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First published online August 24, 2006
Stem Cells Vol. 24 No. 12 December 2006, pp. 2753 -2765
doi:10.1634/stemcells.2006-0108; www.StemCells.com
© 2006 AlphaMed Press

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TISSUE-SPECIFIC STEM CELLS

Presence of Mesenchymal Stem Cells in Human Bone Marrow After Exposure to Chemotherapy: Evidence of Resistance to Apoptosis Induction

Lutz P. Mueller, Jana Luetzkendorf, Thomas Mueller, Katrin Reichelt, Heike Simon, Hans-Joachim Schmoll

Department of Medicine IV, Hematology/Oncology, Centre for Applied Medical and Human-Biological Research, Martin-Luther-University Halle-Wittenberg, Halle, Germany

Key Words. MSCs • Chemotherapy • Resistance • Apoptosis

Correspondence: Lutz P. Mueller, M.D., Department of Medicine IV, Hematology/Oncology, Centre for Applied Medical and Human-Biological Research (ZAMED), Martin-Luther-University Halle-Wittenberg, Ernst-Grube-Strasse 40, D-06120 Halle, Germany. Telephone: 49-345-5577278; Fax: 49-345-5572950; e-mail: lutz.mueller{at}medizin.uni-halle.de

Received February 24, 2006; accepted for publication August 3, 2006.
First published online in STEM CELLS EXPRESS   August 24, 2006.


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 Acknowledgments
 References
 
For various potential clinical applications, the use of autologous human MSCs (hMSCs) would be favorable. In vitro observations suggested that hMSCs are resistant for chemotherapeutic substances; however, no data exist on the characteristics of hMSCs from bone marrow (BM) of chemotherapeutically treated patients. Here, we analyzed the character of hMSCs derived from chemotherapy-exposed BM and the in vitro response of hMSCs to chemotherapeutic substances. Colony-forming units-fibroblast (CFU-Fs) were isolated from BM aspirates of patients after high-dose or standard chemotherapy and of donors with unaffected BM. CFU-Fs from chemotherapy-exposed and unaffected BM contained hMSCs with similar phenotype, proliferation capacity, and differentiation potential. No obvious influence of age, sex, or time since chemotherapy exposure on the presence and characteristics of hMSCs was observed. In vitro, hMSCs showed a significant resistance for cisplatin, vincristine, and etoposide compared with sensitive tumor cell lines, particularly at apoptosis-inducing doses. The phenotype and differentiation potential of hMSCs was not altered by genotoxic treatment under clinically relevant conditions in vitro. hMSCs showed an elevated threshold for cisplatin-induced apoptosis, which was characterized by a lack of caspase-9 activity in apoptotic cells. In vitro exposure of hMSCs to cisplatin, vincristine, and etoposide resulted in an increased p53 expression, independent of apoptosis induction. We conclude that hMSCs can be isolated from chemotherapy-exposed BM in sufficient number and quality for potential clinical applications in chemotherapeutically treated patients. Our data suggest that an elevated apoptotic threshold contributes not only to the persistence of hMSCs in the BM after chemotherapy but also to their lifelong presence in the adult BM.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 Acknowledgments
 References
 
MSCs are currently receiving much attention for their projected clinical use. Originally described as fibroblast-like cells derived from the bone marrow (BM) [1], in vitro identification of human MSCs (hMSCs) relies on their proliferation as colony-forming units-fibroblast (CFU-Fs), their multipotent mesodermal differentiation, and their characteristic phenotype of positive and negative surface markers [2]. These characteristics distinguish hMSCs from the more heterogeneous population of stromal cells [3]. Proposed therapeutic applications involve the transplantation of native [4, 5] or genetically modified [6] hMSCs. Although the transplantation of allogeneic hMSCs appears to be safe [4, 5], the use of autologous hMSCs is favorable in several applications for ethical and practical reasons. No systematic data exist on the availability and quality of hMSCs after chemotherapeutic treatment of the donor.

After allogeneic hematopoietic stem cell transplantation, the majority of hMSCs in the BM seem to be of recipient origin [7], although some engraftment of allogeneic hMSCs may occur [8]. This suggests that the putative in vivo correlate of hMSCs should be resistant against chemotherapeutic damage. This assumption is supported by a recent study reporting an in vitro resistance of hMSCs for chemotherapeutic substances [9]. Considering the proapoptotic effects of most chemotherapeutic drugs, the suggested resistance of hMSCs against drug-caused genotoxic damage will probably involve a resistance to induction of apoptosis. Apoptosis can be mediated by intracellular signals such as DNA damage. Downstream signals include proteolytic cleavage of caspase-9 yielding enzymatically active fragments. These in turn activate caspase-3, which executes apoptotic cell death by inactivating key enzymes of DNA synthesis and repair such as poly(ADP-ribose) polymerase (PARP) [10]. A decisive regulator of apoptosis is p53, which can induce cell cycle arrest or apoptosis upon DNA damage, depending on the intracellular context [11, 12].

An investigation of hMSCs from chemotherapeutically treated patients will help to define criteria regarding the quality of hMSCs and the donor selection for clinical applications. This prompted us to study the character of hMSCs derived from BM exposed to chemotherapy and to analyze the in vitro response of hMSCs to chemotherapeutic substances. Our data demonstrate that hMSCs can be isolated from BM aspirates of patients after high-dose or standard chemotherapy. The in vitro studies show that hMSCs are resistant to chemotherapeutic substances of different mechanisms of action and maintain their characteristics despite chemotherapeutic treatment. hMSCs showed an elevated apoptotic threshold for cisplatin that was characterized by a lack of caspase-9 activity in apoptotic hMSCs. In vitro exposure of hMSCs to chemotherapeutic substances of different mechanisms of action resulted in an increased expression of p53, independent of apoptosis induction. We conclude that hMSCs can be isolated in sufficient number and quality from chemotherapy-exposed BM for potential clinical applications. Our data suggest that a high apoptotic threshold contributes to the persistence of hMSCs in the BM despite chemotherapeutic damage. We hypothesize that similar mechanisms contribute to the survival mechanisms warranting the lifelong presence of hMSCs in the adult human BM.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 Acknowledgments
 References
 
hMSC Isolation and Propagation
Twenty ml of BM was aspirated from the iliac crest using sodium citrate as an anticoagulant. Following centrifugation at 300g, pelleted BM cells were resuspended with phosphate-buffered saline (PBS), overlaid on Biocoll (Biochrom, Berlin, http://www.biochrom.de), and centrifuged for 30 minutes at 300g; the interface containing BM mononuclear cells (BMMNCs) was harvested and resuspended with growth medium, and the cell number was counted. After centrifugation at 300g, cells were plated in growth medium on fibronectin-coated culture flasks at 1–2.5 x 105 BMMNCs per cm2. For each donor, at least two culture flasks were plated so that CFU-F counting could be performed at least in duplicates. Fibronectin coating was achieved by covering culture materials 5 minutes with low-glucose Dulbecco's modified Eagle's medium (DMEM) (Biochrom) containing 0.5 µg/ml fibronectin (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com) followed by sterile air drying. Growth medium was composed following a protocol by Reyes et al. [13] and contained 60% DMEM, 40% modified Czapek Dox broth 201, 5 µg/ml apo-transferrin, 5 ng/ml selenous acid, 5 µg/ml linoleic acid, 5 µg/ml bovine insulin, 100 µM ascorbic acid 2-phosphate, 1 nM dexamethasone, 10 ng/ml platelet-derived growth factor-BB, 10 ng/ml epidermal growth factor (all Sigma-Aldrich), 100 U/ml penicillin/100 µg/ml streptomycin and 2% fetal calf serum (FCS) selected for optimal growth (all Invitrogen, Grand Island, NY, http://www.invitrogen.com). CFU-F number was counted microscopically in at least two flasks of each donor when >50% of colonies had reached at least 80% confluence and was calculated as the mean CFU-Fs per 106 BMMNCs. hMSCs were passaged with 0.05% trypsin/0.02% EDTA (Biochrom) upon reaching 50% confluence and replated at 200 cells per cm2 directly or after cryopreservation. For single cell-derived cultures, 100 hMSCs were suspended after limited dilution in 20 ml of growth medium and plated in a 96-well plate in 200-µl aliquots. Cultures derived from a single cell were selected by regular microscopic assessment and used for further analysis.

Donor Characteristics
Bone marrow samples were obtained from donors with a history of previous chemotherapy (chemotherapy-exposed BM; n = 17) and a donor group with no history of chemotherapy or malignant BM infiltration (unaffected BM; n = 16). All donors underwent diagnostic BM aspiration for various clinical conditions, for diagnostic staging of malignant disease, or in preparation of allogeneic stem cell donation after obtaining informed consent. The study protocol was approved by the Institutional Review/Ethics Board. Donor characteristics are shown in Table 1.


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  		Table 1. Characteristics of investigated donors and number of isolated CFU-Fs

 
Sensitive Tumor Cells
The testicular germ cell tumor (TGCT) cell lines 2102EP and H12.1, which both exhibit characteristics of embryonal carcinoma (EC) cells [14], were cultured in RPMI 1640 (Biochrom)/10% FCS. Medium was exchanged daily, and cells were passaged after 2 days.

Flow Cytometry
The following mouse antihuman antibodies were used: Simultest Control y1/y1, anti-CD11c-phycoerythrin (PE) (clone S-HCL-3), anti-CD13-PE (clone L138), anti-CD14-fluorescein isothiocyanate (FITC) (clone M{Phi}P9), anti-CD45-FITC (clone 2D1), anti-human leukocyte antigen (HLA)-DR-FITC (clone L243) (all from Becton, Dickinson and Company, San Jose, CA, http://www.bd.com); anti-CD29-PE (clone MAR4), anti-CD34-FITC (clone 581), anti-CD44-PE (clone G44–26), anti-CD166-PE (clone 3A6), anti-glycophorin-A-PE (clone GA-R2) (all from BD Pharmingen, San Diego, http://www.bdbiosciences.com/pharmingen); and anti-CD105-FITC (clone SN6) (Serotec Ltd., Oxford, U.K., http://www.serotec.com). The expression of these markers was unaffected by trypsin detachment compared with detachment by 1 mM EDTA/EGTA (all from Sigma-Aldrich). hMSCs were trypsinized, PBS-washed, incubated with primary labeled antibody 15 minutes at 4°C and washed with PBS. Analysis was performed on a FACSCalibur using CellQuest software (all from Becton, Dickinson and Company).

Growth Kinetics of hMSCs
Growth of hMSCs was analyzed after seeding passage 1 cells with 200 cells per cm2 with subsequent passaging and cell counting at 50% confluence for up to 10 passages. Population doubling (PD) was calculated using the formula PD = lg[(n cells harvested)/(n cells initially plated)]/lg2 [15]. Population doubling time (PDT) was calculated for each passage using the formula PDT = t/PD, where t represents the time between initial plating and harvest for the respective passage.

Differentiation Induction in hMSCs
hMSCs were plated at first passage at 200 cells per cm2 and incubated in differentiation medium after reaching confluence. Osteogenic differentiation medium consisted of DMEM, 200 µM ascorbic acid 2-phosphate, 1 µM dexamethason, 10 mM glycerol 3-phosphate (all from Sigma-Aldrich), and 10% FCS. Medium was changed every 3 days. Adipogenic differentiation was induced with adipogenic induction medium consisting of DMEM, 50 µM dexamethasone, 10 µg/ml bovine insulin, 100 µM indomethacin, 500 µM 3-isobutyl-1-methylxanthine (Sigma-Aldrich), and 10% FCS for 2 days, followed by adipogenic differentiation medium consisting of DMEM, 50 µM dexamethason, 10 µg/ml bovine insulin, 5 µM rosiglitazone (Alexis, Corporation, Lausen, Switzerland, http://www.axxora.com), and 10% FCS for 3 days. This cycle was repeated three times. When morphological signs of differentiation were visible, cells were fixed for 20 minutes in 2% formaldehyde (Sigma-Aldrich). After PBS washing, calcium deposition was stained for 20 minutes with alizarin red S, pH 4 (1% in aqua dest; Sigma-Aldrich), followed by washes with distilled water. For staining of intracellular lipid droplets, fixed cells were washed with 50% ethanol followed by 20 minutes of staining with oil red (0.7% in 50% ethanol; Sigma-Aldrich) and subsequently washed with 50% ethanol and distilled water.

Reverse Transcription-Polymerase Chain Reaction
hMSCs were rinsed with PBS and detached with trypsin/EDTA, and poly(A) RNA was isolated from 0.5–10 x 106 cells by centrifugation at 300g and lysis with 1.2 ml of TRIzol (Gibco-BRL, Gaithersburg, MD, http://www.gibcobrl.com) for 5 minutes. After mixing with 200 µl of chloroform (Carl Roth, Karlsruhe, Germany, http://www.carl-roth.de), 3 minutes of incubation at 21°C, and 15 minutes of centrifugation at 16,000g, 4°C, clear supernatant was obtained, mixed with an equal volume isopropanol (Carl Roth), and incubated for 24 hours at –20°C followed by 15 minutes of centrifugation at 16,000g, 4°C. The resulting RNA pellets were washed with 75% ethanol (Carl Roth), diluted in 20–50 µl of diethylprocarbonate-treated water, and treated with DNase using the DNAfree kit (Ambion, Huntingdon, U.K., http://www.ambion.com) following the manufacturer's instructions. Reverse transcription-polymerase chain reaction (RT-PCR) was performed using the GeneAmp RNA PCR kit (Applied BioSystems, Foster City, CA, http://www.appliedbiosystems.com) according to the manufacturer's instructions, with the following modifications: reverse transcription was run in 10-µl samples containing 500 ng of RNA. The transcription program consisted of 10 minutes of initial extension, 23°C; 30 minutes of reverse transcription, 42°C; 5 minutes of denaturation, 99°C. Negative controls were run without reverse transcriptase. Amplification was performed in 50-µl samples by 35 PCR cycles consisting of 60 seconds of denaturation, 95°C; 45 seconds of annealing, Tannealing; 80 seconds of synthesis, 72°C; 5 minutes of initial denaturation, 95°C; 5 minutes of final synthesis, 72°C. The following primer pairs were used: glyceraldehyde-3-phosphate dehydrogenase (195 base pairs [bp]), 5'-ccatggagaaggctgggg/5'-caaagttgtcatggatgacc; osteocalcin (OC) (274 bp), 5'-gtgcagagtccagcaaaggt/5'-ctggagaggagcagaactgg; peroxisome proliferator-activated receptor-{gamma} (PPAR-{gamma}) (351bp), 5'-gctgttatgggtgaaactctg/5'-ataaggtggagatgcaggctc (all Tannealing 63°C). Twenty µl of amplification product was mixed with 5 µl of loading buffer (Fermentas; Vilnius, Lithuania, http://www.fermentas.com) and run on 2% agarose (Cambrex, Walkersville, MD, http://www.cambrex.com) gels including molecular weight markers (Fermentas).

Cytotoxicity Testing
Cytotoxicity was tested using the sulforhodamin-B assay [16]. hMSC and TGCT cells were seeded into 96-well plates on day 1 in growth medium or RPMI/10% FCS, respectively. The cell number that had to be seeded to ensure exponential cell growth during the 96-hour duration of the experiment was determined in three independent growth kinetics analysis for hMSCs and TGCT cells, respectively. Using 96-well plates, 1,000 cells per well for hMSCs and 3,000 cells per well for the TGCT cell lines had to be seeded on day 1. Twenty-four hours after plating, media were removed, and hMSCs and TGCT cells were treated for 24 hours with the respective chemotherapeutic substances dissolved in growth medium or RPMI/10% FCS, respectively, at concentrations of 0.01–100 µM cisplatin (Sigma-Aldrich), 0.01–100 µM etoposide (Sigma-Aldrich), and 0.0003–3 µM vincristine (Sigma-Aldrich). Subsequently, medium was removed, and cells were washed with PBS and incubated with substance-free medium. The percentage of surviving cells relative to untreated controls was determined 72 hours after finishing substance treatment. The concentrations that inhibited cell growth by 50% (IC50) and 90% (IC90) were determined for each treatment schedule from semilogarithmic dose-response plots.

In Vitro Model of Resistance
At 50% confluence hMSCs were treated for 2 hours with 3 µM cisplatin or etoposide dissolved in growth medium, rinsed twice with PBS, and incubated in substance-free growth medium. This treatment was repeated four times every 24 hours. Subsequent analysis or differentiation induction was performed 72 hours after the end of treatment.

Apoptosis Detection by Trypan Blue and Annexin V Staining
hMSCs and TGCT cells were treated for 24 hours with cisplatin at the respective IC90 dose. For trypan blue staining detached cells were collected, rinsed with PBS, stained with trypan blue (Sigma-Aldrich), and analyzed by light microscopy. For annexin V/propidium iodide (PI) staining, cells were detached by trypsin/EDTA after end of treatment, rinsed twice with PBS, and stained with the annexin V-FITC apoptosis detection kit I (BD PharMingen). Analysis was performed on a FACSCalibur using CellQuest software (Becton, Dickinson and Company).

Western Blotting
Attached cells were rinsed with PBS and lysed in RIPA buffer (50 mM Tris-HCl [pH 8.0], 150 mM NaCl, 1% NP40, 0.5% sodium deoxycholate, 0.1% SDS, 0.1 mM dithiothreitol [DTT] [all from Sigma-Aldrich]) supplemented with a protease inhibitor (Sigma-Aldrich). For analysis of detached cells, cell culture supernatant was harvested and centrifuged at 300g, and the pellet was lysed in RIPA buffer. Insoluble components were removed by centrifugation, and protein concentrations were measured (Bio-Rad protein assay; Bio-Rad, Hercules, CA, http://www.bio-rad.com). After 5 minutes of boiling in SDS loading buffer (62.5 mM Tris-HCl [pH 6.8], 20% glycerol, 2% SDS, 100 mM DTT, 0.0025% bromphenol blue [all from Sigma-Aldrich]), 40 µg of protein per lane was separated by SDS-polyacrylamide gel electrophoresis and electroblotted onto nitrocellulose membrane (Bio-Rad). Equal protein loading was controlled by Ponceau S staining (Sigma-Aldrich). Membranes were blocked with 5% nonfat dry milk in PBS/0.1% Tween 20 for 1 hour and probed for 2 hours with the following antibodies diluted in PBS/0.1% Tween 20/5% milk at the respective concentrations: mouse anti-actin (0.5 µg/ml; Santa Cruz Biotechnology Inc., Santa Cruz, CA, http://www.scbt.com), mouse anti-caspase-3 (0.5 µg/ml; MBL International Corp., Woburn, MA, http://www.mblintl.com), mouse anti-caspase-9 (0.5 µg/ml; MBL), mouse anti-PARP (1 µg/ml; BD PharMingen), and mouse anti-p53 (0.1 µg/ml; Santa Cruz Biotechnology). Immunocomplexes were visualized by enhanced chemiluminescence (Amersham Pharmacia Biotech, Little Chalfont, U.K., http://www.amersham.com) using horseradish peroxidase-conjugated anti-mouse IgG (Santa Cruz Biotechnology) and Roti-Lumin (Carl Roth).

Caspase Activity
Relative caspase activity was analyzed in a substrate cleavage assay. Cells were treated for 24 hours with cisplatin at the respective IC90 dose. Detached apoptotic cells were discarded, and only adherent cells were analyzed. Cells were washed with PBS, lysed in cell lysis buffer (50 mM Hepes [pH 7.4] [Biochrom], 1% Triton X-100 [Sigma-Aldrich]) and incubated for 10 minutes on ice. All subsequent steps were performed on ice. After 1 minute of centrifugation at 16,000g, 4°C, cytosolic extracts were transferred to fresh tubes, and protein concentrations were measured using a Bio-Rad protein assay (Bio-Rad). Cell extract containing 100 µg of protein was combined with equal volumes of reaction buffer (50 mM Hepes [pH 7.4], 0.1% CHAPS [3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid, Carl Roth], 5 mM EGTA [Carl Roth], 5% glycerol) in a 96-well plate followed by the addition of 5 µl of peptide substrates Ac-VDVAD-pNA, Ac-DEVD-pNA, Ac-IETD-pNA, and Ac-LEHD-pNA (all from BioSource, Camarillo, CA, http://www.biosource.com) for analysis of caspase-2, -3, -8, or -9 activity, respectively. After overnight incubation in the dark at 37°C, samples were read in a microplate reader at 405 nm. Caspase activities were evaluated by the optical density ratio of treated/untreated samples.

Statistical Analysis
Statistical comparisons were performed using SPSS 10.0.5 software (SPSS, Inc., Chicago, http://www.spss.com). An updated t test including Levene testing or a Mann-Whitney test was used depending on the variance. A p value <.05 was considered significant.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 Acknowledgments
 References
 
CFU-Fs Can Be Isolated from Human BM Exposed to Chemotherapy
First, we investigated whether BM exposed to chemotherapy contains CFU-F. In plastic-adherent BMMNCs from all donors except one, CFU-F formation was observed after 3–7 days, and criteria for counting of CFU-F (more than 50% of CFU-F with at least 80% confluence) were reached after similar cultivation time in preparations from chemotherapy-exposed and unaffected BM (12.0 days ± 2.3 days, range 8–17 days vs. 11.3 days ± 2.2 days, range 8–15 days; p = .4). Counting of CFU-F revealed differences in the mean number of CFU-F/106 BMMNCs (Table 1) isolated from unaffected (13.3 ± 12.0) and chemotherapy-exposed BM (7.0 ± 5.7), which did not, however, reach significance (p = .073). The relevance of subgroup analysis was limited because of the small size of the groups and the large standard deviation of CFU-F numbers. However, analysis of CFU-F/106 BMMNCs in the group of donors with unaffected BM regarding male (n = 10; 11.4 ± 7.5) and female (n = 6; 16.3 ± 17.7) donors (p = .54) and donors older (n = 9; 10.2 ± 7.9) and younger (n = 7; 17.1 ± 15.7) than 40 years (p = .27) suggested a higher CFU-F number in younger donors but did not reach significance. In the group of donors with chemotherapy-exposed BM, analysis of CFU-F numbers regarding male (n = 14; 6.4 ± 5.9) and female (n = 3; 9.7 ± 4.6) donors (p = .39), donors older (n = 9; 7.1 ± 6.4) and younger (n = 8; 6.9 ± 5.3) than 40 years (p = .94), donors with (n = 14; 6.8 ± 5.9) and without (n = 3; 8.0 ± 6.0) malignant BM infiltration (p = .78), and donors with BM aspiration less (n = 9; 6.2 ± 5.5) or more (n = 8; 7.9 ± 6.2) than 3 months after the end of chemotherapy (p = .57) did not reveal any obvious differences.

When BM from pairs of donors with unaffected and chemotherapy-exposed BM with similar demographical characteristics (donors 46 vs. 47, 68 vs. 67, and 75 vs. 76) were isolated in parallel by the same investigator, only minor variances in the CFU-F number were observed. Given the differential clinical conditions leading to BM aspiration in both donor groups and the multiple factors with a putative impact on the CFU-F number, a multivariant analysis was not feasible. We hypothesized that chemotherapy-exposed BM contains hMSCs with a character similar to that of unaffected BM.

hMSCs from Chemotherapy-Exposed and Unaffected BM Show Similar Phenotype, Proliferation Capacity, and Multipotent Differentiation Potential
Next, to prove their hMSC character, we analyzed whether CFU-F cells from chemotherapy-exposed BM show typical hMSC phenotype, proliferation capacity, and multipotent differentiation potential. For CFU-F cells from chemotherapy-exposed BM, flow cytometry revealed the following expression pattern, which is characteristic of hMSCs: CD14–, CD34–, HLA-DR–, CD44+, CD105+, CD166+ (Fig. 1A) and CD11c–, CD45–, glycophorin-A–, CD13+, CD29+ (data not shown). An identical phenotype was observed for CFU-F cells from unaffected BM (Fig. 1A). No influence of age, sex, time since chemotherapy exposure, or plating density on the phenotype was obvious (data not shown).


Figure 1
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  		Figure 1. Human MSCs (hMSCs) from chemotherapy-exposed and unaffected bone marrow (BM) show similar phenotype, proliferation capacity, and multipotent differentiation potential. (A): Colony-forming units-fibroblast (CFU-F) cells from unaffected (donor 95) and chemotherapy-exposed (donor 92) BM were harvested at passage 1 and analyzed by flow cytometry. Data are shown as histograms of mean channel fluorescence. Isotype control (no filling) for the respective instrument settings are overlaid on specific fluorescein isothiocyanate- or PE-conjugated antibody (gray filling). Data are representative of ten independent experiments. (B): Growth kinetics of hMSCs from chemotherapy-exposed (red) (donors 47, 63, and 92) and unaffected (blue) (donors 61, 72, and 77) BM from passage 1 (day 0) to passage 10. Each symbol represents the time of passaging for which population doubling and population doubling time were calculated. Note the logarithmic scale of the y-axis. (C): Nonclonal hMSCs from chemotherapy-exposed BM (donor 92) were incubated with differentiation-inducing media for adipogenic or osteogenic differentiation or cultivated in growth medium. Subsequently, samples were stained with oil red or alizarin, pH 4, at the same time and revealed lipid droplets (unfilled arrows) or calcium deposits (arrows) respectively. Light microscopy (microscope: Nikon Eclipse TS100; original magnification x100; camera: Nikon Coolpix 990; software: Nikon View 3.1; processing: identical automatic color balance for both images, cropped size). (D) Non-clonal hMSCs from chemotherapy-exposed BM (donor 67) were submitted to adipogenic or osteogenic differentiation induction, resulting in signs of differentiation, that is, lipid droplets (unfilled arrows) or calcium deposits (arrows), respectively. Light microscopy (Nikon Eclipse TS100; original magnification x200; Nikon Coolpix 990; Nikon View 3.1; cropped size). (E): In the experiment shown in (C), RNA was isolated from the cultures depicted in D after adipogenic and osteogenic differentiation induction and analyzed by reverse transcription-polymerase chain reaction for the respective transcripts. In addition, RNA was harvested from nonclonal undifferentiated hMSCs of the same donor (i.e., donor 67) on day 0 of differentiation induction and analyzed. In comparison, nonclonal, differentiated, and undifferentiated hMSCs from unaffected BM (donor 77) were investigated. Gel images were acquired (Syngene GeneGenius with Syngene GeneSnap software) and processed (cropped size). Data for (C–E) are representative of at least three independent experiments. Abbreviations: a, adipogenic; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; HLA, human leukocyte antigen; o, osteogenic; OC, osteocalcin; PPAR-{gamma}, peroxisome proliferator-activated receptor-{gamma}; u, undifferentiated.

 
Analysis of growth kinetics revealed similar in vitro proliferation capacity over at least 10 passages (Fig. 1B) for hMSCs from chemotherapy-exposed and unaffected BM resulting in similar PD from passage 2 (3.5 ± 0.5 vs. 5.4 ± 3.0; p = .38) through passage 5 (15.2 ± 2.0 vs. 15.6 ± 3.0; p = .85) and passage 10 (26.4 ± 4.5 vs. 26.6 ± 3.9; p = .97), respectively. This corresponds to similar PDT for hMSCs from chemotherapy-exposed and unaffected BM at passage 2 (1.12 days ± 0.3 days vs. 1.02 days ± 0.2 days; p = .67) through passage 5 (1.29 days ± 0.2 days vs. 1.4 days ± 0.1; p = .47) and passage 10 (2.0 days ± 0.5 days vs. 2.0 days ± 0.2 days; p = .89), respectively. As demonstrated by the increasing PDT, proliferation of hMSCs from both chemotherapy-exposed and unaffected BM decreased in the course of in vitro passaging (Fig. 1B). However, the data show that hMSCs from chemotherapy-exposed BM can be expanded in vitro by a factor of 50–100 x 106.

Upon incubation with the respective differentiation-inducing media in vitro, nonclonal hMSCs from all analyzed donors with chemotherapy-exposed BM (n = 13) showed the typical morphological signs of adipogenic and osteogenic differentiation: intracellular lipid droplets that stained by oil red and extracellular calcium deposition that stained by alizarin, pH 4 (Fig. 1C), as seen in hMSCs from all analyzed donors with unaffected BM (n = 7; data not shown). When such morphological signs were detectable (Fig. 1D), analysis of gene expression revealed expression of characteristic adipogenic and osteogenic genes in nonclonal hMSCs from chemotherapy-exposed and unaffected BM (Fig. 1E). Upon induction of differentiation, similar signs of adipogenic and osteogenic differentiation were observed with staining or RT-PCR analysis in clonal, single cell-derived hMSC populations from chemotherapy-exposed (n = 6) and unaffected (n = 3) BM (data not shown). No obvious influence of age, sex, or time since chemotherapy exposure on the differentiation potential of hMSCs was observed. Thus, hMSCs with typical phenotype, proliferation capacity, and multipotent differentiation potential could be isolated from BM after extensive chemotherapy, and we hypothesized that hMSCs are resistant to the toxic effects of chemotherapeutic substances.

hMSCs Are Resistant to Chemotherapeutic Substances of Different Mechanisms of Action In Vitro
To evaluate this hypothesis, we analyzed the in vitro response of hMSCs from chemotherapy-exposed and unaffected BM to chemotherapeutic substances of different mechanisms of action in comparison with TGCT cell lines of known high sensitivity. hMSCs from unaffected and chemotherapy-exposed BM showed a reduced sensitivity for cisplatin, vincristine, and etoposide compared with the TGCT cell line 2102EP (Fig. 2). The relevance of a statistical comparison of IC values was limited since data from multiple analysis of a single cell line were compared with data from primary cells of multiple sources. However, analysis suggested that the resistance in hMSCs was most pronounced at higher doses, reflected by significantly higher IC90 values for hMSCs (Fig. 2D). A resistance of hMSCs from chemotherapy-exposed and unaffected BM was also seen in comparison with the TGCT cell line H12.1, although the differences here were only significant in comparison with the IC90 values of H12.1 for cisplatin (3.5 µM ± 0.9 µM; p < .006) and vincristine (0.0028 µM ± 0.0002 µM; p < .0001) but not for etoposide (16.7 µM ± 7.5 µM; p = .24 vs. hMSCs from unaffected BM; p < .0001 vs. hMSCs from chemotherapy-exposed BM). Relative to hMSCs from unaffected BM higher IC90 values were observed for hMSCs from chemotherapy-exposed BM for all substances, although these differences only reached significance for etoposide (Fig. 2D). For further analysis, a cisplatin dose of 20 µM was used as the IC90 dose for 24 hours of treatment of hMSCs, representing the mean IC90 for hMSCs from unaffected and chemotherapy-exposed BM (Fig. 2D).


Figure 2
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  		Figure 2. hMSCs are resistant to chemotherapeutic substances of different mechanisms of action in vitro. hMSCs from unaffected ({diamondsuit}) and chemotherapy-exposed ({blacksquare}) BM and the TGCT cell line 2102 EP ({circ}) were treated for 24 hours with cisplatin (A), vincristine (B), and etoposide (C). Seventy-two hours after the end of treatment, cell survival was analyzed in the sulforhodamin-B (SRB) assay. Results of (A–C) are represented as mean ± SD in semilogarithmic dose-response plots. (D): Comparison of the IC50 and IC90 values in hMSCs from unaffected and chemotherapy-exposed BM and in the TGCT cell line 2102EP. IC values were determined from the semilogarithmic dose-response plots of the SRB assay for each experiment and are shown as mean (± SD). If the IC90 dose was not reached, the highest concentration used in the assay is stated and was used for statistical analysis. a p < .05 versus TGCT cell line 2102EP; b p < .05 versus hMSCs from chemotherapy-exposed BM. (A–D): Results from at least three independent experiments for each cell type and substance. Abbreviations: BM, bone marrow; hMSC, human MSC; IC50, concentration that inhibited cell growth by 50%; IC90, concentration that inhibited cell growth by 90%; NA; not available; TGCT, testicular germ cell tumor.

 
These data suggested that hMSCs are resistant to chemotherapy-induced damage. We hypothesized that hMSCs from unaffected BM retain their stem cell characteristics after chemotherapeutic treatment in vitro.

hMSCs Retain Their Stem Cell Character After Genotoxic Treatment In Vitro
In proof of this hypothesis, we analyzed the characteristics of hMSCs from unaffected BM after treatment with cisplatin and etoposide in vitro since both substances exert an apoptosis-inducing, DNA-damaging activity [17, 18]. The treatment (3 µM cisplatin or etoposide for 2 hours, repeated four times every 24 hours) represented a dose corresponding to clinically relevant serum concentrations and a schedule used in current therapies [19–21]. Following a lag phase of 7 days for etoposide and 14 days for cisplatin after the end of treatment (data not shown), treated cells resumed growth at PDs similar to those of untreated cells (Fig. 3A). Flow cytometry revealed an identical phenotype with CD14–, CD34–, CD105+, CD166+ (Fig. 3B) and HLA-DR–, CD11c–, CD45–, glycophorin-A–, CD13+, CD29+, CD44+ (data not shown) before and after in vitro treatment. Upon induction of differentiation, treated hMSCs showed signs of osteogenic and adipogenic differentiation (Fig. 3C).


Figure 3
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  		Figure 3. Human MSCs (hMSCs) retain their stem cell character after genotoxic treatment in vitro. hMSCs from unaffected bone marrow (donor 69) were treated with 3 µM cisplatin or etoposide for 2 hours in vitro, and treatment was repeated four times after 24 hours. Data are representative of independent experiments with hMSCs from two donors. (A): Beginning at the start of treatment (passage 0), growth kinetics was analyzed in comparison with continuously passaged untreated cells, and population doubling was calculated. (B): Phenotype was analyzed by flow cytometry for selected markers in treated and untreated cells 72 hours after end of treatment. Data are shown as histograms of mean channel fluorescence. Isotype control (open) for the respective instrument settings is overlaid on specific fluorescein isothiocyanate- or phycoerythrin-conjugated antibody (filled). (C): Nonclonal treated and untreated hMSCs were incubated with differentiation-inducing media for adipogenic or osteogenic differentiation or cultivated in growth medium, resulting in typical signs of differentiation, that is, lipid droplets (open arrows) stained by oil red or calcium deposits (arrows) stained by alizarin, pH 4, respectively. Light microscopy (Nikon Eclipse TS100; original magnification x100; Nikon Coolpix 990; Nikon View 3.1; cropped size). Abbreviations: a, adipogenic; o, osteogenic; u, undifferentiated.

 
These data supported the hypothesis that hMSCs are resistant to genotoxic damage. As this resistance was most pronounced at high doses (Fig. 2), which typically represent apoptosis-inducing doses, we hypothesized that hMSCs have a higher apoptotic threshold for genotoxic damage compared with sensitive cells.

hMSCs Show an Elevated Threshold for Cisplatin-Induced Apoptosis
To evaluate this assumption, we investigated the induction of apoptosis upon 24 hours of treatment with cisplatin at the respective IC90 doses in hMSCs and in cisplatin-sensitive H12.1 cells. Trypan blue staining of floating cells showed a high amount of unstained cells with membrane blebbing typical of apoptosis (Fig. 4A). The percentage of PI-negative adherent cells expressing the apoptosis marker annexin V increased in treated samples compared with untreated controls (Fig. 4B). In contrast, the percentages of secondary necrotic cells (annexin V- and PI-positive) and primary necrotic cells (PI-positive and annexin V-negative) that result from the enzymatic detachment before staining remained unchanged in treated and untreated samples (Fig. 4B).


Figure 4
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  		Figure 4. hMSCs show an elevated threshold for cisplatin-induced apoptosis. hMSCs and the TGCT cell line H12.1 were treated with cisplatin for 24 hours at the respective IC90 dose, that is, 20 µM for hMSCs (mean of IC90 for hMSCs from unaffected and chemotherapy-exposed bone marrow [BM]) and 3 µM for H12.1. Data represent equal results of three independent experiments. (A): Detached cells were harvested, stained with trypan blue, and analyzed microscopically. Filled arrowheads mark unstained (i.e., apoptotic) cells with typical membrane blebbing. Unfilled arrowheads mark stained (i.e., secondary necrotic) cells. Light microscopy (Nikon Eclipse TS100; original magnification x200; Nikon Coolpix 990; Nikon View 3.1; cropped size). (B): hMSCs from unaffected BM (donor 72) were treated, stained with PI and for annexin V, and analyzed by flow cytometry. Histograms show the cell number in correlation to the mean channel fluorescence for annexin V. Inserted dot blots show the staining for PI and annexin V with the respective percentages of cells in the quadrants. (C): hMSCs from unaffected (donor 95) and chemotherapy-exposed (donor 54) BM, as well as the TGCT cell line H12.1, were treated with cisplatin for 24 hours at the respective doses. Protein lysates of adherent and detached cells were analyzed by Western blotting with the respective antibodies, including ß-actin as a loading control. Molecular weight markers are indicated by arrows. Abbreviations: a, adherent; d, detached; FITC, fluorescein isothiocyanate; hMSC, human MSC; PARP, poly(ADP-ribose) polymerase; PI, propidium iodide; TGCT, testicular germ cell tumor.

 
Western blot analysis revealed that cisplatin treatment at the respective IC90 dose resulted in cleavage of PARP (116 kDa) in hMSCs and TGCT cells, yielding an 85-kDa fragment characteristic for apoptosis execution. Cleavage was most prominent in detached (i.e., apoptotic) cells. Treatment of hMSCs from both unaffected and chemotherapy-exposed BM with cisplatin at the IC90 dose of the cisplatin-sensitive TGCT cell line H12.1 did not result in any PARP cleavage. No differences were observed between hMSCs from chemotherapy-exposed and unaffected BM (Fig. 4C). Treatment with the specific IC90 dose resulted in cleavage of procaspase-9 (46 kDa) and procaspase-3 (32 kDa) into the active fragments of caspase-9 (37 and 35 kDa) and caspase-3 (20 and 17 kDa) in hMSCs and TGCT cells. As for PARP, no active fragments were observed upon treatment of hMSCs with the IC90 dose of the TGCT cell line H12.1. Cleavage of procaspase-9 appeared to be reduced in hMSCs in comparison with the sensitive TGCT cells, whereas cleavage of procaspase-3 was observed at a similar extent. Again, for both hMSCs and TGCT cells, cleavage was most prominent in detached cells, and no differences were observed between hMSCs from chemotherapy-exposed and unaffected BM (Fig. 4C).

These data demonstrated that cisplatin treatment resulted in apoptosis induction in hMSCs, for which, however, a sixfold higher cisplatin concentration was required in comparison with cisplatin-sensitive tumor cells. Conclusively, we hypothesized that an impaired apoptosis induction contributes to the elevated threshold for cisplatin-induced apoptosis in hMSCs.

hMSCs Show Impaired Caspase-9 Activation upon Cisplatin-Induced Apoptosis
To identify possible causes for impaired apoptosis induction, we analyzed the activity of caspases upon apoptosis-inducing cisplatin treatment in a substrate-cleavage assay. For caspase-2 and -3, an increase in the enzymatic activity was similarly observed in hMSCs from unaffected and chemotherapy-exposed BM compared with untreated cells. No increase in the activity of caspase-8 and -9 was seen in hMSCs from unaffected or chemotherapy-exposed BM (Fig. 5A). This is remarkable considering the observed cleavage of procaspase-9 in apoptotic hMSCs (Fig. 4C). The lack of caspase-9 activation in hMSCs upon cisplatin-induced apoptosis was in strong contrast to the increased caspase-9 activity in cisplatin-sensitive H12.1 cells after IC90 cisplatin treatment (Fig. 5B).


Figure 5
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  		Figure 5. Human MSCs (hMSCs) show impaired caspase-9 activation upon cisplatin-induced apoptosis. Caspase activity induced by 24 hours of cisplatin treatment at the respective IC90 dose in hMSCs (20 µM; mean of IC90 for hMSCs from unaffected and chemotherapy-exposed BM) and in the TGCT cell line H12.1 (3 µM) was measured in a colorimetric substrate cleavage assay. Values represent mean + SD of OD ratios of treated versus the respective untreated cells (control) from at least three independent experiments. (A): Activity of caspase-2, -3, -8, and -9 in hMSCs from unaffected and chemotherapy-exposed BM. (B): Comparison of caspase-9 activity in hMSCs from unaffected and chemotherapy-exposed BM with activity in H12.1 cells. Abbreviations: OD, optical density; TGCT, testicular germ cell tumor.

 
These results showed that cisplatin resistance in hMSCs is characterized by an impaired caspase-9 activation upon cisplatin-induced apoptosis compared with cisplatin-sensitive cells. Since caspase-9-dependent apoptosis represents a mechanism not typically activated by other chemotherapeutic substances, we hypothesized that additional mechanisms contribute to the resistance to chemotherapeutic damage in hMSCs.

Exposure of hMSCs to Chemotherapeutic Substances with Different Mechanisms of Action Results in an Apoptosis-Independent Increase of p53 Expression
We investigated the p53 expression in hMSCs and in the TGCT cell line H12.1 upon 24 hours of exposure to cisplatin, vincristine, and etoposide at their respective IC90 doses. Apoptosis-inducing cisplatin treatment resulted in an increased p53 expression in hMSCs from unaffected and chemotherapy-exposed BM, as well as in the sensitive H12.1 cells. Increased p53 expression was similarly observed in attached cells and in detached, apoptotic cells (Fig. 6A). However, treatment of hMSCs with the IC90 dose of the sensitive TGCT cell line H12.1, a dose that does not lead to apoptosis in hMSCs (Fig. 4C) did yield an increase of p53 expression similar to that seen with the higher, hMSC-specific IC90 dose (Fig. 6A). Treatment of hMSCs with etoposide and vincristine at their respective IC90 doses did also result in increased p53 expression. As seen for cisplatin, this increase was already observed in hMSCs after treatment with IC90 doses of the TGCT cell line H12.1, doses well below the hMSC-specific IC90 dose (Fig. 6B). Treatment with the hMSC-specific IC90 dose did not result in an additional increase in p53 levels in hMSCs but rather in stable levels or, as for vincristine, in reduced expression compared with the treatment with the lower dose. Conclusively, p53 expression is increased in hMSCs upon subapoptotic damage by cisplatin, etoposide, and vincristine and does not correlate with the respective apoptosis-inducing IC90 dose.


Figure 6
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  		Figure 6. Exposure of hMSCs to chemotherapeutic substances with a different mechanism of action results in an apoptosis-independent increase of p53 expression. Expression of p53 was analyzed in protein lysates after 24 hours of treatment with the respective chemotherapeutic substances by Western blotting. ß-Actin served as a loading control, and the positions of molecular weight markers are indicated by arrows. Data represent equal results of three independent experiments, respectively. (A): p53 expression in adherent and detached cells after cisplatin treatment of hMSCs from unaffected (donor 95) and chemotherapy-exposed (donor 54) bone marrow (BM), as well as of TGCT H12.1 cells at the respective doses. (B): p53 expression in attached hMSCs (donor 40) after treatment with cisplatin, etoposide, and vincristine at the indicated doses. For each substance, the higher dose represents the IC90 dose of hMSCs (mean of IC90 for hMSCs from unaffected and chemotherapy-exposed BM), and the lower dose represents the IC90 dose of the TGCT cell line H12.1. Abbreviations: a, adherent; d, detached; TGCT, testicular germ cell tumor.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 Acknowledgments
 References
 
The work presented here originated from the question of whether hMSCs can be isolated for potential therapeutic applications from chemotherapy-exposed BM, and the intial findings led us to the investigation into mechanisms of chemotherapy resistance in hMSCs. This study represents the first systematic investigation on the characteristics of hMSCs derived from chemotherapy-exposed BM. Furthermore, it offers first insights into the mechanisms of apoptosis regulation in hMSCs upon genotoxic and cytotoxic damage.

CFU-Fs could be isolated consistently from BM aspirates of chemotherapy-treated and untreated donors. The lower CFU-F numbers in chemotherapy-treated patients as well as the high standard deviation in the CFU-F numbers in both donor groups may indicate an impact of yet unknown conditions, as well as of the specific chemotherapy regimens on the presence of hMSCs in BM aspirates. Similarly, the observed range of cultivation time until defined confluence of CFU-F in preparations from chemotherapy-exposed and unaffected BM suggests interindividual differences in the initial growth kinetics of preparations from different donors due to yet unidentified factors. Our data show that MSCs can be isolated from human BM after chemotherapeutic treatment, which is in accordance with the observation that hMSCs isolated from the BM of recipients of allogeneic BM transplants are of recipient origin and thus must have sustained high-dose chemotherapy [22, 23]. We observed no differences in the proliferation capacity and mesodermal differentiation potential of hMSCs from chemotherapy-exposed and unaffected BM. Our findings are in contrast to studies suggesting a quantitative and qualitative damage of BM stromal cells or hMSCs after hematopoietic stem cell transplantation [24–28]. In contrast to our analysis, most of these studies did focus on the support of hematopoiesis by hMSCs. Moreover, allogeneic hematopoietic stem cell transplantation may cause additional damage, such as extensive radiation and immunological depletion of recipient hMSCs by donor lymphocytes. Several reports demonstrated alterations in hMSCs or their putative in vivo correlate due to malignant BM infiltration [29] or various diseases [30–32], as well as in hMSCs from aged donors [33], although characterization of hMSCs in these studies was only performed to a limited extent. The size of the nonhomogeneous donor population investigated in our study did not allow a multivariant analysis. Thus, influences of age, sex, time since chemotherapy exposure, and other diseases or therapies on the characteristics of hMSCs cannot be fully excluded. The in vitro characteristics of hMSCs are determined by culture conditions [2]. A specific medium composition developed by others [13] was used in our study. However, we have been able to isolate hMSCs from chemotherapy-exposed BM using DMEM low glucose medium with 10% FCS selected for optimal growth and observed no differences regarding tested characteristics compared to the present data. Thus, an influence of the medium composition on specific resistance mechanisms cannot be excluded, but the isolation of hMSCs from chemotherapy-exposed BM and the general in vitro chemotherapy resistance of hMSCs seems not to depend on the growth factors used in the present study.

Our data suggest that hMSCs from chemotherapy-exposed BM may serve as a cell source for potential clinical applications. This may be of particular importance for the use of autologous transgenic hMSCs in cancer treatment [6]. Although allogeneic hMSCs have been successfully transplanted [5, 34], possible risks of tumor development and late rejection have yet to be excluded, and the use of autologous hMSCs would be preferable. Compared with allogeneic transplantation, isolation, in vitro expansion, and genetic manipulation of autologous hMSCs represent an approach with less logistic challenges and ethical restrictions.

The subsequent in vitro experiments demonstrated that hMSCs are resistant to chemotherapeutic substances of different mechanisms of action compared with sensitive cells, particularly at high, apoptosis-inducing doses. This is in accordance with a recent study showing an in vitro resistance of hMSCs to a wide panel of chemotherapeutic substances [9]. Because of the different experimental settings, the IC values of the two studies cannot be compared directly, but for vincristine, both studies suggest a maximum cytotoxic effect in hMSCs at approximately 0.01 µM. To elucidate the yet unknown resistance mechanisms in hMSCs, we investigated the effects of DNA-damaging substances of different mechanisms of action. hMSCs maintained their stem cell character despite repeated exposure to cisplatin or etopside at clinically relevant concentrations. Typically, in cisplatin-sensitive cells such as EC cells, DNA platination by cisplatin results in the induction of the intrinsic pathway of apoptosis via activation of caspase-9 [35, 36]. We showed that induction of apoptosis by cisplatin as detected by trypan blue and annexin V staining required significantly higher doses in hMSCs than in the TGCT cell lines H12.1 and 2102EP, which represent typical EC cells [37]. Moreover, cisplatin-induced apoptosis in hMSCs was characterized by a lack of caspase-9 activity despite cleavage of procaspase-9. Thus, the higher apoptotic threshold for cisplatin in hMSCs could result from a suppression of caspase-9 activity by specific inhibitors [38] or a possible sequestration of caspase-9, which would not only render its activity undetectable but probably also ineffective for apoptosis induction [39]. We had previously observed a similar pattern of elevated apoptotic threshold and lack of caspase-9 activation in cisplatin-resistant EC cells.

A major role in the cellular decision between apoptosis and cell cycle arrest is exerted by p53 [11]. Whereas p53-driven expression of p21 induces a cell cycle arrest, ongoing damage or failed repair leads to induction of apoptosis through p53-driven activation of proapoptotic genes [12]. Our data show that cytotoxic or genotoxic damage at subapoptotic doses leads to p53 upregulation in hMSCs. Considering the resistance of hMSCs against apoptosis induction, we hypothesize that this p53 upregulation signals a cell cycle arrest. This arrest seems to be temporary, since we could isolate proliferating hMSCs from chemotherapy-exposed BM and observed proliferation of hMSCs after repeated genotoxic treatment in vitro. We hypothesize that p53 activation upon intracellular damage contributes to the persistence of the stem cell character in hMSCs. This is supported indirectly by the observation that undifferentiated embryonic stem cells and EC cells that lack a G1 checkpoint, thereby prohibiting a cell cycle arrest despite p53 activation [40, 41], are highly sensitive to genotoxic damage [36, 42, 43]. In contrast, our data show that resistance of hMSCs to genotoxic damage is associated with an increased p53 expression. This leads to the hypothesis that in stem cells, the degree of resistance to genotoxic damage correlates with the relevance of the particular cell type for the generation of progeny. Pluripotent stem cells, which can generate germ cells, show a lack of p53-dependent cell cycle arrest contributing to a high sensitivity for genotoxic damage. In contrast, adult stem cells such as hMSCs, which do not participate directly in the generation of germ cells are probably capable of p53-dependent cell cycle arrest to allow damage repair. This differential response to genotoxic damage could secure genomic integrity in the progeny but also maintain the function of adult stem cells of the organism.

Taken together, our study shows that hMSCs can be isolated from chemotherapy-exposed BM in sufficient number and quality and may thus serve as a source for potential clinical applications in chemotherapeutically treated patients. hMSCs are resistant to chemotherapeutic substances of different mechanisms of action and maintain their stem cell character despite cytotoxic and genotoxic damage in vivo and in vitro. Our data suggest that an elevated apoptotic threshold contributes not only to the persistence of hMSCs in the BM after chemotherapy but also to their lifelong presence in the adult BM despite physiological cytotoxic and genotoxic damage. These findings will help to define criteria for the clinical use of hMSCs and to understand the biology of this cell type.


   DISCLOSURES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
Acknowledgments
 References
 
The authors indicate no potential conflicts of interest.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 Acknowledgments
References
 
We are grateful to the donors who consented the use of their bone marrow aspirates. This study was partially conducted at the Centre for Applied Medical and Human-Biological Research (ZAMED) at the Medical Faculty of the Martin-Luther-University Halle-Wittenberg. This study was supported in part by grants from the Federal State of Saxonia-Anhalt and the German Ministry of Education and Research through the Wilhelm-Roux Program at the Medical Faculty of the Martin-Luther-University Halle-Wittenberg (1/09 3/15 6/08, 3/27) (L.P.M.) and the European Union through BioService Halle GmbH (J.L.).


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 Materials and Methods
 Results
 Discussion
 Disclosures
 Acknowledgments
 References
 

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