Fetal mesenchymal stem cells derived from human umbilical cord sustain primitive characteristics during extensive expansion
Stem cells of fetal origin lie between embryonic and adult stem cells in terms of potentiality. Because of the ethical controversy surrounding embryonic stem cells and the relatively inferior quality of adult stem cells, the use of fetal stem cells would be an attractive option in future therapeutic applications. Here, we have investigated primitive characteristics of human umbilical-cord-derived fetal mesenchymal stem cells (UC fMSCs) during extensive expansion. We have successfully isolated and cultured UC fMSCs from all UC samples, but with two early fungal contaminations. UC fMSCs proliferated without significant evidence of morphological changes, and the average cumulative population-doubling level was over 25 for about 3 months. UC fMSCs showed the positive expression of several CD markers, known to be related to MSCs, including CD73 (SH-3, 4), CD90 (Thy-1), CD105 (SH-2), CD117 (c-kit), and CD166 (ALCAM). They demonstrated primitive properties throughout the expansion period: multilineage differentiation potentials examined by functional assays, a variety of pluripotent stem cell markers including Nanog, Oct-4, Sox-2, Rex-1, SSEA-3, SSEA-4, Tra-1–60, and Tra-1–81, minimal evidence of senescence as shown by β-galactosidase staining, and the consistent expression of telomerase activity. These results suggest that UC fMSCs have more primitive properties than adult MSCs, which might make them a useful source of MSCs for clinical applications.
KeywordsUmbilical cord Mesenchymal stem cells Pluripotent markers Senescence Telomerase Human
Mesenchymal stem cells (MSCs) are characterized by their capacity for self-renewal and the production of multiple lineages (Pittenger et al. 1999). Whereas MSCs were first isolated from bone marrow (Friedenstein et al. 1970), recent studies have demonstrated that MSCs exist in many other tissues, e.g., skeletal muscle (Deasy et al. 2001), fat (Zuk et al. 2001), synovial membrane (Jo et al. 2007), and hepatic tissues (Alison and Sarraf 1998). The selection of an MSC source for clinical use must reflect its balance of pluripotency, ease of isolation, and the ethical issues surrounding its harvest. Embryonic stem cells offer higher pluripotency but remain problematic in clinical use because of ethical issues. In contrast, MSCs harvested from adult organisms are ethically uncomplicated and readily available. However, the harvesting of these cells requires invasive procedures, and adult MSCs have poor quality when compared with embryonic stem cells in several aspects, such as frequency, expansion potential, proliferative rate, and differentiation potential, all of which decrease significantly with age (D’ippolito et al. 1999; Rao and Matton 2001; Mendes et al. 2002; Baxter et al. 2004; Mareschi et al. 2006; Roura et al. 2006).
Stem cells with fetal origins have the potential to offer the ideal balance between quality and ethics. Fetal stem cells isolated from fetal tissue, such as umbilical cord (UC), umbilical cord blood (UCB), or placenta, may lie between embryonic stem cells and adult MSCs with respect to their quantity and quality (Pojda et al. 2005). The frequency of stem cells obtained from fetal sources is smaller than that from embryonic tissues, but the absolute number is greater than that from embryonic tissues (Guillot et al. 2006; Lu et al. 2006). Various characteristics of fetal stem cell quality, such as multilineage differentiation potential, telomere length, and telomerase activity, are inferior to those of embryonic stem cells but are again superior to those of adult stem cells (Izadpanah et al. 2006; Guillot et al. 2007). One of the most attractive aspects of fetal stem cells is that their harvest does not require the destruction of embryos, since the source of the stem cells is considered waste material.
UC itself contains fetal MSCs (UC fMSCs) within Wharton’s jelly (Wang et al. 2004), the perivascular mesenchymal area (Sarugaser et al. 2005), umbilical vein, and subendothelial tissues (Romanov et al. 2003). MSCs isolated from these substructures can be differentiated into adipogenic, osteogenic, chondrogenic, and cardiomyogenic lineages (Wang et al. 2004), and dopaminergic neurons (Fu et al. 2006; Weiss et al. 2006). Recently, a few authors have shown that UC fMSCs possess some superior characteristics to adult MSCs, such as the higher frequency of colony-forming-unit fibroblasts (CFU-F) and shorter population doubling time (PDT; Lu et al. 2006). Some other investigators have demonstrated that UC-derived cells express a few pluripotent stem cell markers including Oct-4, Sox-2, and Nanog in cells from porcine sources (Carlin et al. 2006), Oct-4, SSEA-4, and c-Kit in cells from equine sources (Hoynowski et al. 2007), and SSEA-1, SSEA-4, Tra-1–60, and Tra-1–81 in cells from human sources (Fong et al. 2007), suggesting that they might be more immature than adult MSCs. However, evidence is still insufficient to confirm the primitive characteristics of UC-derived cells. The purpose of this study has been to improve the characterization of UC fMSCs, especially in terms of primitive characteristics and throughout the mid-term expansion period. We have performed growth kinetics studies, surface phenotyping, functional analyses of multilineage differentiation, an investigation of a variety of pluripotent stem cell markers expression at the level of mRNA and protein, and senescence and telomerase activity analysis with respect to increasing passage number.
Materials and methods
Isolation and expansion of cells from human UCs
UCs (n=32) were obtained from full-term delivery patients with informed consent. The Institutional Ethical Review Board approved tissue collection for this research. UCs were washed twice in calcium- and magnesium-free phosphate-buffered saline (DPBS) and were finely minced into 1– to 2-mm fragments by using scissors and scalpels. Cells were released by treating with 0.1% collagenase for 2 h and then 0.25% trypsin for 1 h at 37°C in low-glucose Dulbecco’s modified Eagle medium (LG DMEM) containing antibiotic-antimycotic solution (100 U/ml penicillin, 100 μg/ml streptomycin, and 0.25 μg/ml amphotericin B) with gentle agitation. After the addition of the same volume of DPBS, undigested tissues were removed by using a 100-μm nylon sieve. Cells were collected by centrifugation, washed twice, resuspended in LG DMEM supplemented with 10% fetal bovine serum (FBS), 10 ng/ml basic fibroblast growth factor (bFGF), and antibiotic-antimycotic solution, and plated in 100-mm tissue culture dishes at a density of 1×104 cells/cm2 for expansion at 37°C in a humidified 5% CO2 atmosphere, for 3–4 days to allow cell to adhere. The medium was replaced twice a week. Non-adherent cells were removed by medium changes. When cells reached to 60%–80% confluence, they were detached by incubation for 10 min with trypsin-EDTA (0.25% trypsin, 0.53 mM EDTA; Life Technologies), washed, and then replated at the same density.
Growth kinetics studies
Growth kinetics was analyzed by calculating cumulative population-doubling level (CPDL) and population-doubling time (PDT). To investigate CPDL, cells from UCs (n=20) were counted at each passage during expansion by using the trypan blue excluding method for 3 months. To calculate PDT according to passage, cells from UCs (n=7) at passage 2 (P2), P5, P7, and P9 were plated at a density of 1×103 cells/cm2 and counted every other day for 2 weeks. PDT was calculated by the following formulae: PD=Log(Nf/Ni)/Log2, PDT = CT/PD, where PD is the population-doubling time, CT is cell culture time, Nf is the final number of cells, and Ni is the initial number of cells (Vidal et al. 2006).
CFU-F assays (n=9) were performed by using a previously described method with slight modification (Jo et al. 2007). Briefly, cells were collected after enzymatic digestion, washed twice with DPBS, plated at a density of 1×102 cells/cm2 in 100-mm dishes, and cultured for 14 days. On day 14, the cells were washed twice with DPBS and fixed in 1% paraformaldehyde in DPBS for 20 min, stained with 0.1% crystal violet (in 1% formaldehyde solution) for 1 h, and then rinsed in tap water. Aggregates of 50 cells or more with a fibroblast phenotype were defined as CFU-Fs.
A total of 31 antibodies were used in flow-cytometry (FCM): CD9, CD10, CD11a, CD13, CD14, CD29, CD31, CD34, CD40, CD44, CD45, CD49a, CD49b, CD54, CD56, CD62e, CD66, CD68, CD71, CD73, CD90 (Becton Dickinson), CD105 (Abcam), CD106, CD117, CD120a, CD133, CD166, CD235a, HLA-ABC, HLA-DR (Becton Dickson), and STRO-1 (Developmental Studies Hybridoma Bank). After cells had been detached, aliquots of 2×105 cells at P2, P5, P7, and P9 were washed twice with DPBS, centrifuged, washed in ice-cold DPBS supplemented with 1% bovine serum albumin (FCM buffer), and fixed in 2% paraformaldehyde in FCM buffer, followed by incubation with fluorescein isothiocyanate (FITC)- or phycoerythrin (PE)-conjugated antibodies for 15 min on ice in a dark room. Data were obtained by analyzing 10,000 events on a Becton Dickinson FACSAria with FACSDiva software (Becton Dickinson, San Jose, Calif., USA). All analyses were standardized against negative control cells incubated with isotype-specific IgGs.
Differentiation was induced according to established protocols (Karahuseyinoglu et al. 2007; Secco et al. 2008). All experiments were performed in triplicate with controls (n=3) cultured without induction. Oil red O (ORO) assay, calcium deposition assay, alkaline phosphatase (ALP) activity assay, and glycosaminoglycan (GAG) assay was performed with cells at P2, P5, P7, and P9 (n=3 in all passages).
For adipogenic differentiation, subconfluent cells were incubated with adipogenic differentiation medium consisting of LG DMEM supplemented with 10% FBS, 1 μM dexamethasone, 500 μM 3-isobutyl-1-methylxantine, 60 μM indomethacin, and 5 μg/ml insulin. After 3 weeks, cells were fixed in 10% formalin for 10 min at room temperature, washed, and stained with a working solution of 0.18% ORO for 10 min. For quantification of adipogenic differentiation over passage, ORO assay was performed by using an intracellular lipid extraction solution (Cayman Chemical Company, Ann Arbor, Mich., USA) according to the manufacturer’s instruction. Briefly, 500 μl extraction solution was added to extract dye for 30 min, and its absorbance was measured spectrophotometrically at 540 nm and compared with the ORO standard titration curve by using triglyceride to determine lipid contents. The measured lipid contents were normalized to total DNA amount. DNA quantification was carried out by using the PicoGreen assay (Molecular Probes, Eugene, Ore., USA) on cells cultured in parallel with induced or control cells.
For osteogenic differentiation, subconfluent cells were treated for 3 weeks with osteogenic differentiation medium (LG DMEM supplemented with 10% FBS, 100 nM dexamethasone, 200 μM ascorbic acid 2-phosphate, and 10 mM β-glycerophosphate). Von Kossa staining was performed for the visualization of calcium deposition. Cells were fixed with 10% formalin for 10 min at room temperature, washed, stained with 5% silver nitrate for 1 h under a 60-W lamp, followed by 5% sodium thiosulfate for 5 min. The amount of calcium deposition was determined by using the ortho-cresolphthalein complexone (OCPC) method as previously described with slight modification (Jaiswal et al. 1997). Briefly, 1×105 cells in 6-well plate were washed twice in DPBS to which 500 μM 0.5 N acetic acid had been added and were incubated over night. Samples were stored at -20°C until used. OCPC solution was prepared as follows; 60 mg OCPC was added to 75 ml demineralized H2O with 0.5 ml KOH (1 M) and 0.5 ml 0.5 N acetic acid. Sample solution was prepared as a 5-ml OCPC solution to which 5 ml 14.8 M ethanolamine/boric acid buffer (pH 11), 2 ml 8-hydroxyquinoline (5 g in 100 ml 95% ethanol), and 88 ml demineralized water were added. For every 10 μl sample, 300 μl sample solution was added. A serial dilution of CaCl2 was set up (1–200 μg/ml) to generate a standard curve. Plates were incubated at room temperature for 10 min and then read at 575 nm. The amount of calcium deposition was displayed as micrograms of calcium per DNA amount. ALP activity assay was performed as previously described with slight modification (Yoshimura et al. 2007). Briefly, cells were harvested with lysis buffer (0.1 M TRIS-HCL, 5 mM MgCl2, 2% Triton X-100, and 1 mM phenylmethylsulfonyl fluoride). An aliquot of 10 μl supernatant was added to 100 μl 50 mM p-nitrophenylphosphatase hexahydrate (p-NPP) containing 1 mM MgCl2, and the mixture was incubated at 37°C for 30 min. Absorbance at 405 nm was measured with a spectrophotometer. Relative ALP activity was normalized to DNA amount and represented as the amount of p-NPP produced after 30 min of reaction per DNA amount.
For chondrogenic differentiation, 2.5×105 cells were detached, washed, and resuspended in chondrogenic medium (high-glucose DMEM with 100 nM dexamethasone, 10 ng/ml transforming growth factor-β1, 50 μg/ml ascorbic acid 2-phosphate, 100 μg/ml sodium pyruvate, 40 μg/ml proline, and ITS + premix [6.25 μg/ml insulin, 6.25 μg/ml transferrin, 6.25 μg/ml selenous acid, 5.35μg/ml linoleic acid, 1.25 mg/ml bovine serum albumin)]. Aliquots of 2×105 cells were placed in 15-ml polypropylene conical tubes, centrifuged at 600g for 5 min to form a pellet, and cultured, with caps loosened, for 3 weeks. A total of nine pellets per donor, viz., three for histology, three for reverse transcription with polymerase chain reaction (RT-PCR), and three for GAG assay, were cultured. After 3 weeks, pellets were harvested, fixed in 10% formalin, and washed, and cryosections were stained with 0.1% safranin O. Alternatively, pellets were washed with DPBS and digested with 200 μl papain solution (1 μg/ml papain in 50 mM sodium phosphate, pH 6.5, containing 2 mM N-acetyl cysteine and 2 mM EDTA) for 16 h at 65°C. GAG was measured with the 1,9 dimethylmethylene blue (DMMB) method by using the Blyscan kit (Accurate Chemical & Scientific, Westbury, N.Y., USA) according to the manufacturer’s instruction. Briefly, 100 μl digested solution was mixed with 250 μl Blyscan dye and placed on a mechanical shaker for 30 min. The precipitate was collected by high speed centrifugation (>10,000g) for 10 min. Unbound dye was removed completely, and 600 μl dissociation reagent was mixed with the precipitate for 10 min to release bound dye. The absorbance of re-dissolved dye was measured in 96-well plates by using a Bio-red microplate reader at an absorbance of 656 nm. A standard curve of GAG was generated from the GAG standard solution supplied by the manufacture. The GAG concentration in each sample is presented as the GAG content normalized to the DNA amount.
Extraction of RNA and RT-PCR
Total RNA was extracted, and reverse transcription and amplification were performed as previously described (Jo et al. 2003). Briefly, total RNA was extracted by using Trizol reagent (Gibco-BRL, USA). All samples were treated with DNase I prior to first-strand synthesis. Quantities of RNA were measured by using a NanoDrop ND-100 spectrophotometer (NanoDrop, Wilmington, Del., USA). First-strand complementary DNA (cDNA) was synthesized by using 1 μg total RNA with Superscript II, oligo(dT) 12–18, and 10 mM dNTP mixture (Invitrogen, Carlsbad, Calif., USA). The resulting cDNA was then amplified with an Accupower PCR kit (Bioneer, Korea) in a volume of 20 μl in a thermocycler (Long Gene Scientific, China). Amplification of the β-actin gene was carried out in parallel for normalization. Of the reaction mixtures, 10 μll was electrophoresed in 1.5% agarose gel containing ethidium bromide to evaluate the degree of amplification and the sizes of the generated fragments. A 100-bp DNA ladder (Bioneer) was used as a standard size marker. The following human-specific primer sequences were used: β-actin (500 bp), forward: 5′-TCA TGT TTG AGA CCT TCA A-3′, reverse: 5′-GTC TTT GCG GAT GAT GTC CAC G-3′; fatty acid-binding protein (AP2, 290 bp), forward: 5′-AAG AAG TAG GAG TGG GCT TTG C-3′, reverse: 5′-CCA CCA CCA TGG TAT CAT CCT C-3′; lipoprotein lipase (LPL, 298 bp), forward: 5′-ATG GAG AGC AAA GCC CTG CTC-3′, reverse: 5′-TAC AGG GCG GCC ACA AGT TTT-3′; peroxisome proliferator-activated receptor γ2 (PPARγ2, 300 bp) forward: 5′-TTG GTG ACT TTA TGG AGC CC-3′, reverse: 5′-CAT GTC TGT CTC CGT CTT CTT G-3′; osteocalcin (OC, 100 bp), forward: 5′-GAG CCC CAG TTC CCC TAC CC-3′, reverse: 5′-GCC TCC TGA AAG CCG ATG TG-3′; osteopontin (OP, 350 bp) forward: 5′-GAG ACC CTT CCA AGT AAG TCC A-3′, reverse: 5′-GAT GTC CTC GTC TGT AGC ATC A-3′; alkaline phosphatase (ALP, 453 bp), forward: 5′-TGG AGC TTC AGA AGC TCA ACA CCA-3′, reverse: 5′-ATC TCG TTG TCT GAG TAC CAG TCC-3′; type IIa1 collagen (400 bp), forward: 5′-GAA GCT GGA AAA CCA GGT GA-3′, reverse: 5′-ACT TCT CCC TTC TCG CCA TTA G-3′; type IX collagen (170 bp), forward: 5′-CAC AGA GGT TTC AGT GGT TTG G-3′, reverse: 5′-GCA CCA GTA GCA CCA TCA TTT C-3′; aggrecan (280 bp), forward: 5′-GGA GCA TTC TGG ATT TCT GGA C-3′, reverse: 5′-GAC TCA AAA AGC TGG GGT GTG T-3′; Oct-4 (800 bp), forward: 5′-TGG AGA CTT TGC AGC CTG AG-3′, reverse: 5′-TGA ATG CAT GGG AGA GCC CA-3′; Nanog (360 bp), forward: 5′-AGG GTC TGC TAC TGA GAT GCT CTG-3′, reverse: 5′-CAA CCA CTG GTT TTT CTG CCA CCG-3′; Sox-2 (340 bp), forward: 5′-TAG CAC TTG TTG CCA GAA CG-3′, reverse: 5′-AAG CCG CTC TTC TCT TTT CC-3′; Rex-1 (200 bp), forward: 5′-TGA AAG CCC ACA TCC TAA CG-3’, reverse: 5′-CAA GCT ATC CTC CTG CTT TGG-3′.
Cells were cultured on glass coverslips in 24-well plates, fixed with 4% formaldehyde solution, permeabilized with 0.2% Triton X-100, and blocked with 5% normal goat serum in DPBS. The primary antibodies were rabbit anti-Oct-4, rabbit anti-Nanog, rat anti-SSEA-3, mouse anti-SSEA-4, mouse anti-Tra-1–60, and mouse anti-Tra-1–81 (Abcam) for pluripotent stem cell markers. Secondary antibodies were rabbit anti-goat IgG conjugated to Alexa 488 or 555 (Molecular Probes). All antibodies were diluted 1:150 to 1:200 in blocking solution according to the manufacturer’s instructions and were incubated on the cells for 1 h at room temperature. Nuclei were counterstained with 4’,6-diamino-2-phenylindole (DAPI) and imaged on a Carl Zeiss LSM 510 Meta confocal laser scanning microscope (Carl Zeiss, Jena, Germany). Images were captured with the Axiovision image analysis system (Carl Zeiss).
Senescence-associated β-galactosidase staining
Subconfluent cells at P2 (n=4), P5 (n=4), P7 (n=3), and P9 (n=3) were fixed with 2% formaldehyde/0.2% glutaraldehyde solution for 5 min at room temperature, incubated overnight at 37°C with fresh β-galactosidase (β-gal) staining solution (1 mg/ml 5-bromo-4-chloro-3-indolyl β-D-galactoside, 40 mM citric acid/sodium phosphate buffer pH 6.0, 5 mM potassium ferrocyanide, 5 mM potassium ferricyanide, 150 mM NaCl, and 2 mM MgCl2). MSCs derived from bone marrow were used at P5 as a control. Stained slides were examined, and images were processed with a Leica DFC290 camera and Leica application suite version 2.5.0.R1 (Leica Microsystems, Switzerland). Five random fields containing more than 100 cells were used to calculate the percentage of positive cells.
Telomerase activity assay
Telomerase activity was measured by the telomeric repeat amplification protocol (TRAP) with the telomerase PCR enzyme-linked immunosorbent assay (ELISA) kit (TeloTAGGG Telomerase PCR ELISA, Roche Molecular Biochemicals, Brussels, Belgium) according to the manufacturer’s protocol. Briefly, 2×105 cells at P2–P9 were pelleted at 3000g for 10 min at 4°C, washed twice with cold PBS, incubated for 20 min at 4°C with 200 μl pre-cooled lysis buffer (solution 1 of the kit), and centrifuged at 16,000g for 20 min. An aliquot of supernatant (2 μl) was mixed with the reaction mixture of the kit to perform the TRAP assay. Telomeric repeats were added to a biotin-labeled primer during the first reaction; then, the elongation products were amplified by PCR. An aliquot of the PCR product was denatured, hybridized to the digoxigenin (DIG)-labeled telomeric repeat-specific probe, and immobilized on a streptavidin-coated 96-well plate. Finally, the immobilized PCR product was detected with an anti-DIG-POD antibody, visualized as a color reaction product with the substrate 3,3′,5,5′-tetramethyl benzidine. The absorbance was measured in triplicate at 450 nm, by reading against a blank (reference absorbance at 690 nm). Samples are regarded as telomerase-positive if the difference in absorbance (A450 nm – A690 nm) was higher than 0.2.
All data values are shown as means±SD. Data were analyzed by using the paired t-test and analysis of variance. The significance level was set at P < 0.05.
Yield, morphology, growth kinetics, and CFU-F assay of cells from UC
Surface phenotype characterization
Adipogenic, osteogenic, and chondrogenic differentiation
Presence of pluripotent stem cell markers in human UC fMSC
Senescence and telomerase activity with respect to passage
UC fMSCs did not show significant senescence from P2 through P9 (Fig. 5b-e), whereas MSCs derived from bone marrow demonstrated significant senescence starting at P5 (Fig. 5f). We found that 1.11 ± 0.76% of the P2 UC fMSCs were positive and, similarly, that 1.50 ± 0.30% of the P9 UC fMSCs were positive (P > 0.05; Fig. 5g). Using the TRAP assay, we showed telomerase activitiy in UC fMSCs at all passages examined (Fig. 5h), with no statistically significant differences between passage numbers (P > 0.05).
We have successfully isolated MSCs from human UCs and cultured them for over 3 months without evidence of senescence. Except for two cases of fungal contamination at an early phase of cell harvest, the success rate of our isolation technique was 100%. This compares well with the reported success rate of 0%–63% for isolating cells from UCB (Wexler et al. 2003; Bieback et al. 2004). Our results, which are consistent with a previous report (Lu et al. 2006), demonstrate that UC is a reliable source of fetal MSCs. We have also been able to show that UC fMSCs can be induced to undergo adipogenic, osteogenic, and chondrogenic differentiation throughout a mid-term culture period of 3 months without significant evidence of reduction in differentiation potential. Additionally, no or minimal evidence of senescence has been observed, and telomerase activity has been consistently identified throughout the expansion period. These results are contrary to those reported in adult MSCs, which show a decreased differentiation potential and a decrease in telomerase activity after serial passaging (Izadpanah et al. 2006; Bonab et al. 2006). Although we have not directly compared UC fMSCs with adult MSCs, our results together with previous reports suggest that UC fMSCs might be more “primitive” and have superior quality in terms of both self-renewal and production of multiple lineages to adult MSCs.
We have isolated MSCs from the whole UC without discarding any material (Lu et al. 2006; Secco et al. 2008). Because nearly every component of the UC, including Wharton’s jelly, perivascular mesenchymal area, umbilical vein, and subendothelial tissues, has been reported to contain MSCs (Romanov et al. 2003; Wang et al. 2004; Sarugaser et al. 2005), we have hypothesized that use of the whole cord would be a simpler method and would also maximize the yield of MSC, which may prove crucial in future clinical applications. Indeed, we have found our MSC isolation method from whole UC to be both simple and high–yielding. Our yield of (3.99 ± 5.22)×105 cells/cm compares favorably with the yields from Wharton’s jelly (∼1×104 cells/cm; Weiss et al. 2006; Karahuseyinoglu et al. 2007) or from UC perivascular tissue (2.5–25)×104 cells/cm; Sarugaser et al. 2005). Some cells, however, show endothelial morphology at P0, but they do not significantly interfere with MSC proliferation under our culture conditions, as shown previously (Secco et al. 2008).
Adult MSCs have been widely investigated and found to present several significant drawbacks for use in clinical applications, such as donor site morbidity, lower isolation yield, and inferior proliferation and differentiation potential, all of which, moreover, decrease with age. The aging and senescence characteristics of adult MSCs in terms of their morphology (Stenderup et al. 2003), low frequency of CFU-F (Baxter et al. 2004), decreased maximal life span (Stenderup et al. 2003), lack of telomerase activity (Zimmermann et al. 2004), loss of multilineage differentiation potential (Roura et al. 2006), and senescence marker staining (Stolzing and Scutt 2006) are well documented. Senescence of adult MSCs is also known to increase with serial passage (Izadpanah et al. 2006). The potential for use of adult MSCs seems even less promising, considering that regenerative treatment strategies involving MSCs would be more frequently applied to older patients than younger patients. This study has demonstrated that UC fMSCs exhibit no evidence of senescence at least until P9 (CDPL >25) based on morphological observation, functional analysis of multilineage differentiation potential, senescence-associated β-gal staining, and TRAP assay, for the first time, as far as we know. Although a few authors have expressed concerns about the potential risk of the transformation of telomerase-positive MSCs (Rubio et al. 2005; Wang et al. 2005), others have demonstrated that MSCs from UC matrix expand long-term with no evidence of genetic changes (Karahuseinoglu et al. 2007; Fong et al. 2007). In this study, although we have observed positive telomerase activity from an early period of culture, this does not change over passages, suggesting their safety under our culture conditions.
Pluripotent stem cell markers are reported to be expressed in undifferentiated embryonic stem cells and not expressed in differentiated cells (Guillot et al. 2007). Nanog, Oct-4, Sox-2, and Rex-1 are crucial to pluripotency and self-renewal (Boyer et al. 2005). Previously, some of these pluripotent markers have been identified in UC matrix cells from pigs (Carlin et al. 2006), in which Nanog, Oct-4, and Sox-2 have been detected, and in horses (Hoynowski et al. 2007), in which Oct-4, SSEA-4, and c-Kit are expressed, but SSEA-3 and Tra-1–60 are weakly detected (<10%). In human UCs, only one study has revealed that Wharton’s jelly stem cells stain positively for SSEA-1, SSEA-4, Tra-1–60, and Tra-1–81 and negatively for SSEA-3 (Fong et al. 2007). In the present study, we have shown that UC fMSCs express eight well-known pluripotent stem cell markers at both the gene and protein levels throughout expansion up to P9 without evidence of any reduction in expression levels. Our results are contrary in part to the previous work by Fong et al. (2007) in which SSEA-3 stains negatively. We believe that this difference is caused by the different culture conditions used, including basic media (high versus low glucose DMEM), the concentration of serum, and the presence of bFGF. In addition, our results are in contrast to those from human UCB cells (McGuckin et al. 2005), which appear to be negative for SSEA-1, indicative of restriction in stemness. Considering that adult stem cells tend to lose their multi-differentiation potential by serial passage (D’ippolito et al. 1999; Rao and Matton 2001; Mareschi et al. 2006; Roura et al. 2006), our results revealing the expression of pluripotent stem cell markers, minimal evidence of senescence by β-gal staining, and the maintenance of multilineage differentiation potential and telomerase activity together suggest the advantage of UC fMSCs with regard to sustaining primitive characteristics during in vitro expansion.
In summary, this study provides a thorough characterization of the growth, differentiation, and gene expression properties of UC fMSCs over several passages. Despite the above-mentioned concerns regarding potential risks of transformation of telomerase-positive MSCs (Rubio et al. 2005; Wang et al. 2005), MSCs from UC matrix can be expanded long-term with no evidence of genetic changes (Karahuseinoglu et al. 2007; Fong et al. 2007), and the positive telomerase activity observed during early periods of culture of UC fMSCs does not change over passages, suggesting that they will probably not undergo transformation (present study). Indeed, we believe that several characteristics of UC fMSCs make them an attractive source of stem cells for clinical applications; there are no ethical concerns concerning their harvest, no donor morbidity is required, and UC fMSCs remain immature over several passages. In addition, in comparison with stem cells isolated from the UCB, a counter-part of UC, UC fMSCs are easier to isolate, and their isolation does not involve the expense of hematopoietic stem cells. Although more studies should be carried out for an exhaustive validation of the advantages of UC fMSCs, including in vivo implantation experiments, we suggest that UC MSCs may play an important role in bringing stem cell biotechnology from bench to bedside.
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