ESCs are important as research subjects since the mechanisms underlying cellular differentiation, expansion, and self-renewal can be studied along with differentiated tissue development and regeneration in vitro. Furthermore, human ESCs hold promise for cell and tissue replacement approaches to treating human diseases. The rhesus monkey is a clinically relevant primate model that will likely be required to bring these clinical applications to fruition. Monkey ESCs share a number of properties with human ESCs, and their derivation and use are not affected by bioethical concerns. Here, we summarize our experience in the establishment of 18 ESC lines from rhesus monkey preimplantation embryos generated by the application of the assisted reproductive technologies. The newly derived monkey ESC lines were maintained in vitro without losing their chromosomal integrity, and they expressed markers previously reported present in human and monkey ESCs. We also describe initial efforts to compare the pluripotency of ESC lines by expression profiling, chimeric embryo formation, and in vitro-directed differentiation into endodermal, mesodermal, and ectodermal lineages.
ESCs are important as research subjects since the mechanisms underlying cellular differentiation, expansion, and self-renewal can be studied along with differentiated tissue development and regeneration in vitro. Stem cells also provide the underpinning for the field of regenerative medicine and offer hope for the treatment of a wide range of human conditions that can be attributed to the loss or malfunction of specific cell types. Finally it is becoming increasingly apparent that stem cells could ultimately serve as a source of viable gametes [1, –3]. Nonhuman primates (NHPs), especially old world macaques, are valuable animal models because of their extensive use in biomedical research, their close phylogenetic relationship to humans, and, hence, their clinical relevance. Although the development of cell-based therapies can and does benefit from experimentation in rodent models, potential applications of human ESCs in regenerative medicine must be pioneered in NHP species prior to or in parallel with human research for scientific and ethical reasons.
Although more than 100 human ESC lines have been isolated and are either registered by NIH or in the scientific literature [4, , , –8], fewer than 25 NHP ESC lines have been derived. Among these are 16 that originated from blastocysts recovered by nonsurgical uterine flush procedures from rhesus monkeys (Macaca mulatta) [9, 10] or common marmosets (Callithrix jacchus) . In the latter instance, the derivation of three additional ESC lines was reported recently . In the cynomolgus macaque (Macaca fascicularis), four ESC lines have been derived from in vitro produced embryos , and one line has been derived from a parthenogenetic blastocyst . The ESCs derived from humans and NHPs share similar growth, morphologic characteristics, and marker expression compared with each other and marked differences compared with the mouse. Furthermore, although interline differences are beginning to emerge in primate ESC lines, specific properties have not yet been adequately assessed in any species.
We have been interested in rhesus monkey ESC lines, initially with the intent of comparing lines established from in vitro derived versus in vivo derived blastocysts, as the latter represents a gold standard unavailable in the human . Subsequent studies on epigenetic regulation of gene expression in primates and the production of transgenic animals led to additional derivations [16, 17]. Here, we summarize our experience in the establishment of 18 ESC lines from rhesus monkey preimplantation embryos generated by the application of the assisted reproductive technologies at the Oregon National Primate Research Center. The newly derived monkey ESC lines were maintained in vitro without losing their chromosomal integrity, and they expressed markers previously reported present in human and monkey ESCs. We also describe initial efforts to compare the pluripotency of ESC lines by expression profiling, chimeric embryo formation, and in vitro directed differentiation into endodermal, mesodermal, and ectodermal lineages.
Materials and Methods
Ovarian Stimulation, Recovery of Rhesus Macaque Oocytes, Fertilization by Intracytoplasmic Sperm Injection, and Embryo Culture
Controlled ovarian stimulation and oocyte recovery has been described previously . Briefly, cycling females were subjected to follicular stimulation using twice-daily intramuscular injections of recombinant human follicle-stimulating hormone, as well as concurrent treatment with Antide, a gonadotropin-releasing hormone antagonist, for 8–9 days. Females received recombinant human luteinizing hormone on days 7–9 and recombinant human chorionic gonadotropin (hCG) on day 10. Unless indicated otherwise, all hormones and Antide were from Ares Advanced Technologies, Inc. (Norwell, MA, http://www.serono.com), and other reagents were from Sigma-Aldrich (St. Louis, http://www.sigmaaldrich.com). Cumulus-oocyte complexes were collected from anesthetized animals by laparoscopic follicular aspiration (27–29 h post-hCG) and placed in Hepes-buffered TALP (modified Tyrode solution with albumin, lactate, and pyruvate)  containing 0.3% bovine serum albumin (TH3) at 37°C. Tubes containing follicular aspirates were placed in a portable incubator (Minitube, Verona, WI, http://www.minitube.com/main.html) at 37°C for transport to the laboratory. Hyaluronidase (0.5 mg/ml) was added directly to the tubes containing aspirates followed by incubation at 37°C (30 seconds) before the contents were gently agitated with a serological pipette to disaggregate cumulus and granulosa cell masses and sifted through a cell strainer (Falcon, 70-μm mesh size; Becton Dickinson and Company, Franklin Lakes, NJ, http://www.bd.com). Oocytes were retained in the mesh, whereas blood, cumulus, and granulosa cells were sifted through the filter. The strainer was immediately backwashed with TH3, and the medium containing oocytes was collected. Residual cumulus cells were removed with a small-bore pipette (approximately 125 μm in inner diameter), and oocytes were placed in chemically defined, protein-free hamster embryo culture medium (HECM)-9 medium  at 37°C in 6% CO2, 5% O2, and 89% N2 covered with paraffin oil (Ovoil; Zander IVF, Vero Beach, FL, http://www.zanderivf.com). Fertilization by intracytoplasmic sperm injection (ICSI) and embryo culture were performed as described . Briefly, sperm were diluted with 10% polyvinylpyrrolidone (1:4; Irvine Scientific, Santa Ana, CA, http://www.irvinesci.com), and a 5-μl drop was placed in a micromanipulation chamber. A 30-μl drop of TH3 was placed in the same micromanipulation chamber next to the sperm droplet, and both were covered with paraffin oil. The micromanipulation chamber was mounted on an inverted microscope equipped with Hoffman optics and micromanipulators. An individual sperm was immobilized, aspirated into an ICSI pipette (Humagen, Charlottesville, VA, http://www.humagenivf.com), and injected into the cytoplasm of a metaphase II-arrested oocyte, away from the polar body. After ICSI, injected oocytes were placed in 4-well dishes (Nalge Nunc International Co., Naperville, IL, http://www.nalgenunc.com) containing protein-free HECM-9 medium covered with paraffin oil and cultured at 37°C in 6% CO2, 5% O2, and 89% N2. Embryos at the eight-cell stage were transferred to fresh plates of HECM-9 medium supplemented with 5% fetal bovine serum (FBS) (HyClone, Logan, UT, http://www.hyclone.com) and cultured for a maximum of 9 days, with medium change every other day.
ESC Derivation and Culture
Zonae pellucidae of selected expanded blastocysts were removed by brief exposure (45–60 seconds) to 0.5% pronase in TH3 medium. For immunosurgical isolation of inner cell masses (ICMs) , zona-free blastocysts were exposed to rabbit anti-rhesus spleen serum (a gift from Dr. J.A. Thomson) for 30 minutes at 37°C. After extensive washing in TH3, embryos were incubated in guinea pig complement reconstituted with HECM-9 (1:2, vol/vol) for an additional 30 minutes at 37°C. Partially lysed trophectodermal cells were mechanically dispersed by gentle pipetting with a small-bore pipette (125 μm in inner diameter; Stripper pipette; Midatlantic Diagnostics Inc., Marlton, NJ, http://www.midatlanticdiagnostics.com) followed by the rinsing of ICMs three times with TH3 medium. Isolated ICMs were plated onto Nunc four-well dishes containing mitotically inactivated feeder layers consisting of mouse embryonic fibroblasts (mEFs) and cultured in either Dulbecco's modified Eagle's medium (DMEM) with glucose and without sodium pyruvate (Invitrogen, Carlsbad, CA, http://www.invitrogen.com) supplemented with 1% nonessential amino acids (Invitrogen), 2 mM l-glutamine (Invitrogen), 0.1 mM β-mercaptoethanol, and 20% FBS or DMEM/Ham's F-12 medium (DMEM/F12) (Invitrogen) with the same supplements but 15% FBS. ICMs that attached to the feeder layer and initiated outgrowth were manually dissociated into small-cell clumps with a microscalpel and replated onto new mEFs. After the first passage, colonies with ESC-like morphology were selected for further propagation, characterization, and low temperature storage. The medium was changed daily, and ESC colonies were split every 5–7 days manually or by disaggregation in collagenase IV (1 mg/ml, at 37°C for 2–3 minutes; Invitrogen) and replating collected cells onto dishes with fresh feeder layers. Cultures were maintained at 37°C, 3% CO2, and the balance air or 3% CO2, 5% O2, and 92% N2.
Embryoid Body Formation and In Vitro Differentiation of ESCs
For embryoid body (EB) formation, entire ESC colonies were loosely detached from feeder cells and transferred into feeder-free, 6-well, Ultra Low adhesion plates (Corning Costar, Acton, MA, http://www.corning.com/lifesciences) and cultured in suspension in ESC medium for 5–7 days. To induce further differentiation, EBs were transferred into collagen-coated, 6-well culture dishes (Becton Dickinson and Company) to allow attachment. To induce neuronal differentiation, medium was replaced with serum-free DMEM/F12 containing ITS supplement (insulin, transferrin, and sodium selenite; Invitrogen) and fibronectin (5 μg/ml; Invitrogen) . Cultures were maintained for 7 days, with medium replenishment every 2 days. The resulting cultures were disaggregated with collagenase or trypsin treatment and replated onto polyornithine- and laminin-coated plates or glass coverslips in N2 medium consisting of DMEM/F12 supplemented with laminin (1 μg/ml; Invitrogen), basic fibroblast growth factor (bFGF) (10 ng/ml; R&D Systems Inc., Minneapolis, http://www.rndsystems.com), and N2 supplement (Invitrogen). Cultures were maintained for an additional 7 days with daily medium changes. After 7 days, bFGF was omitted from the medium, and cultures were maintained for an additional 7–12 days to induce differentiation into mature neuronal phenotypes. For pancreatic differentiation (C-peptide positive, endodermal lineage), the initial steps were similar to neuronal differentiation. After expanding progenitor cells, bFGF was omitted, and final differentiation was induced by supplementation of the medium with 10 nM exendin-4 and 10 mM nicotinamide (Stem Cell Technologies, Vancouver, BC, Canada, http://www.stemcell.com) . Differentiation into cardiac cells or retinal pigment epithelium was initiated by EB formation in suspension as described above. EBs were then plated into collagen-coated dishes, and cultures were maintained in ESC medium for 2–4 weeks.
Undifferentiated and differentiated ESCs were fixed in 4% paraformaldehyde for 20 minutes. After permeabilization with 0.2% Triton X-100 and 0.1% Tween-20, nonspecific reactions were blocked with 10% normal goat serum (Jackson Immunoresearch Laboratories, West Grove, PA, http://www.jacksonimmuno.com). Cells were then incubated for 40 minutes in primary antibodies, washed three times, and exposed to secondary antibodies conjugated with fluorochromes (Jackson Immunoresearch Laboratories) before costaining with 2 μg/ml 4′,6-diamidino-2-phenylindole for 10 minutes, whole-mounting onto slides, and examination under epifluorescence microscopy.
Primary antibodies were from Santa Cruz Biotechnology Inc. (Santa Cruz, CA, http://www.scbt.com) (OCT-4, stage-specific embryonic antigen [SSEA]-1, SSEA-3, SSEA-4, tumor-related antigen [TRA]-1-60, TRA-1-81, THY-1, NANOG, FOXD3, bestrophin, and cellular retinaldehyde binding protein [CRALBP]), Chemicon (Temecula, CA, http://www.chemicon.com) (C-peptide, neuron-specific nuclear protein [NeuN], microtubule-associated protein [MAP2C], β-III-tubulin [TujIII], glial fibrillary associated protein [GFAP], troponins I and T [cTnI and cTnT], α-myosin heavy chain protein [α-MHC], slow tonic myosin heavy chain protein [sMHC], sarcoplasmic reticular Ca2+-ATPase [SERCA2], atrial natriuretic peptide [ANP], tropomyosin, α-actinin, myosin light chain 2A and 2V [MLC-2V and MLC-2A], and cardiac transcription factors GATA-4 and myocyte enhancer factor 2), ImmunoStar, Inc. (Hudson, WI, http://www.immunostar.com; serotonin), and R&D Systems Inc. (nestin). Musashi1 was kindly provided by Dr. H. Okano (Keio University, Tokyo, Japan).
Reverse Transcription-Polymerase Chain Reaction
Total RNA was extracted from ESCs and ESC-derived differentiated phenotypes using RNA purification kit (Invitrogen) according to the manufacturer's instructions. Total RNA was treated with DNase I before cDNA preparation using SuperScript III First-Strand Synthesis System for reverse transcription-polymerase chain reaction (RT-PCR) (Invitrogen) according to the manufacturer's instructions. The first strand cDNA was further amplified by PCR using individual primer pairs for specific genes. The sequence, annealing temperature, and cycle number of each pair of primers are listed in supplemental online Table 1. All PCR samples were analyzed by electrophoresis on 2% agarose gel containing 0.5 μg/ml ethidium bromide.
Table Table 1.. Transcriptome analysis of rhesus monkey ESC lines
Total RNA was isolated as indicated above. Labeling, hybridization, and scanning was performed according to standard Affymetrix protocols (more details are given in Affymetrix GeneChip Expression Analysis Technical Manual, http://www.affymetrix.com). A microarray analysis was performed on a rhesus macaque Affymetrix GeneChip with 52,865 probe sets representing over 20,000 genes (protocols and Minimum Information About Microarray Experiment information are given in supplemental online data). The normalized microarray data were further analyzed using GeneChip Operating System (GCOS) 1.2. MAS-5 statistical analysis was performed to calculate the signal log ratio (SLR) for each probe set to determine the percentage of the transcriptome that significantly varied (p < .002) between compared samples. Gene expression fold changes (FCs) between two samples were calculated from the SLR using the formula FC = (2SLR). All normalized microarray data used in this research can be found in supplemental online data (MicroarrayData1.xls and MicroarrayData2.xls).
Mitotically active ESCs in log phase were incubated with 120 ng/ml ethidium bromide for 40 minutes at 37°C, 5% CO2, followed by 120 ng/ml colcemid (Invitrogen) treatment for 20–40 minutes. Cells were then dislodged with 0.25% trypsin and centrifuged at 200g for 8 minutes. The cell pellet was gently resuspended in 0.075 M KCl solution and incubated for 20 minutes at 37°C, followed by fixation with methanol:glacial acetic acid (3:1) solution. Fixed cells were dropped on wet slides, air-dried, and baked at 90°C for 1 hour. G banding was performed using trypsin-EDTA and Leishman stain (GTL) by immersing slides in 1× trypsin-EDTA with two drops of 0.4 M Na2HPO4 for 20–30 seconds. Slides were then rinsed in distilled water and stained with Leishman stain for 1.5 minutes, rinsed in distilled water again, and air-dried. For GTL-banding analysis, 20 metaphases were fully karyotyped under an Olympus BX40 microscope equipped with ×10 and ×100 plan-apo objectives. Images were then captured, and cells were karyotyped using a CytoVysion digital imaging system (Applied Imaging Corporation, San Jose, CA, http://www.aicorp.com).
Chimeric Embryo Production
Disaggregated ESC clumps (2–3 cells per clump) were labeled with the membrane-specific red fluorescent dye PKH-26, according to the manufacturer's directions, to identify and track ESCs within chimeric embryos. Cleavage stage (4–8-cell) rhesus monkey embryos were produced by ICSI as described above. A total of 15–25 labeled cells was injected into the perivitelline space of recipient embryos to ensure close contact with blastomeres. Injected embryos were cultured to the blastocyst stage, and the incorporation of cells into the chimeric embryo was monitored by epifluorescence microscopy every other day throughout preimplantation development. Cell numbers in blastocysts were quantitated by confocal microscopy.
Derivation and Propagation of Novel Rhesus Monkey ESC Lines
Expanded blastocysts with good morphology containing distinct, prominent ICMs were selected for ESC line derivation experiments. The majority (approximately 70%) of individually plated ICMs usually attached to the feeder layer within 2–3 days and initiated three-dimensional outgrowths consisting of tightly packed cells. After 5–7 days of culture, outgrowths were manually excised from the plates, dissociated into 5–10 smaller clumps with a microscalpel, and replated onto fresh mEFs. Approximately 50% of the replated ICMs generated one or two ESC-like colonies within 3–7 days, consisting of cells with a high nuclear to cytoplasmic ratio and prominent nucleoli (Fig. 1A). These colonies were isolated and passaged onto fresh feeders manually or with collagenase treatment for an additional 5–10 passages to generate sufficient cell numbers for cryopreservation, characterization, karyotyping, and in vitro differentiation studies. Eighteen ESC lines (designated Oregon rhesus monkey embryonic stem 1–18 [ORMES-1 to -18]) were isolated in 14 separate experiments conducted between 2001 and 2004 from 94 blastocysts with an overall efficiency of 27% ± 6%. Increased blastocyst age contributed to a progressively increasing efficiency in ESC line derivation (14% ± 0%, 19% ± 7%, 26% ± 7%, and 41% ± 20% for day 6, 7, 8, and 9 blastocysts, respectively; p > .05; one-way analysis of variance and Fisher's protected least significant difference test), perhaps reflecting the increased cell numbers in the ICMs of older blastocysts, although other differences in the derivation conditions also existed. For example, the first six ORMES cell lines were isolated and cultured in high-glucose DMEM without sodium pyruvate supplemented with l-glutamine, nonessential amino acids, β-mercaptoethanol, and 20% FBS, as originally described for monkey and human ESC culture [4, 9]. In an effort to improve ESC culture conditions, DMEM/F12 containing sodium pyruvate and several additional amino acids and vitamins was evaluated . DMEM/F12 supported faster growth rates (passage interval, 5–7 days) than DMEM (8–10 days), even though the FBS supplementation in DMEM/F12 was lower than that of DMEM. The efficiency of ESC line isolation was also significantly higher (p < .05) when conducted in DMEM/F12 (39% ± 10%) compared with DMEM (12% ± 1%) (supplemental online Table 2).
Table Table 2.. Genes with the greatest average fold change in monkey ESCs compared with the somatic cell control
Although DMEM/F12 was associated with higher derivation efficiency and improved ESC growth properties, pH stability was problematic under 5% CO2 conditions. Maintenance of medium pH in the optimal 7.2–7.4 range was achieved, however, when the CO2 level was reduced to 3%. Recognizing that optimal growth and development of preimplantation stage monkey embryos from which ESCs are derived is achieved under reduced oxygen levels , culture in 5% O2 was evaluated. Cells grew at a rate similar to controls; however, a lower incidence of spontaneous differentiation was noted (results not shown). Notable differences in growth rates were observed between ORMES cell lines with passage intervals ranging between 5 and 7 days for fast-growing (ORMES-6, -7, and -13) and between 14 and 21 days for slow-growing lines (ORMES-4 and -12) under comparable conditions. The latter were also characterized by poor survival after conventional freezing. The slow growth rate of ORMES-4 was evident during initial ESC line establishment steps and was maintained at later passages. In contrast, the initial slow growth rate of ORMES-12 was overcome during extensive propagation.
Monkey ESCs are extremely sensitive biological materials, and reproducible propagation of the undifferentiated phenotype is associated with rigorous optimization of culture conditions and protocols. Quality control of media and media components, plasticware, and equipment generally involved with ESC culture was critically important in trouble-free maintenance of ESCs. For instance, toxic effects of several brands or lots of culture dishes were observed uniquely with monkey ESCs (results not shown), as other cell types such as fibroblasts did not display negative effects. ORMES cell lines were also sensitive to mEF quality. Feeder plates prepared from early passage mEFs (passages 2–3) supported better undifferentiated cell growth than those prepared from later passages (passage 5–10). We also found that monkey ESCs were sensitive to FBS preparations, necessitating that each new lot be tested for the ability to support ESC growth before purchase. In contrast to human ESCs , we found no noticeable effect of bFGF on rhesus monkey ESC growth. When growth medium was supplemented with 4 ng/ml bFGF, undifferentiated growth on mEFs and passage intervals of several tested ORMES cell lines were similar to controls. Several ORMES cell lines were adapted to grow in medium supplemented with 20% knockout serum replacement instead of FBS, resulting in slightly slower growth rates and minor changes in morphology.
In Vitro Characterization of Monkey ESC Lines
All established ESC lines expressed key pluripotent markers, including OCT-4, SSEA-3, SSEA-4, TRA-1-60, TRA-1-81, and alkaline phosphatase as detected by immunocytochemistry (ICC) or immunohistochemistry (Fig. 1B). In addition, monkey ESCs also reacted to antibodies against other pluripotency-related products, including NANOG, the forkhead transcriptional regulator FOXD3, and phosphatidyl-anchored cell surface glycoprotein THY-1. SSEA-1 was not expressed in these lines (results not shown). RT-PCR analysis was used to further assess expression of genes characteristic of pluripotent cells, including POU5F1 (OCT4), NANOG, FOXD3, DPPA5, CRIPTO, acidic zinc finger gene REX-1, transcription factor SOX-2, growth factors FGFR-4 and LEFTYA, telomerase-related TERF1, TERF2, and TERT, ABC transporter BCRP-1/ABCG-2, and the gap junctional components CONNEXINS 43 and 45. All ORMES cell lines expressed these genes, with the exception of the REX-1 gene (supplemental online Fig. 1). RT-PCR with four previously published primers  and one set of primers based on the human REX-1 gene sequence failed to amplify any product in monkey ESCs. Overall, our results demonstrate that the ORMES cell lines express markers previously reported present in mouse (except SSEA-1), human, and monkey ESCs.
To further investigate the transcriptional activity of monkey ESCs and begin defining “stemness” genes, we conducted genome-wide expression profiling of several ORMES cell lines. Initially, two samples from a single ESC line (ORMES-6) at passage 41 were collected from the same culture dish (ORMES-6A and ORMES-6B) and compared with each other and with a culture of adult rhesus monkey skin fibroblasts. Only minimal variation (1.2%) was observed between the two ORMES-6 samples. However, significant variation was observed when each of these samples was compared with the somatic cell control (25.4% and 26.1%, respectively).
Next, the transcriptomes of five ORMES cell lines were compared with each other using GCOS and to adult rhesus monkey skin fibroblasts (Table 1). The transcriptome of each ORMES cell line was consistently different from the calculated average by approximately 11%, a value that was remarkably consistent. The transcriptome of each ORMES cell line was different from the control somatic cell line by 25%–29%. The percentage of the transcriptome with a change of more than twofold was only ∼4% for each ORMES cell line compared with the ESC average, but it was still over 17%–18% compared with the somatic control. This filtered result suggests that the majority of the differences observed between ORMES cell lines were relatively small. Despite differences, all analyzed ORMES cell lines consistently and strongly expressed a subset of stem cell marker genes. We calculated the average fold change for each probe set and then selected 25 ontologically identifiable genes with the greatest average fold change compared with the somatic cell control (Table 2).
Several stem cell markers were highly expressed in all five ORMES cell lines, as expected, including NANOG, LIN-28, PODXL, POU5F1, and GDF-3 (Table 2 in bold). Subsequent semiquantitative RT-PCR analysis was used to validate expression of NANOG and POU5F1 in ORMES cell lines (results not shown).
Detailed G-banding analysis of ORMES cell lines revealed that 15 were karyotypically normal, with a diploid set of 42 chromosomes. The anticipated even distribution of male (n = 9) and female (n = 9) lines was observed (Table 3). ORMES-1 carried a stable, balanced 11;16 translocation, and ORMES-2 had balanced 5;19 and 1;18 translocations based on the presence of these abnormalities at early and subsequent passage numbers. ORMES-5 demonstrated a pericentric inversion involving chromosome 1.
Table Table 3.. Derivation conditions and karyotype of rhesus monkey ESC lines 1–18; all lines were cultured on mEFs
ESC Line Pluripotency
To determine whether newly established ESC lines can give rise to cell lineages representative of all three embryonic germ layers, we applied in vitro approaches designed to induce differentiation into neural, retinal (ectodermal), cardiac (mesodermal), and pancreatic (endodermal) lineages via EB formation.
Mesodermal differentiation of several ORMES cell lines involved EB production in suspension culture for 5–7 days followed by plating them onto collagen-coated dishes for adherent culture. Approximately 7–14 days after plating, attachment, and further spontaneous differentiation, contracting cell aggregates were observed in all tested ORMES cell lines (Table 4). Analysis of these aggregates by ICC revealed expression of cardiac-specific markers, including cTnI and cTnT, α-MHC, sMHC, SERCA2, ANP, tropomyosin, α-actinin, MLC-2V, MLC-2A, and cardiac transcription factors GATA-4 and myocyte enhancer factor 2 (supplemental online Fig. 2a–2d) [26, 27].
Table Table 4.. In vitro differentiation of ORMES cell lines
Differentiation into neuronal phenotypes was induced using a stepwise approach originally developed for mouse ESCs and adapted by us for primate cells [16, 23]. Following selection and expansion of progenitor cell populations in the serum-free N2 medium supplemented with bFGF for 2 weeks, immunostaining revealed that >90% of cells were nestin+ and Musashi1+ (Fig. 1C, a). OCT-4 expression was not detected at this stage. For further differentiation, nestin+/Musashi1+ progenitor cells were plated onto polyornithine/laminin-coated surfaces for adherent culture and maintained in N2 medium without bFGF for an additional 2–8 weeks. Withdrawal of bFGF induced robust differentiation into various neuronal phenotypes demonstrating small-cell bodies and long axon-like extensions (Fig. 1C, b). ICC analysis showed that the vast majority of the cells expressed the neuronal markers NeuN, MAP2C, serotonin, and TujIII (Fig. 1C, c) [23, 28]. Between 10% and 30% of cells in the differentiated population expressed the glial cell marker GFAP.
Differentiation into dark pigmented epithelial cells resembling retinal pigment epithelial (RPE) was observed during EB formation in suspension culture for 5–7 days followed by plating EBs onto collagen-coated dishes to allow attachment and adhesion as described above for cardiac differentiation. After approximately 2–3 weeks of adherent culture, clusters of darkly pigmented cells were observed (Fig. 1C, d) that grew in size and eventually become visible to the naked eye at a density of 2–3 clusters per 35-mm plate. Pigmented epithelial cells appeared polygonally shaped, similar to those observed in cultured retinal cells derived from human ESCs (Fig. 1C, e) . These clusters were manually excised, replated onto new dishes, and subcultured for an additional 5–6 months. As colonies of pigmented cells expanded, the cells at the periphery become less pigmented; however, once confluence was reached, some islands become strongly pigmented again. Pigmented and nonpigmented epithelial cells coexisted with other, unidentified nonepithelial cells (Fig. 1C, f). At confluence, cultures became multilayered, with pigmented or nonpigmented epithelial cells in the top layers, and highly organized, nonepithelial cells growing underneath (Fig. 1C, g). To further define the phenotype of these cells, we performed ICC analysis for proteins characteristic of RPE cells including CRALBP and bestrophin. CRALBP is involved in the regeneration of visual pigment and abundant in the RPE and Muller cells of neuroretina but is not expressed in photoreceptors. Bestrophin is considered a very late marker of RPE differentiation during normal development and is localized to the basolateral plasma membrane of the RPE . Both pigmented and nonpigmented epithelial cells in these cultures expressed CRALBP (Fig. 1C, h), whereas bestrophin expression was correlated with the level of pigmentation.
Pancreatic β-Cell Differentiation
Populations of progenitor cells expressing nestin were subjected to protocols using exendin-4 and nicotinamide to redirect the differentiation into endocrine phenotypes . Identification of β-cell phenotypes was performed by ICC detection of C-peptide, a cleavage byproduct of insulin biosynthesis from proinsulin. We have consistently produced C-peptide-positive cells at efficiency of 70% using this protocol with various monkey ESC lines (Table 4) (supplemental online Fig. 2e) [16, 24].
In an effort to further define the potency of monkey ESCs, we documented the participation of ORMES-1 cells in the preimplantation development of chimeric monkey embryos. Approximately 10–15 ESCs labeled with PKH-26 were injected into monkey embryos at the 4–8-cell stage produced by ICSI, and the resulting chimeric embryos were cultured in vitro to the blastocyst stage. In five replicates involving 29 embryos, development progressed to the compact morula (19 of 29; 65%) and blastocyst (12 of 29; 41%) stages at rates comparable to nonmanipulated control counterparts. Incorporation of fluorescent cells was observed in all chimeric embryos throughout the preimplantation development period up to the day 8 expanded blastocyst stage with evidence of integration into both the trophectoderm and ICM (Fig. 2). As determined by confocal microscopy, total cells numbers in chimeric (n = 8) and noninjected control (n = 6) blastocysts were not significantly different (527 ± 45 [SEM] and 401 ± 41 [SEM], respectively; p > .07; Mann-Whitney test). An ESC contribution to chimeric embryos was evidenced by the presence of 49 ± 7 (SEM) fluorescent cells. One clinical pregnancy was established following the transfer of 15 chimeric embryos at various preimplantation stages into five recipients that aborted at 5 weeks.
ESC line banking efforts provide several insights into factors affecting the isolation and propagation of primate ESC lines. Although the average efficiency of derivation (27 ± 6) was comparable to the reported experience with human and nonhuman primate ESCs [7, 12, 13], we found success to be dependent on the culture media employed and blastocyst age. The latter may simply reflect the size of the ICM and hence the number of ICM cells. Interestingly, this isolation rate was higher than implantation rates experienced with ICSI-produced embryos at 10%–20% . Unquestionably, embryo quantity is another factor in the equation for success.
ICC analysis confirmed expression of well-defined pluripotent markers in these novel monkey ESC lines, corroborated by PCR analysis. These genes include POU5F1, NANOG, TERT, FOXD3, SOX2, TERF1, TERF2, and LEFTYA; however, in contrast to mouse ESCs, REX-1 gene expression was not detected in undifferentiated ORMES cell lines using several primer sets designed for the human sequence. Microarray expression data confirmed the absence of REX-1 transcripts in ORMES cell lines, and it is worth noting the inconsistency in REX-1 expression among several human ESC lines, suggesting that expression of this gene is not essential for self-renewal of primate ESCs .
Another characteristic of ESCs is their ability to maintain normal karyotype during prolonged culture in vitro. Cytogenetic analysis of ORMES cell lines revealed that the majority of lines maintained a normal karyotype with a diploid set of 42 chromosomes. Both male (n = 9) and female (n = 9) cell lines were equally represented, in agreement with the conclusion that primate pluripotent cell lines from both sexes can be efficiently derived and maintained in vitro . In contrast, female mouse ESC lines are unstable during culture and extended in vitro propagation, resulting in loss of an inactive X chromosome . The establishment of stable XX mouse ESC lines is associated with global DNA hypomethylation of both repetitive and differentially methylated regions that regulate expression of parentally imprinted genes . Parenthetically, we have shown that several ORMES cell lines display aberrant imprinting in the H19/IGF2 domain . Three ORMES cell lines displayed chromosomal abnormalities in the form of balanced translocations and a pericentric inversion. These rearrangements appeared to be stable, since they were detected at both early and late passage cells. Moreover, the directed differentiation of ORMES-1 appeared to be comparable to other ORMES cell lines. Interestingly, parthenogenetically derived ESCs in the cynomolgus macaque also seem to be multipotent . In vitro propagation of human ESCs may give rise to abnormal karyotypes; specifically, trisomy of chromosomes 17q and 12 has been observed [35, –37]. Notably, these changes were associated with exposure to an extended feeder-free culture system on Matrigel or to enzymatic passaging using collagenase, trypsin, or cell-dissociation buffer. Thus, it is possible that the collagenase-based splitting technique used for ORMES-1 to -6 may have contributed to the karyotypic abnormalities in ORMES-1, -2, and -5.
Despite close similarities in the morphology, marker expression properties, and karyotypes of ORMES cell lines, important differences are also apparent. Because of the implications of this observation to the long-term objective of cell-based therapeutic applications, we have begun a comparison of monkey ESC lines. Differences in growth rates and survival following cryopreservation were noted during the establishment of some ORMES cell lines that persisted during more extensive propagation. In contrast, growth rates of other cell lines changed over time at later passages, perhaps reflecting adaptation to culture conditions, inadvertent selection of a different phenotype, or even genetic and/or epigenetic changes.
With regard to the gene expression profile, ORMES cell lines exhibited similar but not identical transcriptomes. Expression profiling by rhesus monkey Affymetrix oligonucleotide array revealed that 11%–12% of the transcriptomes from the five different ORMES cell lines demonstrated significant changes in gene expression compared with the pooled ESC average (a shared significant homology in gene expression of ∼88%–89%). Approximately 4% of the compared transcriptomes demonstrating a fold change greater than two (Table 1). The majority of this transcriptional variation must be biologically significant, as we observed only 1.2% transcriptional variation between biological replicates that was likely introduced during the RNA purification and microarray analysis process (RNA amplification, cDNA processing, washing, scanning, etc.). The shared significant homology in gene expression between the five different rhesus ESC lines examined at ∼88%–89% is comparable with microarray comparisons between human ESC lines, where the shared significant homology in gene expression is ∼85% . Despite the observed transcriptome differences between ESC lines, all of the ORMES cell lines examined strongly expressed key markers necessary to maintain a stem cell state, including POU5F1, NANOG, LIN-28, PODXL, and GDF-3 (Table 2). RT-PCR analysis confirms that NANOG and POU5F1 are present only in undifferentiated and not in differentiated ORMES cells. This suggests that it may not be the entire transcriptome that dictates the embryonic stem cell state, but only a subset of key stemness genes. Perhaps as long as key stemness genes, such as NANOG, LIN-28, PODXL, POU5F1, and GDF-3, are strongly expressed, the ESC transcriptomes possess a degree of biological plasticity that can accommodate a limited degree of transcriptional variation while still maintaining an undifferentiated embryonic stem cell state.
The ultimate measure of pluripotency is demonstrating ESC participation in chimeric animals, a well-defined and frequently exercised option in mice . The pluripotency of human ESCs is presumed by expression of specific markers and the ability to differentiate into various somatic tissues in vitro or in vivo by teratoma formation in SCID mouse. However, the creation of chimeric human embryos is affected by bioethical concerns. Since rhesus monkey ESCs share a number of properties with human ESCs, they serve as a primate model for in vivo studies involving chimeric embryos. Here, the fluorescent lipophilic carbocyanine dye PKH-26 was used as a harmless, noninvasive cell lineage marker . Our results demonstrated that chimeric blastocysts could be produced readily in the rhesus monkey by ESC injection, setting the stage for future potency determination in chimeric fetuses or offspring. Although the fluorescent signal from PKH-26 made it impossible to trace ESC fate in chimeric fetuses or offspring during long-term studies, creating genetically modified ESC lines carrying reporter genes, such as eGFP, would appear feasible given adequate resources.
In conclusion, we have summarized our experience in generating 18 new ESC lines from rhesus monkey blastocysts generated in vitro. This bank of ESC lines, augmented by in vivo-derived lines  that can serve as a gold standard, represents a valuable resource. Future plans include continued interline comparative studies and creating additional MHC-typed and xeno-free monkey ESC lines in anticipation of transplantation trials.
D.W. has acted as a consultant for the NIH within the last 2 years.
We acknowledge the Division of Animal Resources, the Endocrine Services Core, and the Molecular Biology Core at the Oregon National Primate Research Center and Affymetrix Microarray Core at the Oregon Health and Science University for their assistance and technical services. We are grateful to Cathy Ramsey, Christine Gagliardi, Carrie Greenberg, Michelle Sparman, and Ching-Yu Chuang for technical assistance; Dr. John Fanton and Darla Jacobs for laparoscopic oocyte retrieval; Julianne White for administrative support; and Joel Ito for help with illustration materials. The Assisted Reproductive Technologies/ESC core facility assisted by providing semen samples, media, and mEF feeder layers for ESCs. Rabbit anti-rhesus spleen whole serum was developed by and obtained from Dr. J.A. Thomson, Wisconsin National Primate Research Center, University of Wisconsin-Madison. This study was supported by NIH Grants RR00163 (Dr. D. Dorsa, Oregon Health & Science University), HD18185 (Dr. R.L. Stouffer, Oregon Health & Science University), and RR15199 (D.W.) and by an Academia Sinica of Taiwan grant (H.-C.K.).