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First published online October 11, 2007
Stem Cells Vol. 26 No. 1 January 2008, pp. 135 -145
doi:10.1634/stemcells.2007-0098; www.StemCells.com
© 2008 AlphaMed Press

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THE STEM CELL NICHE

Lysophosphatidic Acid Protects Mesenchymal Stem Cells Against Hypoxia and Serum Deprivation-Induced Apoptosis

Jinghai Chena, Anwar R. Baydounb, Ruixia Xua, Linzi Denga, Xuebin Liua, Weiquan Zhua, Linhui Shia, Xiangfeng Conga, Shengshou Hua, Xi Chena

aResearch Center for Cardiovascular Regenerative Medicine, The Ministry of Health of China, Cardiovascular Institute and Fu Wai Hospital, Chinese Academy of Medical Sciences and Peking Union Medical College, Beijing, People's Republic of China;
bUniversity of Hertfordshire, Faculty of Health and Human Sciences, School of Life Sciences, College Lane, Hatfield, United Kingdom

Key Words. Lysophosphatidic acid • Mesenchymal stem cells • Hypoxia/serum deprivation • Apoptosis • Survival

Correspondence: Correspondence: Xi Chen, Ph.D. and Shengshou Hu, M.D., Research Center for Cardiovascular Regenerative Medicine, the Ministry of Health of China, 167 Beilishilu, Beijing 100037, People's Republic of China. Telephone: 86-10-88398584; Fax: 86-10-88398584; e-mail: chenxifw{at}yahoo.com.cn and e-mail: huss{at}vip.sohu.com

Received on February 25, 2007; accepted for publication on September 25, 2007.

First published online in STEM CELLS EXPRESS  October 11, 2007.

   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Bone marrow-derived mesenchymal stem cells (MSCs) have shown great promise for cardiac repair. However, poor viability of transplanted MSCs within the ischemic heart has limited their therapeutic potential. Our previous studies have documented that hypoxia and serum deprivation (hypoxia/SD), induced MSCs apoptosis through the mitochondrial apoptotic pathway. Since serum lysophosphatidic acid (LPA) levels are known to be significantly elevated after acute myocardial infarction and that LPA enhanced survival of other cell systems, we embarked on determining whether LPA protects MSCs against hypoxia/SD-induced apoptosis. We have also investigated the potential mechanism(s) that may mediate such actions of LPA. All experiments were carried out on rat bone marrow MSCs. Apoptosis was induced by exposure of cells to hypoxia/SD in a sealed GENbox hypoxic chamber. Effects of LPA were investigated in the absence and presence of inhibitors that target either Giproteins, the mitogen activated protein kinases ERK1/2, or phosphoinositide 3-kinase (PI3K). The data obtained showed that hypoxia/SD-induced apoptosis was significantly attenuated by LPA through Gi-coupled LPA1 receptors linked to the downstream ERK1/2 and PI3K/Akt signaling pathways that function in parallel. Additional studies have demonstrated that hypoxia/SD-induced activation of mitochondrial dysfunction was virtually abolished by LPA treatment and that inhibition of the LPA1 receptor, Gi proteins, the PI3K/Akt pathway, or ERKs effectively reversed this protective action of LPA. Taken together, our findings indicate that LPA is a novel, potent survival factor for MSCs and this may prove to be of considerable therapeutic significance in terms of exploiting MSC-based therapy in the infracted myocardium.

Disclosure of potential conflicts of interest is found at the end of this article.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Bone marrow-derived mesenchymal stem cells (MSCs) are self-renewing and able to differentiate into osteoblasts [1], chondrocytes [1], astrocytes [2], neurons [3], skeletal muscle cells [4] and cardiomyocytes both in vitro [5] and in vivo [6, 7]. The ease of isolation and ex vivo expansion in culture, combined with their multipotency, makes MSCs a promising source of stem cells for cardiac regeneration [8]. However, the clinical exploitation of MSCs is hampered by the fact that transplanted stem cells do not survive efficiently well within the diseased myocardium. For instance, more than 99% of MSCs injected into the left ventricle of CB17 SCID/beige adult mice die within 4 days of injection [9]. In a recent studies, we demonstrated that hypoxia and serum deprivation (Hypoxia/SD), both components of ischemia [10, 11], induce MSCs apoptosis through the mitochondrial pathway [12]. These findings indicate that the ischemic microenvironment of the infracted myocardium may not be conducive to MSCs survival and protection of these cells from apoptosis and together with enhancing their viability and survival in ischemic conditions, may be crucial for their successful utilization in cellular therapy.

Lysophosphatidic acid (LPA) is a naturally occurring bioactive lipid with multiple functions in biological systems that include its ability to induce apoptosis in various cells such as neurons [13], smooth muscle cells [14], and myeloid progenitor TF-1 and D2 cells [15]. Paradoxically, LPA also promotes survival of hypoxia-challenged neonatal cardiomyocytes [16], rescues intestinal epithelial cells from camptothecin-induced apoptosis [17], protects B-cell lines and chronic lymphocytic leukemia (CCL) cells against fludarabine- and etoposide-induced apoptosis [18], and prevents apoptosis in serum-deprived fibroblasts [19], Schwann cells [20], renal tubular cells [21] and macrophages [22]. In skeletal muscle cells, it has been reported that the same concentration of LPA could activate mitogenic and apoptotic signaling pathway simultaneously [23] while in human CD4+8+3low T lymphoblasts, LPA protected against Fas, CD2, or a combination of CD3 and CD28 induced apoptosis, but promoted the latter when applied in the absence of other pro-apoptotic agents [24]. These contradictory findings highlight the complex actions of LPA and reflect its diverse cellular actions, which are still not very well-defined. Moreover, despite these diverse reports, very little is known about the effects of LPA in MSCs or, indeed, the underlying mechanisms that may mediate its actions in these cells.

Most of the potent actions of LPA are mediated through the G-protein-coupled receptors (GPCRs) of which there are five currently identified. Three of these, referred to as LPA1, LPA2, and LPA3, belong to the endothelial differentiation genes (Edg) family formerly named Edg2, Edg4, and Edg7 [25]. The fourth receptor referred to as LPA4 and originally cloned in humans [26], appears to be distinct and with only ~24% amino acid homology with the other three proteins. Moreover, LPA4 is weakly expressed in human tissues, contrasting with the ubiquitous nature of LPA1 and the strong expression of LPA2 and LPA3, in select human organs [25]. The fifth protein, which has only been partially characterized [27], may have a distinct amino acid sequence, distribution pattern and physiological functions to the other receptors.

The LPA receptors are coupled to at least three different heterotrimeric G-proteins: G{alpha}i, G{alpha}q, G{alpha}12/13 [28]. LPA activation of the pertussis toxin (PTX)-sensitive G{alpha}i inhibits adenylyl cyclase and also activates a series of signaling molecules including Ras, the mitogen activated protein kinases (MAPKs) and phosphoinositide 3-kinase (PI3K). Activation of G{alpha}i and G{alpha}q results in the stimulation of phospholipase C activity, leading to phosphoinositide (PI) hydrolysis and the subsequent activation of protein kinase C (PKC). Some of these events initiated via LPA receptor activation may induce cell proliferation and enhance cell survival. However, the precise pathway may be cell type-specific since in Hela and CCL cells, LPA protects against apoptosis through activation of the PI3K/Akt signaling pathway [18, 29], while in fibroblasts the same process appears to be mediated via the ERKs [19]. Furthermore, both PI3K/Akt and ERK1/2 are required for LPA induced cell survival in other systems, including intestinal epithelial cells [17, 30, 31].

Since it is unclear whether LPA promotes survival of MSC, we embarked on investigating the effects of this compound on hypoxia/SD-induced apoptosis in these cells. More importantly, our studies were extended to identify the LPA receptor(s) and the signaling mechanisms that may mediate such actions, focusing particularly on the pathways discussed above. In addition, we have also examined whether the ERKs and/or PI3K/Akt protect against mitochondrial apoptotic events in these cells. Our current data demonstrated for the first time that LPA protects MSCs from hypoxia/SD-induced apoptosis and that LPA exerts its antiapoptotic effects in these cells through Gi-coupled LPA1 receptor via its the downstream ERK and PI3K/Akt pathways. Moreover, hypoxia/SD-induced activation of mitochondrial dysfunction was virtually abolished by LPA with inhibition of LPA1, Gi protein, PI3K/Akt or the ERKs effectively reversing the protective effects of LPA and strongly implicating the latter as a potent survival factor for MSCs.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Materials
Iscove's modified Dulbecco's medium (IMDM) and fetal bovine serum were from Gibco (Grand Island, NY, http://www.invitrogen.com). Trizol reagent was purchased from Invitrogen (Carlsbad, CA, http://www.invitrogen.com). M-MLV reverse transcriptase was from Promega (Madison, WI, http://www.promega.com) and rTaq polymerase from Takara (Otsu, Shiga, Japan, http://www.takara-bio.com). LPA (oleoyl C: 18:1) and DGPP lglycerol pyrophosphate [8:0]) were from Avanti Polar Lipids (Alabaster, AL, http://www.avantilipids.com). Hoechst 33342, Rhodamine 123, and mouse monoclonal anti-rat β-actin antibody were from Sigma-Aldrich (St. Louis, http://www.sigmaaldrich.com). The annexin V-FITC Apoptosis Detection Kit was purchased from Oncogene (San Diego, http://www.oncogene.com) whereas pertussis toxin (PTX), Wortmannin, and U0126 were from Biomol Research Labs (Plymouth Meeting, PA, http://www.biomol.com). PD98059 was from Calbiochem (San Diego, http://www.emdbiosciences.com). LY294002, rabbit polyclonal anti-rat caspase-3, Bax, Phospho-p44/42 MAP kinase (Thr202/Tyr204, p-ERK1/2), p44/42 MAP kinase (Erk1/Erk2), Phospho-Akt (Ser473), Akt, and the horseradish peroxidase-conjugated secondary antibodies to rabbit or mouse were obtained from Cell Signaling Technology (Danvers, MA, http://www.cellsignal.com). Caspase-3/cpp32 Colorimetric Assay Kit, the Cytochrome c Releasing Apoptosis Assay Kit and mouse monoclonal anti-rat cytochrome c antibody from BioVision (Palo Alto, CA, http://www.biovision.com). Nitrocellulose membrane was purchased from Amersham (Piscataway, NJ, http://www.amersham.com) and the chemiluminescence detection kit from Santa Cruz Biotechnology (Santa Cruz, CA, http://www.scbt.com).

Cell Culture and Treatment
The MSCs were isolated from Sprague-Dawley rats (Vital River Laboratory Animal Inc., Beijing, http://www.vitalriver.com.cn) as previously described [12, 32]. All procedures in the present study were approved by the Animal Care Committee of Cardiovascular Institute and Fuwai Hospital. Bone marrow was harvested from the tibia and femur of 80-g rats, plated in IMDM supplemented with 15% inactivated fetal bovine serum and 100 units/ml penicillin/streptomycin and incubated at 37°C in a humidified tissue culture incubator containing 5% CO2 and 95% air. The medium was replaced 4 hours after plating and 24 hours later to discard nonadherent hematopoietic cells. Adherent MSCs were further grown in medium replaced every 2 days. All cells used in the assay were of passages 1 to 3.

Induction of apoptosis in vitro by hypoxia and serum deprivation (hypoxia/SD), designed to mimic the in vivo conditions of the ischemic myocardium, was initiated as previously reported [12]. Cells exposed to hypoxia/SD alone were used as the apoptotic controls and this was induced by incubating MSCs washed in serum-free media in a sealed hypoxic GENbox jar fitted with a catalyst (BioMé'rieux, Marcy l'Etoile, France, http://www.biomerieux.com) to scavenge free oxygen. Cells cultured in complete medium alone were used as the nonischemic controls. In preliminary experiments, a time-course study involving preincubation of cells with LPA for 0, 0.5, 1.0, or 2.0 hours was carried out and a 1 hour preincubation period was determined as the optimum time point for LPA pretreatment prior to exposure to hypoxia/SD. In subsequent studies, LPA (1–25 µM) was initially preincubated with MSCs for this time period in complete medium before washing the cells in serum-free IMDM and reincubating in the latter for 6 hours in the absence or continued presence of different concentrations of LPA under hypoxic conditions. In additional control studies, cells were exposed to LPA only when being subjected to hypoxia/SD with no prior preincubations.

When used, 50 µM DGPP (LPA1/LPA3 antagonist), 100 nM wortmannin or 25 µM LY294002 (PI3K/AKT inhibitors), 20 µM U0126 or 50 µM PD98059 (ERK inhibitors) were preincubated with cells in complete medium for a predetermined period of 80 minutes before exposure to hypoxia/SD whereas LPA (10 µM) was added in the presence of each drug for 1 hour prior to exposure to hypoxia/SD. Cells were subsequently washed in serum-free IMDM and exposed to hypoxia/SD in the continued presence of LPA (10 µM) and a select inhibitor. In parallel experiments carried out to determine whether Gi proteins are involved in the antiapoptotic actions of LPA, MSCs were initially incubated with 200 ng/ml PTX for 16 hours in complete medium. This period of incubation was determined from pilot experiments and is well within the time frame reportedly used in several other studies [27, 33–36]. LPA (10 µM) was added for the last hour of preincubation when required. Cells were eventually washed in serum-free IMDM and exposed to hypoxia/SD for 6 hours in the continued presence of PTX (200 ng/ml) and LPA (10 µM). In studies aimed at exploring the effects of LPA on kinase phosphorylation, cells were initially serum starved for 12 hours prior to exposure to LPA (10 µM) for a designated period in serum-free medium. When used, PTX (200 ng/ml) was added during the serum deprivation phase and incubated with cells for 12 hours. This is the maximum period of incubation that could be tolerated by the cells without any adverse cytotoxic effects.

RT-PCR Analysis of LPA Receptors Subtype Expression
Total RNA was extracted from MSCs using Trizol reagent according to manufacturer's instructions. Following the preparation of RNA, the cDNA was generated from 2 µg of total RNA using M-MLV reverse transcriptase and oligo(dT) 18 primer. For the PCR, the following gene-specific extron primers were used: LPA1: 5'-TCTTCTGGGCCATTTTCAA-3' and 5'-GCCGTTGGGGTTCTCGTT-3'; LPA2: 5'-CCTACCTCTTCCTCATGTTC-3' and 5'-AATGATGACAACCGTCTTGACTA-3'; LPA3: 5'-TGTCAACCGCTGGCTTCT-3' and 5'-CAGTCATCACCGTCTCATTAG -3'; β-Actin: 5'-CCTAGCACCATGAAGATCAA-3' and 5'-TTTCTGCGCAAGTTAGGTTTTGTCAA-3'. Amplification of the cDNA was performed using rTaq polymerase with an initial denaturation at 94°C for 5 minutes followed by 32 cycles of 95°C for 30 seconds, 59°C for 30 seconds, 72°C for 45 seconds and a final extension step at 72°C for 10 minutes. PCR products were run and imaged on 1.2% agarose gels stained with ethidium bromide. Expected fragment sizes are 386 bp for LPA1, 488 base pairs (bp) for LPA2, 437 bp for LPA3 and 221 bp for β-Actin. cDNA templates reverse transcribed from neonatal rat cardiac myocytes were used as positive controls for each set of primers [34].

Assessment of Morphological Changes
MSCs treated with various agents and hypoxia/SD were observed by phase-contrast microscope and photographed (Olympus IX70; Tokyo http://www.olympus-global.com). Nuclear condensation and fragmentation were assayed using chromatin dye Hoechst 33342 as previously described [12]. Briefly, MSCs were washed with phosphate buffered saline (PBS) and stained by 5 µg/ml Hoechst 33342 before visualized using fluorescent microscopy (Olympus IX70. Apoptotic cells were characterized by morphological alteration as condensed apoptotic nuclei and cell shrinkage.

Flow Cytometric Analysis of Cell Apoptosis and Mitochondrial Membrane Potential
Apoptosis was determined by detecting phosphatidylserine (PS) exposure on cell plasma membrane with the fluorescent dye Annexin V-FITC Apoptosis Detection Kit according to the manufacturer's protocols. This assay discriminates intact (Annexin V–/PI–), early apoptotic (Annexin V+/PI–), late apoptotic (Annexin V+/PI+) and necrotic cells (Annexin V–/PI+). In brief, cells were harvested and washed in ice-cold PBS, resuspended in 200 µl of binding buffer and incubated with 10 µl of Annexin V-FITC solution for 30 minutes at 4°C in the dark. This was followed by a further incubation with 5 µl of propidium iodide (PI) for 5 minutes and then immediately analyzed by bivariate flow cytometry using a FACScan-LSR (BD, San Jose, CA, http://www.bd.com) equipped with CellQuest (BD) software. Approximately 1–2 x 104 cells were analyzed in each of the samples.

The loss of mitochondrial membrane potential ({Delta}{psi}m) was assessed using the {Delta}{psi}m-specific stain rhodamine 123 [37]. In brief, after treatment with various agents and exposure to hypoxia/SD, MSCs were harvested and 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 employing a flow cytometer EPICS XL from Beckman Coulter (Fullerton, CA, http://www.beckmancoulter.com): 1–2 x 104 cells were analyzed in each of samples.

Protein Extraction and Western Blot Analysis
For analysis of cellular protein levels, stimulated cells were rinsed twice with ice-cold PBS and then lysed in ice-cold lysis buffer (20 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% TritonX-100, 2.5 mM sodium pyrophosphate, 1 mM β-glycerolphosphate, 1 mM Na3VO4, 1 mM PMSF, and 10 µg/ml each of Leupeptin, Aprotinin, and Pepstatin) for 30 minutes. Cell lysates were centrifuged at 13,000 g for 10 minutes at 4°C and the protein concentration determined by the Bradford assay. Amounts for equal loading of proteins per lane were determined and mixed with 5 x SDS sample buffer, boiled for 5 minutes and separated on 8%–15% SDS-PAGE gels before transferring the proteins onto a nitrocellulose membrane by semi-dry transfer. After blocking in 5% skim milk for 1 hour, membranes were rinsed and incubated overnight at 4°C with the appropriate diluted primary antibody in 5% BSA, 1 x TBS, and 0.1% Tween-20 (TBS/T), with gentle shaking. Excess antibody was removed by washing the membrane in TBS/T and subsequently incubated for 1 hour with HRP-conjugated secondary antibody at room temperature. After further washes in TBS/T, bands were visualized with an enhanced chemiluminescence detection kit and exposed to radiographic film.

For the analysis of cytochrome c release from mitochondria and Bax translocation to mitochondria, we used Cytochrome c Releasing Apoptosis Assay Kit according to the manufacturer's protocols. In brief, 5 x 107 cells were harvested and washed with ice-cold PBS. Cells were incubated with 1.0 ml of 1 x Cytosol Extraction Buffer Mix for 10 minutes, and then homogenized using an ice-cold Dounce tissue grinder. The homogenates were centrifuged at 700 g and then the supernatants further centrifuged at 10,000 g for 30 minutes at 4°C. The cytosolic supernatants were decanted and the pellets resuspended in 0.1 ml mitochondrial extraction buffer mix. Both the mitochondrial and cytosolic fraction were subjected to standard Western blotting and probe with a mouse monoclonal anti-rat cytochrome c antibody and a rabbit polyclonal anti-rat Bax antibody.

Measurement of Caspase-3 Activation
Activity of caspase-3 was detected by using Caspase-3/CPP32 Colorimetric Assay Kit according to the manufacturer's instructions. In brief, cells were lysed in 50 µL of chilled cell lysis buffer and incubate on ice for 10 minutes before centrifuging for 1 minute at 10,000 g. The supernatant was collected and 50 µL of the cell lysate containing 50–200 µg protein was added to 50 µL of 2 x reaction buffer and 5 µL of DEVD-pNA substrate (200 µM final concentration). After incubation at 37°C for 1–2 hours, DEVD-pNA cleavage was monitored by enzyme-catalyzed release of pNA, by determining the absorbance at 405 nm in a microtiter plate reader.

Statistical Analysis
Data are expressed as mean ± SEM. Differences among groups were tested by one-way ANOVA. Comparisons between two groups were evaluated using Student's ttest. A value of p < .05 was considered as significantly different.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
LPA Protects MSCs from Hypoxia/SD-Induced Apoptosis
We have previously demonstrated that hypoxia/SD induced MSCs apoptosis with a maximal induction of early apoptosis at 6 hours [12]. We have now extended these studies by examining whether LPA could protect MSCs from this process. In the first series of experiments, the data obtained show a clear antiapoptotic action of LPA (Fig. 1A, 1B). This was however, only evident when the compound was present throughout the 6 hours exposure to hypoxia/SD. Removal of LPA after an initial 1 hour pretreatment period failed to prevent the process occurring, indicating that LPA is required during the initiation of apoptosis to prevent the later from occurring. Significant inhibition of apoptosis was also evident when LPA was added at the point of subjecting the cells to hypoxia/SD. The inhibitions were, however, lower but not statistically different from those seen following the 1 hour pretreatment (Fig. 1B). All subsequent experiments were therefore carried out by preincubating cells with LPA for 1 hour prior to any further manipulations. Moreover, LPA was maintained in the incubations throughout each experimental procedure.


Figure 1
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  		Figure 1. Effects of lysophosphatidic acid (LPA) pretreatment on hypoxia and serum deprivation (hypoxia/SD)-induced apoptosis in mesenchymal stem cells (MSCs). Apoptosis was determined by flow cytometry in MSCs preincubated with LPA (10 µM) for 1 hour prior to exposure to hypoxia/SD for 6 hours in the continued presence (LPA Pre. 1 h+ 6 h) or absence (LPA Pre 1 h) of LPA. In other experiments, LPA was added to cells at the point of exposure to hypoxia/SD and was maintained in the medium throughout the latter phase (LPA 6 h). MSCs cultured in complete medium under non-hypoxic conditions were used as the non-apoptotic controls and cells exposed to hypoxia/SD alone without any prior or sustained exposure to LPA (No LPA) represents the apoptotic controls. (A) represents FACScan Flow Cytometric Analysis of apoptotic cells after Annexin V-FITC and propidine iodide (PI) staining. Viable cells are Annexin V–/PI–. The Annexin V+/PI– cells are in the early apoptotic phase, whereas the Annexin V+/PI+ cells are in the late apoptotic phase. Necrotic cells are shown as Annexin V–/PI+. (B) is the mean data and is presented as fold changes when compared with the corresponding non-apoptotic control cells. Each column represents the mean ± SEM of three independent experiments; *, p < .001 versus hypoxia/SD (6 h); NS means no difference versus hypoxia/SD (6 h).

 
In further studies, MSCs were exposed to increasing concentrations of LPA (1–25 µM) followed by exposure to hypoxia/SD for 6 hours. Cell death was determined morphologically and by flow cytometry. As shown in Figure 2A and 2B, control cells had the normal elongated MSCs morphology with large regular nuclei. After hypoxia/SD treatment (0 µM LPA), there were clear apoptotic characteristics of shrinkage in cell size and cell loss together with clear chromatin condensation and typical fragmented nuclei. Treatment with varying concentrations of LPA (1–25 µM) efficiently blocked the apoptotic process with cells maintaining their elongated morphology and large nuclei (Fig. 2A, 2B). The degree of apoptosis induced was determined by monitoring positive Annexin V-FITC stained cells at the end of the incubation period. Exposure of MSCs to hypoxia/SD resulted in approximately a 6 to 7 fold increase in Annexin V+/PI– when compared to the control and this is consistent with our previous findings [12]. LPA (1–25 µM) significantly attenuated hypoxia/SD-induced Annexin V+/PI– accumulation, with the maximum reduction just marginally higher than those detected in controls (Fig. 2C, 2D).


Figure 2
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  		Figure 2. LPA protects MSCs against hypoxia/SD-induced apoptosis. MSCs were washed and then exposed to hypoxia/SD for 6 hours. MSCs cultured in complete medium, under non-hypoxic conditions, were used as control. When used, LPA (1–25 µM) was preincubated with the cells for 1 hour in complete medium before exposure to hypoxia/SD and was maintained in the incubation medium throughout the hypoxia/SD treatment period. Induction of apoptosis was determined by morphological changes using phase-contrast microscope (A) and by apoptotic nuclear condensation using fluorescence microscopy following Hoechst 33342 staining (B). C is representative of 3 FACScan Flow Cytometric analyses of apoptotic cells after Annexin V and propidine iodide (PI) staining and (D) is presented as fold changes compared with the corresponding control cells. Each data point represents the mean ± SEM of three independent experiments. *, p < .001 versus control; Figure 2, p < .001 versus Hypoxia/SD alone.

 
Caspase-3 is one of the key mediators of apoptosis and to further evaluate the antiapoptotic effects of LPA, MSCs were examined for caspase-3 activation following exposure to hypoxia/SD in the presence of LPA. The data in Figure 3A show that a 6 hours exposure to hypoxia/SD induced a 3.3-fold increase in caspase-3 activity which was inhibited in a concentration-dependent manner by LPA, with maximal effects obtained at 10 µM and 25 µM (1.5-fold vs. Control). In parallel, Western blot analysis using caspase-3 selective antibody, confirmed that LPA-elicited significant inhibitions of caspase-3 cleavage as indicated by the reduction in hypoxia/SD-induced conversion of the 35-kDa procaspase-3 to the 17 kDa cleaved caspase-3 (Cl.caspase-3) (Fig. 3B). These changes correlate well with the reduction of caspase-3 activation shown in Figure 3A.


Figure 3
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  		Figure 3. LPA inhibits hypoxia/SD-induced caspase-3 activation in MSCs. MSCs were washed and then exposed to hypoxia/SD for 6 hours. When used, LPA (1–25 µM) was preincubated with the cells for 1 hour in complete medium before exposure to hypoxia/SD and was maintained in the incubation medium throughout the hypoxia/SD treatment period. Caspase-3 activity was determined by using the Caspase-3/CPP32 Colorimetric Assay Kit according to the manufacturer's instructions (BioVision). Changes in activation of caspase-3 is expressed in (A) as fold increase relative to the control values and represents the mean ± SEM of at least three independent experiments. *, p < .001 versus control; Figure 3, p < .001 versus hypoxia/SD alone. (B) shows representative Western blots of procaspase-3 cleavage induced by hypoxia/SD and its inhibition by LPA using anticaspase-3 antibody. The blots shown are representative of three independent experiments.

 
These data demonstrate that hypoxia/SD induces apoptosis in MSCs. More importantly, LPA was found to noticeably promote MSCs survival under these conditions, with the most pronounced effects observed at 10–25 µM. Based on these results, all subsequent experiments were performed with 10 µM LPA.

LPA Promotes MSCs Survival through a LPA1/Gi-Coupled Pathway
Prior to exploring the signaling responses activated by LPA, we performed RT-PCR analysis to determine the expression profile of subtypes of LPA receptor. As shown in Figure 4A, PCR primers for LPA1–3 receptor subtypes detected their respective transcripts in total RNA isolated from cardiac myocytes and used as positive controls. In contrast, analysis of total RNA from MSCs showed only the expression of LPA1 indicating that these cells only express the LPA1 receptor subtype.


Figure 4
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  		Figure 4. LPA promotes MSCs survival through LPA1/Gi-coupled pathway. The expression profile of LPA receptor subtypes was determined by RT-PCR analysis as described in the Methods. cDNA templates reverse transcribed from neonatal rat cardiac myocytes (CMs) were used as positive controls for each set of LPA receptor primers. The determined bp for the LPA receptor subtypes were: LPA1 386bp, LPA2 488bp, LPA3 437bp. In the data shown in (B), cells were preincubated in complete medium for either 80 minutes with the LPA1/3 antagonist DGPP (50 µM) or for 16 hours with the Gi protein inhibitor PTX (200 ng/ml) before exposure to hypoxia/SD. When present, LPA (10 µM) was added in the presence of each drug for 1 hour prior to exposure to hypoxia/SD. All drugs were maintained in the incubation medium throughout the hypoxia/SD treatment period. Apoptosis was quantified by flow cytometric analysis after staining with Annexin V and PI and the data are presented as fold changes compared with the corresponding control cells. Each data point represents the mean ± SEM of three independent experiments. *, p < .001 versus Hypoxia/SD + LPA; Figure 4, p < .001 versus Hypoxia/SD alone.

 
To determine the relevance of the LPA1 receptor in LPA-mediated inhibition of apoptosis, MSCs were treated with DGPP, a specific antagonist of LPA1 and LPA3 [38], prior to exposing cells to LPA and hypoxia/SD. As shown in Figure 4B, 50 µM DGPP was able to partially reverse the antiapoptotic effects of LPA, resulting in a significant increase in apoptotic cells. Parallel studies carried out to determine whether Gi proteins are involved in the antiapoptotic effects of LPA, revealed that preincubation of MSCs, with the potent Gi protein inhibitor, PTX at a concentration of 200 ng/ml, also reverse the protection offered by LPA against hypoxia/SD-induced apoptosis (Fig. 4B). Interestingly, when added in the absence of LPA, both DGPP and PTX attenuated hypoxia/SD induced apoptosis. This was, however, less pronounced when compared to the responses seen with LPA alone (Fig. 4B).

LPA Prevents MSCs from Hypoxia/SD-Induced Apoptosis Through PTX-Sensitive ERK1/2 and PI3K/Akt Pathways
As highlighted earlier, the MAPK and PI3K/Akt signaling pathways, are reported to be important in promoting survival in other cell systems. We therefore investigated whether these pathways mediate the antiapoptotic effects of LPA in MSCs. Western blot analysis showed low detectable levels of phosphor-ERK 1/2 in controls. More importantly, LPA time-dependently induced a pronounced increase in ERK1/2 phosphorylation, and this appeared to have reached a peak at 5 minutes and sustained for up to 10 minutes. Phosphorylation was, however, virtually back to basal levels 30 minutes after the initial stimulation (Fig. 5A). To implicate ERK1/2 in LPA-induced protection against hypoxia/SD, MSCs were treated with PD98059 and U0126, both potent inhibitors of this pathway. Analysis by flow cytometry revealed that PD98059 (50 µM) and U0126 (20 µM) completely blocked the antiapoptotic effect of LPA and neither compound altered the apoptotic actions of hypoxia/SD when used independently of LPA (Fig. 5B).


Figure 5
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  		Figure 5. LPA prevents MSCs from hypoxia/SD-induced apoptosis through PTX-sensitive ERK1/2 and PI3K/Akt pathways. Serum starved (12 h) MSCs were either incubated in serum-free medium alone or stimulated with 10 µM LPA in serum-free medium for the indicated times and the expressions of total p44/42 MAP kinase protein (ERK1/2), phospho-p44/42 MAP kinase (Thr202/Tyr204 p-ERK1/2) (A), Akt or phospho-Akt (Ser473) (C) determined by Western blotting as described in the Methods. To determine the role of the respective kinase pathways in the antiapoptotic actions of LPA, cells were pretreated with either the MEK/ERK1/2 inhibitors, PD98059 (PD, 50 µM) and U0126 (20 µM) (B) or the PI3k inhibitors, wortmannin (Wort., 100 nM) and LY294002 (LY, 25 µM) (D) for 80 minutes in complete medium before exposure to hypoxia/SD while LPA (10 µM) was added in the presence of each drug for 1 hour prior to exposure to hypoxia/SD. All drugs were maintained in the incubation medium throughout the hypoxia/SD treatment period. Apoptosis was quantified by flow cytometric analysis after staining with Annexin V and PI and the data are presented as fold changes compared with the corresponding control cells. Each data point represents the mean±SEM of three independent experiments. *, p < .001 versus hypoxia/SD+ LPA; #, p < .001 versus hypoxia/SD alone. (E) shows representative Western blots of phosphorylated ERK1/2, total ERK1/2, phosphorylated Akt, and total Akt in serum starved (12 h) cells that were treated with either LPA (10 µM) alone for 5 minutes or pretreated with PTX (200 ng/ml) for 12 hours, PD98059 (PD; 50 µM), U0126 (20 µM), LY294002 (LY; 25 µM), or Wortmannin (Wort; 100 nM) for 1 hour during serum starvation before exposure to LPA for 5 minutes in the serum-free medium. The blots are each representative of three separate experiments.

 
In parallel with the phosphorylation of ERK1/2, LPA was also found to cause a significant increase in Akt phosphorylation within 5 minutes of exposure to the compound and, in this case, the phosphorylation was sustained for up to 180 minutes (Fig. 5C). Administration of the potent PI3K inhibitors, LY294002 (25 µM) and wortmannin (100 nM), markedly attenuated the antiapoptotic actions of LPA. When added independently of LPA, both compounds unexpectedly caused a partial, but significant inhibition of apoptosis induced by hypoxia/SD (Fig. 5D).

Since the above data strongly suggest a role for both ERKs and PI3K/Akt, we went on to investigate whether there was cross-talk between the two pathways by investigating whether there was any cross inhibition of ERK1/2 phosphorylation by LY294002 or wortmannin and of Akt phosphorylation by PD98059 or U0126. LPA-induced stimulation of ERK1/2 phosphorylation was, however, unaffected by 25 µM LY294002 or 100 nM wortmannin (Fig. 5Ei) but inhibited significantly by 200 ng/ml PTX and 50 µM PD98059. Similarly, LPA-induced phosphorylation of Akt was also prevented by PTX (200 ng/ml) and LY294002 (25 µM), but not by PD98059 (50 µM) or U0126 (20 µM) (Fig. 5Eii).

Collectively, these observations indicate that LPA-mediated protection against apoptosis in MSCs is dependent on both ERK1/2 and PI3K/Akt signaling pathways, which act in parallel and downstream of the PTX-sensitive Gi protein.

LPA Elicits Antiapoptotic Effect via Inhibition of the Mitochondria Pathway
The translocation of proapoptotic Bax plays an important role in the loss of {Delta}{psi}m and in the regulation of mitochondrial cytochrome c release leading to activation of caspase-3. The effect of LPA (10 µM) on hypoxia/SD-induced Bax translocation was therefore examined. Western blot analysis revealed that the level of Bax in the mitochondrial fraction notable increased after treatment with hypoxia/SD whereas LPA significantly inhibited Bax translocation into mitochondria. When used, the LPA1/3 antagonist DGPP (50 µM), the Gi protein inhibitor PTX (200 ng/ml), the ERK1/2 inhibitor PD98059 (50 µM), and the PI3K/Akt inhibitor LY294002 (25 µM), all potently prevented the inhibition caused by LPA (Fig. 6A) and elevated Bax levels in the mitochondrial fraction.


Figure 6
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  		Figure 6. LPA exerts antiapoptotic effects via inhibition of mitochondrial dysfunction. MSCs were washed and then treated with hypoxia/SD for 6 hours. In parallel experiments, cells were pretreated with either PTX (200 ng/ml) for 16 hours or DGPP (50 µM), PD98059 (PD, 50 µM), or LY294002 (LY, 25 µM) for 80 minutes before exposure to hypoxia/SD for 6 hours. When present, LPA (10 µM) was added in the presence of each drug for 1 hour prior to exposure to hypoxia/SD. All drugs were maintained in the incubation medium throughout the hypoxia/SD treatment period. (A) is a representative Western blot of BAX expression in mitochondrial and cytosolic fractions of MSCs in the absence and presence of the different drugs listed above. (B) shows changes in {Delta}{psi}m detected by flow cytometry of Rhodamine 123-stained cells and (C) is a representative Western blot of hypoxia/SD-induced translocation of cytochrome c (Cyt C) from the mitochondria into the cytosol in the absence and presence of LPA ± the different drugs. Changes in caspase-3 activity, determined by colorimetric assay, and are shown in (D) as fold increase relative to the control values. The data in the graph are the mean ± SEM of at least three independent experiments. *, p < .001 versus control; Figure 6, p < .001 versus Hypoxia/SD alone; {dagger}, p < .001 versus Hypoxia/SD + LPA. (E) shows a representative Western blot of hypoxia/SD-induced procaspase-3 cleavage to Cl-caspase-3 in the absence and presence of LPA ± the different drugs. All blots shown are representative of three individual experiments.

 
In parallel with the above studies, we ascertained whether LPA influenced the loss of {Delta}{psi}m in MSCs subjected to hypoxia/SD. As shown in Figure 6B, hypoxia/SD induced changes in {Delta}{psi}m was clearly reversed by LPA, where as cotreatment with DGPP, PTX, PD98059 or LY294002 significantly increased the loss of {Delta}{psi}m.

Mitochondrial dysfunction provokes release of cytochrome c from the mitochondria into the cytosol. Western blot analysis revealed that exposure of MSCs to hypoxia/SD induced a significant increase in cytosolic cytochrome c levels. This was accompanied by a parallel decrease of cytochrome c in the mitochondrial fraction with LPA pretreatment inhibiting these changes. When applied, DGPP, PTX, PD98059, and LY294002 individually attenuated the effects LPA and increased cytochrome c accumulation in the cytosol whereas decreasing levels in the mitochondrial fraction (Fig. 6C).

Examination of changes in caspase-3 activity revealed that exposure to hypoxia/SD significantly increased enzymatic activity by 3.3 ± 0.1-fold when compared with control. LPA markedly inhibited hypoxia/SD-induced activation of caspase-3–1.5 ± 0.1-fold. Cotreatment with DGPP, PTX, PD98059, or LY294002 significantly reversed LPA inhibitions of caspase-3 activity with the latter, respectively, increasing to 3.0, 3.1, 2.4, or 2.2-fold of the control basal values (Fig. 6D). In addition, Western blot analysis of the cleavage of procaspase-3 to Cl.caspase-3 confirmed that LPA was able to inhibit hypoxia/SD-induced activation of caspase-3 and that cotreatment with the various inhibitors abrogated the effects of LPA (Fig. 6E).


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosure of Potential...
 Acknowledgments
 References
 
Mesenchymal stem cells have shown great therapeutic potential because of their ability to regenerate and repopulate the injured myocardium, restoring its function after transplantation into ischemic or infarcted heart [7, 39, 40]. In addition to being easily obtained and expanded, MSCs are an ideal source of cells for the repair of a damaged myocardium. However, although MSCs transplantation therapy has attracted considerable interest in recent years, their therapeutic exploitation has been limited by the fact that most of the transplanted MSCs do not fully establish and are readily lost, presumably due to cell death induced by the ischemic environment into which they are introduced in vivo [9, 41, 42].

In a recent study we demonstrated that serum deprivation and hypoxia, both components of ischemia [10, 11], induced programmed MSCs death through the mitochondrial apoptotic pathway [12]. As a result we are now interested in identifying molecules that could prevent this process and promote survival of MSCs in the ischemic myocardium. In the current studies we have focused on LPA, a simple and natural phospholipid which accumulates in patients with acute myocardial infarction, increasing to levels of ~10 mg/l within 48–72 hours of the onset of the disease [43]. Although this increase may be considered pathological, in view of the potential pro-apoptotic actions of LPA [44–46], it is in fact possible that its accumulation in the infracted myocardium may offer cardioprotection through its ability to influence cell proliferation or survival, and potentially enhance MSC survival via the activation of distinct signaling pathways. There is indeed precedence that enhancement of LPA levels in other disease states, such as ovarian [47] and prostate cancer [48] and multiple myeloma [49], may promote cell proliferation and survival. Should this also be true in MSCs, then LPA could influence MSC survival within the injured myocardium and thereby potentially facilitate cardiac regeneration. Experiments were therefore carried out to examine the antiapoptotic actions of LPA on MSCs and further explore the potential cellular mechanisms that may mediate its actions. Studies relating to the latter initially focused on identifying the LPA receptor(s) expressed in MSCs using RT-PCR and on the requirement of Gi proteins which may be coupled to these receptors. Furthermore, since signaling for LPA in other cell systems involves the MAPK and PI3Ks [17–19, 29–31], we extended our studies by examining whether activation of these two pathways are critical for the actions of LPA in MSCs.

In the present investigation, we demonstrate for the first time that LPA protects MSCs against hypoxia/SD-induced apoptosis and that this effect required the continued presence of LPA during the hypoxia/SD phase in order for it to inhibit the apoptotic process. More importantly, the antiapoptotic actions of LPA were observed at concentrations that are well within the range found in serum (2–20 µM) [50], thus making our findings physiologically and therapeutically relevant.

The above actions of LPA appear to be mediated via the activation of the Gi protein-linked LPA1 which belongs to Edg family of proteins. This was confirmed by RT-PCR using LPA receptor subtype specific primer and the potent LPA1/3 antagonist, DGPP. In the first instance, the results obtained from the RT-PCR analysis show again, for the first time, that MSCs express LPA1 but not LPA2 or LPA3. The latter two proteins may not be expressed in these cells as their transcripts were not detected even though their respective primers identified expression in total RNA isolated from rat cardiomyocytes that were used as a positive control. The data obtained with DGPP, fairly conclusively implicates the LPA1 receptor since this antagonist reversed the antiapoptotic effects of LPA and at a concentration that is selective for the LPA1/3 proteins. As LPA3 mRNA was not detectable, we can therefore conclude that the target receptor for LPA in our studies is LPA1.

The implication of the LPA1 receptor in the antiapoptotic actions of LPA in MSC is of some interest since in a previous study, we indicated that activation of this receptor subtype may mediate LPA-induced apoptosis in neonatal rat cardiac fibroblasts (CFs) [37]. However, this was based on data obtained using DGPP and since CFs also expressed the LPA3 receptor, we cannot be certain that the latter was in fact not responsible for our previous observations. Although this still remains to be validated using a specific LPA1 or LPA3 antagonist, it is likely that LPA signaling in MSCs mediated through LPA1 may protect against apoptosis. Indeed, this may also be true in Schwann cells where LPA acting through LPA1 has been shown to protect these cells from programmed cell death [20, 51–53].

In our studies, activation of LPA1 may signal through the Gi protein because pretreatments of MSCs with PTX markedly attenuated responses to LPA. The down stream transduction cascades linked to the Gi protein-coupled LPA1 receptor, are likely to involve the ERK1/2 and PI3K/Akt pathways which have both been reported to contribute to LPA-induced survival of various cells [17, 30, 31]. Both these pathways play a critical role in mediating the antiapoptotic actions of LPA in MSCs. In the first instance, our data revealed that LPA induced rapid and sustained phosphorylation of both ERK1/2 and Akt in MSCs. Moreover, application of inhibitors of either ERK1/2 or PI3K significantly inhibited the protection offered by LPA against hypoxia/SD-induced apoptosis. Thus, taken together, these findings strongly indicate that both kinase pathways are critically required and inhibition of either cascade would result in the reversal of the antiapoptotic actions of LPA. This is in contrast to Hela and CCL [18, 29] or fibroblasts [19] where PI3K/Akt and ERK1/2 mediate LPA-induced protections respectively. What is even more interesting from our studies is the fact that there does not appear to be cross-talk between the ERK1/2 and PI3K pathways, but instead act in parallel as demonstrated in experiments using potent inhibitors of each kinase against the phosphorylation of the other. The data obtained showed clearly that the ERK1/2 inhibitors, PD98059 and U0126, did not alter LPA-induced Akt phosphorylation. Similarly, neither wortmannin nor LY294002 (both potent inhibitors of PI3K) altered ERK1/2 phosphorylation. These observations strongly demonstrate that the two pathways act in parallel. It is, however, possible that the signal from ERK1/2 and PI3K converge on a common downstream target which is yet to be identified in MSCs.

Given the cell survival activity demonstrated for LPA and PI3K, it is intriguing that DGPP, PTX, LY294002, and wortmannin would independently inhibit hypoxia/SD-induced apoptosis. The precise reason for this unexpected finding is unclear and warrants further investigation. However, these effects may reflect an action of the drugs on other cellular events that are essential for cell survival, but independent of the LPA1/Gi pathway and/or PI3K activation. Consistent in part with this notion, is a recent report demonstrated that LY294002 may inhibit apoptosis in rat kidney epithelial cells by increasing bcl-2 protein levels in the mitochondria through a PI3K-independent mechanism [54]. PTX on the other hand has been shown to promote survival of bone marrow-derived macrophages through activation of PI3K [55]. In the context of our studies, it is feasible that PTX is able to reverse the antiapoptotic actions of LPA through blockade of Gi protein but, on its own, promote MSC survival through its ability to enhance PI3K activation. Similarly, DGPP is reported to enhance secretion of arachidonate metabolites in PD388D1/MAB macrophage cell line through activation of the p42/44 MAPKs [56], a pathway which we have demonstrated to be essential for the ant-apoptotic actions of LPA in MSCs. Whether any of these mechanisms account for the independent actions of the inhibitors in MSCs now remains to be determined.

Our previous study has shown that hypoxia/SD induced apoptosis in MSCs through the mitochondrial apoptotic pathway by inducing Bax protein translocation to the mitochondria, loss of {Delta}{psi}m, release of cytochrome c and activation of caspase cascades [12]. We now report that these processes may be prevented by LPA, which inhibits hypoxia/SD induced mitochondria-dependent apoptosis by preventing Bax translocation to mitochondria, loss of {Delta}{psi}m, release of cytochrome c into the cytosol, and activation of caspase-3. These effects are mediated via the Gi coupled LPA1 receptor linked to both ERK1/2 and PI3K. This conclusion is based on the fact that DGPP, PTX, PD98059, and LY294002, all effectively blocked the effects of LPA on the mitochondrial apoptotic pathway.

In summary, we have presented data which, to our knowledge, demonstrates for the first time that LPA can promote MSCs survival under conditions that mimic those in the ischemic myocardium. This potentially beneficial effect of LPA appears to converge on the mitochondria, acting presumably to protect mitochondrial integrity and function through the Gi coupled LPA1 receptor that is linked in parallel to both the ERK1/2 and PI3K signaling pathways. Since these effects occur at concentrations of LPA that are reported to exist in serum [50], it is feasible that LPA acting through the LPA1 receptor can exert its pro-survival actions on MSCs in vivo. Our findings are therefore of considerable therapeutic significance and provide the potential of now exploiting LPA and MSCs clinically in cardiac regeneration therapies.


   DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
Acknowledgments
 References
 
The authors indicate no potential conflicts of interest.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
References
 
This work was supported by the grants from National Natural Science Foundation of China (30370524 and 30271290) and Doctoral Program of Higher Education of China (20050023016) and Beijing Municipal Science and Technology Commission (D0906004040391) and the Major National Basic Research Program in the People's Republic of China (Program 973, 2007CB512108).


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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 

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