidence was obtained to suggest differentiation into specific lineages, including hepatocytes and hematopoietic cells. However, a greater number of hPMCs migrated to the placenta than to the fetus, thus limiting the degree of cell engraftment in fetal organs.
CONCLUSIONS: We conclude that hPMCs are mutipotent cells that can be engrafted long-term in immunocompetent rats after in utero transplantation.
Key words: engraftment/in utero transplantation/mesenchymal stem cells/placenta/rat
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Introduction |
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With advances in prenatal diagnostic techniques, many congenital diseases can be identified prior to the full development of the fetal immune system. The early fetus is uniquely tolerant to foreign antigens, accepting allogeneic or xenogeneic cells without the need to match major histocompatibility complex (MHC) antigens or induce immunosuppression (Muench, 2005
). Thus, the possibility arises of introducing foreign progenitor or stem cells to the unborn patient to correct an inherited defect such as an immunodeficiency disorder, hemoglobinopathy or enzyme deficiency (Le Blanc et al., 2005; Muench, 2005).
Cells of different origins have been used for in utero transplantation in a number of models. Human bone marrow-derived mesenchymal stem cells have been transplanted into fetal sheep and shown to persist for as long as 13 months with multilineage differentiation potential (Liechty et al., 2000). In utero transplantation of 1 x 108/kg CD34+ paternal canine bone marrow-derived cells in a canine model achieved a low level of microchimerism (<1%) in various tissues (Blakemore et al., 2004). Transplantation of human cord blood-derived CD34+-enriched stem cells into the peritoneal cavity of 45- to 60-day-old ovine fetuses achieved 18% engraftment at 1–3 months after birth (Young et al., 2003). Human fetal liver mononuclear cells or fetal bone marrow-derived CD34+ cells transplanted into NOD/SCID (non-obese diabetic/severe combined immunodeficiency) mice on Day 13 or 14 of gestation resulted in multilineage human hemopoietic engraftment in 10 and 12% of recipients at 8 weeks of age, respectively (Turner et al., 1998). Transplantation of human fetal mesenchymal stem cells into fetal mice with Duchenne muscular dystrophy on day 14–16 resulted in wide-spread long-term (19 weeks) engraftment in multiple organs (Chan et al., 2007). Embryonic stem cells transplanted into murine fetuses achieved low level (<0.4%) chimerism (Moustafa et al., 2004). Human amnion and chorion cells from term placenta had successfully engrafted in neonatal swine and rats 90 days after transplantation (Bailo et al., 2004).
These studies have shown that both hematopoietic and non-hematopoietic progenitor or stem cells have the capacity to home to the appropriate tissue microenvironment and contribute to some degree of chimerism. Even if the low level of chimerism (0.3%) achieved is not curative, this approach may be useful in severe mesenchymal or enzyme deficiency syndromes, where low-level protein expression may ameliorate disease pathology (Muenzer, 2004; Le Blanc et al., 2005; Chan et al., 2007). Alternative, and perhaps more convenient sources of human stem cells include umbilical cord (Wang et al., 2004), amniotic membranes (Bailo et al., 2004; Miki et al., 2005; Ilancheran et al., 2007) and the placenta (Fukuchi et al., 2004; Yen et al., 2005). Placenta-derived mesenchymal stem cells have been reported to proliferate in vitro, maintaining a homogenous morphology and consistent phenotype, as well as differentiating into bone, cartilage, adipose tissue, hepatocytes or insulin secretion cells (Fukuchi et al., 2004; Yen et al., 2005; Chien et al., 2006; Zhang et al., 2006; Chang et al., 2007). These placenta- derived mesenchymal stem cells are generally negative for CD34, CD45 and HLA-DR expression and positive for CD29, CD44, CD73, CD90, CD105 and CD166 (Barry et al., 1999, 2001; Parolini et al., 2008). In addition to their multipotency, they have a direct immunosuppressive effect on the proliferation of CD4+ and CD8+ lymphocytes from human peripheral blood and umbilical cord blood in vitro, and are expected to have a potential application in allograft transplantation (Li et al., 2007a). Proinflammatory cytokines, such as RANTES (regulated upon activation, normally T-expressed, and secreted), interleukin (IL)-1, IL-6 and IL-8 can stimulate the proliferation of these cells (Li et al., 2007b). However, their fate in vivo after systemic administration in not known.
In this study, we established a rat model for in utero transplantation of human placental mesenchymal stem cells (hPMCs) to investigate if these cells would effect long-term, organ-specific engraftment.
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Materials and Methods |
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Isolation, culture and labeling of placenta-derived cells
Clinically normal human term placentas (37–40 weeks of gestation, n = 28) were collected after Cesarean section. Tissue was obtained after informed consent of the women, and all experiments were approved by the Institutional Review Board of Mackay Memorial Hospital, Taipei. The animal portion of this study was specifically approved by the ethics committee for animal experimentation.
Isolation of hPMCs was modified from previously reported methods (Zhang et al., 2004; Yen et al., 2005; Portmann-Lanz et al., 2006). About 100 g tissue from the central placental cotyledons was minced, trypsinized (0.25% trypsin-EDTA solution; Invitrogen, Carlsbad, CA, USA) and treated with 10 U/ml DNAse I (Sigma, St Louis, MO, USA) in Dulbecco’s modified Eagle’s medium (DMEM; GIBCO, Grand Island, NY, USA) at 37°C for 5 min several times, and filtered through a 70-mm cell strainer (BD Biosciences, San Jose, CA, USA). The supernatants were pooled and spun at 1000 g for 10 min. Mononuclear cells in the medium were recovered by Percoll density gradient fractionation (1.073 g/ml, Sigma) (Pittenger et al., 1999). The cells were resuspended and seeded in a 25 cm2 flask. Cultures were maintained in DMEM with 10% fetal bovine serum (FBS; Hyclone, Road Logan, UT, USA) at 37°C with 5% CO2. Approximately 2–3 weeks later, some colonies consisting of fibroblast-like cells were observed. These cells were trypsinized and replated for expansion. In order to obtain single cell-derived hPMC clones, cells were serially diluted in 96-well culture plates (BD Biosciences) at a final density of 60 cells/plate. Colonies that grew with homogeneous bipolar morphology were expanded.
To track cell migration and protein expression in vivo, cells destined for in utero transplantation were incubated with 1 µg/ml bisbenzimide (blue nuclear fluorescence; Hoechst 33342; Sigma) for 24 h at 37°C before trypsinization. The viability of hPMCs was determined by trypan blue exclusion assay.
Flow cytometry
The cell surface phenotype of hPMCs was characterized using a panel of phycoerythrin (PE)- or fluorescein isothiocyanate (FITC)-conjugated antibodies purchased from Serotec (Oxford, UK), Chemicon (Temecula, CA, USA), or BD Biosciences-Pharmingen (NJ, USA) using standard fluorescence-activated cell sorting analysis.
Chimerism in the blood or bone marrow was evaluated in mononuclear cells isolated by a Ficoll density gradient (Histopaque 1083; Sigma) from peripheral blood or bone marrow from fetal rats after in utero transplantation. Mouse anti-human HLA-ABC (W6/32) PE-conjugated monoclonal antibody was used as a marker of hPMCs. Mouse anti-human CD45 (F10-89-4) FITC-conjugated monoclonal antibody was used to detect human hematopoietic cells. The cells were further counted by 2-color flow cytometry (Becton Dickinson) to show the hematopoietic lineage engraftment of hPMC in recipient rats. Ten thousand cells were counted from each sample. An aliquot of unstained cells from each cell suspension was used as the negative control to determine background autofluorescence and to set a compensation gate. Cell suspensions from rats of the same age that had not received human cells were used as additional negative controls in each experiment. Test samples were evaluated by subtracting the negative control values.
Induction of osteogenic or adipogenic differentiation
hPMC cells in passages 6 through 10 (n = 7) were cultured in one of two media: (i) osteogenic medium consisting of DMEM containing 10% FBS (Hyclone), 0.1 µM dexamethasone, 10 mM β-glycerol phosphate and 500 µM ascorbic acid (Sigma) or (ii) adipogenic medium consisting of DMEM containing 10% FBS (Hyclone), 1 µM dexamethasone, 5 µg/ml insulin, 500 µM isobutylmethylxanthine, 200 µM indomethacin and 10 µg/ml insulin (Sigma) (Zhang et al., 2004). The medium was changed every 3 days. After 2 weeks of culture, the cells were fixed with methanol, stained either with 1% Alizarin Red S (Sigma) to assess for calcium phosphate deposition indicating osteogenic differentiation or with Oil Red O (Sigma) to look for lipid droplets indicating adipogenic differentiation (Reyes et al., 2001).
Induction of endothelial cell differentiation
hPMCs were seeded at a density of 3 x 103 cells/cm2 in Petri dishes and cultured in Endothelial Cell Growth Medium 2 (EGM2; Promocell, Heidelberg, Germany) with Supplement Mix (Promocell) and 2% FBS (Hyclone). Another 50 ng/ml vascular endothelial growth factor (VEGF, Chemicon) was added to the medium to induce differentiation. The cultures were maintained for 3 weeks, with the culture medium being replaced every 7 days.
Induction of hepatocyte differentiation
hPMCs (5 x 105) were plated on 6-cm culture dishes coated with poly-L-lysine (Sigma) and cultured in differentiation medium consisting of 60% low glucose DMEM and 40% MCDB201 (Sigma) supplemented with 1x Insulin–Transferrin–Selenium, 4.7 µg/ml linoleic acid, 1 mg/ml bovine serum albumin, 10–4 M ascorbic acid, 10–8 M dexamethasone (Sigma), 10 ng/ml epidermal growth factor (R&D Systems, Minneapolis, MN, USA) and 10 ng/ml platelet-derived growth factor-BB (R&D Systems). After 16 h of incubation, the cells were further cultured with differentiation medium containing 20 ng/ml hepatocyte growth factor (PeproTech, Rocky Hill, NJ, USA) and 10 ng/ml fibroblast growth factor-4 (PeproTech) for 28 days. The medium was changed once a week (Chien et al., 2006).
In vitro angiogenesis
Induction of capillary tube formation was performed by using an In Vitro Angiogenesis kit (Chemicon) as recommended by the manufacturer. The cells were harvested and resuspended to 1 x 105 cells/ml in EGM2, after which 100 µl of the cell suspension was added onto the surface of the polymerized ECMatrixTM in each well. The cells were incubated for 6 h at 37°C for full development of capillary-like networks, which were assessed under a light microscope at x50 magnification (Axiovert 200; Zeiss Müchen-Hallbergmoos, Germany).
Mixed lymphocyte reaction
To evaluate whether the hPMCs elicit an immunomodulatory effect, the mixed lymphocyte reaction was assessed as previously described (Ringden and Berg, 1977). Briefly, purified Sprague–Dawley (SD) rat peripheral blood mononuclear cells (PBMCs) were prepared by density gradient centrifugation of heparinized blood over Histopaque 1083 (Sigma). Separated cells were cultured in RPMI-1640 medium supplemented with 10% FBS (Hyclone) for 24 h to remove monocytes by adherence. The remaining PBMCs were then collected and counted. hPMCs were then treated with 50 µg/ml mitomycin C (Sigma) for 4 h and plated 24 h ahead of co-culture with PBMCs. Responding PBMCs (2 x 105/well) were cultured in triplicate mixed at different ratios with hPMCs in a flat-bottomed 96-well plate to ensure efficient cell–cell contact for 4 days in 0.2 ml RPMI-1640 containing 10% FBS. An equal number of mitomycin C-treated PBMCs (2 x 105/well) were used as stimulating cells. All plates were pulsed with 10 µM 5-bromo-2'-deoxy-uridine (BrdU)/well during the last 18 h of the 4-day culture. Lymphocyte proliferation was determined using BrdU Labeling and Detection Kit III (Roche Diagnostics GmbH, Mannheim, Germany) according to the manufacture’s instructions.
Reverse–transcription polymerase chain reaction
The following primers were used for PCR reactions: human osteopontin sense, 5'-GTGCCATACCAGTTAAACA-3'; antisense, 5'-CTTACTTGGAAGGGTCTCT-3'; human peroxisome proliferator-activated receptor gamma2 (PPAR
2) sense, 5'-TGTCAGTACTGTCGGTTTC-3'; antisense, 5'-AATGGTGATTTGTCTGTTG-3'; human von Willebrand factor (vWF) sense, 5'-ACGTGATCCTTCTCCTGGATG-3'; antisense, 5'-TTCACCACGTTGGAGTCGCCT-3'; CD31 sense, 5'-GACGTGCTGTTTTACAACATCTC-3'; antisense, 5'-CCTCACGATCCCACCTTGG-3'; vascular endothelial (VE)-cadherin sense, 5'-GGGAGACCACGCCTCTGTC-3'; antisense, 5'- GGAGGCCCTGGGCATCTC-3': human albumin sense, 5'-CAACTATGTCCGTGAGCTTCCA-3'; antisense, 5'-GTGGTCGGTGCTGGTCTATATG-3'; 18S sense, 5'-TAGAGCTAATACATGCCGACGG-3'; antisense, 5'-GGGCCTCGAAAGAGTCCTGTATT-3'. The RT–PCR were performed as previously described (Feng et al., 1999; Hong et al., 2005; Jiang et al., 2006).
Immunohistochemistry
Sections (5 µm thickness) were air-dried and fixed in ice-cold acetone or 4% paraformaldehyde for 10–20 min. Immunohistochemistry was performed as previously described (Chen and Aplin, 2003).
Primary antibodies against specific proteins included CD31 (1:50; Dako, Carpinteria, CA, USA), VE-cadherin (1:100; Chemicon), vWF (1:200; Sigma), troponin I (1:200; a marker of cardiac muscle; Santa Cruz, Santa Cruz, CA, USA), surfactant protein D (1:100; a marker of alveolar type II cells; abcam), albumin (1:10; a marker of expressed liver protein; R&D Systems), CD45 (1:200; a marker of leucocytes; Chemicon) and Ki-67 (1:100; expressed in all active phases of the human cell cycle; Dako). For cytokeratin staining, monoclonal antibody against cytokeratin 18 (1:100; Chemicon) was mixed with monoclonal antibody against keratin 8 (1:100; Chemicon) in a 1:1 dilution.
The double-immunofluorescence technique with specific antibodies against bisbenzimide-labeled cells was further analyzed with a Zeiss LSM510 laser-scanning confocal microscope as previously described (Shyu et al., 2008). The green (FITC) fluorochromes on the slides were excited by the laser beam at 488 nm, and emissions were acquired sequentially with a photomultiplier tube through 500- to 535-nm emission filters.
Animal experiments
The care of animals complied with the institutional guidelines. On embryonic day (E) 17, all dated pregnant SD rats were anesthetized with an i.p. injection of chloral hydrate (0.35 g/kg), and underwent midline laparotomy to expose the uterine horns. Using a 30-gauge needle (Hamilton, Reno, Nevada) under a dissection microscope (x10 magnification; Olympus, Tokyo, Japan), each fetus was injected i.p. through the uterine wall with 1 x 106 hPMCs in 5 µl of normal saline. The uterus was replaced, and the incision was closed.
On E21, a low abdominal midline incision was made and the number of live fetuses in each uterine horn was recorded. Then, placenta, fetal blood and fetal organs including brain, heart, lung, liver, spleen and bone marrow were collected. For studies of blood, bone marrow and various other organs in live offspring, rats that had received cell transplants during fetal life, were sacrificed post-natally at 3 or 12 weeks. Peripheral blood was collected into EDTA tubes from the rat fetuses after decapitation at E21 and from femoral vessels at 3 or 12 weeks post-natally. A solution containing 150 mM ammonium chloride and 10 mM sodium bicarbonate with 0.1 mM EDTA (pH 7.0) was added to lyse erythrocytes. After centrifugation, the cell pellets were washed and resuspended in 2% bovine serum in phosphate-buffered saline (PBS). Bone marrow was harvested from E21- or 3- or 12-week-old recipients by flushing the tibias and femurs with PBS through a 23-gauge needle. A single-cell suspension was made by three gentle passes through the needle. Cell suspensions were passed through a 70 µm nylon mesh filter, and mononuclear cells were isolated by Ficoll gradient separation as described earlier.
Fluorescence in situ hybridization
Cryosections (5 µm thickness) of rat tissue were probed using an X chromosome probe (labeled with SpectrumOrange; CEP X, Xp11.1-q11.1, locus DXZ1, Vysis, Bergisch-Gladbach, Germany) with or without a Y chromosome probe (labeled with SpectrumGreen; CEP Y, DYZ1, Yq12, Vysis) as recommended by the manufacturer. The tissue sections were then counterstained with 10 µl 4',6- diamidino-2-phenylindole dihydrochloride (DAPI; Sigma), and viewed using a Zeiss microscope (Müchen-Hallbergmoos, Germany).
FISH in combination with immunohistochemistry was performed as previously reported (Donadoni et al., 2004). After post-hybridization washes, sections were incubated with antibody against human-specific albumin (HAS-11; 1:20; Sigma) at room temperature for 1 h, followed by washing in 2x saline sodium citrate and exposure to a FITC-conjugated secondary antibody (Dako) for 1 h.
Real-time PCR
Numbers of hPMCs in rat tissue were estimated based on the quantitative detection of human β2-microglobulin-specific sequences present in rat tissue (Zijlstra et al., 2002; Chen et al., 2008) Genomic DNA was extracted from mononuclear cells from 200 µl peripheral blood, bone marrow or frozen tissues using a spin-column extraction kit (Qiagen, Hilden, Germany). The rat organs were weighed and the genomic DNA of total heart, spleen and 100 mg of other organs extracted and pooled. Human-specific β2-microglobulin primers (sense: 5'-ACCCCCACTGAAAAAGATGAGTATG-3' and antisense: 5'-ACTATCTTGGGCTGTGACAAAGTC-3'), and an internal TaqMan detection probe (5'-CCTGCCGTGTGAACCA-3'; Applied Biosystems) were used. PCR reactions contained 250 ng template DNA, 10 µl 2xTaqMan universal PCR Master Mix, 1 µl 20xTaqMan Gene Expression Assay (18 µM each primer and 5 µM probe; Applied Biosystems) in a 20 µl reaction. All sequences were amplified by a first step of 15 s at 95°C, followed by 1 min at 60°C for 50 cycles. No signal amplification was observed in rat tissue without human genomic DNA.
Primers and probes for 18S were obtained from Applied Biosystems (18S, Hs99999901_s1; MOL1980000, 198000). Relative quantification of target gene expression was calculated by the comparative Ct method. When indicated, the β2-microglobulin signal was normalized against the relative quantity of 18S and expressed as 
Ct=Ct β2-microglobulin–Ct 18S.
A standard curve was generated through quantitative amplification of genomic DNA extracted from a serial dilution of hPMCs mixed with the DNA of individual rat placenta homogenates. Total human cell DNA was prepared from 1 x 106 hPMCs. Serial dilutions of 25 ng of this DNA preparation in 250 ng rat tissue genomic DNA were used and the corresponding cell numbers calculated. This calibration curve was created by plotting the number of hPMCs in rat tissue DNA corresponding to the amount of each standard DNA versus the value of its Ct. The number of transplant human β2-microglbulin-positive cells for all of the tested samples was expressed as hPMC equivalents per milligram of total rat tissue, as determined by the standard calibration curve. By interpolating the β2-microglobulin signal from experimental samples with the standard curve, the actual number of hPMCs could be estimated over a range of 9–35 526 cells per milligram of rat tissue.
Statistical analysis
The data are described as mean ± SD. Differences were assessed by using the independent-samples t-test, paired-samples t-test or Mann–Whitney U-test when appropriate. A P-value of <0.05 was considered significant.
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Results |
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Isolation and characterization of mesenchymal cells from term placenta
To determine whether hPMC in term placentas are multipotent, we extracted 5.5 ± 1.0 x 107 nucleated cells per 100 g of wet tissue weight from 28 placentas delivered at a mean gestational age of 38.6 ± 0.5 weeks. There was no significant correlation between gestational age, number of mononuclear cells collected and success in generation of hPMCs. After 7–21 days (mean 14 days), adherent cells with fibroblastic morphology were detected.
Independent of gestational age, hPMCs were negative for CD14, CD34, CD45, HLA-DR, HLA-G and cytokeratin 7; expressed moderate levels of CD13 and CD54; and had high levels of CD29, CD44, CD49e, CD73, CD90, CD105, CD166 and HLA-ABC (Fig. 1B). These characteristics are consistent with mesenchymal stem cells. To exclude variation arising from the possibility of a heterogeneous starting cell population, five clones were expanded after culture at limiting dilution and the surface marker repertoire profiled. We observed phenotypes similar to the parental cells. The hPMCs were cultured for more than 20 passages without any spontaneous differentiation (data not shown). However, the ability of hPMCs to differentiate into multilineage cells decreased after passage 20. To confirm the absence of maternal cell contamination, FISH analysis was applied using a green fluorescent Y-probe associated with an orange fluorescent X-probe in hPMCs from the placenta of a male fetus. The cells were Y chromosome-positive, indicating that they were of fetoplacental origin (Fig. 1A).
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Multilineage differentiation potential of hPMC in vitro
Individual hPMC preparations (n = 7) from passages 6–10 were analyzed for their potential to differentiate into adipocytes, osteocytes, endothelial cells or hepatocytes.
Osteogenic differentiation of hPMCs was demonstrated by nodule formation and staining with Alizarin Red S, indicating calcium salt crystallization (Fig. 1D). Very little staining was observed in control (Fig. 1C). mRNA for osteopontin, an osteogenic marker, was also expressed in induced but not control cells (Fig. 1E).
Similarly, hPMCs differentiated to form adipocytes as demonstrated by Oil Red O-positive cytoplasmic lipid droplets (Fig. 1G), not seen in control cells (Fig. 1F). The adipocyte marker PPAR2 mRNA was expressed in induced but not untreated cells (Fig. 1H).
hPMCs treated with 50 ng/ml VEGF for 21 days underwent endothelial differentiation. There was no difference of cell morphology between the differentiated and undifferentiated cells, but differentiated cells had markedly enhanced vWF (Fig. 1J), CD31 (Fig. 1L) and VE-cadherin (Fig. 1M) staining after 21 days of cultivation compared with undifferentiated cells (Fig. 1I). Similarly, the differentiated cells expressed mRNA encoding these endothelial markers, while undifferentiated cells did not (Fig. 1K). Capillary formation in vitro was assayed by plating hPMCs on ECMatrixTM gel. VEGF-treated cells developed tube-like structures after 6 h of cultivation (Fig. 1O), while undifferentiated cells did not (Fig. 1N). Under inducing conditions, hPMCs expressed albumin protein and mRNA, which mark functional hepatocytes (Fig. 1Q and R).
Mixed lymphocyte reaction
To test whether the hPMCs inhibited proliferation of SD rat PBMCs, the two types of cells were co-cultured at various ratios. Native PBMCs showed a strong proliferative response to allogeneic mitomycin C-treated cells (absorbance for BrdU incorporation: 0.32 ± 0.04) (Fig. 2). There was a significant reduction in PBMC proliferation when mixed cultures of PBMCs stimulated by mitomycin C-treated allogeneic PBMCs (1:1 ratio) were grown in the presence of mitomycin C-treated hPMCs. Proliferation was suppressed in a dose-dependent manner as the number of hPMCs added to the culture was increased. There was relatively low suppression at a dose of 102 hPMCs, whereas it increased significantly when 2 x 104 hPMCs were added (absorbance for BrdU incorporation: 0.27 ± 0.06 versus 0.06 ± 0.04, P < 0.0001).
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Engraftment of hPMCs in fetal organs
Preliminary trials with 0.7% methylene blue demonstrated that microinjection into the peritoneal cavity could be reliably performed at E17, with the injected material rapidly appearing in the fetal abdomen without leakage into the amniotic cavity. The fetal loss rate of 47% (Table I) after transplantation of xenogeneic hPMCs is similar to prior reports (Turner et al., 1998
; Rio et al., 2005).
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To show that hPMCs injected in utero on E17 engrafted in fetal organs, we collected fetal organ samples at E21 (term is E21 ± 1 days) as well as adult organ specimens collected 3 weeks, or 12 weeks post-natally (Table I). Most fetal tissues had demonstrable hPMC engraftment at E21. Although the distribution pattern and numbers of cells in individual fetuses varied, hPMCs were detectable in more than 60% of the fetal rats and persisted for 12 weeks post-natally (Table I). There was no evidence of rejection of these xenogeneic cells.
We assessed the presence of hPMCs in various fetal rat tissues. Hematoxylin-eosin staining and imunofluorescence of serial sections are shown in Fig. 3. Bisbenzimide-labeled hPMC nuclei were found in rat myocardium with immunostaining for troponin I (Fig. 3B–D). Laser-scanning confocal microscopy of lung tissue stained with antibody to surfactant protein D clearly showed co-localization with nuclear bisbenzimide, indicating cells of hPMC origin (Fig. 3F). Standard immunofluorescence images were also gathered and demonstrated co-localization of the surfactant protein D with transplanted hPMCs in the recipient lung (Fig. 3H–J). In areas of lung containing abundant hPMCs, the nuclei of bisbenzimide-labeled hPMCs were also positive for Ki67, indicating that these cells were proliferating (Fig. 3K). Bisbenzimide-labeled hPMCs were also present in fetal liver and spleen immunostained for albumin and leukocyte marker CD45 (Fig. 3M–S). Unexpectedly, the greatest number of hPMCs (49 ± 12 hPMCs/mg tissue) was observed engrafted in the placenta (Table I). From the image taken from the area around a large vessel that enters the placenta near the cord insertion, a large number of bisbenzimide-labeled hPMCs were identified within the labyrinth in close proximity to cytokeratin-positive trophoblasts (Fig. 3U–W). These cells are presumed to have migrated to the placenta through the fetal circulation. The identity of hPMCs present in fetal tissues was further confirmed by detection of human β2-microglobulin-specific sequences through real-time PCR (Table I).
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hPMCs persisted in post-natal tissues including brain, lung, heart, liver and spleen for 12 weeks post-natally (Table I). FISH demonstrated two positive signals in each cell, indicating that they contained two X chromosomes (Fig. 4A). FISH and immunofluorescence were combined to determine if hPMCs underwent differentiation after engraftment. A few cells that expressed human X chromosome-positive signals were stained for human-specific albumin in the sinusoids of rat liver at 12 weeks after birth (Fig. 4C–F), indicating the engrafted hPMCs may differentiate into functional liver cells. However, limited by the low level of engrafted hPMCs and lack of availability of other human-specific antibodies, we were not able to confirm that hPMCs which persisted elsewhere for 12 weeks had differentiated into the respective lineages.
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Using flow cytometry we evaluated the frequency and level of human HLA-ABC-positive hPMCs in the peripheral blood and bone marrow of recipient rats after in utero transplantation. Because there was a detectable and not negligible cell fraction recognized by anti-human HLA-ABC or CD45 antibody in the blood or bone marrow of rats without cell transplantation, a group of control rats was used when measuring the level of detectable engraftment at E21 and 3 or 12 weeks post-natally.
Rat pups that received prenatal hPMC transplantation had hPMCs in the peripheral blood or bone marrow with an overall frequency of engraftment of 27.3 and 18.2% at E21, respectively (Table I). However, prenatal transplantation of these rat at E17 with 1 x 106 hPMCs resulted in about 5% (after subtracting control values) of peripheral blood engraftment (7.4 ± 1.0 versus control 2.1 ± 1.1%; n = 3) at E21 (5 days after transplantation) and about 2% at 3 weeks after birth (6.7 ± 0.6 versus control 4.1 ± 0.2%; n = 3), decreasing to a non-detectable level at 12 weeks after birth (not shown). Similarly, about 1% of bone marrow engraftment (5.2 ± 0.1 versus control 4.3 ± 0.4%; n = 3) was seen at E21, decreasing to a non-detectable level at 3 weeks after birth (1.8 ± 0.1 versus control 1.9 ± 0.1%; n = 3). Representative data were shown in Fig. 5A and B.
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To determine if the engrafted hPMCs underwent hematopoietic differentiation in the peripheral blood and bone marrow of chimeric recipients, the presence of HLA-ABC-positive hPMCs that co-expressed CD45 was examined. About 0.6% of HLA-ABC-positive cells in peripheral blood co-expressed the hematopoietic cell marker CD45 (1.0 ± 0.3 versus control 0.4 ± 0.2%; n = 3) at E21, indicating the hematopoietic lineage differentiation of hPMCs. The level of transplantated hPMC that co-expressed HLA-ABC and CD45 decreased to 0.4% (1.6 ± 0.1 versus control 1.2 ± 0.1%; n = 3) at 3 weeks and was non-detectable at 12 weeks (not shown) post-natally. In the bone marrow, we found that <0.3% of HLA-ABC-positive cells co-expressed the hematopoietic cell marker CD45 (1.4 ± 0.4 versus control 1.1 ± 0.4%; n = 3) at E21, indicating the very low hematopoietic lineage differentiation of hPMCs. The level of transplanted hPMC that co-expressed HLA-ABC and CD45 decreased to 0% (0.4 ± 0.1 versus control 0.5 ± 0.1%; n = 3) at 3 weeks and was not detectable at 12 weeks post-natally (not shown). A representative data were shown in Fig. 5C and D.
A real-time quantitative PCR assay for human β2- microglbulin was developed in order to evaluate hPMC numbers engrafted in the organs of recipient rat fetuses. DNA extracted from rat tissue was mixed with DNA extracted from hPMC cells (Supplementary material, Fig. S1A–D) to produce a highly precise and reproducible standard curve over a wide linear range down to nine hPMC cells in 250 ng genomic DNA.
Postmortem analysis of the organs from E21 fetal rats confirmed that hPMCs engrafted in more than 60% of fetal rats after in utero transplantation. Engrafted hPMC persisted for 12 weeks post-natally without significant differences in the cell numbers in the brain, heart, lung, liver and spleen (P > 0.05; Table I). However, hPMCs were not detectable in the recipient blood or bone marrow at 12 weeks after birth by real-time quantitative PCR assay, which was consistent with the result of flow cytometry assay.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionSupplementary materialFundingAuthor contributionsReferences |
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We have demonstrated that hPMCs are multipotent in vitro. We further used cell tracking (Frangioni and Hajjar, 2004
), FISH and real-time PCR to demonstrate that hPMCs are capable of engrafting in multiple fetal tissues (brain, heart, lung, liver and spleen) and can survive post-natally for at least 12 post-natal weeks after in utero transplantation. The data suggest that these cells may undergo differentiation into different phenotypes after engraftment.
The phenotype and capacity for differentiation of our hPMC preparations were similar to those of human bone marrow-derived mesenchymal stem cells (Pittenger et al., 1999; Orlic et al., 2001). Our cells did not express hematopoietic markers (CD34, CD45) but they did express a number of adhesion molecules and mesenchymal markers, including CD29, CD44, CD73, CD105 and CD166 (Barry et al., 1999, 2001). In agreement with previous reports (Fukuchi et al., 2004; Zhang et al., 2004; Yen et al., 2005; Chien et al., 2006; Portmann-Lanz et al., 2006), they were able to proliferate in vitro, maintaining a homogeneous morphology, and, under appropriate stimulation, differentiated into multiple cell lineages including osteocytes, adipocytes, hepatocytes and endothelial cells. The endothelial cells could also be induced to undergo tube formation in vitro. These observations are supported by the report of Fukuchi et al. (2004) that hPMCs express many of the genes derived from mesoderm, ectoderm and endoderm. Those authors also demonstrated hematopoietic/endothelial cell-related genes in hPMCs (Fukuchi et al., 2004).
Engraftment involves many complex steps to allow transiently circulating hPMCs to become resident within the tissue of various organs (Nilsson and Simmons, 2004). hPMCs express human leukocyte antigen class I (HLA-ABC) but not class II antigens (HLA-DR), which may limit the immune response and antigen recognition by the host (Le Blanc and Ringden, 2005). Furthermore, the major MHC appears to have little adverse influence on engraftment at an early gestational age. High rates (>50%) of long-term engraftment are thus theoretically achievable, although the degree of engraftment (i.e. the percentage of donor cells in any particular type of tissue) may be very low (0.0001–1%) (Carrier et al., 1995). However, even when donor cells are fewer than 0.01% in recipient blood or tissues, they appear to be sufficient to induce and maintain tolerance (Carrier et al., 1995). Mesenchymal progenitors isolated from amnion and chorion did not induce allogeneic or xenogeneic lymphocyte proliferation responses (Bailo et al., 2004). Culture-expanded hPMCs were observed to inhibit proliferation of cord blood lymphocytes triggered by allogeneic peripheral blood lymphocytes or phytohemagglutinin in a dose-dependent manner (Li et al., 2005, 2007a). We found a similar effect of hPMCs on allogeneic lymphocyte proliferation, again confirming the findings of others (Le Blanc and Ringden, 2005). The apparent immunomodulatory properties of hPMCs may play a major role in the induction of tolerance to allogeneic or xenogeneic transplantation. Immunosuppression induced by mesenchymal stem cells is a complex process which may involve IL-2 and IL-10 signaling or prostaglandins (Rasmusson et al., 2005). Additionally, it is thought that contact between the two types of cells, particularly of prolonged duration, is required for full inhibition (Puissant et al., 2005).
Trans-species animal models have been widely used in the study of stem cell migration and engraftment (Liechty et al., 2000; Saito et al., 2002). It has been shown that human cord blood-derived cells can differentiate into hepatocytes in the mouse liver without evidence of cellular fusion (Newsome et al., 2003). Human microchimerism was observed in various organs and tissues at 4 months after transplantation of human amnion and chorion mesenchymal progenitors in neonatal swine and rats (Bailo et al., 2004). Human mesenchymal stem cells colonized multiple fetal sheep tissues for as long as 13 months after in utero transplantation (Liechty et al., 2000). Differences observed in cell numbers may be due to colonization efficiency in different tissue environments or the rate of cell turnover in each organ (Krause et al., 2001). Our study adds to this body of work by establishing an in utero (E17) model of xenogeneic hPMC transplantation in immunocompetent rats. More than 60% of the adult rats demonstrated hPMC engraftment at 12 weeks after birth, although the degree of engraftment was low, usually less than 20 cells per milligram of rat tissue. In fact, most of the injected hPMCs ended up in the placenta after i.p. transplantation. This phenomenon suggests a cell homing effect by which hPMCs migrate to the placenta, perhaps resulting in a lower degree of engraftment in the fetal organs. The fate of mesenchymal stem cells after in utero transplantation is still largely unknown. We demonstrated engraftment of transplanted human cells in various fetal tissues, and found some evidence that they underwent specific cell lineage differentiation. For example, human-specific albumin staining of human X chromosome-bearing cells engrafted in liver tissue suggests hepatocytic differentiation. Furthermore, co-expression of human HLA-ABC and CD45 by hPMCs in the rat circulation may indicate that hPMCs differentiate into hematopoietic cells. The developmental environment of the fetus may enable a wider spectrum of cell fates than in an adult tissue, but further work will be required to investigate other cell lineages. Additionally, we cannot exclude the possibility that hPMCs present in host tissues either remain undifferentiated or adopt other so far uncharacterized phenotypes. Thus, the actual function of such differentiated cells must be demonstrated before we can be confident that they have therapeutic potential.
Identifying sources of donor cells is a major factor for the success of in utero transplantation. It has been suggested an immune barrier exists to allogeneic engraftment and that stable, long-term, multilineage chimerism can only be achieved after transplantation of congenic cells (Peranteau et al., 2007). The apparent low frequency of cell engraftment may be further explained by the xenogeneic barrier in this model. In the observations of Schoeberlein et al. (2004), the frequency of xenogeneic stem cells had rapidly diminished to <1% by 2 days after in utero transplantation. However, the low frequency of cells indicated by real-time PCR in our study may be misleading. The number of hPMCs per milligram of rat tissues was not significantly decreased from birth to 12 weeks after birth. This finding indicates that engrafted hPMCs may proliferate in host rat organs as has been reported in other animal models (Zanjani et al., 1996; Liechty et al., 2000). The presence of even a few stem cells that proliferate or differentiate could have a dramatic rescue effect on certain congenital diseases (Frattini et al., 2005). The low degree of engraftment produced by in utero transplantation might also be increased by a post-natal boost with even a small number of donor cells (Peranteau et al., 2002). Even if the cells did not differentiate enough to be functional, the establishment of low-level chimerism with MHC-mismatched cells in utero could render the patient tolerant to subsequent post-natal therapeutic transplantation without the need for conditioning therapy (Peranteau et al., 2002).
Mesenchymal stem cells are mainly derived from bone marrow (Orlic et al., 2001), but it may be difficult to obtain sufficient autologous cells from some patients, particularly those who are older or who have malignancies. Therefore, alternative sources are needed. It appears that hPMCs from an allogeneic donor might constitute such a source. A further potential benefit is the exposure of the fetus to allogeneic cells, inducing tolerance such that future treatment with allogeneic cells would not constitute an immunologic challenge (Muench, 2005). Our study supports the possibility that hPMCs are transplantable, at least in utero, and thus hold promise for cellular and gene therapy applications. Much work remains to be done before this approach can be used clinically.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionSupplementary materialFundingAuthor contributionsReferences |
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Supplementary material is available at http://humrep.oxfordjournals.org/.
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This work was supported by grant from Mackay Memorial Hospital (MMH-E 96001 to C.-P. Chen).
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C.-P.C. conceived, designed the experiments and wrote the paper. S.-H.L., Y.-H.W., P.-C.C., C.-S.H., C.-C.K., M.-Y.L. and C.-Y.C. performed the experiments. C.-P.C., J.-P.H. and J.-D.A. analyzed the data. C.-P.C. and J.-P.H. contributed reagents/materials/analysis tools.
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Submitted on August 14, 2007; resubmitted on August 17, 2008; accepted on August 21, 2008.
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