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First published online October 27, 2005
Stem Cells Vol. 24 No. 2 February 2006, pp. 416 -425
doi:10.1634/stemcells.2005-0121; www.StemCells.com
© 2006 AlphaMed Press

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THE STEM CELL NICHE

Hypoxia and Serum Deprivation-Induced Apoptosis in Mesenchymal Stem Cells

Weiquan Zhu, Jinghai Chen, Xiangfeng Cong, Shengshou Hu, Xi Chen

Research Center for Cardiovascular Regenerative Medicine, Cardiovascular Institute and Fu Wai Hospital, Chinese Academy of Medical Sciences and Peking Union Medical College, Beijing, People’s Republic of China

Key Words. Mesenchymal stem cells • Hypoxia/serum deprivation • Apoptosis • Mitochondria

Correspondence: Xi Chen, Ph.D., and Shengshou Hu, M.D., Research Center for Cardiovascular Regenerative Medicine, Cardiovascular Institute and Fu Wai Hospital, Chinese Academy of Medical Sciences & Peking Union Medical College, Beijing, 100037, People’s Republic of China. Telephone: 086-10-68331762; Fax: 086-10-68331762; e-mail: chenxifw{at}yahoo.com.cn and huss{at}163bj.com


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In recent years, the understanding that regeneration progresses at the level of the myocardium has placed stem cell research at the center stage in cardiology. Despite an increasing interest in cell transplant research, relatively little is known about the biochemical regulation of the stem cell itself after transplantation into an ischemic heart. We demonstrated here, using rat mesenchymal stem cells (MSCs), that cells undergo caspase-dependent apoptosis in response to hypoxia and serum deprivation (SD), which are both components of ischemia in vivo. In particular, the treated cells exhibited mitochondrial dysfunction, including cytochrome C release, loss in {Delta}{Psi}m, and Bax accumulation, but in a p53-independent manner. Although the cells treated by hypoxia/SD possess the activity of caspase-8, zIEDT-fmk, a specific caspase-8 inhibitor, failed to inhibit cell apoptosis induced in our system. Taken together, our findings indicate that MSCs are sensitive to hypoxia/SD stimuli that involve changes in mitochondrial integrity and function but are potentially independent of caspase-8.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Mesenchymal stem cells (MSCs) are self-renewing, clonal precursors of nonhematopoietic tissues [1], which can be cultured in vitro while retaining the potential to give rise to osteoblasts [2], chondrocytes [2], astrocytes [3], neurons [4], and skeletal muscle [5]. Several groups have reported that putative MSCs derived from bone marrow can differentiate into cardiac myocytes in vitro [6] and in vivo [7, 8]. However, it is evident that transplanted stem cells do not survive well, with, for instance, more than 99% of MCSs injected into the left ventricle of CB17 SCID/beige adult mice dying within 4 days of injection. This may reflect the harsh, proapoptotic microenvironment of the infarcted heart, which may not be conducive of MSC survival [9]. Thus, protection of stem cells against apoptosis is critical for successful cellular therapy. It is therefore becoming essential to have a clear understanding of the events and factors that may predispose MSCs to undergo apoptosis in ischemia tissue. Although unclear, it is feasible that poor blood supply and low oxygen tension, together with as-yet-unidentified cellular events, may lead to MSC apoptosis in the ischemic myocardium [10].

Although a fully comprehensive model of apoptosis has not yet been assembled, two general pathways have been delineated [11]. The mitochondrial, or intrinsic pathway, involves the release of proteins, such as cytochrome C and Smac/DIABLO from the intermembrane space of mitochondria [12]. This is usually in association with alterations of mitochondrial membrane proteins, such as the Bcl-2 family of proteins [13]. Generally, cytochrome C binds Apaf-1 in the cytosol, leading to the oligomerization and activation of pro-caspase-9. This, in turn, cleaves and activates pro-caspase-3 [14], which is critical for the effector stage of apoptosis. In the death receptor (DR) (e.g., TNFR60, Fas/Apo1, or TRAIL-R/Apo2) or extrinsic pathway, ligand binding to DR leads to the recruitment and activation of pro-caspase-8 through specific adaptor molecules, such as Fas-associated death domain protein (FADD) [15]. In different types of cells, activation of caspase-8 either cleaves and activates pro-caspase-3 directly or cleaves the proapoptotic Bcl-2 protein Bid to tBid, which then activates the mitochondrial pathway, resulting in the activation of caspase-3 and other effector caspases [16, 17].

Several studies have shown that embryonic and perinatal progenitor cells undergo apoptosis via the caspase pathway and/or involvement of the Fas receptor [18]. Recently, Tamm et al. [19] demonstrated that neural stem cells exposed to apoptosis inducers, such as oxidative stress, stauroaporine, or 2,3-dimethoxy-1,4-naphthoquinone, activates the mitochondrial apoptotic pathway, with cytochrome C release and subsequent activation of caspase-3, resulting in apoptosis. However, exposure to Fas monoclonal antibody (mAb) failed to induce apoptosis despite the presence of the Fas receptor in adult stem cells.

To date, studies of MSCs have focused on characterizing restoration of heart function, the potential of differentiation, and migration. However, little is known about the mechanisms that result in the essential processes of cell response during cell transplantation. In the present study, we have therefore investigated the effects of hypoxia and serum deprivation, two components of ischemia [20–22], and demonstrated that MSCs undergo caspase-dependent apoptosis induced by hypoxia plus serum deprivation (SD) in vitro. Hypoxia/SD-induced cell apoptosis is involved in mitochondrial dysfunction but caspase-8-independent.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Iscove’s modified Dulbecco’s medium (IMDM) and fetal bovine serum were from Gibco (Grand Island, NY, http://www.invitrogen.com). Antibodies used were as follows: mouse monoclonal antibodies, anti-rat Bax, and anti-cytochrome c (Santa Cruz Biotechnology, Santa Cruz, CA, http://www.scbt.com); rabbit polyclonal antibody, anti-rat Fas (Santa Cruz Biotechnology); rabbit monoclonal antibody, anti-rat caspase-3 (NeoMarkers, Fremont, CA, http://www.labvision.com). Horse-radish peroxidase-conjugated secondary antibodies to mouse or rabbit were obtained from Santa Cruz Biotechnology. Ac-DEVD-pNA and Ac-IEDT-pNA were from BIOMOL Research Labs (Plymouth Meeting, PA, http://www.biomol.com). General caspase inhibitor zVAD-fmk and caspase-8 inhibitor zI-ETD-fmk were from R&D systems (Minneapolis, http://www.rndsystems.com).

In Vitro Model of MSC Apoptosis
MSCs were prepared from 80-g Sprague-Dawley rats as described by Fridenshtein et al. [23] and other investigators [24–26] based on their adherence to tissue culture surfaces. All MSCs were grown in IMDM containing 15% heat-inactivated fetal bovine serum and 100 U/ml penicillin-streptomycin and were incubated at 37°C in a humidified atmosphere containing 5% CO2 and 95% air. All cells in the testing were first passage. Select conditions that may be observed in ischemia were simulated by serum deprivation of the culture medium and by hypoxia. Cells were washed with serum-free IMDM and placed in serum-free medium and then incubated in a sealed, hypoxic GENbox jar fitted with a catalyst (BioMérieux, Marcy l’Etoile, France, http://www.biomerieux.com) to scavenge free oxygen. Oxygen tension in the medium was measured using a blood gas analyzer and was found to be 33.5 mm Hg within 0.5 hour after being transferred into the hypoxic chamber and maintained at approximately 22 to 24 mm Hg over the experimental time.

Assessment of Morphological Changes
Chromosomal condensation was assessed using the chromatin dye Hoechst 33342 (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com). Cells were fixed for 30 minutes in phosphate-buffered saline (PBS) containing 1% glutaraldehyde. After fixing at room temperature, the cells were washed twice with PBS and then exposed to 5 µg/ml Hoechst 33342 in PBS for 30 minutes at room temperature. All samples were observed using a fluorescence microscope. Apoptotic cells were characterized by morphological alteration such as condensed nuclei and cell shrinkage.

Measurement of Cell Apoptosis and Mitochondrial Membrane Potential
Phosphatidylserine (PS) exposure on the outer leaflet of the plasma membrane was detected using the fluorescent dye Annexin V-FITC Apoptosis Detection Kit (Oncogene, San Diego, http://www.oncogene.com) according to the manufacturer’s instructions. In brief, cells were rinsed with ice-cold PBS and then resuspended in 200 µl of binding buffer. Ten microliters of Annexin V stock solution was added to the cells and incubated for 30 minutes at 4°C. The cells were then further incubated with 5 µl propidine iodide (PI) and were immediately analyzed on a FACSC-LSR (Becton, Dickinson and Company, San Jose, CA, http://www.bd.com) equipped with CellQuest (Becton, Dickinson and Company) software; approximately 1 to 2 x 104 cells were analyzed in each of the samples.

The mitochondrial transmembrane potential ({Delta}{Psi}m) was analyzed using {Delta}{Psi}m-specific stain Rhodamine 123 (Sigma-Aldrich). In brief, at the indicated time points, 105 cells were stained in a solution containing 0.1 µmol/l Rhodamine 123 for 30 minutes at 37°C. Staining was quantified by scatter characteristic using a flow cytometer EPICS XL from Beckman Coulter (Fullerton, CA, http://www.beckmancoulter.com); approximately 1 to 2 x 104 cells were analyzed in each of the samples.

Cell Extract Preparation and Western Blot Analysis
For analysis of protein levels, stimulated cells were rinsed twice with ice-cold PBS and then lysed with ice-cold lysis buffer (1% Triton X-100, 20 mmol/l HEPES [pH 7.5], 5 mmol/l MgCl2, 1 mmol/l EDTA, 1 mmol/l EGTA, 1 mmol/l DTT, 1 mmol/l phenylmethane-sulfonylfluoride, and 1 mg/ml each of leupeptin, aprotinin, and pepstatin for 30 minutes). Cell lysates were centrifuged at 14,000g for 10 minutes at 4°C; the supernatant was then mixed with 5 x SDS sample buffer, boiled for 5 minutes, and separated through 8% to 15% SDS-PAGE gels. After electrophoresis, the proteins were transferred to nylon membranes by electrophoretic transfer. The membranes were blocked in 5% skim milk for 2 hours, rinsed, and incubated overnight at 4°C with primary antibody in 5% skim milk. Excess antibody was then removed by washing the membrane in PBS/0.1% Tween 20, and the membranes were incubated for 2 hours with horseradish peroxidase-conjugated secondary antibodies. After washes in PBS/0.1% Tween 20, bands were visualized by enhanced chemoluminescence and exposed to radiography film.

For the analysis of cytochrome C release from mitochondria, control and treated cells were harvested and washed twice with ice-cold PBS. Cells were resuspended in 50 µl lysis buffer (0.01% digitonin, 10 mmol/l HEPES, pH 7.4, 70 mmol/l sucrose, 210 mmol/l D-mannitol, 5 mmol/l succinate, 0.2 mmol/l EGTA, 0.15% bovine serum albumin). After 5 minutes, the cells were centrifuged at 15,000g for 5 minutes at 4°C, and the cytosolic fraction contained in the supernatant was removed and kept at –20°C. The residual pellet was resuspended in 50 µl of the digitonin lysis buffer described above, snap-frozen in liquid nitrogen, and quick thawed to complete the lysis. The membrane fraction and cytosolic fraction were then subjected to Western blot analysis for cytochrome C release.

Immunocytochemistry
Cells were fixed with 2% paraformaldehyde buffer containing 78 mmol/l L-lysine and 10 mmol/l NaIO4 for 20 minutes at room temperature and washed with PBS three times. Cells were then incubated in 0.3% Triton X-100 for 5 minutes and blocked in 10% goat serum albumin for 20 minutes, rinsed, and incubated for 60 minutes at room temperature with primary antibody against cytochrome C. Cells were subsequently rinsed with PBS and incubated with fluorescein isothiocyanate (FITC)-conjugated secondary antibody for 45 minutes at room temperature. After washing, cells were stained with Hoechst 33342 and examined using an Olympus FV500 (FX500+IX81) laser confocal microscope system (Tokyo, http://www.olympus-global.com).

Determination of Caspase Activity
Caspase activity was measured using the following pNA-derived chromogenic substrates for caspases: DEVD and IETD were used as preferred substrates for caspase-3 and -8, respectively. In brief, cells were lysed on ice in 1% Triton X-100, 20 mmol/l HEPES (pH 7.5), 5 mmol/l MgCl2, 1 mmol/l EDTA, 1 mmol/l EGTA, 1 mmol/l DTT, 1 mmol/l phenylmethane-sulfonylfluoride, and 1 mg/ml each of leupeptin, aprotinin, and pepstatin for 30 minutes and centrifuged at 14,000g for 3 minutes. Protease assays included 20 mg of protein (in 20 ml of lysis buffer), 80 ml of reaction buffer (100 mmol/l HEPES, pH 7.6, 20% glycerol, 1 mmol/l DTT, and 0.1 mmol/l EDTA), and 1 µl (100 µmol/l final concentration) of 10 mmol/l pNA peptide substrates. Samples were then incubated for an additional 2 hours at 37°C, and DEVD/IETD cleavage was monitored by enzyme-catalyzed release of pNA by determining absorbance at 405 nm in a microtiter plate reader.


   RESULTS
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 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effects of Serum Deprivation and Hypoxia in MSC Apoptosis
During ischemia, multiple changes contribute to cellular death. Among these are deprivation of nutrients, growth and survival factors, and oxygen. To study the effect of these stimuli, MSCs were exposed to culture conditions represented by hypoxia and SD over different periods of time and characteristics of cell death were analyzed by morphological means—Hoechst 33342 and flow cytometric analysis. As shown in Figure 1AGo, most cells from the control group (normoxia) had big, regular nuclei, with only a few showing apoptotic nuclei with condensed chromatin. In the cells exposed to hypoxia/SD, there was clear evidence of chromatin condensation together with a clear decrease in cell size (Fig. 1BGo), with both features being characteristic of apoptosis. Cell death was also measured using fluorescent dye Annexin V-FITC, which binds to phosphatidylserine residues that are redistributed from the inner to the outer leaflet of the cell membrane as an early event in apoptosis. After loss of membrane integrity, PI can enter the cell and intercalate into DNA. Figures 1C and 1DGo show the percentages of Annexin V-stained and PI-stained cells in response to hypoxia/SD treatment (3 to 24 hours). Processing of Annexin V+/PI– occurred within 3 hours of SD/hypoxia and reached its maximal level within 6 hours. However, with a longer treatment of hypoxia/SD (24 hours), the population of Annexin V+/PI+ cells was much greater than that of Annexin V+/PI– or Annexin V–/PI+ cells.


Figure 1
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  		Figure 1. Serum deprivation (SD)/hypoxia triggers apoptosis in mesenchymal stem cells (MSCs). MSCs were incubated for 3, 6, 9, and 24 hours in hypoxia/SD. Apoptosis was then determined by detection of cells with apoptotic nuclear morphology using fluorescence microscopy upon Hoechst 33342 staining (A) and by fluorescence-activated cell sorter (FACS) analysis using light-scatter characteristics (B). (C, D): Apoptosis was then quantified by FACS analysis after staining with Annexin V and propidine iodide (PI). Viable cells are Annexin V–/PI–. The Annexin V+/PI– cells are early in the apoptotic process, whereas the Annexin V+/PI+ cells have lost cell membrane integrity and have taken up PI. Necrotic cells show Annexin V–/PI+. The results are presented as fold changes compared with corresponding control cells. Each data point represents mean ± standard deviation of three independent experiments. *p < .01 vs. control.

 
These data suggest that serum deprivation and hypoxia, both components of ischemia in vivo, induced apoptosis in MSCs. As a result of these findings, we adopted in the following experiments a model of simulated ischemia consisting of MSCs cultured in the conditions of hypoxia and serum deprivation.

Role of Caspase in Hypoxia/SD-Induced MSC Apoptosis
We tested the ability of serum and oxygen deprivation to induce caspase activation, an early marker of apoptosis. The combination of SD and hypoxia induced a time-dependent cleavage of the 35-kDa pro-caspase-3, resulting in the appearance of the 19-kDa active form of this caspase (Fig. 2AGo). Moreover, to further strengthen that hypoxia/SD-induced processing of caspase-3 was associated with an increase in activity of caspase-3, we monitored caspase-3 activity with the substrate DEVD. The proteolytic cleavage of Ac-DEVD-pNA yields a signal that can be used to assess the level of caspase-3 activity in MSCs during SD/hypoxia. As shown in Figure 2BGo, caspase-3 activity increased after the hypoxia/SD treatment of cells, plateauing within 6 hours thereafter. To determine the functional importance of caspase activation in hypoxia/SD-induced MSCs apoptosis, we examined the effect of the general caspase inhibitor zVAD-fmk on hypoxia/SD-induced PS exposure. The treatment of MSCs with zVAD-fmk at concentrations ranging from 10 to 100 µmol/L inhibited hypoxia/SD-induced apoptosis, as evident from flow cytometric analysis (Figs. 2C, 2DGo). Treatment with 100 µmol/l zVAD-fmk inhibited hypoxia/SD-induced cell apoptosis, completely confirming the functional importance of caspase activation in this model.


Figure 2
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  		Figure 2. Serum deprivation (SD)/hypoxia-induced cell apoptosis is caspase dependent. Mesenchymal stem cells (MSCs) were incubated in hypoxia/SD. Analysis for procaspase-3 cleavage detected by immunoblotting (A) and caspase-3 activity determined by colorimetric assay (B). (C, D): Cells were incubated with zVAD-fmk for 1 hour before hypoxia/SD treatment for 6 hours. Apoptosis was quantified by fluorescence-activated cell sorter analysis after staining with Annexin V and propidine iodide. **p < .01 vs. control.

 
To delineate the individual contribution of serum deprivation and hypoxia to MSC apoptosis, MSCs were subjected to hypoxia and SD alone or in combination. As shown in Figure 3AGo, SD induced significant early apoptosis (Annexin V+/PI–) within 6 and 24 hours, even in the absence of hypoxia. The combination of hypoxia and SD modestly increased the level of PS exposure compared with SD MSCs alone. Figure 3BGo shows that within 24 hours, late apoptosis induced by SD alone (Annexin V+/PI+) was more pronounced than that induced by hypoxia alone. In the presence of serum, incubation of cells for up 6 hours under hypoxic conditions failed to induce caspase-3 activation (Fig. 3CGo). These results suggest that serum deprivation is the stronger one in MSCs between the two stimuli in this system.


Figure 3
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  		Figure 3. Effect of hypoxia of serum deprivation (SD) alone or a combination of both on cell apoptosis. Cultured mesenchymal stem cells (MSCs) were subjected to hypoxia and SD alone or in combination for 6 hours or 24 hours, and then cell apoptosis was quantified by fluorescence-activated cell sorter analysis after staining with Annexin V and propidine iodide (A, B). (C): Caspase-3 activity was examined after 6-hour incubation. The results are presented as fold changes compared with corresponding control cells. Each data point represents mean ± standard deviation of three independent experiments. *p < .05; **p < .01 vs. control.

 
Hypoxia/SD Induces MSC Mitochondrial Dysfunction
Apoptosis may involve mitochondrial dysfunction, and as a result we have evaluated whether hypoxia/SD provokes mitochondrial release of cytochrome C. Immunostaining was performed with an antibody against cytochrome C. The results obtained reveal the typical mitochondrial cytochrome C release pattern documented by others. Untreated cells exhibited clear dot-like patterns distributed distinctly throughout the cell, indicating that cytochrome C was localized in the mitochondria under normal conditions (Figs. 4Aa–4AcGo). In contrast, treated cells showed an apoptotic nuclear morphology and revealed a diffuse fluorescence, indicating the presence of cytochrome C throughout the cytosol due to its release from mitochondria (Figs. 4Ad–4AfGo). To further verify these findings, we performed a fractionation of treated MSCs into cytosolic and membrane fractions and subjected these to Western blot analysis. Western blot analysis showed that cytochrome C was detected in the cytosolic fraction after treatment with hypoxia/SD. These data confirm our immunocytochemical results on the cytochrome C release.


Figure 4
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  		Figure 4. Hypoxia/serum deprivation (SD) induces mesenchymal stem cell (MSC) mitochondrial dysfunction. (A): Immunocytochemistry shows (a) control cells with big, regular nuclei stained by Hoechst 33342 and (b) cytochrome C immunoreactive dot-like structure. (c): Result of superimposing (a) and (b). After exposure to hypoxia/SD for 6 hours, (d) apoptotic nuclear morphology was found and (e) cytochrome C localization was visualized in the cytosol. (f): Result of superimposing (d) and (e). (B): Western blot of cytosolic and membrane fractions of MSCs showing a clear cytochrome C release after treatment with hypoxia/SD. (C, D): Changes in {Delta}{Psi}m detected by flow cytometry of Rhodamine 123-stained cells. Each data point represents mean ± standard deviation of three independent experiments. **p < .01. MSCs were incubated for 3, 6, 9, or 24 hours in hypoxia/SD. (E): Western blot of cytosolic and membrane fractions of MSCs shows Bax translocation to the mitochondrial after treatment by hypoxia/SD. (F): Analysis of p53 expression and phosphorylation level of p53 Ser 15 were detected by Western blot.

 
Because changes in mitochondrial function resulting from opening of the mitochondrial PT and loss in {Delta}{Psi}m, can initiate apoptosis, we next ascertained whether the observed hypoxia/SD-induced cell apoptosis was associated with a loss in MSC {Delta}{Psi}m. For these experiments, we use the potential sensitive dye Rhodamine 123 to monitor changes in {Delta}{Psi}m. In contrast to control group, cells treated with hypoxia/SD did not display any changes (p = .149) in membrane potential within 3 hours of incubation. Prolonged incubations resulted in a time-dependent decrease of {Delta}{Psi}m (Figs. 4C, 4DGo). Those results suggest that mitochondrial dysfunction may be involved in the apoptotic process of MSCs induced by hypoxia/SD.

Effects of Hypoxia/SD on Translocation of Bax in MSCs
The translocation of proapoptotic Bax plays important roles in the regulation of mitochondrial cytochrome C release and loss in {Delta}{Psi}m. The effect of hypoxia/SD on the translocation of this protein was therefore examined. Western blot analysis revealed that levels of Bax in the membrane fraction continued to increase after treatment with hypoxia/SD (Fig. 4EGo). This was accompanied by a parallel decrease in Bax in the cytosolic components of cell extracts.

Hypoxia in some experimental models may increase the activation of proapoptotic p53, and p53 itself can induce Bax translocation. To identify possible alterations in the function of p53 in response to hypoxia/SD in MSCs, we examined p53 phosphorylation at Ser 15, which reflects its functional response to cellular stress, such as DNA damage, and leads to apoptosis. We found that p53 phosphorylation occurred in the control cells and did not change after treatment with hypoxia/SD (Fig. 4FGo). These data suggest that in the present model of simulated ischemia, the induction of MSC apoptosis is associated with the activation of the mitochondrial pathway and may involve Bax but may be independent of p53.

Role of Caspase-8 in Hypoxia/SD-Induced MSC Apoptosis
Activation of the downstream caspase cascade may take place within death receptor complexes of the cytoplasmic membrane involving the activation of the initiator caspase-8. Therefore, we examined the expression of the death receptor-Fas by Western blot analysis. In Figure 5AGo, hypoxia/SD induced Fas and Fas-L expression under conditions where significant apoptosis was induced. Moreover, we measured the activity of caspase-8 by the cleavage of the specific peptide substrates in MSCs treated by hypoxia/SD. Figure 5BGo shows that hypoxia/SD induced a time-dependent increase in caspase-8 activity that was first evident after 3 hours of incubation, reached a maximal level within 6 hours, but declined back to basal levels at 24 hours of incubation.


Figure 5
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  		Figure 5. Mesenchymal stem cells (MSCs) were incubated for 3, 6, 9, and 24 hours in hypoxia/serum deprivation (SD). Analysis of Fas and Fas-L expression were detected by immunoblotting (A), and caspase-8 activity was detected by colorimetric assay (B). Cells were incubated with zIETD-fmk for 1 hour before hypoxia/SD, after 6 hours of treatment. (C): Analysis for pro-caspase-3 cleavage was detected by immunoblotting. (D): Apoptosis was detected by cell size and quantified after staining with Annexin V and propidine iodide by fluorescence-activated cell sorter analysis. (E): Caspase-8 activity was detected by colorimetric assay. Each data point represents mean ± standard deviation of three independent experiments. **p < .01.

 
To further clarify the functional contribution of the death receptor signaling in our model, we used a specific caspase-8 inhibitor, zIEDT-fmk. Interestingly, neither cell apoptosis nor caspase-3 cleavage was affected by zIEDT-fmk (Figs. 5C, 5DGo). To test whether the concentration of caspase-8 inhibitor used in this study was optimal, the activity of caspase-8 was evaluated. In Figure 5EGo, treatment with zIEDT-fmk (50 µmol/l) significantly reduced hypoxia/SD-induced caspase-8 activity.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
MSCs have shown great promise in regenerating and repopulating the damaged myocardium, restoring its function [27, 28]. However, the use of this approach is limited by the fact that most of the transplanted MSCs are readily lost, potentially triggered by the ischemic environment in vivo. However, the precise events leading to MSC loss are unclear, and very little is known about the cellular mechanisms that may mediate this process. We were therefore interested in unraveling why transplanted MSCs fail to survive in ischemic tissue and identifying the factors and examining the mechanisms that may trigger this response. Because of poor blood supply and low oxygen tension in the infarcted heart, cell death due to survival growth factor withdrawal and hypoxia is of considerable interest and significance [20, 21]. The MSCs in this study were treated with serum deprivation and hypoxia. To our knowledge, the present study is the first to describe the dynamic characteristic of MSC death after treatment with hypoxia/SD and to explore the mechanism of the apoptosis.

Our experiments show that hypoxia/SD triggers apoptotic MSC death, as detected by nuclear shrinkage, chromatin condensation, and decrease in cell size, in a time-dependent manner. Future evidence of apoptosis is the PS translocation to the cell surface and loss of membrane integrity. These were confirmed with Annexin V and PI labeling. These are in agreement with the study by Mangi et al. [28] indicating that transplanted MSCs may be lost undergoing apoptosis induced by extremely harsh microenvironment in the infarcted heart. Mangi et al.’s evidence was obtained by DNA ladder, TUNEL, and Bax/Bcl-2 expression of MSCs after 24 hours of hypoxia in vitro and injection into an infarcted heart in vivo. Further, we described the dynamic process in the different appearances of cell death: early apoptosis (Annexin V+/PI–) occurs mainly before 9 hours of treatment, and late apoptosis (Annexin V+/PI+) increases markedly at 24 hours. Moreover, our data demonstrated caspase-3 activation and the presence of activated caspase-3 fragments in response to hypoxia/SD. Additionally, pretreatment with the general caspase inhibitor zVAD-fmk completely blocked apoptotic cell death in MSCs. These combined findings confirm that hypoxia/SD-induced apoptosis occurs via a caspase-dependent mechanism in MSCs.

The apoptotic model of cultured MSCs used in this study combines the following two components of ischemic injury [20–22]: survival growth factor withdrawal and hypoxia. However, serum deprivation seems to be the more predominant influencing factor because apoptosis was more pronounced under this condition than in the presence of hypoxia alone. Thus, MSCs may undergo apoptosis in the absence of defined growth factors rather than as a consequence of hypoxia. In this regard, several trophic factors could be used to pretreat the cells before transplantation or could indeed be applied together with the cells during transplantation. For instance, factors such as interleukin-6 and insulin-like growth factor 1 [20, 29], which have been shown to be cardioprotective and to inhibit cardiac myocyte apoptosis, may be used to protect not only transplanted MSCs but also cardiac myocytes. These studies are pending and may prove beneficial in future MSC therapy.

In addition to being crucial for energy production, mitochondria are also known to act as key regulators of apoptosis in several cell types [30]. The release of proteins from mitochondria, including cytochrome C, has been demonstrated to be involved in the subsequent activation of caspase cascade during apoptosis [31]. Using an antibody specific for cytochrome C, we found that hypoxia/SD induced cytochrome C release in MSCs. We also discovered the loss of mitochondrial membrane potential and Bax translocation from the cytosol to the mitochondria during apoptosis. Hypoxia/SD may lead to mitochondrial translocation of Bax, which may associate with components of the mitochondrial permeability transition pore, causing loss of the mitochondrial membrane potential. These results are consistent with several earlier studies showing that perturbation of mitochondrial functions could contribute to apoptosis [32]. In our apoptosis model, the accumulation of Bax and dysfunction of mitochondria were completely independent of p53 protein as there were no changes observed on the phosphorylated level of p53 Ser 15 during exposure of MSCs to hypoxia/SD. The role of p53 in apoptosis is highly dependent on cell type and cell context [20, 33], and several pathways of apoptosis are independent of p53 [34–36].

It has been suggested that caspase-8 signaling is necessary and sufficient for apoptosis of cardiomyocytes subjected to hypoxia/SD [21]. However, the fact that inhibition of the activity of caspase-8 did not affect the ischemia-induced apoptosis in MSCs indicated that the death receptor pathway may not be involved or, if it is, may act independently of caspase-8 in the process of cell apoptosis.

Multiple death receptors (e.g., TNFR60, Fas/Apo1, or TRAIL-R/Apo2) and their adapter FADD can lead to cell apoptosis as well as other physiological processes (e.g., cell outgrowth and acceleration of functional recovery) after cell injury in a caspase-8-independent manner. This may occur through activation of the nuclear factor (NF)-{kappa}B pathway [37, 38], serine protease-dependent pathway [39], or mitogen-activated protein kinase (MAPK) pathway [19]. In this study, we actually detected cellular changes, which are characteristic of the death receptor pathway. For instance, we were able to demonstrate caspase-8 activity and show a close correlation between this and changes in caspase-3 activity together with the translocation of PS.

It is clear now that once the caspase network becomes fully activated, the processing of an individual caspase is not indicative of the processing of an upstream apoptosis-signaling pathway [40, 41]. The finding that caspase-8 is activated during hypoxia/SD-induced apoptosis may simply reflect a secondary activation of caspase-8 during the execution phase of apoptosis [42], analogous to the observation made for radiation- and drug-induced apoptosis, where caspase-8 acts during execution of apoptosis rather than during its induction [43, 44]. Thus, the possibility that extrinsic pathway contributes to the specific activity measurements presented in the current study is difficult to completely exclude, and this should be the focus of future studies.

Other limitations of this study should also be acknowledged. We did not identify other pathways (e.g., NF-{kappa}B pathway, MAPK pathway, or protein kinase C pathway) that can modulate cell apoptosis under some circumstances [45–48]. Although defining the functional importance of these various pathways will be a complex undertaking, these studies may prove valuable. Finally, we have also focused on a simple in vitro model of apoptosis induced by hypoxia/SD in MSCs. The relevance of these findings to the transplanted MSCs in an intact heart remains to be demonstrated.

In conclusion, the data presented strongly suggest that survival growth factor withdrawal and hypoxia cause apoptosis via the caspase-dependent manner in transplanted MSCs. In this hypoxia/SD-induced apoptosis model, the mitochondrial release of cytochrome C and loss in {Delta}{Psi}m, which is regulated by translocation of Bax, result ultimately in the activation of caspase, nuclear condensation, and other apoptotic features. The data also imply that caspase-8 may not be involved in the hypoxia/SD-induced cell apoptosis, although the cellular events characteristic of death receptor pathway, such as Fas and Fas-L expression, were observed.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
This work was supported by grants from the National Natural Science Foundation of China (nos. 30370524 and 30271290) and from the National Science Fund for Distinguished Young Scholars (no. 30125039). The authors are indebted to Dr. Anwar Y. Baydoun for expert review of the manuscript.

DISCLOSURES
The authors indicate no potential conflicts of interest.


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Received March 17, 2005; accepted for publication August 8, 2005.



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