Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2013 Jun 17.
Published in final edited form as: Stem Cells. 2012 May;30(5):975–987. doi: 10.1002/stem.1069

Atmospheric Oxygen Inhibits Growth and Differentiation of Marrow-Derived Mouse Mesenchymal Stem Cells via a p53 Dependent Mechanism: Implications for Long-Term Culture Expansion

Siddaraju Boregowda 1,2, Veena Krishnappa 1,2, Jeremy Chambers 2, Phillip V LoGrasso 2, Wen-Tzu Lai 3, Luis A Ortiz 4, Donald G Phinney 2
PMCID: PMC3683654  NIHMSID: NIHMS482521  PMID: 22367737

Abstract

Large scale expansion of human mesenchymal stem cells (MSCs) is routinely performed for clinical therapy. In contrast, developing protocols for large scale expansion of primary mouse MSCs has been more difficult due to unique aspects of rodent biology. Currently, established methods to isolate mouse MSCs select for rapidly dividing subpopulations that emerge from bone marrow cultures following long-term (months) expansion in atmospheric oxygen. Herein, we demonstrate that exposure to atmospheric oxygen rapidly induced p53, TOP2A and BAX expression and mitochondrial ROS generation in primary mouse MSCs resulting in oxidative stress, reduced cell viability and inhibition of cell proliferation. Alternatively, procurement and culture in 5% oxygen supported more prolific expansion of the CD45−ve/CD44+ve cell fraction in marrow, produced increased MSC yields following immuno-depletion, and supported sustained MSC growth resulting in a 2300-fold increase in cumulative cell yield by 4th passage. MSCs cultured in 5% oxygen also exhibited enhanced tri-lineage differentiation. The oxygen-induced stress response was dependent upon p53 since siRNA mediated knockdown of p53 in wild type cells or exposure of p53−/− MSCs to atmospheric oxygen failed to induce ROS generation, reduce viability, or arrest cell growth. These data indicate that long-term culture expansion of mouse MSCs in atmospheric oxygen selects for clones with absent or impaired p53 function, which allows cells to escape oxygen-induced growth inhibition. In contrast, expansion in 5% oxygen generates large numbers of primary mouse MSCs that retain sensitivity to atmospheric oxygen, and therefore a functional p53 protein, even after long-term expansion in vitro.

Keywords: Mesenchymal stem cells, mesenchymal stromal cells, p53, oxidative stress, reactive oxygen species

INTRODUCTION

Friedenstein et al [1] first demonstrated that mesenchymal stem cells (MSCs) can be enriched from bone marrow by attachment to tissue culture plastic. Although protocols based on this methodology have been developed for large-scale expansion of human MSCs [2–4] the procurement of primary MSCs from mouse bone marrow has proven more challenging due to species specific differences in cell biology. For example, plastic adherent cultures derived from mouse bone marrow exhibit sustained granulopoiesis and B cell lymphopoiesis in the absence of exogenously added cytokines [5–7]. Moreover, hematopoietic stem cells capable of repopulating bone marrow in vivo have been shown to bind to stromal cell layers in vitro [8–10]. Therefore, attachment to plastic is ineffective in removing hematopoietic cell lineages that persist in these cultures even after serial passage [6, 11–13]. Alternative methods such as plating at limiting dilution [14], use of metabolic poisons [15] and cell sorting [16, 17] have been employed with limited success to enrich MSCs from mouse bone marrow.

Previously we developed a method based on immuno-depletion to fractionate MSCs from hematopoietic cell lineages in early passage bone marrow cultures [13, 18, 19]. However, this approach is constrained by the fact that immuno-depleted MSCs exhibit limited growth when cultured under standard conditions in vitro. Consequently a growing number of laboratories have adopted purification schemes that select for rapidly proliferating subpopulations which emerge from plastic adherent marrow cultures following long-term expansion in atmospheric oxygen [20–26]. These subpopulations typically exhibit variable capacities for tri-lineage differentiation, express hematopoietic lineage markers, and survive in culture for over 50 passages in vitro [22, 23, 25] and therefore share traits common to immortalized marrow stromal cell lines [27]. Indeed, the emergence of rapidly dividing subpopulations following long-term expansion is indicative of cell immortalization, which occurs at a much higher frequency in rodent vs. human populations due to differences in checkpoint control mechanisms [28, 29]. For example, growth restrictive conditions have been shown to select for cells with inactivating mutations in p53, a protein mutated in the vast majority of immortalized rodent cell lines [30, 31].

In this report we demonstrate that exposure to atmospheric oxygen rapidly induces p53 and BAX protein expression, mitochondrial reactive oxygen species (ROS) generation, and inhibits proliferating cell nuclear antigen (PCNA) protein expression in primary mouse MSCs resulting in reduced cell viability and rapid growth arrest. We also show that culture of mouse bone marrow cells in 5% oxygen promotes expansion of the CD45−ve/CD44+ve cell fraction and supports sustained growth of MSCs enriched from this fraction by immuno-depletion. Indeed, procurement and expansion up to 4th passage in 5% vs. 21% oxygen resulted in a 2300-fold increase in cumulative MSC yields. Culture in low oxygen also significantly enhanced the trilineage differentiation capacity of MSCs. Finally, we show that exposure of p53−/− MSCs to atmospheric oxygen did not result in ROS production, reduced cell viability, or growth inhibition indicating that the oxygen-induced stress response is p53-dependent. Collectively, these data demonstrate that mouse MSCs are uniquely sensitive to atmospheric oxygen and that long-term expansion of cells under standard culture conditions selects for subpopulations with reduced or absent p53 activity, thereby allowing escape from oxygen-induced growth inhibition. Based on these findings, we describe a purification scheme that incorporates immuno-depletion coupled with culture in low oxygen to isolate large numbers of primary mouse MSCs, which retain a functional p53 protein and sensitivity to atmospheric oxygen after extensive sub cultivation.

MATERIALS AND METHODS

MSC isolation, cultivation, transfection and irradiation

MSCs were isolated from FVB/n, C57BL/6 and B6.129S2-Trp53<tm1Tyj> mice (The Jackson Laboratory, Bangor, ME, http://jaxmice.jax.org) by immuno-depletion as previously described [19]. MSCs were cultured for <10 days prior to immuno-depletion and thereafter designated as 1st passage. Populations were split into two fractions at the time of harvest and manipulated identically except that one was cultured at 37°C with 5% CO2 in a humidified chamber in atmospheric (21%) oxygen and the other in a modular airtight chamber (BioSpherix Ltd., Lacona, NY, http://www.biospherix.com/) flushed with 5% O2 balanced with N2. Where indicated MSCs initially cultured in 5% oxygen were switched to 21% oxygen and vice versa. Some experiments were conducted using media supplemented with 5 mM N-acetylcysteine (NAC). Population doubling times were calculated as PD = t log2 / (log Nt - log No) where t is time period, Nt is the number of cells at time t, and No is the initial number of cells plated. Cumulative cell numbers were determined from initial plating density and total population doublings for each passage.

Delivery of siRNA into MSCs was accomplished using the Lipofectamine™RNAiMAX Transfection Reagent (Invitrogen, Carlsbad, CA, http://invitrogen.com). Briefly, transfection reagent was prepared by mixing Lipofectamine™ RNAiMAX (1:2000) and the appropriate RNAi duplex (10nM, Santa Cruz Biotechnology Inc., Santa Cruz, CA, http://www.scbt.com) in Opti-MEM® Reduced Serum Medium (1:5). A single cell suspension of MSCs (P1) was then plated on top of the transfection reagent at a final density of 1000 cells/cm2. Cultures were fed with fresh media 2h hours later and every 2–3 days thereafter for up to 7 days. In some studies MSCs (P1) maintained in 5% oxygen were plated at 2000 cells/cm2 in a T-25 flask, incubated in 5% or 21% oxygen for 3 additional days, and then irradiated at the dose of 4Gy using a GammaCell-40 irradiator. Cells were cultured for additional 4 days during which time growth kinetics and viability (tryphan blue exclusion) were quantified. Cultures were photographed using a Leica DMI3000 B upright fluorescent microscope attached to a DFC295 digital camera (Micro Optics of Florida, Inc., Davie, FL, http://www.microopticsfl.com).

Flow cytometry

To analyze the cell surface phenotype, MSCs (2.5 × 105 cells/ml) were suspended in Hank's balanced saline solution (HBSS) and incubated with antibodies against CD29, CD73, CD31, Sca1, SSEA-1 and SSEA-4 (R&D Systems Inc., Minneapolis, MN, http://www.rndsystems.com) for 30 minutes at 4°C. Cells were washed twice with HBSS, incubated with the appropriate fluorochrome-conjugated secondary antibody for 30 minutes at 4°C and analyzed using an Epics FC 500 FACS scanner equipped with CXP software (Beckman Coulter, Inc., Indianapolis, IN, http://www.beckmancoulter.com). To evaluate cell proliferation MSCs were incubated with 10μM CFSE (Invitrogen) at 37° C for 10 minutes, quenched by addition of five volumes of complete culture media (CCM; α minimal essential media containing L-glutamine and supplemented with 10% FBS, 100U/ml penicillin, and 100U/m; streptomycin) then plated at 1000 cells/cm2 and cultured for 7 days at 37°C in 5% or 21% oxygen. Media was changed every 2–3 days. To evaluate p53 function MSCs were washed in PBS and fixed in 70% ethanol at −20°C for 2h. Cells were collected by centrifugation, then treated with 0.1M HCl at 37°C for 12 min to extract low molecular weight DNA with minimally denature intact chromosomal DNA. Cells were washed in PBS and stained using propidium iodide (5μg/ml) for 20 minutes. MSCs stained with CFSE or PI were analyzed using a LSR II Flow Cytometer System (Beckton Dickinson, Franklin Lakes, NJ, http://www.bd.com) and FlowJo software (Tree Star, Inc., Ashland, OR, http://www.treestar.com).

Fluorescent detection of ROS and mitochondrial superoxide

MSC were stained with 2',7'-di-chlorofluorescein derivative, H2DCF-DA (DCF-DA) to detect whole cell ROS. DMEM was exchanged with DMEM without phenol red two hours prior to the start of the experiment. Cells were stained prior to anisomycin treatment with DCF-DA or after anisomycin treatment depending on the experiment. Cells were stained with 10μM DCF-DA for 10 minutes, washed in DMEM without phenol red and with 2.5% FBS to reduce the fluorescent contributions of the dye and the serum, and then visualized microscopically in phenol red-free DMEM with 0.25% FBS. Alternatively, MSCs were stained with MitoSOX-Red (Invitrogen) to quantify mitochondrial superoxide production. For microscopic analysis 10μM MitoSOX-Red was used while 100nM was used for fluorescent quantification. To confirm mitochondrial localization of the dye, cells were counterstained with 5μM Mito-Tracker Green (Invitrogen) for 10 minutes. Normalization of fluorescent data was done using counterstaining with Hoechst 33342 (1μM) for 5 minutes. Fluorescent wavelength pairs for the individual dyes were 568nm/600nm for DHE, 510nm/580nm for MitoSOX-Red, and 490nm/516nm for Mito-Tracker Green.

Mitochondrial membrane potential

Mitochondrial membrane potential was determined using JC-1 staining. Cells were grown to ~80% confluence in 96-well plates and stained with 2μM JC-1 for 10 minutes following anisomycin treatment. Green fluorescence (depolarization) was monitored at 488nm and red fluorescence (polarized) at 590nm on a Spectromax M5e plate reader (Molecular Devices, Inc, Sunnyvale, CA, http://www.moleculardevices.com). Actively growing cells were used as a negative control and cells treated with 2mM FCCP and 1mM valinomycin as the positive control. Normalization to cell number was achieved as described above. Data are presented as percent viable cells based on presumed 95–100% depolarization observed in FCCP/valinomycin-treated cells.

Oxidative stress, cell viability, and determination of ATP concentration

Protein carbonyls, lipid peroxides, and aconitase activity were quantified using the Protein Carbonyl Assay Kit, Lipid Hydroperoxide Assay Kit, and the Aconitase Assay kit, respectively, (Cayman Chemical, Ann Arbor, MI, http://www.caymanchem.com) and reduced Glutathione levels were determined using the GSH-Glo™ Glutathione Assay (Promega, Corp., Madison, WI, http://www.promega.com). For each assay, a minimum of 10,000 MSCs were used for each replicate measurement. Cell viability was monitored via SYTOX Green (Invitrogen) nucleic acid staining in a 96-well plate according to manufacturer's instructions. Intracellular ATP concentration was determined using the ATP Determination Kit (Invitrogen). Briefly, cells were washed twice in PBS, lysed by addition of boiling H2O, and the suspension cleared by centrifugation at 14,000×g for 15 minutes at 4°C. ATP quantification was carried out immediately following lysis.

Western blot

Protein lysates were prepared using the Qproteome Mammalian Protein Prep Kit (Qiagen, Valencia, CA, http://www.qiagen.com) and protein concentrations were determined using the Bradford assay (Bio-Rad, Hercules, CA, http://www.bio-rad.com). Protein samples (20 μg) were prepared in Laemmli sample buffer (Bio-Rad) containing 5% β-mercaptoethanol, denatured at 95°C for 5min, electrophoresed on NuPAGE 10% Bis-Tris gels using 1X NuPAGE MES SDS Running Buffer (Invitrogen), and then transferred to 0.45um nitrocellulose membranes in 1X NuPAGE transfer buffer containing 10% methanol. Membranes were washed with Tris-buffered saline (TBS) for 5 min, incubated in TBS with 0.1% Tween-20 (TBST) and Odyssey® blocking buffer (LI-COR Biosciences, Lincoln, NE, http;//www.licor.com) overnight at 4°C, washed an additional 3× in TBST, and then incubated with anti-p53, anti-MDM2, anti-p16, anti-p21, anti-β-actin (1:200, Santa Cruz Biotechnology Inc.), anti-TOP2A, anti-BAX (1:500, Cell Signaling Technology, Inc., Danvers, MA, http://www.cellsignal.com), anti-PCNA (1:1000, Abcam Inc., Cambridge, MA, http://www.abcam.com) or anti-GAPDH (1:1000, Imgenex Corp., San Diego, CA, http://www.imgenex.com) antibodies in Odyssey® blocking buffer for 2 hours at room temperature with gentle agitation. Membranes were washed 5× in TBST and probed with a fluorescent-labeled secondary antibody at a 1:10000 dilution in Odyssey® blocking buffer for one hour. Blots were scanned using Odyssey® infrared image system (LI-COR Biosciences).

Cell differentiation

MSCs (0.125×104 cells/cm2) were cultured expanded in 21% or 5% oxygen and at 2nd passage their tri-lineage differentiation potential quantified as described previously [21] with the following exceptions. Adipogenic induction media consisted of CCM supplemented with 0.5uM dexamethasone, 0.5mM isobutylxanthine and 50uM indomethacin and lipid accumulation was quantified by staining cells with AdipoRed Assay Reagent (Lonza Rockland, Inc., Rockland, ME, http://www.lonza.com) for 10 minutes and quantifying fluorescence using a fluorescent plate reader (excitation 485nm; emission 535). Chondrogenesis was induced by incubating cell pellets (250,000 cells) for 21 days successively in chondrogenic induction media and hypertrophic media. The glycosaminoglycan (GAG) content of the pellets was then quantified as described previously [33].

Statistical analysis

All data were expressed as mean ± standard deviation. Data from treated groups were compared to untreated groups using the Student's t test. Differences between treatment groups were considered significant if the p-value was <0.05.

RESULTS

Exposure to atmospheric oxygen inhibits growth of MSCs enriched from mouse bone marrow by immuno-depletion

Immuno-depletion provides a reproducible means to fractionate MSCs from plastic adherent cultures of mouse bone marrow [18, 19] but the proliferative capacity of the purified cells rapidly diminishes as a function of passage under standard conditions in vitro irrespective of initial plating density (Figure 1A, B). Culture in FGF2-supplemented media significantly (p<0.01) stimulated the growth of immuno-depleted MSCs at first passage (P1) but had no significant effect at P2. All growth factors tested (FGF2, IGF-1, and LIF) significantly stimulated growth at P3 compared to untreated cells but the magnitude of the effect was minimal (Figure 1C). Therefore, the low proliferative capacity of immuno-depleted MSCs does not appear to result from nutrient or growth factor deprivation.

Figure 1. Atmospheric oxygen levels inhibit growth of CD45−ve/CD44+ve bone marrow cells and marrow-derived MSCs purified by immuno-depletion.

Figure 1

Open in a new tab

A–F) Immuno-depleted MSCs plated at the indicated cell densities (500–13,000 cells/cm2) were culture expanded in 21% (A) or 5% (D) oxygen and cumulative cell yields as a function of passage was determined by counting. B) Plotted is the fold increase in MSC number (5000 cell/cm2) as a function of passage for data in (A) and (D). C) MSCs plated at 5000 cells/cm2 were cultured in CCM alone or CCM supplemented with 20 ng/ml FGF2 (FGF2–20), 50 ng/ml FGF2 (FGF2–50), 100ng/ml FGF2 (FGF2–100), 20 ng/ml IGF-1 (IGF-1), or 1000U/ml LIF (LIF) and growth rates (cells/day) in 21% oxygen were quantified as a function of passage by counting. E, F) Population doubling times at P1 and P4 (E) and the total number of population doublings accumulated over four passages (F) were calculated from data in A and D. Doubling times (DT) were determined as DT = (T1–T2)*(Log(2)/Log(Q1/Q2)) where Q1 is the number of cells at T1 and Q2 is the number of cells at T2. Total population doublings at each passage were determined as 2n where n = Log(Q1/Q2)/Log(2). The plotted data (mean ± SD) were determined from experiments run in triplicate. G, H) Two different preparations of whole bone marrow were suspended in CCM supplemented with 10% FBS, expanded in 21% vs. 5% oxygen for 7 days and analyzed by flow cytometry to quantify the percentage of CD45+ve (G, gate illustrated by horizontal line) and CD44+ve/CD45−ve (H, denoted by rectangular gates) subpopulations. Samples were analyzed in triplicate. I) The total yield of plastic adherent cells from bone marrow cultures expanded in 21% vs. 5% oxygen for seven days was determined by counting. J) Plotted are the percentages of CD45+ve, CD45−ve, and CD44+ve/CD45−ve cells in bone marrow cultures quantified by flow cytometry as illustrated in G and H. Note that most CD45+ve cells are also CD44+ve. K). The yield of primary MSCs obtained following immuno-depletion of plastic adherent marrow cultures expanded for 8 days in 21% vs. 5% oxygen was determined by counting. Plotted data (mean ± SD) represent results from at least 5 marrow preparations from each experimental condition. *, p<0.05; **, p<0.01; #, p<0.005; +, p<0.001.

Several recent studies have documented that oxygen levels profoundly affect the growth of rodent fibroblasts [34], neural progenitor cells [35, 36] and induced pluripotent and cancer stem cells [37]. Considering MSCs are derived from bone marrow, a low oxygen environment, we postulated that atmospheric oxygen levels may impede cell growth in vitro. As shown in Figure 1D immuno-depleted MSCs exhibited robust and sustained growth up to P4 irrespective of plating density when cultured in a closed system in 5% oxygen, which was chosen to represent physiologic concentrations of oxygen in bone marrow. As anticipated, population doubling times of MSCs grown in 21% oxygen were significantly (p<0.01) longer at P4 vs. P1 for all plating densities evaluated (Figure 1E) and increased by 54.4, 12.3, and 10.5-fold for cells plated at 1000, 5000, and 13,000 cells/cm2, respectively. Doubling times of MSCs cultured in 5% oxygen were also significantly (p<0.005) longer at P4 vs. P1 but increased by only 1.3, 1.6, and 1.2-fold for cells plated at 500, 1000, and 5000 cells/cm2, respectively (Figure 1E). Moreover, doubling times for MSCs grown in 21% oxygen were significantly (p<0.01) longer at all passages and plating densities then compared to cells grown in 5% oxygen (Figure 1E). MSCs cultured in 21% oxygen completed 4.4, 1.9, and 1.5 cumulative population doublings over a 50 day period when plated at 1000, 5000, or 13,000 cells/cm2, respectively, whereas MSCs cultured at 5% oxygen underwent 12.4, 11.2, and 5.7 cumulative population doublings in an average of 18 days when plated at 500, 1000, or 5000 cells/cm2 (Figure 1F). Difference in cumulative population doublings between 21% and 5% oxygen cultures were also highly significant (p<0.001). Therefore, oxygen appears to be the principle factor restricting the growth of immuno-depleted MSCs.

Culture in 5% oxygen promotes expansion of the CD45−ve/CD44+ve fraction in bone marrow resulting in significantly enhanced MSC yields following immuno-depletion

To determine the effect of oxygen on MSC growth prior to purification via immuno-depletion, bone marrow preparations from two separate cohorts of mice were divided into two fractions and maintained in 21% or 5% oxygen. Flow cytometric analysis confirmed that the percentage of CD45+ve cells, most of which were also CD44+ve, was significantly greater (p<0.005) in cultures expanded in 21% vs. 5% oxygen (59.1% vs. 48.9%, respectively) but the percentage of CD45−ve (51.0±5.3% vs. 41.05±3.1%, p<0.005) and CD45−ve/CD44+ve cells (37.1±6.0% vs. 29.7±3.1%, p<0.05) were significantly more abundant in 5% vs. 21% oxygen cultures (Figure 1G, H, J). In addition, yields of plastic adherent marrow cells at 7 days post-plating were 1.5-fold greater (p<0.001) (Figure 1I) and overall yields of MSCs following immuno-depletion were 2.7-fold greater (11.7% vs. 4.3% for 5% or 21% oxygen, p<0.005) when cultured in 5% vs. 21% oxygen (Figure 1K). Based on differences in growth rates, procurement and expansion up to four passages in 5% vs. 21% oxygen resulted in a ~2300-fold increase in immuno-depleted MSC yields.

To verify the effect of oxygen on cell proliferation CFSE-labeled MSCs were expanded in 21% vs. 5% oxygen for one passage and the distribution of label analyzed by flow cytometry. As shown in Figure 2A, the majority of CFSE was retained in the 7th vs. 2nd cell generation for populations cultured in 5% vs. 21% oxygen, respectively, which was consistent with differences in growth rates illustrated in Figure 1. Populations expanded in 5% vs. 21% oxygen were also readily distinguished by differences in their physical characteristics. For example, the majority of proliferating cells (61.5%) in 5% oxygen cultures were characterized by low forward and side scatter. However, only 3.8% of cells in 21% oxygen cultures were contained in the low forward and side scatter gate (Figure 2B). This difference was exemplified by analyzing the CFSELow and CFSEHigh fractions from each population (Figure 2C). Populations expanded in 5% vs. 21% oxygen also were readily distinguished based on morphological differences visualized microscopically. Specifically, 5% oxygen cultures mostly contained small, round cells with a high nuclear to cytoplasmic ratio and 21% oxygen cultures predominantly consisted of large, flattened cells with a high cytoplasmic to nuclear ratio (Figure 2C, photomicrographs).

Figure 2. Atmospheric oxygen arrests cell proliferation but does not induce cellular senescence in primary mouse MSCs.

Figure 2

Open in a new tab

A, B) Immuno-depleted MSCs (1000 cells/cm2) were procured and expanded in 5% (A) or 21% (B) oxygen for one passage (P1) and analyzed by flow cytometry for CFSE staining (A) or forward and side scatter (B). C) MSCs cultured as in (A) were sorted based on CSFE label intensity and then analyzed for forward and side scatter. Photomicrographs visually illustrate the distinct morphological differences in populations expanded in 5% vs. 21% oxygen detected by flow cytometric analysis. D) MSCs procured and expanded for four passages (P4) in 5% oxygen were passed an additional time (P5) while maintained in 5% oxygen or switched from 5% to 21% oxygen and then analyzed by flow cytometry for CFSE labeling or forward and side scatter. E) MSCs expanded for four passages (P4) in 21% oxygen were passed an additional time (P5) in 21% oxygen or switched from 21% to 5% oxygen and analyzed as in (D). F, G) Growth rates (F) and population doubling times (G) were calculated for MSCs expanded in 21% oxygen, 5% oxygen, or switch cultures as described above. Plotted values (mean ± SD) represent duplicates from a single experiment. *, p<0.05; **, p<0.01, #, p<0.005.

To confirm that atmospheric oxygen induced growth arrest we performed a series of switch experiments. As shown in Figure 2D, MSCs cultured in 5% oxygen and labeled with CFSE at P4 continued to divide if maintained in 5% oxygen for one additional passage (P5) but exhibited a rapid cessation in cell proliferation when switched to 21% oxygen. This change in proliferative capacity following the switch from 5% to 21% oxygen was coincident with a decrease (14.6% to 7.0%) in the gated subpopulation characterized by low forward and side scatter. Conversely, MSCs labeled with CFSE at P4 and maintained in 21% oxygen for one additional passage remained quiescent but began to rapidly proliferate when switched to 5% oxygen, which also resulted in a net increase (0.8% to 5.8%) in the gated subpopulation of proliferating cells (Figure 2E). These results were confirmed by quantifying the growth rate and doubling time of populations in switch cultures. For example, growth rates of cells cultured in 21% oxygen significantly (p<0.05) declined as a function of passage but following a switch to 5% oxygen at P3 increased significantly (p<0.05) and remained elevated for up to P9. Moreover, growth rates of cells procured and expanded in 5% oxygen were significantly (p<0.005) greater at all passages as compared to that measured at P1 for cells expanded in 21% oxygen (Figure 2F). Similar trends were seen with respect to population doubling times (Figure 2G). For example, population doubling times gradually increased in cells cultured in 21% oxygen but rapidly decreased when cells were shifted to 5% oxygen and were significantly shorter at all passages for MSCs cultured in 5% vs. 21% oxygen. Therefore, exposure to atmospheric oxygen induced rapid growth arrest of mouse MSCs.

Atmospheric oxygen induces p53, TOP2A, and BAX expression and mitochondrial ROS production resulting in oxidative stress and reduced cell viability

We next performed a series of biochemical experiments to determine if exposure of MSCs to atmospheric oxygen induced oxidative stress. MSCs cultured in 21% vs. 5% oxygen were found to contain significantly (p<0.0005) higher levels of intracellular ROS (Figure 3A) and staining with MitoSOX-Red confirmed that increased ROS levels were generated predominantly by mitochondria (Figure 3B). Staining cells with the membrane permeable dye JC-1 further demonstrated a significant (p<0.05) increase in mitochondrial membrane permeability in cells exposed to 21% oxygen (Figure 3C). Consistent with these data, flow cytometric analysis demonstrated that populations cultured in 21% vs. 5% oxygen contained a higher percentage of Annexin V-positive (11.1% vs. 1.63%) and Annexin V/PI double-positive cells (16.2% vs. 4.3%) indicative of higher rates of cellular apoptosis and necrosis (Figure 3D). MSCs maintained in 21% oxygen were also stained to a significantly (p<0.001) greater degree with the vital dye SyTox Green as compared to cells cultured in 5% oxygen (Figure 3E). Significant increases in the concentration of protein carbonyls and lipid peroxides (Figure 3E) and significant decreases in aconitase activity, reduced glutathione levels, and ATP levels (Figure 3F) were also evident in MSCs exposed to 21% vs. 5% oxygen.

Figure 3. Exposure to atmospheric oxygen induces p53, TOP2A, and BAX expression and mitochondrial ROS production resulting in oxidative stress and reduced cell viability.

Figure 3

Open in a new tab

Immuno-depleted MSCs were culture expanded in 5% or 21% oxygen for 7 days and biochemical analyses were used to quantify whole cell ROS (A), mitochondrial ROS (B), mitochondrial membrane permeability (C), cell viability (D), protein carbonyl and lipid peroxidase levels (E), and aconitase activity, reduced glutathione and ATP levels (F). All biochemical analyses were performed as described in the experimental procedures and each experiment was conducted using three distinct replicate cultures from each experimental group (5% vs. 21% oxygen cultures). Plotted values (mean ± SD) were determined from four replicates for each sample. D) MSCs were stained with an anti-Annexin V antibody and PI and analyzed by flow cytometry to determine the percentage of early apoptotic, late apoptotic and necrotic cells. G) Cell extracts were prepared from immuno-depleted MSCs that were isolated from bone marrow cultures procured and expanded in either 5% or 21% oxygen for the indicated passage numbers and analyzed by Western blot using an anti-p53 and anti-β-actin antibodies. H) Western blot of the indicated proteins was performed on cell extracts harvested from MSCs expanded in 5% or 21% oxygen at P1 or P2. Western blots were also performed on cell extracts obtained from switch cultures wherein MSCs were expanded for four passages (P4) in 5% oxygen and one additional passage (P5) in 5% oxygen (5/5) or 21% oxygen (5/21). Alternatively, MSCs were expanded to P4 in 21% oxygen and then one additional passage (P5) in 21% oxygen (21/21) or switched to 5% oxygen (21/5). I) MSCs were stained with anti-PCNA and anti-p53 antibodies and analyzed by flow cytometry to determine the percentage of single and double positive cells in populations cultured in 5% or 21% oxygen. *, p<0.05; **, p<0.01, #, p<0.005; +, p<0.001; ++, p<0.0005.

Western blot analysis of cell extracts revealed that MSCs procured and expanded in 21% oxygen expressed high levels of p53 protein, which increased as a function of passage up to P4 (Figure 3G). In contrast, p53 protein expression was negligible until P4 in MSCs expanded in 5% oxygen. In addition to p53, levels of TOP2A and BAX were also elevated in cells cultured in 21% vs. 5% oxygen at both P1 and P2 (Figure 3H). In contrast, PCNA levels were dramatically lower at P2 in MSCs cultured in 21% vs. 5% oxygen, consistent with the fact that higher oxygen levels block cell proliferation (Figure 1A, D and Figure 2A). Similar results were also observed in switch cultures. For example, when MSCs expanded to P4 in 5% oxygen were switched to 21% oxygen, levels of p53, TOP2A and BAX were increased as compared to cells maintained in 5% oxygen (Figure 3H). Alternatively, switching cells from 21% to 5% oxygen reduced expressed levels of these proteins but increased PCNA levels. These data were consistent with flow cytometric analyses, which showed that MSCs maintained in 21% vs. 5% oxygen expressed higher p53 levels and lower levels of PCNA (Figure 3I). Western blot analysis also revealed that p16 and p21 expression levels were significantly higher in populations cultured in 5% vs. 21% oxygen, which reflected the higher proliferative rate of the former (Supplemental Figure 1). These results demonstrate that oxygen-induced up regulation of p53 expression were inversely correlated with changes in growth rate and population doubling times (Figure 1A, B and Figure 2D, E).

Culture with the ROS scavenger N-acetylcysteine (NAC) spared MSCs from some of the detrimental effects of atmospheric oxygen. For example, NAC inhibited oxygen-induced increases in intracellular ROS levels (Supplemental Figure 2A), mitochondrial membrane permeability (Supplemental Figure 2B), protein carbonyls (Supplemental Figure 2C) and lipid peroxidases (Supplemental Figure 2D) as well as oxygen-induced decreases in reduced glutathione levels (Supplemental Figure 2E) and aconitase activity (Supplemental Figure 2F). Western blot analysis confirmed that NAC treatment largely mitigated oxygen-induced increases in BAX expression but resulted in overall higher p53 expression levels in cells cultured in both 5% and 21% oxygen, which corresponded with reduced MDM2 levels (Supplemental Figure 2G). Consistent with these results, NAC treatment produced a modest but significant (1.3-fold, p<0.05) increase in cell growth in 21% oxygen (Supplemental Figure 2H), which was confirmed by CFSE labeling (Supplemental Figure 2I), and reduced oxygen-induced apoptosis by greater than 2-fold (p<0.05) (Supplemental Figure 2J). Therefore, NAC was more effective at abrogating ROS-mediated effects on viability as compared to oxygen-mediated effects on cell growth, which is regulated by nuclear functions of p53.

MSCs expanded in 21% vs. 5% oxygen exhibited slight differences in surface phenotype but a reduced capacity for tri-lineage differentiation potential

Flow cytometric analysis confirmed that culture expansion in 5% vs. 21% oxygen failed to alter the expression profile of most surface antigens on MSCs with a few exceptions. For example, both populations uniformly expressed CD29, CD44, and SSEA-4 and lacked expression of CD31 and SSEA-1 (Table 1). However, MSCs cultured in 5% vs. 21 oxygen expressed higher levels of Sca-1 antigen (94% vs. 65%, respectively) and lower levels of SSEA-4 (88.5% vs. 99.3%, respectively) (Table 1). Oxygen also significantly affected the tri-lineage differentiation potential of MSCs. For example, osteogenic differentiation as quantified by Alizarin Red S staining was significantly greater (p<0.005) in populations procured and expanded in 5% vs. 21% oxygen (4.11±0.04 vs. 2.57±0.01, respectively) (Table 1). Moreover, MSCs maintained in 5% vs. 21% oxygen also exhibited a 5-fold greater capacity for adipogenic differentiation (36.5±4.7 vs. 7.11±0.09, respectively) and a 6-fold greater capacity for chondrogenic differentiation (43.9±2.9 vs. 7.08±1.6, respectively) and these differences were highly significant (p<0.005) (Table 1).

Table 1.

Effect of oxygen on the cell surface phenotype and tri-lineage differentiation potential of mouse MSCs.

Antibody 21% Oxygen 5% oxygen
CD29 99.2% 97.6%
CD31 0.3% 0.39%
CD44 96.7% 98.3%
Sea1 67.8% 94.3%
SSEA1 1.23% 1.27%
SSEA4 99.3% 88.5%
Cell Lineage 21% Oxygen 5% Oxygen p-value
Osteoblast (Alizarin Red S Absorbacne) 2.57 ± 0.0125 4.11 ± 0.041 p=0.0023
Adipogenesis (AdipoRed Fluorescence) 7.11 ± 0.096 36.51 ± 4.79 p=0.0012
Chondrogenesis (μg GAG/μg DNA) 7.08 ± 1.64 43.99 ± 2.95 p=0.00018
Open in a new tab

Primary mouse MSCs were procured and expanded in 21% or 5% oxygen as described and their cell surface phenotype was evaluated at P2 via flow cytometry using the indicated antibodies. The percentage of immuno-reactive cells in each population is reported. Data are representative of a typical MSC preparation. Isotype controls were run in parallel for each antibody examined. The capacity of cells to differentiate into osteoblasts, adipocytes, and chondrocytes was also quantified as described in the experimental methods. Differentiation data (mean ± SD) were calculated from experiments performed in triplicate.

Loss of p53 function abrogates the inhibitory effect of atmospheric oxygen on growth of primary mouse MSCs

Several groups have reported that MSCs derived from p53-deficient mice exhibit incremental increases in growth rate when compared to wild type cells isolated from marrow by long-term culture expansion [38, 39]. In contrast, atmospheric oxygen had profoundly different effects on the growth rates of MSCs isolated via immuno-depletion from p53+/+ and p53−/− mice. For example, flow cytometric analysis following CSFE labeling confirmed that p53+/+ and p53−/−MSCs rapidly proliferated in 5% oxygen but that growth of only p53+/+ MSCs was significantly inhibited in 21% oxygen (Figure 4A, B). Differences in growth rates were also evident based on the light scattering properties and morphology of populations (Figure 4A–C) as shown previously for MSCs derived from FVB/n mice. For example, p53+/+ MSCs cultured in 5% oxygen contained a higher proportion of small cells with low granularity and those cultured in 21% oxygen contained mostly large, granular cells (Figure 4C). In contrast, the size and granularity of p53−/− MSCs were essentially indistinguishable between the two culture conditions (Figure 4C). Population doubling times of p53+/+ MSCs were also significantly (p<0.0005) longer at all passages examined when cultured in 21% vs. 5% oxygen and doubling times in 5% oxygen were significantly (p<0.0005) longer than compared to p53−/− MSCs cultured in either 21% or 5% oxygen (Figure 4D). Importantly, population doubling times of p53−/− MSCs were slightly faster in 5% vs. 21% oxygen at all passages and this difference was statistically significant (p<0.05) (Figure 4D). Similarly, p53−/− MSCs cultured in 5% oxygen underwent a significantly greater number of population doublings at each passage as compared to cells expanded in 21% oxygen (Figure 4E), which resulted in slightly higher overall cell yields following expansion in vitro (Figure 4F). Cumulative population doublings and cell yields were significantly less for p53+/+ MSCs cultured in 21% vs. 5% oxygen and as expected p53+/+ MSCs expanded in 21% oxygen cultures generated the lowest cumulative MSC yield (Figure 4F). Collectively, these data indicate that p53 is the dominant factor modulating oxygen-induced growth inhibition of primary mouse MSCs.

Figure 4. Growth inhibition of MSCs following exposure to atmospheric oxygen is p53 dependent.

Figure 4

Open in a new tab

A, B) Immuno-depleted MSCs (1000 cells/cm2) derived from wild type (A) or p53−/− (B) mice were expanded in 5% or 21% oxygen for one passage (P1) and analyzed by flow cytometry for CFSE staining. C) MSCs from wild type and p53−/− mice were sorted based on CSFE fluorescence and the CSFElow and CSFEHigh fractions were evaluated by flow cytometry based on forward and side scatter. Photomicrographs visually illustrate the distinct morphological differences in populations expanded in 5% vs. 21% oxygen detected by flow cytometric analysis. D) Population doubling times at P1–P5 were calculated for wild type and p53−/− MSCs in 5% or 21% oxygen from data in A. E, F) Cumulative population doublings (E) and cumulative cell number (F) were calculated for from data in D. Doubling times (DT) were determined as DT = (T1–T2)*(Log(2)/Log(Q1/Q2)) where Q1 is the number of cells at T1 and Q2 is the number of cells at T2. Total population doublings at each passage were determined as 2n where n = Log(Q1/Q2)/ Log(2). Plotted values (mean ± SD) represent duplicates from a single experiment. *, p<0.05; **, p<0.01, #, p<0.005.

Functional p53 is necessary for ROS production in response to oxygen exposure

Biochemical-based studies further confirmed that p53 function mediates oxygen-induced stress responses in primary mouse MSCs following exposure to atmospheric oxygen. For example, intracellular and mitochondrial ROS levels were significantly elevated in p53+/+ MSCs cultured in 21% vs. 5% oxygen but no such change was evident for p53−/− MSCs (Figure 5A, B). Similarly, exposure to 21% oxygen significantly increased mitochondrial membrane permeability of p53+/+ but not p53−/− MSCs (Figure 5C). Consistent with these results, high oxygen levels adversely affected the viability of p53+/+ but not p53−/− MSCs based on flow cytometric analysis of Annexin V and PI (Figure 5D) or SyTox Green staining (Figure 5E). This analysis also revealed that MSCs derived from the C57BL/6 strain are intrinsically more sensitive to atmospheric oxygen as as evaluated by PI and Annexin V staining as compared to cells derived from the FVB/n strain (compare Figures 3D and 5D). The p53+/+ MSCs also exhibited significant increases in protein carbonyls and lipid peroxides (Figure 5E) and significant decreases in aconitase activity, reduced glutathione levels, and ATP levels (Figure 5F) when cultured in 21% vs. 5% oxygen but these changes were not evident in p53−/− MSCs. Western blots of cell extracts further demonstrated that culture in 21% vs. 5% oxygen results in up regulation of BAX and MDM2 expression in p53+/+ but not p53−/− MSCs (Figure 5G). As anticipated, p53−/− MSCs also showed no significance difference in tri-lineage differentiation potential (data not shown).

Figure 5. MSCs derived from p53 null mice fail to exhibit increased ROS production and oxidative stress upon exposure to 21% oxygen.

Figure 5

Open in a new tab

Immuno-depleted MSCs from wild type or p53−/− mice were culture expanded in 5% or 21% oxygen for 7 days and biochemical analyses were used to quantify whole cell ROS (A), mitochondrial ROS (B), mitochondrial membrane permeability (C), cell viability (D), protein carbonyl and lipid peroxidase levels (E) and aconitase activity, reduced glutathione and ATP levels (F). All biochemical analyses were performed as described in the experimental procedures and each experiment was conducted using three distinct replicate cultures from each experimental group (5% vs. 21% oxygen cultures). Plotted values (mean ± SD) were determined from four replicates for each sample. D) Immuno-depleted MSCs treated as in A were stained with an anti-Annexin V antibody and PI and analyzed by flow cytometry to determine the percentage of early apoptotic, late apoptotic and necrotic cells. G) Cell extracts were prepared from immuno-depleted MSCs isolated from wild type and p53−/− mice expanded in 5% or 21% oxygen and analyzed by Western blot using anti-p53, anti-MDM2, anti-BAX, and anti-GAPDH antibodies. *, p<0.05; **, p<0.01, #, p<0.005; +, p<0.001; ++, p<0.0005.

To confirm that effects attributed to loss of p53 function are not strain specific, siRNA mediated knockdown of p53 was performed in MSCs derived from the FVB/N strain and resulting changes in oxygen sensitivity were quantified. As anticipated, transfection of MSCs cultured in 21% oxygen with a p53 specific but not a scrambled siRNA resulted in a significant (p<0.001) decrease in total cellular ROS levels (Figure 6A), which was commensurate with a significant (p<0.001) decrease in mitochondrial membrane permeability (Figure 6B). Other cellular measures of oxidative stress including changes in protein carbonyls and reduced glutathione levels were also normalized by knockdown of p53 (Figures 6C, D). Western blot analysis confirmed that the p53 specific but not scrambled siRNA reduced overall p53 protein levels in cells, as well as levels of MDM2 and BAX (Figure 6E). Importantly, knockdown of these proteins was clearly evident at 3 days but not by 7 days post-transfection, consistent with the kinetics of siRNA decay in cells. Nevertheless, p53 knockdown resulted in a significant (p<0.001) increase in cumulative cell yield over a 7 day time course (Figure 6F) and this change in proliferation was also clearly evident based on CFSE staining (Figure 6G). However, these cells exhibited oxygen-induced growth inhibition at later passages as anticipated based on the transient nature of the knockdown (not shown). Knockdown of p53 also abrogated oxygen-induced increases in apoptosis as evidenced by Annexin V and PI staining (Figure 6H). Finally, to confirm function of p53 under low oxygen conditions, MSCs derived from the all three mouse strains were culture-expanded in 5% or 21% oxygen and then dosed with 4 Gy of radiation. As anticipated, radiation exposure had a profound negative effect on the growth and viability of wild type MSCs from both FVB/n and C577BL/6 strains and this effect was independent of oxygen concentration (Supplemental Figure 3). In contrast, growth and viability of MSCs derived from p53 null mice were not significantly affected by radiation exposure.

Figure 6. Knockdown of p53 ameliorates oxygen-induced changes in ROS production, cell growth, and viability in wild type MSCs.

Figure 6

Open in a new tab

Immuno-depleted MSCs from FVB/N mice were cultured in 5% oxygen and transfected with a p53-specific or scrambled (control siRNA) siRNA. Transfected and control (mock-transfected) cells were then switched to 21% oxygen and 7 days later biochemical analyses were performed to quantify whole cell ROS (A), mitochondrial membrane permeability (B), protein carbonyl (C) and reduced glutathione (D) levels. Each experiment was conducted using three distinct replicate cultures from each experimental group and plotted values (mean ± SD) were determined from four replicates for each sample. E) Cell extracts were prepared from each experimental group at 3 and 7 days post-transfection and analyzed by Western blot using anti-p53, anti-BAX, anti-MDM2 and anti-GAPDH antibodies. F) Cumulative cell number after 7 days expansion of the indicated populations was calculated by counting. G, H) Mock transfected (red line, control) cells and those transfected with a p53-specific (black line) or control siRNA (green line) were labeled with CFSE (G) or Annexin V and PI (H) and examined by flow cytometry. #, p<0.005; +, p<0.001.

DISCUSSION

Mammalian cells possess a number of mechanisms to regulate production and consumption of oxygen in response to changes in its availability within the microenvironment. However, effects of oxygen appear to be cell-type specific. For example, hypoxic conditions have been shown to enhance paracrine activity, proliferation, differentiation potential and survival of human MSCs but induce apoptosis of aortic endothelial cells [40, 41]. In this study we demonstrate that exposure to atmospheric oxygen induced mitochondrial ROS production and p53 stabilization in mouse MSCs resulting in reduced cell viability and arrest of cell proliferation. These effects are similar to those reported in embryonic kidney cells in response to hypoxia, which included enhanced mitochondrial ROS production and Rho kinase activation resulting in p53 and HIF-1α stabilization [42]. The unique sensitivity of mouse MSCs to atmospheric oxygen may reflect the fact that the cells reside in bone marrow, which is known to be a low oxygen environment [43]. Indeed, several recent studies have showed that hypoxia promotes expansion of human MSCs [44] and maintains rodent MSCs in an undifferentiated state [45]. In our hands mouse MSCs expanded in 21% oxygen exhibited a reduced capacity for multi-lineage differentiation (not shown), which was attributed to reduced viability. Atmospheric oxygen also strongly induced BAX expression in MSCs, which is up regulated in response to cellular stress [46] and functions to inhibit the death repressor activity of BCL2 thereby promoting apoptosis [47]. BAX induction in response to oxygen stress in MSCs was strongly correlated with a decrease in cell viability, consistent with its known mechanisms of action.

Growth inhibition of MSCs following exposure to atmospheric oxygen was rapidly reversed in most cells following switch to 5% oxygen indicating that oxidative stress does not exclusively induce cellular senescence as recently suggested [48]. Moreover, the cellular response to oxidative stress was shown to be p53 dependent since oxygen-induced increases in ROS generation and decreases in cell growth and viability were not evident in MSCs derived from p53−/− mice or wild type cells following siRNA-mediated knockdown of p53. Importantly, growth rates of p53−/− MSCs were slightly faster in 5% vs. 21% oxygen indicating that other factors in addition to p53 may contribute to oxygen-induced growth regulation. The fact that oxygen-induced increases in ROS production and BAX were not evident in p53−/− MSCs or following p53 knockdown is also consistent with previous studies indicating that mitochondrial ROS homeostasis, membrane potential, and p53 activity are coordinately regulated [49]. Because the molecular mechanisms linking p53 activity to mitochondrial ROS generation are not fully elucidated, the cell culture system described herein may represent a useful tool to explore these mechanisms in more detail.

Established methods to isolate mouse MSCs yield cell populations that exhibit robust growth in atmospheric oxygen [20–26]. In contrast, immuno-depletion yields cells that are sensitive to oxygen-induced growth inhibition [18, 19]. A critical difference between these two isolation schemes is the period of time that bone marrow cells are culture expanded in vitro prior to MSC enrichment. For example, in our laboratory marrow cells are expanded for <10 days prior to immuno-depletion. In contrast, other labs reportedly cultivate plastic adherent marrow cells for weeks to months in order to enrich a subpopulation of MSCs [20–26]. Data presented herein explain why these two approaches yield MSC populations with greatly different sensitivities to atmospheric oxygen. Specifically, rodent cells exhibit a much higher frequency of immortalization in vitro as compared to human cells and p53 is mutated in the vast majority of immortalized rodent cell lines. Our results indicate that long-term expansion of bone marrow cells in atmospheric oxygen selects for MSC clones with reduced or absent p53 function, which allows escape from oxygen-induced growth inhibition. This conclusion is consistent with studies demonstrating that p53−/− MSCs exhibit only an incremental increase in growth rate as compared to wild type cells that were initially procured by culture expansion for 80 days prior to analysis [38] and that late passage mouse MSCs express high levels of mutant p53 protein [50].

CONSLUSION

We contend that established methods to isolate mouse MSCs select for clonally-derived subpopulations with reduced or absent p53 function, which facilitates rapid cell growth in atmospheric oxygen. Consequently, oxygen sensitivity provides a surrogate measure of p53 function in primary mouse MSCs and should be included as an important metric to evaluate cell quality. Moreover, since isoaltion methods that alter oxygen sensitivity may also affect other MSC functions, comparative studies of mouse MSCs that employ different isolation schemes should be cautiously interpreted.

Supplementary Material

Suppl Figure 1
Suppl Figure 2
Suppl Figure 3

ACKNOWLEDGEMENTS

This research was supported in part by a grant from the National Institute of Health to D.G.P. (R01 NS052301-01A2).

Footnotes

Author contribution S.B. performed research, collected and analyzed data; V. K. performed research, collected and analyzed data; J. C. performed research, collected and analyzed data; P. L. interpreted data and reviewed the manuscript; W-Z. L. performed research and collected data; L.A.O. interpreted data and reviewed the manuscript; D.G. P. conceived the study, designed research, analyzed data, supervised all experimentation, funded research, and wrote the paper.

Disclaimer: The authors indicate no potential conflicts of interest

REFERENCES

  • 1.Friedenstein AJ, Chailakhjan RK, Lalykina KS. The development of fibroblast colonies in monolayer cultures of guinea-pig bone marrow and spleen cells. Cell Tissue Kinet. 1970;3:393–403. doi: 10.1111/j.1365-2184.1970.tb00347.x. [DOI] [PubMed] [Google Scholar]
  • 2.Digirolamo CM, Stokes D, Colter D, et al. Propagation and senescence of human marrow stromal cells in culture: a simple colony-forming assay identifies samples with the greatest potential to propagate and differentiate. Br J Hematol. 1999;107:275–281. doi: 10.1046/j.1365-2141.1999.01715.x. [DOI] [PubMed] [Google Scholar]
  • 3.Sotiropoulou PA, Perez SA, Salagianni M, et al. Characterization of the optimal culture conditions for production of human mesenchymal stem cells. Stem Cells. 2006;24:42–47. doi: 10.1634/stemcells.2004-0331. [DOI] [PubMed] [Google Scholar]
  • 4.Schallmoser K, Rohde E, Reinisch A, et al. Rapid large-scale expansion of functional mesenchymal stem cells from unmanipulated bone marrow without animal serum. Tissue Eng Part C Methods. 2008;14:185–196. doi: 10.1089/ten.tec.2008.0060. [DOI] [PubMed] [Google Scholar]
  • 5.Iguchi A, Okuyama R, Koguma M, et al. Selective stimulation of granulopoiesis in vitro by established bone marrow stromal cells. Cell Struct Funct. 1997;22:357–364. doi: 10.1247/csf.22.357. [DOI] [PubMed] [Google Scholar]
  • 6.Witte PL, Robinson M, Henley A, et al. Relationship between B-lineage lymphocytes and stromal cells in long-term bone marrow cultures. Eur J Immunol. 1987;17:1473–1484. doi: 10.1002/eji.1830171014. [DOI] [PubMed] [Google Scholar]
  • 7.Obinata M, Okuyama R, Matsuda K-I, et al. Regulation of myeloid and lymphoid development of hematopoietic stem cells by bone marrow stromal cells. Leukemia & Lymphoma. 1998;29:61–69. doi: 10.3109/10428199809058382. [DOI] [PubMed] [Google Scholar]
  • 8.Gordon MY, Bearpark AD, Clarke D, et al. Haemopoietic stem cell subpopulations in mouse and man: Discrimination by differential adherence and marrow repopulating ability. Bone Marrow Transpl. 1990;5(Suppl):6–8. [PubMed] [Google Scholar]
  • 9.Kiefer F, Wagner EF, Keller G. Fractionation of mouse bone marrow by adherence separates primitive hematopoietic stem cells from in vitro colony-forming cells and spleen colony-forming cells. Blood. 1991;78:2577–2582. [PubMed] [Google Scholar]
  • 10.Winemann JP, Nishikawa S-I, Muller-Sieburg CE. Maintenance of high levels of pluripotent hematopoietic stem cells in vitro: effect of stromal cells and c-kit. Blood. 1993;81:365–372. [PubMed] [Google Scholar]
  • 11.Friedenstein AJ, Chailakhyan RK, Gerasimov UV. Bone marrow osteogenic stem cells: in vitro cultivation and transplantation in diffusion chambers. Cell Tissue Kinet. 1987;20:263–272. doi: 10.1111/j.1365-2184.1987.tb01309.x. [DOI] [PubMed] [Google Scholar]
  • 12.Simmons PJ, Masinovsky B, Longenecker BM, et al. Vascular cell adhesion molecule-1 expressed by bone marrow stromal cells mediates the binding of hematopoietic progenitor cells. Blood. 1992;80:388–395. [PubMed] [Google Scholar]
  • 13.Phinney DG, Kopen G, Isaacson RL, et al. Plastic adherent stromal cells from the bone marrow of commonly used inbred strains of mice: Variations in yield, growth, and differentiation. J Cell Biochem. 1999;72:570–585. [PubMed] [Google Scholar]
  • 14.Wang Q-R, Wolf NS. Dissecting the hematopoietic microenvironment. VIII. Clonal isolation and identification of cell types in murine CFU-F colonies by limiting dilution. Exp Hematol. 1990;18:355–359. [PubMed] [Google Scholar]
  • 15.Modderman WE, Vrijheid-Lammers T, Lowik CW, et al. Removal of hematopoietic cells and macrophages from mouse bone marrow cultures: isolation of fibroblast-like stromal cells. Expt Hematol. 1994;22:194–201. [PubMed] [Google Scholar]
  • 16.Short B, Brouard N, Driessen R, et al. Prospective isolation of stromal progenitor cells from mouse BM. Cytotherapy. 2001;3:407–408. doi: 10.1080/146532401753277265. [DOI] [PubMed] [Google Scholar]
  • 17.Gang EJ, Bosnakovski D, Fiqueiredo CA, Visser JW, Perlingeeiro RC. SSEA-4 identifies mesenchymal stem cells from bone marrow. Blood. 2007;109:1743–1751. doi: 10.1182/blood-2005-11-010504. [DOI] [PubMed] [Google Scholar]
  • 18.Baddoo M, Hill K, Wilkinson R, et al. Characterization of mesenchymal stem cells isolated from murine bone marrow by negative selection. J Cell Biochem. 2003;89:1235–1249. doi: 10.1002/jcb.10594. [DOI] [PubMed] [Google Scholar]
  • 19.Phinney DG. Isolation of mesenchymal stem cells from murine bone marrow by immunodepletion. Methods Mol Biol. 2008;449:171–186. doi: 10.1007/978-1-60327-169-1_12. [DOI] [PubMed] [Google Scholar]
  • 20.Al-Khaldi A, Eliopoulos N, Martineau D, et al. Postnatal bone marrow stromal cells elicit a potent VEGF-dependent neoangiogenic response in vivo. Gene Therapy. 2003;10:621–629. doi: 10.1038/sj.gt.3301934. [DOI] [PubMed] [Google Scholar]
  • 21.Sun S, Guo Z, Xiao X, et al. Isolation of mouse marrow mesenchymal progenitors by a novel and reliable method. Stem Cells. 2003;21:527–535. doi: 10.1634/stemcells.21-5-527. [DOI] [PubMed] [Google Scholar]
  • 22.Meirelles Lda S, Nardi NB. Murine marrow-derived mesenchymal stem cell: isolation, in vitro expansion, and characterization. Br J Haematol. 2003;123:702–711. doi: 10.1046/j.1365-2141.2003.04669.x. [DOI] [PubMed] [Google Scholar]
  • 23.Peister A, Mellad JA, Larson BL, et al. Adult stem cells from bone marrow (MSCs) isolated from different strains of inbred mice vary in surface epitopes, rates of proliferation, and differentiation potential. Blood. 2004;103:1662–1668. doi: 10.1182/blood-2003-09-3070. [DOI] [PubMed] [Google Scholar]
  • 24.Tropel P, Noel D, Platet N, et al. Isolation and characterization of mesenchymal stem cells from adult mouse bone marrow. Expt Cell Res. 2004;295:395–406. doi: 10.1016/j.yexcr.2003.12.030. [DOI] [PubMed] [Google Scholar]
  • 25.Li Y, Zhang C, Xiong F, et al. Comparative study of mesenchymal stem cells from C57BL/10 and mdx mice. BMC Cell Biol. 2008;9:24. doi: 10.1186/1471-2121-9-24. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Wu X, Chen S, Orlando SA, et al. p85α regulates osteoblast differentiation by cross-talking with MAPK pathway. J Biol Chem. 2011;286:13512–13521. doi: 10.1074/jbc.M110.187351. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Deryugina EI, Muller-Sieburg CE. Stromal cells in long-term cultures: Keys to the elucidation of hematopoietic development. Crit Rev Immun. 1993;13:115–150. [PubMed] [Google Scholar]
  • 28.Prowse KR, Greider CW. Developmental and tissue-specific regulation of mouse telomerase and telomere length. Proc Natl Acad Sci USA. 1995;92:4818–4822. doi: 10.1073/pnas.92.11.4818. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Wadhwa R, Sugihara T, Taira K, et al. The ARF-p53 senescence pathway in mouse and human cells. Histol. Histopathol. 2004;19:311–316. doi: 10.14670/HH-19.311. [DOI] [PubMed] [Google Scholar]
  • 30.Harvery DM, Levine AJ. P53 alteration is a common event in the spontaneous immortalization of primary BALB/c embryo fibroblasts. Genes Dev. 1991;5:2375–85. doi: 10.1101/gad.5.12b.2375. [DOI] [PubMed] [Google Scholar]
  • 31.Sherr CJ, DePinho RA. Cellular Senescence: Mitotic Clock or Culture Shock. Cell. 2000;102:407–410. doi: 10.1016/s0092-8674(00)00046-5. [DOI] [PubMed] [Google Scholar]
  • 32.Chiou M, Xu Y, Longaker MT. Mitogenic and chondrogenic effects of fibroblast growth factor-2 in adipose-derived mesenchymal cells. Biochem Biophys Res Commun. 2006;343(2):644–652. doi: 10.1016/j.bbrc.2006.02.171. [DOI] [PubMed] [Google Scholar]
  • 33.Russell KC, Phinney DG, Lacey MR, et al. In vitro high capacity assay to quantify the clonal heterogeneity in trilineage potential of mesenchymal stem cells reveals a complex hierarchy of lineage commitment. Stem Cells. 2010;28:788–798. doi: 10.1002/stem.312. [DOI] [PubMed] [Google Scholar]
  • 34.Busuttil RA, Rubio M, Dollé ME, et al. Oxygen accelerates the accumulation of mutations during the senescence and immortalization of murine cells in culture. Aging Cell. 2003;2:287–94. doi: 10.1046/j.1474-9728.2003.00066.x. [DOI] [PubMed] [Google Scholar]
  • 35.Chen HL, Pistollato F, Hoeppner DJ, et al. Oxygen tension regulates survival and fate of mouse central nervous system precursors at multiple levels. Stem Cells. 2007;25:2291–301. doi: 10.1634/stemcells.2006-0609. [DOI] [PubMed] [Google Scholar]
  • 36.Mazumdar J, O'Brien WT, Johnson RS, et al. O2 regulates stem cells through Wnt/β-catenin signaling. Nat Cell Biol. 2010;12:1007–1013. doi: 10.1038/ncb2102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Mohyeldin A, Garzon-Muvdi T, Quinones-Hinojosa A. Oxygen in stem cell biology: a critical component of the stem cell niche. Cell Stem Cell. 2010;7:150–161. doi: 10.1016/j.stem.2010.07.007. [DOI] [PubMed] [Google Scholar]
  • 38.Armesilla-Diaz A, Elvira G, Silva A. p53 regulates the proliferation, differentiation and spontaneous transformation of mesenchymal stem cells. Exp Cell Res. 2009:3598–3610. doi: 10.1016/j.yexcr.2009.08.004. [DOI] [PubMed] [Google Scholar]
  • 39.Rubio R, Garcia-Castro J, Gutierrez-Aranda I, et al. Deficiency in p53 but not retinoblastoma induces the transformation of mesenchymal stem cells in vitro and initiates leiomyosarcoma in vivo. Cancer Res. 2010;70:4185–94. doi: 10.1158/0008-5472.CAN-09-4640. [DOI] [PubMed] [Google Scholar]
  • 40.Das R, Jahr H, van Osch GJ, et al. The role of hypoxia in bone marrow-derived stem cells: considerations for regenerative medicine approaches. Tissue Eng Part B Rev. 2010;16:159–168. doi: 10.1089/ten.TEB.2009.0296. [DOI] [PubMed] [Google Scholar]
  • 41.Hung SC, Pochampally RR, Chen SC, et al. Angiogenic effects of human multipotent stromal cell conditioned medium activate the PI3K-Akt pathway in hypoxic endothelila cells to inhibit apoptosis, increase survival, and stimulate angiogenesis. Stem Cells. 2007;25:2363–2370. doi: 10.1634/stemcells.2006-0686. [DOI] [PubMed] [Google Scholar]
  • 42.Chandel NS. Mitochondrial regulation of oxygen sensing. Adv Exp Med Biol. 2010;661:339–354. doi: 10.1007/978-1-60761-500-2_22. [DOI] [PubMed] [Google Scholar]
  • 43.Chow DC, Wenning LA, Miller WM, et al. Modeling pO(2) distributions in the bone marrow hematopoietic compartment. II. Modified Kroghian models. Biophys J. 2001;81:685–696. doi: 10.1016/S0006-3495(01)75733-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Hung SP, Ho JH, Shih YR, Lo T, Lee OK. Hypoxia promotes proliferation and osteogenic differentiation potentials of human mesenchymal stem cells. J Orthop Res. 2011 Aug 1; doi: 10.1002/jor.21517. Doi: 10.1002/jor.21517 [Epub ahead of print] [DOI] [PubMed] [Google Scholar]
  • 45.Basciano L, Nemos C, Foliguet B, de Isla N, de Carvalho M, Tran N, Dalloul A. Long term culture of mesenchymal stem cells in hypoxia promotes a genetic program maintaining their undifferentiated and multipotent states. BMC Cell Biol. 2011;12:1–12. doi: 10.1186/1471-2121-12-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Chipuk JE, Kuwana T, Bouchier-Hayes L, et al. Direct activation of Bax by p53 mediates mitochondrial membrane permeabilization and apoptosis. Science. 2004;303:1010–1014. doi: 10.1126/science.1092734. [DOI] [PubMed] [Google Scholar]
  • 47.Oltvai ZN, Milliman CL, Korsmeyer SJ. Bcl-2 heterodimers in vivo with a conserved homolog, Bax, that accelerates programmed cell death. Cell. 1993;74:609–619. doi: 10.1016/0092-8674(93)90509-o. [DOI] [PubMed] [Google Scholar]
  • 48.Coutu DL, Francois M, Galipeau J. Inhibition of cellular senescence by developmentally regulated FGF receptors in mesenchymal stem cells. Blood. 2011;117:6801–6812. doi: 10.1182/blood-2010-12-321539. [DOI] [PubMed] [Google Scholar]
  • 49.Lebedeva MA, Eaton JS, Shadel G. Loss of p53 causes mitochondrial DNA depletion and altered mitochondrial reastic oxygen species heomeostasis. Biochim Biophys Acta. 2009;1787:328–334. doi: 10.1016/j.bbabio.2009.01.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Li H, Fan X, Kovi RC, et al. Spontaneous expression of embyronic factors and p53 point mutations in aged mesenchymal stem cells: A model of age-related tumorigenesis in mice. Cancer Res. 2007;67:10899–10898. doi: 10.1158/0008-5472.CAN-07-2665. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Suppl Figure 1
NIHMS482521-supplement-Suppl_Figure_1.tif (7MB, tif)
Suppl Figure 2
NIHMS482521-supplement-Suppl_Figure_2.tif (7.4MB, tif)
Suppl Figure 3
NIHMS482521-supplement-Suppl_Figure_3.tif (16.3MB, tif)

RESOURCES