Stem Cell Rev and Rep (2011) 7:17–31 DOI 10.1007/s12015-010-9165-y

Growth and Differentiation Properties of Mesenchymal Stromal Cell Populations Derived from Whole Human Umbilical Cord Ingrida Majore & Pierre Moretti & Frank Stahl & Ralf Hass & Cornelia Kasper

Published online: 2 July 2010 # Springer Science+Business Media, LLC 2010

Abstract Up to 2.8×107 fibroblast-like cells displaying an abundant presence of mesenchymal stem cell (MSC) markers CD73, CD90, CD105 and a low level of HLA-I expression can be isolated from one whole human umbilical cord (UC) using a simple and highly reproducible explant culture approach. Cells derived from whole UC, similar to cells collected from separate compartments of UC, display a distinct chondrogenic and adipogenic potential. Therefore they are potential candidates for cartilage and adipose tissue engineering. Cell differentiation along the osteogenic pathway is, however, less efficient, even after the addition of 1.25-dihydroxyvitamin D3, a potent osteoinductive substance. Isolated cells are highly proliferative, tolerate cryopreservation with an average survival rate of about 75% and after thawing can be propagated further, at least over 20 population doublings before their proliferative activity begins to decline. More importantly, they synthesize numerous trophic factors including neurotrophins and factors I. Majore : P. Moretti : F. Stahl : C. Kasper (*) Institute of Technical Chemistry, Leibniz University of Hannover, Callinstraße 5, Hannover 30167, Germany e-mail: [email protected] I. Majore e-mail: [email protected] P. Moretti e-mail: [email protected] F. Stahl e-mail: [email protected] R. Hass Laboratory of Biochemistry and Tumor Biology, Department of Obstetrics and Gynecology, Medical University, Hannover, Carl-Neuberg-Straße 1, Hannover 30625, Germany e-mail: [email protected]

which facilitate angiogenesis and hematopoiesis. In conclusion, cells isolated from whole UC satisfies all requirements essential for the generation of stem cell banks containing permanently available cell material for applications in the field of regenerative medicine. Nevertheless, further studies are needed to improve and adjust the methods which are already employed for adult MSC expansion and differentiation to specific properties and requirements of the primitive stem cells collected from UC. So, our data verify that the choice of individual parameters for cell propagation, such as duration of cell expansion and cell seeding density, has a substantial impact on the quality of UC-derived cell populations. Keywords Umbilical cord . Multipotent mesenchymal stromal cells . Proliferation . Differentiation . Cytokine expression

Introduction Mesenchymal stem cells (MSC), first isolated from bone marrow (BM) stem cell niche in the middle 60s [1], are of particular importance for applications in the field of regenerative medicine. These somatic stem cells (SC) on the one hand possess the capability for extensive selfrenewal, on the other hand the potential to differentiate into various highly specialized cell types of mesodermal [2, 3] and even ectodermal [4, 5] or endodemal origin [6, 7]. In contrast to embryonic stem cells, MSC do not form teratomas after reimplantation and exhibit strong immunosuppressive and immunomodulatory properties [8–10]. Despite an extensive research, an unique cell surface marker for unambiguous identification and separation of MSC is not available up to now [11]. Therefore, in regard to encourage a

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more uniform classification of this type of stem cells, in 2006 the International Society for Cellular Therapy recommended a set of minimal criteria for MSC. To satisfy these criteria collected cells should display adherence to plasticsurfaces, should exhibit the expression of CD73, CD90 and CD105 with a concomitant absence of CD14, CD19, CD34, CD45 and HLA-DR expression and must differentiate at least to chondroblasts, osteoblasts and adipocytes [12]. Currently the most frequently used source of MSC is BM. Bone marrow derived MSC have been extensively investigated and, in particular cases, already successfully used in clinical trials, e.g. to heal large bone defects [13, 14], cartilage lesions [13, 15], spinal cord injuries [16], cardiovascular diseases [17, 18], hematological pathologies [19, 20], osteogenesis imperfecta [21] or attenuate graft versus host disease (GvHD) [22, 23]. The therapeutical effects observed after bone marrow MSC administration are not exclusively restricted to their direct differentiation into cells of appropriate host tissue. Of equal importance are the bioactive substances secreted by these stem cells, e.g. chemokines, growth and differentiation factors, which advance the migration, proliferation, differentiation of neighboring, e.g. tissue-resident progenitor cells, diminish apoptosis, promote angiogenesis or support hematopoiesis [24–26]. Nevertheless, BM aspiration is an invasive procedure and the portion of MSC in the BM mononuclear cell fraction is very small, i.e. apparently varies between 0.001– 0.01% [27]. In addition, the number and quality of MSC in this tissue decline with progressive age [28] and in the presence of degenerative diseases [29]. Therefore, searching for readily accessible, more consistent and richer sources of MSC than BM is a subject that recently attracts more and more attention. Research over the last decade has demonstrated that cells with characteristics similar to bone marrow MSC reside in virtually all post-natal organs [30] as well as in extraembryonic tissues available after child birth [31]. Especially, umbilical cord (UC) is already considered as a useful alternative to BM [32]. Tissue of UC seems to be an ethically non-controversial and reliable source of cells with MSC-like properties [33, 34]. UC is covered by a squamous epithelium and comprises two arteries and one vein, surrounded by mucous connective tissue rich in hyaluronic acid, so called matrix or Wharton’s jelly. Capillaries and lymphatics are not found in UC tissue [33]. MSC-like cells have been isolated from different compartments of UC with various degrees of reproducibility, i.e. from UC epithelium [35], subendothelium of umbilical vein [36], perivascular region [37], as well as from Wharton’s jelly [38, 39]. However, to harness UC derived stem cells, further investigation should be performed to survey their proliferative and differentiation potentials as well as safety of their clinical utilization.

Stem Cell Rev and Rep (2011) 7:17–31

The main objective of the present study was a detailed characterization of cell populations isolated from whole human UC tissue regarding their immunophenotype, purity, proliferative capacity, cytokine expression profile and differentiation potential along osteogenic, chondrogenic and adipogenic lineages.

Materials and Methods MSC Isolation and Cryopreservation For cell isolation from whole UC an explant culture approach was employed. Human UCs (MK 240707, HD 140509, NS 010408, NS 190109) were obtained from termdelivery (38–40 weeks) by Cesarean section patients (n=4) after informed written consent as approved by the Institutional Review Board, project #3037 in an extended permission on 17th June, 2006. Blood from UC vessels was removed and the UC was placed in PBS (phosphate buffered saline) enriched with 5 g/l glucose (Sigma Aldrich), 50 μg/ml gentamicine (PAA Laboratories), 2.5 μg/ml amphotericin B (Sigma Aldrich), 100 U/ml penicillin and 100 μg/ml streptomycin (PAA Laboratories). At the laboratory UC was cut into approx. 10 cm large segments which further were minced in ca. 0.5 cm3 large pieces and placed in 175-cm2 tissue culture flasks (Sarstedt). Then these pieces were incubated in αMEM (Invitrogen) enriched with 15% of allologous human serum (provided by the Division of Transfusion Medicine, Medical University Hannover, Germany) and 50 μg/ml gentamicine at 37°C in a humidified atmosphere with 5% CO2. The medium change was carried out every second day. A beginning outgrowth of an adherent cell layer from single tissue pieces was observed after approx. 10 days. After 2 weeks, the tissue pieces were removed and the adherent cells were harvested by accutase (PAA Laboratories) treatment according to the manufacturer’s protocol. The cell suspension was centrifuged at 200×g for 5 min and the received cell pellet was resuspended in growth medium, i.e. αMEM supplemented with 10% human serum and 50 μg/ml gentamicine. Cells were subcultured at the density of 4.000 cells/cm2 in 175-cm2 tissue culture flasks and grown until 80% of confluence. Subsequently cells were harvested as already described and used for immunophenotype analysis or cryopreserved. Cell cryopreservation was performed in liquid nitrogen using cryomedium containing 10% (v/v) growth medium, 10% (v/v) DMSO (Sigma Aldrich), 80% (v/v) allologous human serum and a freezing rate of 1°C/ min at a average cell density of 1.5×106 cells/ml. For the following experiments cells of passage 2 (after thawing) were used.

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Immunophenotypic Analysis by Flow Cytometry Primary UC-derived cells were harvested by accutase treatment, washed twice in cold PBS supplemented with 2% FCS (PAA Laboratories) and resuspended to a concentration of 1×106 cells/ml. Staining was performed by adding 20 µL of titrated antibody solution to 100 µL of the cell suspension. Mouse anti human antibodies were used for the analyses. PE-conjugated anti MSCA-1 and anti CD271 as well as FITC- anti CD31 antibodies were purchased from Miltenyi Biotec. PECy5- anti CD34, PEanti CD44, PECy5- anti CD45, PE- anti CD73, FITC- anti CD90 antibodies were obtained from BD Biosciences and PE- anti CD105 from Invitrogen. Negative control staining was performed using matched isotype control antibodies. After storage for 20 minutes at room temperature in the dark, 400 µL of PBS supplemented with 2% FCS was added and cells were analyzed in a EPICS XL/MCL flow cytometer (Beckman Coulter). At least, 10.000 gated events were acquired on a LOG fluorescence scale. Positive staining was defined as the emission of a fluorescence signal that exceeded levels obtained by >99% of cells from the control population stained with matched isotype antibodies. Neural ganglioside (GD2) analysis was performed using a mouse primary antibody detected by a secondary goat anti-mouse PE-labeled antibody (all BD Biosciences). For this analysis, more than 5·105 living single cells (according to propidium iodide exclusion and pulse high and area measurements) were acquired. Histograms were generated using the software WinMDI 2.8 (Joseph Trotter). Determination of the Basal Level of Alkaline Phosphatase (AP) Activity UC-derived cells were thawed, seeded in 6-well plates (Sarstedt) at a density of 4.000 cells/cm2 and cultivated in growth medium until 80–90% of confluence was reached. Thereafter, the cell layer was rinsed once with PBS and 1,0 ml/well of AP substrate prepared from SigmaFast™ pNitrophenyl Phosphate Tablet set (Sigma Aldrich) was added. Following an incubation period of 4 h at 37°C, 50 μl of reaction product were transferred from each well into a 96-well plate and the absorbance at 405 nm was measured. AP activity was calculated based on a standard curve created by the use of p-nitrophenole (Sigma Aldrich). Thereafter, the calculated values of AP activity were normalized to the average number of cells per well. Cell Proliferative Capacity After thawing the cells were cultivated over one passage in growth medium applying the seeding density of 4.000 cells/ cm2. At 80% of confluence cells were harvested and

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reseeded in tissue culture flasks (Sarstedt) using seeding densities of 4.000 and 500 cells/cm2. The change of growth medium was performed three times a week. The cells were always harvested at about 80% of confluence and repeatedly reseeded at the same cell density (4.000 or 500 cells/ cm2) until cells reached the complete growth inhibition. The numbers of intact and dead cells within the individual passages were determined in triplicate by Trypan Blue exclusion. From these data, the numbers of cell population doublings, cumulative population doublings, population doubling time, increase in the cell number as well as the percentage of dead cells in the present cell cultures were calculated. The total number of obtained intact cells was computed based on the assumption that 1×105 cells were initially seeded. Cell Cycle Analysis Cell cycle analysis was performed using propidium iodide (PI) as previously described [40]. Determination of Caspase 3/7 Activity In passage 2, 5, 9, 13 and 17 of UC-derived cells, the specific caspase 3/7 activity was measured in triplicate using ApoOne® Homogenous Caspase-3/7 Assay (Promega). Thus, cells were seeded in 24-well plates (Sarstedt) at a density of 4.000 or 500 cells/cm2 and cultivated in growth medium until 80% of confluence. Thereafter cell layer was rinsed once with PBS and 0.2 ml of caspase 3/7 substrate was added to each well. The plates were incubated at 37°C for 2 h and the intensity of fluorescence signals was detected at an excitation wavelength of 480 nm and an emission wavelength of 530 nm. The intensity of fluorescence signals was normalized to the average cell number per well. Determination of Cell Senescence The portion of senescent cells was determined in passages 2, 5, 9, 13 and 17 by the Senescence-associated β-Galactosidase (SA-β-gal) Staining Kit (Cell Signaling Technology) and DAPI (Roche Diagnostics) fluorescence counterstain. Cells were seeded in 6-well plates (Sarstedt) at a density of 4.000 or 500 cells/cm2 and cultivated in growth medium until 80% of confluence. SA-β-gal and DAPI staining was performed in accordance to the manufacturers’ instructions. After completion of the assay procedures, 6 representative images were taken from diverse areas of each cell culture using phasecontrast microscopy, fluorescence microscopy and CellB Imaging Software (Olympus GmbH). For the calculation of the percentage of senescent cells the total number of cell nuclei and number of cell nuclei surrounded by cyan dye were enumerated.

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Stem Cell Rev and Rep (2011) 7:17–31

Chondrogenic Differentiation

Osteogenic Differentiation

For the examination of the chondrogenic potential cells of the first passage were revitalized and over one passage cultivated in growth medium applying the seeding density of 4.000 cells/cm2. At about 80% of confluence cells were harvested and reseeded in the fibronectin (extracted from human plasma, Roche Applied Science) coated (5 μg/cm2) 6-well plates and 75-cm2 tissue culture flasks at the density of 2,000 cells/cm2 and cultivated in growth medium with reduced (5%) serum content. In this way cell proliferative activity was decreased and cells were adapted to low-serum culture conditions prior the initiation of differentiation. At a confluence the growth medium was replaced by the chondrogenic medium, i.e. DMEM (high glucose, Invitrogen) consisting 1% human serum (provided by the Division of Transfusion Medicine, Medical University Hannover, Germany), 1 mM sodium pyruvate, 0.2 mM L-ascorbate-2-phosphate, 100 nM dexamethasone, 0.35 mM L-proline, ITS+1 supplement (all from Sigma Aldrich), 10 ng/ml TGF-β3 (PeproTech) and 50 μg/ml gentamicine (PAA Laboratories). The medium change was performed every 2-3 days. Cells cultivated in corresponding medium without differentiation factors served as negative control. Three weeks afterwards the presence of mRNA for Sox9, COMP (cartilage oligomeric matrix protein), aggrecan and collagen X genes, typically expressed during chondrogenic differentiation, was determined by RT-PCR. The presence of collagen II as well as the accumulation of acidic proteoglycans and aggrecan in the extracellular matrix (ECM) were additionally identified via Alcian Blue staining and immunostaining.

To induce cell differentiation along the osteogenic lineage, cells were revitalized, expanded and seeded in the fibronectin coated 24-well plates and 75-cm2 tissue culture flasks at a density of 2.000 cells/cm2 as already described. One day later the growth medium was replaced by the osteogenic medium, i.e. DMEM (low glucose, Sigma Aldrich) containing 5% human serum (provided by the Division of Transfusion Medicine, Medical University Hannover, Germany), 0.2 mM L-ascorbate-2-phosphate, 5 mM β-glycerophosphate, 100 nm dexamethasone and 50 nm 1.25-dihydroxyvitamin D3 (all from Sigma Aldrich). Adipose tissue (AT)-derived MSC (kindly provided by Prof. Martijn van van Griensven from the Ludwig Boltzmann Institute for Experimental and Clinical Traumatology, Vienna, Austria), with a pronounced capacity for osteogenic differentiation were cultivated under the same conditions and employed as positive control. The osteogenic medium was replaced three times a week. AP activity, collagen I secretion as well as Ca2+ accumulation in the ECM were monitored in both experimental settings over 39 days. In addition, the presense of mRNA for genes relevant to the bone tissue formation—RUNX2, collagen I, AP, osteopontin and osteocalcin—were examined by RT-PCR four weeks after the initiation of osteogenesis. At the end of the cultivation period cells positive for AP and Ca2+ deposits in the ECM were visualized using SIGMA FAST™ BCIP/NBT substrate and calcein stain, respectively (both from Sigma Aldrich).

Adipogenic Differentiation To verify the adipogenic potential cells of the first passage were revitalized, expanded, reseeded in the fibronectin coated 6-well plates at a density of 2.000 cells/cm2 and grown in serum reduced medium as described in the previous section. Starting from the confluence the adipogenic medium, i.e. DMEM (high glucose, Invitrogen) enriched with 5% human serum (provided by the Division of Transfusion Medicine), 100 nm dexamethasone, 1,7 μM insulin, 500 μM IBMX (all from Sigma Aldrich), 1 μM U0126 (Calbiochem), 1 μM rosiglitazone (Biozol) and 50 μg/ml gentamicine (PAA Laboratories) was applied. Medium was changed three times a week. Cells cultivated in the same medium without the addition of adipogenesisinducing substances served as negative control. The intracellular accumulation of lipid granules was visualised using Oil Red O stain (Sigma Aldrich) 4 weeks after induction of adipogenesis.

Determination of Collagen I Synthesis and AP Activity in Osteogenic-Induced Cell Cultures To determinate the intensity of collagen I synthesis, at each medium change the cell culture supernatants from three related wells were pooled, centrifugated at 10.000 rpm for 5 min and the cell-free supernatant stored at -20°C until analysis. The amount of secreted Collagen I was determined by use of Cterminal Propeptide of Type I Collagen (CICP) enzyme immunoassay (Quidel) in accordance to the manufacturers’ instructions. The cumulative release of CICP, i.e. the amount of collagen I matrix accumulated up to a particular point in time, was calculated by summing up the individual CICP-values. The changes in the activity of AP were determined in triplicate as described in the section Determination of the basal level of alkaline phosphatase (AP) activity. To each well of 24-well plate 0.2 ml of AP substrate were added. Calcein Test and Calcein Staining in Osteogenic-Induced Cell Cultures The calcium accumulation in the ECM was tracked by calcein (Sigma), a fluorescent dye which displays a high affinity to

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Ca2+. In brief, the cell layer was rinsed twice with PBS, fixed in ice cold 75% ethanol for 15 min at room temperature, washed with PBS and covered with appropriate volume of calcein solution (5 μg/ml in H2O). Samples were incubated overnight at 4°C and thereafter extensively washed with distilled water. The fluorescence of the bounded calcein was detected in triplicate at an excitation wavelength of 480 nm and an emission wavelength of 530 nm.

were an initial denaturation at 95°C for 5 min followed by 35 cycles of denaturation at 95°C for 30 s, annealing at 48– 62°C for 30 s and extension at 72°C for 30 s. The reaction was completed with a final extension step at 72°C for 10 min. The PCR products were analyzed in Agilent 2100 Bioanalyser using Agilent DNA 7500 Kit in accordance to manufacturer’s protocol. The gene specific primer sequences, annealing temperatures as well as the size of PCR products are listed in the Tables 1, 2 and 3.

Reverse Transcription-Polymerase Chain Reaction (RT-PCR)

Immunostaining and Staining of AP Positive Cells

To determine the expression of lineage-specific genes and housekeeping gene GAPDH, cells were cultivated in fibronectin coated 75-cm2 cell culture flasks under chondrogenic or osteogenic conditions as described in previous sections. The total RNA was extracted using RNeasy® Plus mini kit (Qiagen) according to manufacturer’s instructions. First strand cDNA synthesis was carried out from 1 μg of total RNA by use of QuantiTect® Reverse Transcription kit (Qiagen). The PCR amplifications were performed in a 50-μl reaction mix containing ∼ 25 ng of cDNA, 1.25 units Taq DNA polymerase (OLS), 160 μM of each dNTP (Fermentas), 10 pM of each gene specific primer (MWG), 1x PCR Buffer (OLS) and 35.75 μl of RNase-free water in iCycler (BioRad). The conditions used for amplification

For immunostaining cells were seeded in 6-well plates and cultivated until 80% of confluence (AP immunostaining) or over three weeks (collagen II and aggrecan immunostaining). Afterwards cell layer was rinsed twice with PBS and fixed in 4% paraformaldehyde for 30 min at room temperature. After an extensive rinsing with PBS cells were covered with blocking/permeabilization buffer containing 5% horse serum (Invitrogen) and 0.3% Triton X100 (Sigma Aldrich) in PBS and incubated for 1 h at room temperature. Thereafter the blocking buffer was replaced by mouse monoclonal antibody against the common epitope of AP (ab58958, Abcam), mouse monoclonal antibody against collagen II (sc-59958, Santa Cruz Biotechnology) or mouse monoclonal antibody against aggrecan (MAB19310, Milli-

Table 1 Primer sequences, primer annealing temperatures and sizes of PCR products Lineage

Gene/gene accession number

Direction

Sequence (5’- 3’)

Annealing Tº (º C)

Amplicon size (bp)

Osteogenic

RUNX 2 NM_004348 Coll I α-2 NM_000089 AP NM_000352 Osteopontin NM_000352 Osteocalcin NM_000711 SOX9 NM_000346 Coll X, α-1

forward reverse forward reverse forward reverse forward reverse forward reverse forward reverse forward

TCC TATGACCAGTCTTACCCCT GGCTCTTCTTACTGAGAGTGGAA AATTGGAGCTGTTGGTAACGC CACCAGTAAGGCCGTTTGC GCTGAACAGGAACAACGTGA CCACCAAATGTGAAGACGTG CTCCATTGACTCGAACGACTC CAGGTCTGCGAAACTTCTTAGAT CACTCCTCGCCCTATTGGC GCCTGGGTCTCTTCACTACCT AGCGAACGCACATCAAGAC GCTGTAGTGTGGGAGGTTGAA ATGCTGCCACAAATACCCTTT

62.0

190

62.0

125

51.0

267

62.0

230

62.0

138

60.7 61.7 60.2

110

NM_000493 COMP NM_000095

reverse forward reverse

GGAATGAAGAACTGTGTCTTGGT GTGTGAGACCGGGCAACATAA GGGAAGCCGTCTAGGTCAGT

60.4 62.3 62.8

Aggrecan

forward reverse forward reverse

CCCAACCAGCCTGACAACTTT GTACCGCACCAGGGAATTGAT TGTTGCCATCAATGACCCCTT CTCCACGACGTACTCAGCG

62.8 62.2 62.0

Chondrogenic

Control

X17406 GAPDH NM_002046

134 279 216 202

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Table 2 Immunophenotype of cells collected from whole UC Cell surface marker

were fixed in 4% paraformaldehyde and rinsed twice with PBS. Then 200 μl of AP substrate prepared from SigmaFAST™ BCIP®/NBT tablets (Sigma) were added to each well. Following an incubation at 37°C for 30 min, cell layer was rinsed twice with PBS and samples were analyzed as described above.

Percentage of positive cells Mean value

Standard deviation

CD31 CD34 CD44 CD45 CD73

0.9 1.4 99.9 0.4 98.9

± ± ± ± ±

0.8 0.2 0.2 0.4 1.8

CD90 CD105 GD2 HLA-I

99.9 98.9 0,08 2.5

± ± ± ±

0.2 2.3 0,07 1.9

Determination of Growth Factor Expression Profile The cytokine expression profile of human UC-derived cells was analyzed via RayBio® Human Cytokine Antibody Array (RayBiotech) in accordance to the manufacturers’ guidelines. On each membrane 500 μg of total protein obtained by cell lysis were loaded. The bounded antibodies were detected using ImmPACT™ DAB peroxidase substrate (Vector Laboratories). Thereafter membranes were scanned and image analysis was carried out with Imagene 5 software (BioDiscovery). The detected signal intensities were corrected for background and normalized using positive control included on the array. The extent of bFGF secretion was quantified by Quantikine® Human FGF basic Immunoassay (R&D System) in supernatants of confluent cell cultures 48 h after the last medium change in accordance with the manufacturers’ protocol. Afterwards the calculated bFGF values were corrected for bFGF content in culture medium. In this experimental setting cells were seeded in 6-well plates at a density of 4.000 cells/cm2 and 1.5 ml of growth medium was applied per well at each medium change.

pore), respectively. Per well 0.5 ml of primary antibody, previously diluted to a final concentration of 10 μg/ml in PBS containing 1% human serum albumin (provided by blood bank, Springe, Germany) and 0.3% Triton X were applied. Following overnight incubation at 4°C, cell layer was rinsed three times with PBS and the bound primary antibodies were detected by use of ImmPRESS™ Reagent Anti-Mouse Ig, ImmPACT™ DAB peroxidase substrate or Vector® NovaRED substrate kit for peroxidase (all from Vector Laboratories) in accordance to the manufacturers’ instructions. The negative control samples were incubated with the equal amount of IgG1 (MAB002, R&D Systems) instead of monoclonal antibodies. Samples and negative controls were analyzed using phase-contrast microscopy and CellB Imaging Software (Olympus GmbH) immediately after completion of staining procedure. The portion of AP positive cells was examined in osteogenic-induced cultures at the end of cultivation time. Cells seeded in 24-well plates at a density of 2.000 cells/cm2

Statistical Analysis All data are represented as mean value±standard deviation. The differences between the mean values were examined for statistical significance using the Student’s t-test.

Table 3 Changes in growth kinetics and in number of obtained intact cells during long-term expansion of UC-derived cells Cell seeding density

4,000 cells/cm2

500 cells/cm2

a

Parameters

Days in culture PD time (h) cumulative PD obtained intact cellsa Days in culture PD time (h) cumulative PD obtained cellsa

Passage 2

5

9

13

16

3 27.5±1.6 2,6±0.2 6.0±0.6×105 6 24.5±1.2 5.9±0.3 4.5±1.1×106

14 29.4±2.2 14.2±0.3 1.5±0,2×109 28 36.1±1.1 25.2±0.4 2.0±0.4×1012

31 38.3±0.5 27.9±0.3 1.3±0.1×1013 66 95.1±5,5 44.8±0.4 9.9±2.1×1017

49 42.5±2,5 41.5±0.4 1.0±0.2×1017 – – – –

63 78.9±6.3 47.0±0.4 2.7±0.2×1018 – – – –

The number of obtained intact cells at the particular points in time was calculated based on the assumption that 1×105 cells were initially seeded

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individual UC donors ranged between 0.319±0.061 and 0.822±0.006 mU/106 cells, respectively (Fig. 1a).

Results General Properties of Cell Populations Derived from Whole UC Tissue The UCs from 4 different donors (MK 240707, HD 140509, NS 010408, NS 190109) were prepared and up to 2.8×107 cells displaying a predominantly fibroblast-like morphology (Figs. 1c, 5e) were obtained from each of these individual UC preparations. Collected cell populations were cryopreserved with an average cell recovery rate of 75.0± 12.8% after thawing. Moreover, the UC-derived cell cultures displayed features of at least two different subpopulations exhibiting clear differences in cell size, morphology and in the nucleus-to-cytoplasm ratio [41]. Characterization of Cell Immunophenotype via Quantitative Flow Cytometry The majority of collected UC-derived cells showed a prominent presence of the MSC surface markers CD73, CD90, CD105 as well as the expression of CD44, a receptor for hyaluronic acid (Tab. 2). A small fraction of isolated cells stained positive for neural ganglioside GD2, a novel marker for the identification of MSC [42]. The data demonstrated furthermore, that HLA-I proteins were present in 2.5±1.9% of the cells and there was little if any expression detectable of CD31, CD34 and CD45, respectively, suggesting the absence of endothelial and hematopoietic cells types. Moreover, the immunocytology revealed in UC-derived cell cultures a high portion of cells displaying AP expression (Fig. 1b). The specific AP activity in the cell cultures obtained from 4 different

Analysis of Cell Differentiation Potential The isolated UC-derived cell populations exhibited a potential to differentiate along chondrogenic, adipogenic and, less extended, osteogenic lineage. Thus, a successive cell condensation in nodule like structures exhibiting high intensity of ECM formation was observed in UC-derived cell cultures after the induction of chondrogenesis (Fig. 2a). These nodules were particularly rich in collagen II (Fig. 2c), acidic proteoglycans (Fig. 2d) and aggrecan (Fig. 2e), a proteoglycan typically found in the ECM of hyaline cartilage, as detected by Alcian Blue staining and immunostaining three weeks post induction. RT-PCR confirmed these findings and revealed the transcription of further cartilage specific genes such as SOX9, COMP and collagen X at this moment (Fig. 2b). The expression of collagen X gene suggested the presence of prehypertrophic or hypertrophic chondrocytes in the induced cell cultures. In the adipogenic-stimulated cultures a significant changes in the cell morphology together with continuous accumulation of intracellular lipid granules was observed, which represent unique features for an ongoing adipogenic differentiation. Four weeks after the initiation of adipogenesis many large, flattened, often oval cells exhibited differentially-sized lipid granules (Fig. 3a). Cells displaying unilocular morphology, i.e. mature adipocytes, were not observed at this time. In contrast, cells in the nonstimulated negative control cultures maintained their original spindle-shaped morphology and the formation of lipid granules remained undetectable (Fig. 3b).

A AP activity (mU/106 cells)

1,00

0,75

B

200 µm

C

200 µm

0,50

0,25

0,00

MK240707

NS010408

NS190109

Fig. 1 Expression of AP in UC-derived cell cultures. a specific AP activity in cultures of four UC-derived cell strains; b immunological detection of AP-positive, i.e. dark red colored cells, in a cultures of cell strain MK240707 using monoclonal antibody against common epitope of AP; similar level of AP expression was detected in all UC-

HD140509

derived cell populations; c corresponding control cell cultures incubated with mouse IgG1 instead of AP did not show any staining. For visualisation of cell nuclei hematoxylin counterstain was performed

300

B

A

Aggrecan

COMP

SOX 9

GAPDH

2.0 cm

Collagen X

Stem Cell Rev and Rep (2011) 7:17–31 DNA ladder

24

100

C

200 µm

D

200 µm

E

200 µm

C1

200 µm

D1

200 µm

E1

200 µm

Fig. 2 Chondrogenic potential of UC-derived cells. After the initiation of chondrogenesis a gradual cell condensation in nodulelike structures was observed. Cell condensation sites were visualized using hematoxylin nuclear stain three weeks after the induction of chondrogenesis a. At the same time the expression of GAPDH gene and chondrogenesis-related genes was determinated using RT-PCR b,

presence of collagen II c as well as the accumulation of acidic proteoglycans d and aggrecan e in the cell layer cultures were recognized via Alcian Blue staining d and immunostaining c, e. Cells cultivated in corresponding medium without the addition of chondrogenic factors did not show any or showed less pronounced coloring (c1, d1, e1). Data are representative of four experiments

Collected cell differentiation along the osteogenic lineage was, however, less efficient, even after the addition of 1.25dihydroxyvitamin D3 (Fig. 4). Despite the expression of RUNX2, a master gene for osteogenic differentiation (Fig. 4f), and massive generation of collagen I matrix (Fig. 4a), only a rare presence of strongly AP-positive cells (Fig. 4d) and the first mineralization centers (Fig. 4e) were

observed in UC-derived cultures at the end of the cultivation period of nearly 6 weeks. At the same time MSC cultures from adipose tissue, employed as positive control, showed high AP activity (Fig. 4b, d1) and strong mineralization of the ECM (Fig. 4c, e1). Moreover, RT-PCR analysis revealed in UC-derived cell populations little if any expression of OC gene after 4 weeks of osteogenic induction (Fig. 4f).

A

50 µm

Fig. 3 Adipogenic potential of UC-derived cells. The intracellular accumulation of lipid granules (indicated by white arrows) was visualised using Oil Red O stain a 4 weeks after initiation of adipogenic differentiation. Cells cultivated in corresponding medium without

B

50 µm

adipogenic substances showed presence of small intracellular lipid droplets b. For the visualisation of cell nuclei hematoxylin counterstain was performed. Data are representative of four experiments

25 40

5000 UC-derived MSC AT-derived MSC

35

4000 AP-activity (U/l)

Cumulative release of CICP (ng/Well)

Stem Cell Rev and Rep (2011) 7:17–31

3000 2000

30 25 20 15 10

1000 5 0

0 0

5

10 15 20 25 30 Duration of cell cultivation (days)

35

0

40

A

5

10 15 20 25 30 Duration of cell cultivation (days)

35

40

B UC-derived cells

200

D

150

E

200 µm

AT-derived MSC

50

40

AT-derived MSC

200 µm

Osteopontin

Osteocalcin

RUNX 2

GAPDH

Osteopontin

Collagen I

DNA ladder

D1

E1

200 µm

AP

C

Osteocalcin

10 15 20 25 30 35 Duration of cell cultivation (days)

RUNX 2

5

GAPDH

0

Collagen I

0

F

200 µm

100

AP

Calcein fluorescence (FU)

250

UC-derived cells

Fig. 4 Verification of the osteogenic potential of UC cells and adipose tissue (AT)-derived MSCs. a accumulation of the collagen I matrix (cumulative release of CICP in the cell culture supernatant) ; b changes in the activity of AP; c progress in the extracellular matrix mineralization (calcein fluorescence); d, d1 portion of strongly APpositive cells in cultures of AT- and UC-derived cells at the end of the

cultivation period; e, e1 ECM mineralization (calcein fluorescence) in cultures of UC- and AT-derived cells at the end of the cultivation period. f expression of GAPDH gene and genes relevant to bone tissue formation in cultures of AT- and UC-derived cells 4 weeks after induction of osteogenesis

Analysis of Cell Proliferative Capacity and Viability

Cell seeding at a lower density of 500 cells/cm2 allowed to avoid a frequent enzymatic disruption of cell-cell contacts and reduced the cell loss connected to the additional steps of cell harvesting and reseeding. Independently from the seeding density, cell proliferative activity remained unchanged over approx. 20 population doublings (PD) under the applied cell culture conditions (Fig. 5a). Thereafter, the cell growth rate declined gradually and after approx. 47 PD (seeding density 4,000 cells/cm2) or 45 PD (seeding density 500 cells/cm2) cells entered the phase of complete inhibition of cell division (Fig 5a). At this point in time all investigated cell cultures contained a large number of big, flattened cells and significantly increased portion of

Cells collected from whole human UC exhibited remarkable proliferative activity. So, the average cell population doubling time in the cultures of passage 5 cells corresponded to 29.4±2.2 h (Tab. 3), when seeding density of 4.000 cells/cm2 was applied. Early passage cells reached 80% of confluence within 3-4 days under these culture conditions. Nevertheless, the highest intensity of cell division was observed two days after seeding, i.e. at about 40% of confluence. Flow cytometry yielded 8.9±1.6% of cells located in the S phase and31.1±1.3% of cells in the G2/M phase of cell cycle at that time (Fig. 5b).

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50

60.0 ± 2.9 % (day 2) 94.9 ± 1.7 % (day 6)

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Fig. 5 Proliferative activity and viability of UC-derived MSCs during their long-term expansion. a cumulative population doublings; b cell cycle analysis in subconfluent (2 days after cell seeding) and confluent (six days after cell seeding) cultures of passage 4 cells using seeding density of 4,000 cells/cm2; c changes in the caspase 3/7 activity and in the percentage of dead cells steadily applying cell seeding density of 4,000 cells/cm2; d changes in the caspase 3/7 activity and in the percentage of dead cells steadily applying cell seeding density of 500

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cells/cm2; e, f SA-β-gal-positive cells in the cultures of passage 2 and passage 17 cells (seeding density 4,000 cells/cm2). g changes in the percentage of senescent cells (seeding density 4,000 cells/cm2). In figures c and d the numbers of cumulative population doublings (PD) achieved at the time of analysis are indicated. Asterisks denote statistically relevant differences in comparison to cultures of second passage: * p<0.05, ** p<0.01, *** p<0.001

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and 4.87 ± 1.46 pg/ml as detected by human bFGF immunoassay (Fig. 6b).

cells positive for SA-β-gal in comparison to cultures of early passage cells (Fig. 5e, f, g). Furthermore, cells steadily seeded at a density of 500 cells/cm2 forfeited their proliferative activity and viability during long term expansion more rapidly than cells seeded at a density of 4.000 cells/cm2 (Fig. 5c, d). Fast and strong increases in the caspase 3/7 activity and in the percentage of dead cells (Fig. 5d) were detected in these cell cultures. Accordingly, at a seeding density of 4.000 cells/cm2 1.0±0.1×1011 intact cells and at a seeding density of 500 cells/cm2 6.0±0.4× 1010 intact cells were obtained after approx. 20 PD, i.e. before the cell proliferative activity in both experimental settings began to decline.

Discussion A high number of adherent cells displaying a mainly fibroblast-like morphology as well as the presence of cell surface markers CD44, CD73, CD90 and CD105 can be isolated from whole human UC tissue using an explant culture. This cell isolation procedure is simple, highly reproducible, non-invasive and yields an immunophenotypically homogenous cell population without enzymatic digestion of UC tissue or utilization of additional purification steps associated with blood and endothelial cell elimination. In addition to abundant expression of typical MSC surface markers a low basal level of the AP activity was detected in all examined cell cultures. This enzyme is highly expressed in embryonic stem cells [45] and has also been recorded in cultures of MSC harvested from adult tissues [46]. More importantly, in cell populations collected from whole UC only a small subset of cells carries HLA-I proteins. This finding strongly disagrees with data published by other research groups [47, 48], which reported that the majority of cells isolated from human UC are positive for HLA-I. This discrepancy raises the question, how the employed in vitro culture conditions, in particular the presence of xenogenic sera, influence the extent of

Cytokine Expression Profile of UC-Derived Cells The analysis of cytokine expression pattern via RayBio® Human Cytokine Antibody Array yielded in UC-derived cell lysates the presence of various trophic mediators, including growth factors bFGF, EGF, IGF-I, IGF-II, PDGFs, TGF-βs, neurotrophins (b-NGF, GNDF, NT-3, NT-4) and factors which augment haematopoiesis (G-CSF, GM-CSF, HGF, M-CSF, SCF) or angiogenesis (PIGF, VEGF; VEGF-D) (Fig. 6a). The most prominent expression levels, however, were detected for IGF-II, GM-CSF, FGF-4, G-CSF, TGF-β and bFGF, respectively. At the confluence the concentration of bFGF, a growth factor which particularly prolongs the life span of MSC and supports the maintenance of their multilineage potential [43, 44], ranged in the cell culture supernatant between 1.47±0.41

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Fig. 6 Cytokine expression profile of human UC-derived cells. a expression levels of 24 cytokines detected in lysates of UC-derived cells using RayBio® Human Cytokine Antibody Array (data are representative of three experiments); b quantification of bFGF in the supernatants of confluent cell cultures 48 h after medium change via human bFGF immunoassay

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HLA-I expression [49]. We are the first, who generally avoided the use of any xenogenic media supplements during UC cell isolation, expansion and differentiation. Furthermore, our study demonstrates that cells derived from whole UC tissue, equal to cells harvested from discrete compartments of human UC [32, 50–52], display a distinct chondrogenic potential and the capability to differentiate along the adipogenic pathway. Therefore they are potential candidates for cartilage and adipose tissue engineering. We believe that UC-derived cell differentiation into cartilage-forming cells is strongly supported by the structural properties of UC tissue. Human UC is very rich in hyaluronic acid (HA), a native component of articular cartilage [33]. The presence of HA substantially increases the commitment of BM- and AT-derived MSC to undergo chondrogenesis, as described by Chung & Burdick [53] and Wu et al. [54]. So, in monolayer cultures of UC-derived cells after the induction of chondrogenesis a spontaneous cell aggregation displaying abundant presence of collagen II, aggrecan and acidic proteoglycans was observed. The chondrogenesis of MSC derived from adult tissues is often performed in high-density pellet cultures, which mimic mesenchymal condensation during early stages of skeleton development and therefore facilitate chondrogenesis. In contrast to adipogenic and chondrogenic differentiation potentials, which are confirmed by several scientific groups, the statements regarding the osteogenic capacity of UC-derived cells are widely different. Whereas some investigators describe an osteogenic potential comparable to bone marrow MSC [55, 56] and even higher [32], others show that cells collected from human UC are poorly osteogenic and only some of them are capable to undergo osteogenic differentiation [57, 58]. Our data corroborate the last thesis. The majority of cells derived from whole UC tissue did not complete the osteogenic differentiation, even after the addition of such a potent osteoinductive substance as 1.25-dihydroxyvitamin D3. Despite the expression of the most important osteogenic marker genes and massive secretion of collagen I, the mineralization of the ECM, a reliable marker for ongoing bone tissue formation, remained nearly undetectable even 6 weeks after the initiation of osteogenesis. The very small portion of strongly AP-positive cells and the absence of osteocalcin gene expression suggested that only a few cells in the investigated cell cultures were able to reach the developmental stages of pre-osteoblasts or mature osteoblasts within this time frame. The reasons which cause these discrepancies still need to be clarified by future experiments. It is, however, conceivable that the differentiation potential of UC-derived cells strongly depends on their location within the UC tissue. Thus, Suzdal’tseva et al. [57] and Girdlstone et al. [58] reported that only a moderate number of cells isolated from

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vascular and Wharton’s jelly fractions of UC was able to differentiate into osteoblasts. In contrast, cells harvested from perivascular regions showed even higher osteogenic capacity than bone marrow MSC in a comparative study [32]. Another imaginable cause is the position of UCderived stem cells within the stem cell hierarchy. Similar to cells from other extra-embryonic tissues, they represent intermediates between embryonic and adult stem cells [31] and, perhaps for this reason, the most primitive of them prefer the differentiation along the chondrogenic pathway and lack the potential to undergo intramembranous ossification under existing environmental conditions. Both hypotheses are corroborated by the finding that UC contains stem cells of various maturation stages. Whereas the most primitive of them are distributed in subepithelial and intervascular regions, the largest parts of UC, more mature and fully differentiated cells are found in close proximity to the blood vessels [33]. Evidently, cell populations isolated from different compartments of UC tissue contain a various number of primitive stem cells and therefore exhibit unequal proliferative and differentiation capacities [56–59]. However, it cannot be excluded that the development and utilization of novel differentiation procedures adjusted to the specific properties of these primitive stem cells will markedly enhance the efficiency of the osteogenic differentiation. As the majority of isolated cells were not able to complete the osteogenic differentiation under conditions traditionally used for the induction of osteogenesis, we currently propose, in accordance with the statement of International Society for Cellular Therapy [60], to define stem cells harvested from whole UC tissue as multipotent mesenchymal stromal cells. One of the greatest challenges of regenerative medicine is the creation of large stem cell banks containing quality controlled, permanently available and well characterized cell material. Cells harvested from whole human UC are highly proliferative, tolerate cryoconservation with an average survival rate of about 75% in our cell freezing conditions and after thawing can be propagated further, at least over 20 PD before they proliferative activity begins to decline. Therefore, they satisfy all requirements essential for the generation of such stem cell banks. Due to the differences in the procedures of cell isolation and, most notably, cultivation, a correct comparison of the proliferative activity of cells isolated from whole UC and various UC compartments is difficult. Nevertheless, our data corroborate the general assumption [32, 33, 37, 39] that UC-derived cells proliferate markedly faster than MSC harvested from BM, which exhibit an average population doubling time of approx. 4 days [61]. Accordingly, UC cells can be more rapidly expanded to quantities sufficient for cell based therapies or tissue engineering without their

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prolonged exposure to in vitro conditions which increases the risks of cell transformation or evolution of chromosomal abnormalities [62]. The average dose of MSC which is currently applied in clinical trials for the enhancement of hematopoietic engraftment or treatment of graft versus host disease (GvHD) corresponds to 1-2×106 MSC/kg [63]. In our culture conditions this dose, based on the assumption that the average body weight of an adult man corresponds to ca. 80 kg, can be produced within 10–14 days, if 1×105 cells are initially seeded. Furthermore, our data verify that the choice of individual parameters for cell propagation, e.g. duration of cell expansion and cell seeding density, has a substantial impact on the quality of obtained cell populations. Previous reports, evaluating critical parameters for bone marrow MSC expansion, have proposed that very low seeding density of 10 and 50 cells/cm2 preserves the proliferative capacity and stemness of these stem cells [64, 65]. Our data indicate, however, that the expansion of UC-derived cells using seeding density of even 500 cells/cm2 is not recommendable over more than 2 passages (approx. 12 PD), mainly because of the rapid loss of cell viability. We believe that cells collected from UC need, in contradistinction to bone marrow MSC, a higher frequency of cell-cell contacts or, more likely, an appropriate concentration of secreted bioactive substances acting in an autocrine or paracrine manner to survive under the ex vivo culture conditions over a prolonged time. The high concentrations of trophic growth factors, such as EGF, PDGFs, aFGF, bFGF, IGF-I and TGF-βs found in the Wharton’s jelly of human UC [66] and the data of our experiments support this assumption. Cells derived from whole UC tissue synthesize the above-named growth factors as well as numerous other trophic mediators including neurotrophins and factors which facilitate angiogenesis and hematopoiesis as detected by RayBio® Human Cytokine Antibody Array and human bFGF immunoassay. This finding is also consistent with data published by Lu-Lu et al. [67] and Hiroyama et al. [68] which show that human UC-derived cells support hematopoiesis in co-cultures with BM-derived CD34+ cells and maintain primate embryonic stem cells in a state capable of producing hematopoietic cells. In conclusion, cells collected from whole UC are highly proliferative, possess at least a distinct chondrogenic and adipogenic differentiation potential and synthesize a broad spectrum of trophic factors. All these features make them prime candidates for future applications in the field of regenerative medicine. Nevertheless, further studies are needed to improve and adjust the protocols which are already employed for adult MSC expansion and differentiation to specific properties and requirements of the primitive stem cells collected from UC. More importantly, these protocols should be standardized to prevent the

29

discrepancies between the data collected by different research groups and to encourage a rapid translation of UC cell-based therapies into clinical practice. Acknowledgements The authors would like to thank Martin Pähler for the technical assistance and accomplishment of RT-PCR analysis, Dr. Johanna Walter for the help in the evaluation of Human Cytokine Antibody Array and Prof. DDr. Martijn van Griensven from the Ludwig Boltzmann Institute for Experimental and Clinical Traumatology (Vienna, Austria) for kindly provided adipose tissue-derived MSC. This study was supported by a grant from the German Research Foundation (Project number KA 1784/5).

References 1. Friedenstein, A. J., Piatetzky-Shapiro, I. I., & Petrakova, K. V. (1966). Osteogenesis in transplants of bone marrow cells. Journal of Embryology & Experimental Morphology, 16(3), 381–390. 2. Pittenger, M. F., Mackay, A. M., Beck, S. C., et al. (1999). Multilineage potential of adult human mesenchymal stem cells. Science, 284, 143–147. 3. Dennis, J. E., Merriam, A., Awadallah, A., Yoo, J. U., Johnstone, B., & Caplan, A. I. (1999). A quadripotential mesenchymal progenitor cell isolated from the marrow of an adult mouse. Journal of Bone and Mineral Research, 14, 700–709. 4. Choong, P. F., Mok, P. L., Cheong, S. K., Leong, C. F., & Then, K. Y. (2007). Generating neuron-like cells from BM-derived mesenchymal stromal cells in vitro. Cytotherapy, 9(2), 170–183. 5. Xu, R., Jiang, X., Guo, Z., et al. (2008). Functional analysis of neuron-like cells differentiated from neural stem cells derived from bone marrow stroma cells in vitro. Cellular and Molecular Neurobiology, 28(4), 545–558. 6. Sun, Y., Chen, L., Hou, X. G., et al. (2007). Differentiation of bone marrow-derived mesenchymal stem cells from diabetic patients into insulin-producing cells in vitro. Chinese Medical Journal (Engl), 120(9), 771–776. 7. Saulnier, N., Lattanzi, W., Puglisi, M. A., et al. (2009). Mesenchymal stromal cells multipotency and plasticity: induction toward the hepatic lineage. European Review for Medical and Pharmacological Sciences, 13(1), 71–78. 8. Glennie, S., Soeiro, I., Dyson, P. J., Lam, E. W., & Dazzi, F. (2005). Bone marrow mesenchymal stem cells induce division arrest anergy of activated T cells. Blood, 105(7), 2821–2827. 9. Kode, J. A., Mukherjee, S., Joglekar, M. V., & Hardikar, A. A. (2009). Mesenchymal stem cells: immunobiology and role in immunomodulation and tissue regeneration. Cytotherapy, 11(4), 377–391. 10. Siegel, G., Schäfer, R., & Dazzi, F. (2009). The immunosuppressive properties of mesenchymal stem cells. Transplantation, 87(9), 45–49. 11. Jones, E., & McGonagle, D. (2008). Human bone marrow mesenchymal stem cells in vivo. Rheumatology (Oxford), 47, 126–131. 12. Dominici, M., Le Blanc, K., Mueller, I., et al. (2006). Minimal criteria for defining multipotent mesenchymal stromal cells. The international society for cellular therapy position statement. Cytotherapy, 8, 315–317. 13. Hui, J. H., Ouyang, H. W., Hutmacher, D. W., Goh, J. C., & Lee, E. H. (2005). Mesenchymal stem cells in musculoskeletal tissue engineering: a review of recent advances in National University of Singapore. Annals of the Academy of Medicine, Singapore, 34(2), 206–212. 14. Kraus, K. H., & Kirker-Head, C. (2006). Mesenchymal stem cells and bone regeneration. Veterinary Surgery, 35(3), 232–242.

30 15. Mobasheri, A., Csaki, C., Clutterbuck, A. L., Rahmanzadeh, M., & Shakibaei, M. (2009). Mesenchymal stem cells in connective tissue engineering and regenerative medicine: applications in cartilage repair and osteoarthritis therapy. Histology and Histopathology, 24(3), 347–366. 16. Syková, E., Jendelová, P., Urdzíková, L., Lesný, P., & Hejcl, A. (2006). Bone marrow stem cells and polymer hydrogels-two strategies for spinal cord injury repair. Cellular and Molecular Neurobiology, 26(7-8), 1113–1129. 17. Perin E. (2004). Transendocardial injection of autologous mononuclear bone marrow cells in end-stage ischemic heart failure patients: one-year follow-up. International Journal of Cardiology, 95(Suppl 1), S45–S46. 18. Stamm, C., Kleine, H. D., Westphal, B., et al. (2004). CABG and bone marrow stem cell transplantation after myocardial infarction. Thoracic and Cardiovascular Surgeon, 52(3), 152–158. 19. Lazarus, H. M., Koc, O. N., Devine, S. M., et al. (2005). Cotransplantation of HLA-identical sibling culture-expanded mesenchymal stem cells and hematopoietic stem cells in hematologic malignancy patients. Biology of Blood and Marrow Transplantation, 11(5), 389–398. 20. Fouillard, L., Chapel, A., Bories, D., et al. (2007). Infusion of allogeneic-related HLA mismatched mesenchymal stem cells for the treatment of incomplete engraftment following autologous haematopoietic stem cell transplantation. Leukemia, 21(3), 568–570. 21. Horwitz, E. M., Prockop, D. J., Gordon, P. L., et al. (2001). Clinical responses to bone marrow transplantation in children with severe osteogenesis imperfecta. Blood, 97(5), 1227–1231. 22. Le Blanc, K., Rasmusson, I., Sundberg, B., et al. (2004). Treatment of severe acute graft-versus-host disease with third party haploidentical mesenchymal stem cells. Lancet, 363(9419), 1439–1441. 23. Ringdén, O., Uzunel, M., Rasmusson, I., et al. (2006). Mesenchymal stem cells for treatment of therapy-resistant graft-versushost disease. Transplantation, 81(10), 1390–1397. 24. Majumdar, M. K., Thiede, M. A., Haynesworth, S. E., Bruder, S. P., & Gerson, S. L. (2000). Human marrow-derived mesenchymal stem cells (MSCs) express hematopoietic cytokines and support long-term hematopoiesis when differentiated toward stromal and osteogenic lineages. Journal of Hematotherapy & Stem Cell Research, 9(6), 841–848. 25. Caplan, A. I., & Dennis, J. E. (2006). Mesenchymal stem cells as trophic mediators. Journal of Cellular Biochemistry, 98(5), 1076–1084. 26. Lda Meirelles, S., Fontes, A. M., Covas, D. T., & Caplan, A. I. (2009). Mechanisms involved in the therapeutic properties of mesenchymal stem cells. Cytokine & Growth Factor Reviews, 20 (5-6), 419–427. 27. Pittenger, M. F., Mackay, A. M., Beck, S. C., et al. (1999). Multilineage potential of adult human mesenchymal stem cells. Science, 284(5411), 143–147. 28. Caplan, A. I. (2009). Why are MSCs therapeutic? New data: new insight. The Journal of Pathology, 217(2), 318–324. 29. Kuo, C. K., Li, W. J., Mauck, R. L., & Tuan, R. S. (2006). Cartilage tissue engineering: its potential and uses. Current Opinion in Rheumatology, 18(1), 64–73. 30. da Silva Meirelles, L., Chagastelles, P. C., & Nardi, N. B. (2006). Mesenchymal stem cells reside in virtually all post-natal organs and tissues. Journal of Cell Science, 119(Pt 11), 2204–2213. 31. Marcus, A. J., & Woodbury, D. (2008). Fetal stem cells from extra-embryonic tissues: do not discard. Journal of Cellular and Molecular Medicine, 12(3), 730–742. 32. Baksh, D., Yao, R., & Tuan, R. S. (2007). Comparison of proliferative and multilineage differentiation potential of human mesenchymal stem cells derived from umbilical cord and bone marrow. Stem Cells, 25, 1384–1392.

Stem Cell Rev and Rep (2011) 7:17–31 33. Can, A., & Karahuseyinoglu, S. (2007). Concise review: human umbilical cord stroma with regard to the source of fetus-derived stem cells. Stem Cells, 25(11), 2886–2895. 34. Weiss, M. L., & Troyer, D. L. (2008). Wharton’s jelly-derived cells are a primitive stromal cell population. Stem Cells, 26(3), 591–599. 35. Kita, K., Gauglitz, G. G., Phan, T. T., Herndon, D. N., Jeschke, M. G. (2009). Isolation and characterization of mesenchymal stem cells from the sub-amniotic human umbilical cord lining membrane. Stem Cells Dev, Jul 27, [Epub ahead of print]. 36. Covas, D. T., Siufi, J. L., Silva, A. R., & Orellana, M. D. (2003). Isolation and culture of umbilical vein mesenchymal stem cells. Brazilian Journal of Medical and Biological Research, 36, 1179–1183. 37. Sarugaser, R., Lickorish, D., Baksh, D., Hosseini, M. M., & Davies, J. E. (2005). Human umbilical cord perivascular (HUCPV) cells: a source of mesenchymal progenitors. Stem Cells, 23, 220–229. 38. Weiss, M. L., Medicetty, S., & Bledsoe, A. R. (2006). Human umbilical cord matrix stem cells: preliminary characterization and effect of transplantation in a rodent model of Parkinson's disease. Stem Cells, 24(3), 781–792. 39. La Rocca, G., Anzalone, R., Corrao, S., et al. (2009). Isolation and characterization of Oct-4+/HLA-G + mesenchymal stem cells from human umbilical cord matrix: differentiation potential and detection of new markers. Histochemistry and Cell Biology, 131, 267–282. 40. Brockhoff, G. (2007). Grundlagen, Methoden und klinische Anwendungen der Durchflusszytometrie. In U. Sack, A. Tárnok, & G. Rothe (Eds.), Zelluläre Diagnostik (Vol. 1, pp. 604–646). Basel: Karger. 41. Majore, I., Moretti, P., Hass, R., Kasper, C. (2009). Identification of subpopulations in mesenchymal stem cell-like cultures from human umbilical cord. Cell Commun Signal. http://www.bio signaling.com/content/7/1/6. Accessed 20 March 2009. 42. Martinez, C., Hofmann, T. J., Marino, R., Dominici, M., & Horwitz, E. M. (2007). Human bone marrow mesenchymal stromal cells express the neural ganglioside GD2: a novel surface marker for the identification of MSCs. Blood, 109(10), 4245–4248. 43. Tsutsumi, S., Shimazu, A., Miyazaki, K., et al. (2001). Retention of multilineage differentiation potential of mesenchymal cells during proliferation in response to FGF. Biochemical and Biophysical Research Communications, 288(2), 413–419. 44. Bianchi, G., Banfi, A., Mastrogiacomo, M., et al. (2003). Ex vivo enrichment of mesenchymal cell progenitors by fibroblast growth factor 2. Experimental Cell Research, 287(1), 98–105. 45. Thomson, J. A., Itskovitz-Eldor, J., Sander, S., et al. (1998). Embryonic stem cell lines derived from human blastocysts. Science, 282(5391), 1145–1147. 46. Gronthos, S., Fitter, S., Diamond, P., et al. (2007). A novel monoclonal antibody (STRO-3) identifies an isoform of tissue nonspecific alkaline phosphatase expressed by multipotent bone marrow stromal stem cells. Stem Cells and Development, 16(6), 953–963. 47. Qiao, C., Xu, W., Zhu, W., Hu, J., Qian, H., Yin, Q., et al. (2008). Human mesenchymal stem cells isolated from the umbilical cord. Cell Biology International, 32, 8–15. 48. Zhang, Z. Y., Teoh, S. H., Chong, M. S., Schantz, J. T., Fisk, N. M., Choolani, M. A., et al. (2009). Superior osteogenic capacity for bone tissue engineering of fetal compared with perinatal and adult mesenchymal stem cells. Stem Cells, 27, 126–137. 49. Raval, A., Puri, N., Rath, P. C., & Saxena, R. K. (1998). Cytokine regulation of expression of class I MHC antigens. Experimental & Molecular Medicine, 30(1), 1–13. 50. Wang, H. S., Hung, S. C., Peng, S. T., Huang, C. C., Wei, H. M., Guo, Y. J., et al. (2004). Mesenchymal stem cells in the Wharton's jelly of the human umbilical cord. Stem Cells, 22, 1330–1337.

Stem Cell Rev and Rep (2011) 7:17–31 51. Jo, C. H., Kim, O. S., Park, E. Y., Kim, B. J., Lee, J. H., Kang, S. B., et al. (2008). Fetal mesenchymal stem cells derived from human umbilical cord sustain primitive characteristics during extensive expansion. Cell and Tissue Research, 334, 423–433. 52. Sarugaser, R., Ennis, J., Stanford, W. L., & Davies, J. E. (2009). Isolation, propagation, and characterization of human umbilical cord perivascular cells (HUCPVCs). Methods in Molecular Biology, 482, 269–279. 53. Chung, C., & Burdick, J. A. (2009). Influence of three-dimensional hyaluronic acid microenvironments on mesenchymal stem cell chondrogenesis. Tissue Engineering. Part A, 15(2), 243–254. 54. Wu, S. C., Chang, J. K., Wang, C. K., Wang, G. J., & Ho, M. L. (2010). Enhancement of chondrogenesis of human adipose derived stem cells in a hyaluronan-enriched microenvironment. Biomaterials, 31(4), 631–640. 55. Diao, Y., Ma, Q., Cui, F., & Zhong, Y. (2008). Human umbilical cord mesenchymal stem cells: Osteogenesis in vivo as seed cells for bone tissue engineering. Journal of Biomedical Materials Research. Part A, 91(1), 123–131. 56. Hou, T., Xu, J., Wu, X., et al. (2009). Umbilical cord Wharton's Jelly: a new potential cell source of mesenchymal stromal cells for bone tissue engineering. Tissue Engineering. Part A, 15(9), 2325–2334. 57. Suzdal'tseva, Y. G., Burunova, V. V., Vakhrushev, I. V., Yarygin, V. N., & Yarygin, K. N. (2007). Capability of human mesenchymal cells isolated from different sources to differentiation into tissues of mesodermal origin. Bulletin of Experimental Biology and Medicine, 143, 114–121. 58. Girdlestone, J., Limbani, V. A., Cutler, A. J., & Navarrete, C. V. (2009). Efficient expansion of mesenchymal stromal cells from umbilical cord under low serum conditions. Cytotherapy, 11(6), 738–748. 59. Ishige, I., Nagamura-Inoue, T., Honda, M. J., et al. (2009). Comparison of mesenchymal stem cells derived from arterial, venous, and Wharton's jelly explants of human umbilical cord. International Journal of Hematology, 90(2), 261–269.

31 60. Horwitz, E. M., Le Blanc, K., Dominaci, M., et al. (2005). Clarification of the nomenclature for MSC: The international society for cellular therapy position statement. Cytotherapy, 7(5), 393–395. 61. Suva, D., Garavaglia, G., Menetrey, J., et al. (2004). Nonhematopoietic human bone marrow contains long-lasting, pluripotential mesenchymal stem cells. Journal of Cellular Physiology, 198(1), 110–118. 62. Rubio, D., Garcia-Castro, J., Martín, M. C., et al. (2005). Spontaneous human adult stem cell transformation. Cancer Research, 65(8), 3035–3039. 63. Battiwalla, M., & Hematti, P. (2009). Mesenchymal stem cells in hematopoietic stem cell transplantation. Cytotherapy, 11(5), 503–515. 64. Colter, D. C., Sekiya, I., & Prockop, D. J. (2001). Identification of a subpopulation of rapidly self-renewing and multipotential adult stem cells in colonies of human marrow stromal cells. Proceedings of the National Academy of Sciences of the United States of America, 98(14), 7841–7845. 65. Sotiropoulou, P. A., Perez, S. A., Salagianni, M., Baxevanis, C. N., & Papamichail, M. (2006). Characterization of the optimal culture conditions for clinical scale production of human mesenchymal stem cells. Stem Cells, 24(2), 462–471. 66. Małkowski, A., Sobolewski, K., Jaworski, S., & Bańkowski, E. (2008). TGF-beta binding in human Wharton's jelly. Molecular and Cellular Biochemistry, 311(1-2), 137–143. 67. Lu, L. L., Liu, Y. J., Yang, S. G., et al. (2006). Isolation and characterization of human umbilical cord mesenchymal stem cells with hematopoiesis-supportive function and other potentials. Haematologica, 91(8), 1017–1026. 68. Hiroyama, T., Sudo, K., Aoki, N., et al. (2008). Human umbilical cord-derived cells can often serve as feeder cells to maintain primate embryonic stem cells in a state capable of producing hematopoietic cells. Cell Biology International, 32(1), 1–7.