- Email: [email protected]
ER stress contributes to ischemia-induced cardiomyocyte apoptosis
BBRC Biochemical and Biophysical Research Communications 349 (2006) 1406–1411 www.elsevier.com/locate/ybbrc
ER stress contributes to ischemia-induced cardiomyocyte apoptosis Eva Szegezdi a, Angela Duffy c, Martin E. O’Mahoney a, Susan E. Logue a, Louise A. Mylotte c, Timothy O’Brien b,c, Afshin Samali a,c,* a
c
Department of Biochemistry, National University of Ireland, Galway, Ireland b Department of Medicine, National University of Ireland, Galway, Ireland Regenerative Medicine Institute (REMEDI), National Centre for Biomedical Engineering Science, National University of Ireland, Galway, Ireland Received 1 September 2006 Available online 12 Septemeber 2006
Abstract Myocardial ischemia is a severe stress condition that leads to loss of cardiomyocytes. The cell loss is attributed to apoptosis, although the exact mechanisms involved are only partially defined, which limits therapeutic opportunities. Here, we show caspase activation and apoptosis in neonatal rat cardiomyocyte cultures subjected to simulated ischemia by serum, glucose, and oxygen deprivation (SGO). Caspase activation was preceded by endoplasmic reticulum (ER) stress and the activation of the unfolded protein response (UPR), detected by the induction of Grp78, induction and splicing of XBP1, and phosphorylation of eukaryotic initiation factor 2-a (eIF2a). At a later time the ER stress response switched from UPR and cytoprotective response to a pro-apoptotic response as demonstrated by the upregulation of CHOP and processing of pro-caspase-12. Thus, we provide evidence that the ER can generate and propagate apoptotic signals in response to ischemic stress and this pathway is therefore a novel target for prevention of ischemia-mediated cardiomyocyte loss. 2006 Elsevier Inc. All rights reserved. Keywords: Cardiomyocytes; Ischemia; Unfolded protein response; ER stress; Apoptosis; PERK; Ire1; ATF6; CHOP; Caspase-12
Myocardial ischemia is a severe trauma for cardiomyocytes. The simultaneous drop in cellular oxygen and glucose supply leads to ATP depletion, which triggers profound molecular alterations. Initially, the Na+/K+ pump, which is the major Na+ extrusion pathway in cardiomyocytes, comes to a halt leading to the accumulation of Na+ in the cytosol. High intracellular Na+ concentration in turn blocks or reverses the function of the plasma membrane Na+/Ca2+ exchanger, causing Ca2+ influx. Simultaneously, the intracellular K+ and Mg2+ concentration drops [1,2]. In addition to the ionic changes, inability to resynthesize ATP by oxidative phosphorylation also leads to acidification due to accumulation of inorganic phosphate and the glycolytic end-product lactate [3]. As a result, ischemia causes extensive biochemical changes in the cytosol, but also affects the cytoskeleton *
Corresponding author. Fax: +353 91 494596. E-mail address: [email protected] (A. Samali).
0006-291X/$ - see front matter 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2006.09.009
and other cellular organelles. For example, the intermediate filaments of the cytoskeleton collapse into large perinuclear aggregates, the actin fibres relocalize around the nucleus, and microtubules break up. Mitochondria swell and lose structure, leading to cessation of the oxidative phosphorylation [4]. To cope with the stress, both cytosolic and mitochondrial heat shock proteins, including Hsp70, 60, and 27, are induced [5]. In the brain, ischemia has been shown to also target the endoplasmic reticulum (ER) and ischemic neurons activate the ER stress response (unfolded protein response, UPR) to protect themselves [6]. Generally, the UPR is triggered by accumulation of protein aggregates in the ER lumen. The ER lumen has a high redox potential to maintain an oxidizing environment, which, together with the high protein and Ca2+ concentration, provides the ideal milieu for protein folding. Physiological or pathophysiological conditions that alter this milieu hinder protein folding and cause aggregation of nascent polypeptide chains [7]. Three ER
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transmembrane receptors, namely PERK, ATF6, and Ire1, monitor the homeostasis of the ER and trigger the UPR. The UPR acts on several levels; it rapidly attenuates general protein synthesis, induces the expression of ER chaperone proteins, and enhances the degradation of malfolded proteins [7]. Although the UPR is primarily an adaptive response, if the stress persists, the ER stress receptors can also trigger pro-apoptotic pathways to initiate cell death [8]. The sarco-endoplasmic reticulum of cardiomyocytes forms an extensive network that interweaves the cell. In addition to its general cellular functions, the sarcoplasmic reticulum is also vital for contractibility of cardiomyocytes. Nevertheless, to date little is known as to how ischemia affects the homeostasis of the sarcoplasmic reticulum, and only indirect evidence is available to suggest that ischemia in the heart induces ER stress. It has been shown that reduction of ER Ca2+ activates ATF6 and induces Grp78 in cardiomyocyte cultures [9]. Also, ER preconditioning with low dose tunicamycin protects H9c2 neonatal cardiomyocyte cells against prolonged ATP depletion or oxidative stress [10]. A DNA microarray analysis of the heart from MPC-1 (monocyte chemotactic protein-1) overexpressing mice, an experimental model of ischemia, identified induction of ER chaperones [11]. The involvement of apoptosis in cardiomyocyte death during ischemia is a relatively new concept and to date most studies have focused on the role of the death receptor and mitochondrial death pathways [12,13]. Here, we examined whether ischemia induces the UPR and more importantly, whether induction of the UPR during ischemia activates the ER stress death pathway. We show for the first time that ischemia induces all three arms of the UPR: PERK, ATF6, Ire1, and their downstream targets. In addition to the UPR, induction of CHOP and cleavage of pro-caspase-12 were also detected. These data demonstrate activation of UPR during ischemia in cardiomyocytes and that ER stress is involved in the onset of ischemic cell death. Materials and methods Cell culture and treatments. Primary cultures of neonatal cardiomyocytes were isolated from 1 to 4-day-old Sprague–Dawley rats, as described previously [14]. Briefly, rats were euthanized and hearts excised. After scalpel homogenization, overnight trypsin digestion at 4 C, and a collagenase treatment for 20 min at 37 C, cardiomyocytes were enriched by Percoll gradient centrifugation (Amersham) and plated at a density of 1 · 105/ml in DMEM/F12 medium supplemented with 10% newborn calf serum, 100 U/ ml penicillin, 100 lg/ml streptomycin, 1 mM sodium pyruvate (GibcoBRL), 5% insulin transferrin selenite (ITS) liquid supplement media, and 100 lM 5-bromo-2-deoxyuridine on culture plates coated with 0.2% gelatin. Cells were cultured at 37 C and 5% CO2. To mimic endogenous ischemia, cultures were exposed to hypoxic conditions (O2/N2/CO2, 0.5:94.5:5), using a hypoxia gas chamber (Russkin) in the absence of glucose and serum, using glucose-free DMEM (Gibco-BRL) supplemented with 10 mM 2-deoxyglucose, 100 U/ml penicillin, 100 lg/ml streptomycin, 1 mM sodium pyruvate, and 5% ITS liquid supplement media. Thapsigargin (2 lM) in normal culture medium was used to induce ER stress. All animal experiments were performed in accordance with the ethical regulations of NUI, Galway. All reagents were from Sigma, unless otherwise stated.
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Western blot analysis. Cells were lysed in whole cell lysis buffer (20 mM Hepes, pH 7.5, 350 mM NaCl, 1 mM MgCl2, 0.5 mM EDTA, 0.5 mM EGTA, 1% Igepal-630, 0.5 mM dithiothreitol (DTT), 100 lM PMSF, and 1 lg/ml pepstatin), except for detection of eIF2a phosphorylation, where cells were directly lysed in Laemmli buffer. Cellular proteins were separated by electrophoresis on 8–12% SDS–polyacrylamide gels and transferred onto nitrocellulose membranes. After blocking (5% non-fat milk, 0.05% Tween 20 in PBS), blots were incubated with antibodies to caspase12 (Sigma, rat monoclonal), PKCd (Santa Cruz, rabbit polyclonal), caspase-3 (Cell Signalling Technology, rabbit polyclonal), sarcomeric a-actinin (Sigma, mouse monoclonal), Grp78 (StressGen, rabbit polyclonal), XBP-1 (Santa Cruz, rabbit polyclonal), phospho-eIF2a, and total eIF2a (both from Cell Signalling Technology, rabbit monoclonal) and CHOP (Santa Cruz, rabbit polyclonal). To verify equal protein loading, blots were probed for actin (Sigma, rabbit polyclonal). All primary antibodies were diluted 1:1000 and the appropriate horseradish peroxidise-conjugated goat secondary antibodies (Pierce) were used at a 1:5000 dilution. Protein bands were detected with SuperSignal Ultra Chemiluminescent Substrate (Pierce) on X-ray film (Agfa). Analysis of caspase activity. Cell lysates (25 ll) and 50 lM carbobenzoxy-Asp-Glu-Val-Asp-7-amino-4-methyl-coumarin (DEVD-AMC) in reaction buffer (100 mM Hepes, 10% sucrose, 5 mM DTT, 0.0001% Igepal-630, and 0.1% 3-[(3-cholamidopropyl) dimethylammonio] propane-1sulphonic acid (CHAPS), pH 7.25) were added in duplicate to a microtiter plate. Liberated AMC was measured using 355 nm excitation and 460 nm emission wavelengths. Fluorescence was measured over time and the data analysed by linear regression to determine enzyme activity. Enzyme activity was expressed as nmoles AMC liberated by 1 mg total cellular protein per minute. Hoechst staining. Following treatment, cells grown on coverslips were fixed in 3.7% formaldehyde for 10 min at room temperature and permeabilized with methanol for an additional 10 min. After a PBS wash, cells were stained with 100 lg/ml Hoechst 33342. The excess stain was removed by a series of PBS washes. The stained nuclei were visualized using an Olympus BX51 fluorescent microscope. Haematoxylin–eosin staining. Cells were seeded onto 35 mm dishes at a density of 5 · 104 cells/ml. After treatment the cells were fixed in 3.7% formaldehyde for 5 min at room temperature. The cytosol was stained with eosin Y for 4 min, rinsed in tap water, and stained in Harris haematoxylin solution for 5 min. Pictures were taken using an Olympus BX51 microscope at a final magnification of 100·. Immunocytochemical staining for a-actinin. Cells were fixed in 3.7% formaldehyde for 5 min at room temperature and permeabilized with 0.2% Triton X-100 for an additional 5 min. After rinsing with PBS, coverslips were blocked in 1% BSA containing 5% goat serum for 1 h at room temperature. Cells were incubated with 1:1000 dilution of sarcomeric antia-actinin antibody in 1% BSA for 1 h at room temperature. The excess antibody solution was removed by 3 series of PBS washes. Cells were then incubated in 1% BSA containing 1:100 dilution of FITC-conjugated antimouse IgG (Sigma) and 100 lg/ml Hoechst 33342 stain in darkness for 45 min at room temperature. The excess antibody solution was removed and cells were mounted using Vectashield mounting medium. The stained cells were visualized using an Olympus BX51 fluorescent microscope. RT-PCR. Total RNA from cells exposed to thapsigargin or ischemic treatment was isolated using the GenElute mammalian total RNA extraction kit (Sigma) according to the manufacturer’s instructions. Reverse transcription was carried out with 2 lg total RNA and oligo(dT) (Invitrogen) using 20 U/25 ll reaction AMV reverse transcriptase (Sigma) according to the manufacturer’s instructions. ATF4-, XBP1-, spliced XBP1 (sXBP1), and GADD34-specific sequences were amplified during 32 cycles of 30 s denaturing at 94 C, 60 s annealing at 56 C, and 60 s extension at 72 C, with the following primers: ATF4 forward 5 0 -C CGAGATGAGCTTCCTGA-3 0 ; ATF4 reverse 5 0 -CTCCTTGCCGGT GTCTGA; XBP1 forward 5 0 -CAGACTACGTGCGCCTCTGC-3 0 ; XBP1 reverse 5 0 -CTTCTGGGTAGACCTCTGGG; sXBP1 forward 5 0 -TCT GCTGAGTCCGCAGCAGG-3 0 ; sXBP1 reverse 5 0 -CTCTAAGACTAG AGGCTTGG; GADD34 forward: TTTCTAGGCCAGACACATGG; G ADD34 reverse: TGTTCCTTTTTCCTCCGTGG. GAPDH was used as
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a loading control, its cDNA was amplified during 26 cycles of 30 s denaturing at 94 C, 60 s annealing at 56 C, and 60 s extension at 72 C, with the following primers: GAPDH forward 5 0 -ACCACAGTCCA TGCCATC-3 0 ; GAPDH reverse 5 0 -TCCACCACCCTGTTGCTG.
Results Ischemia initiates apoptosis in cardiomyocytes All experiments were carried out on primary neonatal rat cardiomyocyte preparations. The purity of the preparations was between 85% and 97% cardiomyocytes, based on sarcoplasmic a-actinin immunostaining (data not shown). To mimic in vivo ischemic conditions, cells were subjected to a combination of serum, glucose, and oxygen deprivation (SGO) as described in Materials and methods. After 24 h ischemia shrunken cells with condensed or fragmented nuclei, characteristic of apoptotic cells, were detectable in cardiomyocyte cultures (Fig. 1A). The apoptotic mode of cell death was confirmed by detecting activation and activity of caspase proteases. Cardiomyocyte cultures subjected to 4 h ischemia expressed a 3-fold increase in DEVDase activity, which was associated with processing of pro-caspase-3 and cleavage of the caspase-3 substrate, PCKd (Fig. 1B). Primary cardiomyocyte cultures may contain a small percentage of fibroblasts or other non-cardiomyocyte cells. Immunostaining for a-actinin, a cardiomyocyte-specific caspase-3 substrate [15], revealed its breakdown and loss of staining following 4 h ischemia (Fig. 1C). Western blot analysis identified 45 and 25 kDa processed fragments, confirming cleavage of a-actinin in ischemic cardiomyocytes (Fig. 1C). Collectively, these data confirmed that exposure of primary neonatal cardiomyocytes to ischemia induces apoptotic cell death. Ischemia triggers the unfolded protein response in primary cardiomyocytes In order to examine the effect of ischemia on the ER, induction of Grp78 (Bip), a marker of ER stress was determined. Ischemia (2–4 h) induced Grp78 both at mRNA and protein levels (Fig. 2A and B). To compare the kinetics and the extent of Grp78 induction, cardiomyocyes were treated with a specific ER stressor, thapsigargin (an inhibitor of the SERCA pump). Thapsigargin caused a similar increase in Grp78 expression as ischemia, although the kinetics of mRNA induction was faster, between 0 and 2 h treatment (Fig. 2A and B). Since induction of Grp78 is indicative of the activation of the UPR, we examined which of the three branches of the UPR are activated by ischemia in cardiomyocytes. The activation of the UPR receptors Ire1, ATF6, and PERK was detected by monitoring their target molecules. In primary cardiomyocytes, induction of a sXBP1 mRNA (frame-shift splice variant of XBP1) occurred during the first 2 h of exposure to ischemia, and the level and the kinetics of induction were similar to those of thapsigar-
Fig. 1. Ischemia induces apoptotic cell death in primary rat cardiomyocytes. To mimic ischemia, cardiomyocytes were exposed to serum, oxygen, and glucose deprivation for the times indicated. (A) Morphological changes were determined in cardiomyocytes subjected to 24 h ischemia by both Haematoxylin–Eosin staining (upper panel) and Hoechst staining (lower panel). (B) DEVDase activity in ischemic cardiomyocytes was measured in whole cell lysates. The graph shows the average of three separate experiments ± SD. (C) Caspase-mediated degradation of sarcomeric a-actinin in ischemic cardiomyocytes detected by immunostaining (upper panel) and by Western blotting of whole cell lysates (lower panel).
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Fig. 2. Ischemia induces expression of Grp78 at both mRNA and protein levels. Cardiomyocyte cultures were subjected to ischemia or thapsigargin (Tg, 2 lM) for the indicated times. (A) Total RNA was isolated from ischemic (upper panel) and thapsigargin-treated (lower panel) cardiomyocytes and the Grp78 message was amplified by RT-PCR. GAPDH was used as a loading control. (B) Induction of Grp78 protein expression in ischemic (upper panel) and thapsigargin-treated (lower panel) cardiomyocytes detected by Western blotting. Actin was used as a loading control. The images are representatives of three separate experiments.
gin-treated cells, indicating that ATF6 and Ire1 are both activated during ischemia (Fig. 3A and B). Activation of ATF6 transcription factor induces expression of XBP1 mRNA [16]. Ire1 has an endoribonuclease domain, by which it splices a 26 nucleotide fragment out of the XBP1 mRNA. The generated frame-shift splice variant (sXBP1) codes for a transcription factor that induces the expression of ER chaperones. Therefore, accumulation of sXBP1 mRNA in the cell is the result of the coordinated action of ATF6 and Ire1.
Phosphorylation of eIF2a was clearly detectable as early as 1 h after induction of ischemia and the level of phosphorylation was comparable to that induced by thapsigargin (Fig. 3C and D). Phosphorylation of eIF2a was followed by accumulation of both ATF4 and GADD34 mRNA, detected by RT-PCR (Fig. 3E and F). These results indicate that in addition to the Ire1 and ATF6 pathways, the PERK pathway is also activated early during ischemia. Activated PERK phosphorylates and inhibits the a-subunit of the eukaryotic initiation factor 2 (eIF2a), which leads to atten-
Fig. 3. Ischemia activates all three arms of the UPR. Cardiomyocyte cultures were subjected to ischemia or treated with thapsigargin (Tg, 2 lM) for the times indicated. Accumulation of spliced XBP1 (sXBP1) mRNA in response to (A) ischemia, and (B) thapsigargin treatment was detected by RT-PCR in total RNA samples. GAPDH mRNA was also amplified and used as a loading control. Phosphorylation of eIF2a in response to (C) ischemia and (D) thapsigargin treatment was detected in whole cell lysates by Western blotting using p-eIF2a antibody. Expression of total eIF2a and actin was detected to serve as loading controls. Induction of the PERK-targets ATF4 and GADD34 mRNAs during (E) ischemia, and (F) thapsigargin treatment was determined by RT-PCR on total RNA samples. GAPDH message was also amplified and used as a loading control. All images are representative of three independent experiments.
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Fig. 4. Prolonged ischemia leads to induction of an ER stress-mediated apoptotic pathway. Cardiomyocytes were exposed to ischemia or thapsigargin (Tg, 2 lM) for 0–24 h and whole cell lysates were prepared. (A) Expression of CHOP in ischemic (upper panel) and thapsigargin-treated (lower panel) cardiomyocytes was detected by Western blotting. (B) Prolonged ischemia (upper panel) and thapsigargin treatment (lower panel) induce pro-caspase-12 processing detected on Western blots of whole cell lysates. Membranes were probed for actin and used as a loading control.
uation of general (cap-dependent) protein translation. Inhibition of the cap-dependent protein synthesis enables the translation of ATF4 transcription factor and GADD34 protein phosphatase 1 (PP1)-interacting protein, through an alternative, cap-independent translation pathway [17]. The ER stress-induced apoptotic pathway is activated during prolonged ischemia During prolonged or severe ER stress the cytoprotective UPR can switch to a pro-apoptotic response to initiate cell death. Therefore, we questioned whether the ER stress-mediated apoptotic pathway is a component of ischemiainduced cell death in primary cardiomyocytes. So far, the ER stress-mediated apoptotic pathway is only partially characterized, although some ER stress-specific components of the pathway have been identified. For instance, the transcription factor CHOP has been shown to play a central role by altering the balance of pro- and anti-apoptotic Bcl-2 proteins to promote apoptosis [18]. Another specific component of the pathway is caspase-12, which is localized to the ER and is processed only in response to ER stress [8]. To confirm that these proteins are actual markers of ER stress-induced apoptosis in cardiomyocytes, both proteins were examined in thapsigargin-treated cardiomyocytes. Induction of CHOP as well as processing of pro-caspase12 was detectable in response to thapsigargin treatment (Fig. 4A and B). Exposure of primary cardiomyocyte cultures to ischemia also led to induction of CHOP, which was detectable following 2 h ischemia (Fig. 4A). Ischemia also triggered processing of pro-caspase-12, which occurred between 4 and 8 h of ischemia (Fig. 4B). These results point out that not only the protective UPR is prompted by ischemic stress, but prolonged ischemia induces severe damage that initiates the ER stress-mediated apoptotic pathway. Discussion While there is mounting evidence emphasizing the role of the UPR and ER stress in cerebral ischemia, there is
only circumstantial evidence available implying involvement of the ER stress response in cardiac ischemia [19]. It was recently demonstrated that Grp78 induction and XBP1 splicing occur in hypoxic cardiomyocytes [20,21]. Zhang et al. also demonstrated that pre-induction of Grp78 by tunicamycin, an ER stressor, protects cardiomyocytes from lethal injury due to prolonged ATP depletion or excessive oxidative stress [10]. There is no information, however, regarding the activation of the UPR, in particular which branches of the UPR are activated, or the kinetics of their activation. Here, we show that ischemia, without reperfusion, is sufficient to activate all three branches of the UPR in primary neonatal rat cardiomyocytes. We detected marked induction of sXBP1 mRNA, a marker for the coordinated action of active ATF6 and Ire1. The early appearance of sXBP1 mRNA, during the first 2 h of ischemia, and the level of induction were comparable to those induced by thapsigargin, the classical ER stressor. Activation of the third branch of the UPR, the PERK pathway also occurred. The PERK substrate eIF2a was phosphorylated during the first hour of ischemia, to a similar extent as it was phosphorylated by thapsigargin. Phosphorylation of eIF2a was followed by the induction of the transcription factor ATF4 and the PP1-interacting protein GADD34, both genes specific targets of the PERK/eIF2a pathway. The UPR is primarily an adaptive response, aiming to restore ER homeostasis and protect the cell from stress. The importance of the UPR in protecting stressed cardiomyocytes was first shown by Vitadello et al., who demonstrated that overexpression of Grp94 reduces ischemic cell death in cardiomyocytes [22]. However, if the stress is prolonged and the adaptive response fails, the UPR receptors can launch a pro-apoptotic response and initiate cell suicide. This apoptotic programme originates from the ER, independent of the mitochondria, and may contribute to cardiomyocyte loss. Although the ER stress-mediated apoptotic pathway is only partially characterized, two specific hallmarks of the process have been identified [23]. These are the induction of the pro-apoptotic transcription
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factor CHOP and activation of the ER-localized caspase, caspase-12. Prolonged ischemia led to induction of CHOP, which occurred between 4 and 8 h of exposure. Processing of pro-caspase-12 was also detectable, and occurred in the same time frame, between 4 and 8 h of ischemic exposure. These results emphasize that ischemia not only induces the UPR, but if prolonged, the UPR receptors launch a proapoptotic response which probably drives the cell towards death. Although, we have shown that cardiomyocytes respond to an ischemic insult by activating all three UPR receptors to counteract the cellular stresses evoked by SGO deprivation, in the setting of prolonged ischemia the UPR fails to restore cellular homeostasis and a proapoptotic ER stress pathway is initiated. The regulation of this effect may be via upregulation of CHOP and cleavage of caspase-12. The activated ER stress death pathway, in concert with the mitochondrial death pathway, causes caspase activation, cytoskeletal breakdown, and nuclear fragmentation, leading to the final demise of the cell. The ER stress pathway is therefore a novel target for prevention of ischemia-mediated cardiomyocyte loss. Acknowledgments This work was supported by grants from Higher Education Authority of Ireland (PRTLI-III), SFI CSET grant to REMEDI and SFI PI award to A.S. References [1] M. Avkiran, Protection of the ischaemic myocardium by Na+/H+ exchange inhibitors: potential mechanisms of action, Basic Res. Cardiol. 96 (2001) 306–311. [2] W.E. Cascio, T.A. Johnson, L.S. Gettes, Electrophysiologic changes in ischemic ventricular myocardium: I. Influence of ionic, metabolic, and energetic changes, J. Cardiovasc. Electrophysiol. 6 (1995) 1039–1062. [3] L.H. Opie, Myocardial metabolism and heart disease, Jpn. Circ. J. 42 (1978) 1223–1247. [4] T. Iwai, K. Tanonaka, R. Inoue, S. Kasahara, N. Kamo, S. Takeo, Mitochondrial damage during ischemia determines post-ischemic contractile dysfunction in perfused rat heart, J. Mol. Cell. Cardiol. 34 (2002) 725–738. [5] A. Samali, S. Orrenius, Heat shock proteins: regulators of stress response and apoptosis, Cell Stress Chaperones 3 (1998) 228–236. [6] T. Hayashi, A. Saito, S. Okuno, M. Ferrand-Drake, R.L. Dodd, P.H. Chan, Oxidative injury to the endoplasmic reticulum in mouse brains after transient focal ischemia, Neurobiol. Dis. 15 (2004) 229–239. [7] D.T. Rutkowski, R.J. Kaufman, A trip to the ER: coping with stress, Trends Cell Biol. 14 (2004) 20–28.
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[8] E. Szegezdi, U. Fitzgerald, A. Samali, Caspase-12 and ER-stressmediated apoptosis: the story so far, Ann. N.Y. Acad. Sci. 1010 (2003) 186–194. [9] D.J. Thuerauf, H. Hoover, J. Meller, J. Hernandez, L. Su, C. Andrews, W.H. Dillmann, P.M. McDonough, C.C. Glembotski, Sarco/endoplasmic reticulum calcium ATPase-2 expression is regulated by ATF6 during the endoplasmic reticulum stress response: intracellular signaling of calcium stress in a cardiac myocyte model system, J. Biol. Chem. 276 (2001) 48309–48317. [10] P.L. Zhang, M. Lun, J. Teng, J. Huang, T.M. Blasick, L. Yin, G.A. Herrera, J.Y. Cheung, Preinduced molecular chaperones in the endoplasmic reticulum protect cardiomyocytes from lethal injury, Ann. Clin. Lab. Sci. 34 (2004) 449–457. [11] A. Azfer, J. Niu, L.M. Rogers, F.M. Adamski, P.E. Kolattukudy, Activation of endoplasmic reticulum stress response during the development of ischemic heart disease, Am. J. Physiol. Heart Circ. Physiol. 291 (2006) H1411–H1420. [12] J.L. Reeve, A.M. Duffy, T. O’Brien, A. Samali, Don’t lose heart–therapeutic value of apoptosis prevention in the treatment of cardiovascular disease, J. Cell. Mol. Med. 9 (2005) 609–622. [13] T.M. Scarabelli, R.A. Gottlieb, Functional and clinical repercussions of myocyte apoptosis in the multifaceted damage by ischemia/ reperfusion injury: old and new concepts after 10 years of contributions, Cell Death Differ. 11 (Suppl. 2) (2004) S144–S152. [14] P. Simpson, S. Savion, Differentiation of rat myocytes in single cell cultures with and without proliferating nonmyocardial cells. Crossstriations, ultrastructure, and chronotropic response to isoproterenol, Circ. Res. 50 (1982) 101–116. [15] C. Communal, M. Sumandea, P. de Tombe, J. Narula, R.J. Solaro, R.J. Hajjar, Functional consequences of caspase activation in cardiac myocytes, Proc. Natl. Acad. Sci. USA 99 (2002) 6252–6256. [16] A.H. Lee, N.N. Iwakoshi, L.H. Glimcher, XBP-1 regulates a subset of endoplasmic reticulum resident chaperone genes in the unfolded protein response, Mol. Cell. Biol. 23 (2003) 7448–7459. [17] J.D. Blais, V. Filipenko, M. Bi, H.P. Harding, D. Ron, C. Koumenis, B.G. Wouters, J.C. Bell, Activating transcription factor 4 is translationally regulated by hypoxic stress, Mol. Cell. Biol. 24 (2004) 7469–7482. [18] K.D. McCullough, J.L. Martindale, L.O. Klotz, T.Y. Aw, N.J. Holbrook, Gadd153 sensitizes cells to endoplasmic reticulum stress by down-regulating Bcl2 and perturbing the cellular redox state, Mol. Cell. Biol. 21 (2001) 1249–1259. [19] D.J. DeGracia, H.L. Montie, Cerebral ischemia and the unfolded protein response, J. Neurochem. 91 (2004) 1–8. [20] K. Terai, Y. Hiramoto, M. Masaki, S. Sugiyama, T. Kuroda, M. Hori, I. Kawase, H. Hirota, AMP-activated protein kinase protects cardiomyocytes against hypoxic injury through attenuation of endoplasmic reticulum stress, Mol. Cell. Biol. 25 (2005) 9554–9575. [21] D.J. Thuerauf, M. Marcinko, N. Gude, M. Rubio, M.A. Sussman, C.C. Glembotski, Activation of the unfolded protein response in infarcted mouse heart and hypoxic cultured cardiac myocytes, Circ. Res. 99 (2006) 275–282. [22] M. Vitadello, D. Penzo, V. Petronilli, G. Michieli, S. Gomirato, R. Menabo, F. Di Lisa, L. Gorza, Overexpression of the stress protein Grp94 reduces cardiomyocyte necrosis due to calcium overload and simulated ischemia, Faseb J. 17 (2003) 923–925. [23] E. Szegezdi, S.E. Logue, A.M. Gorman, E. Samali, Mediators of ER stress-induced apoptosis, EMBO Rep. 7 (2006) 880–885.
ER stress contributes to ischemia-induced cardiomyocyte apoptosis Eva Szegezdi a, Angela Duffy c, Martin E. O’Mahoney a, Susan E. Logue a, Louise A. Mylotte c, Timothy O’Brien b,c, Afshin Samali a,c,* a
c
Department of Biochemistry, National University of Ireland, Galway, Ireland b Department of Medicine, National University of Ireland, Galway, Ireland Regenerative Medicine Institute (REMEDI), National Centre for Biomedical Engineering Science, National University of Ireland, Galway, Ireland Received 1 September 2006 Available online 12 Septemeber 2006
Abstract Myocardial ischemia is a severe stress condition that leads to loss of cardiomyocytes. The cell loss is attributed to apoptosis, although the exact mechanisms involved are only partially defined, which limits therapeutic opportunities. Here, we show caspase activation and apoptosis in neonatal rat cardiomyocyte cultures subjected to simulated ischemia by serum, glucose, and oxygen deprivation (SGO). Caspase activation was preceded by endoplasmic reticulum (ER) stress and the activation of the unfolded protein response (UPR), detected by the induction of Grp78, induction and splicing of XBP1, and phosphorylation of eukaryotic initiation factor 2-a (eIF2a). At a later time the ER stress response switched from UPR and cytoprotective response to a pro-apoptotic response as demonstrated by the upregulation of CHOP and processing of pro-caspase-12. Thus, we provide evidence that the ER can generate and propagate apoptotic signals in response to ischemic stress and this pathway is therefore a novel target for prevention of ischemia-mediated cardiomyocyte loss. 2006 Elsevier Inc. All rights reserved. Keywords: Cardiomyocytes; Ischemia; Unfolded protein response; ER stress; Apoptosis; PERK; Ire1; ATF6; CHOP; Caspase-12
Myocardial ischemia is a severe trauma for cardiomyocytes. The simultaneous drop in cellular oxygen and glucose supply leads to ATP depletion, which triggers profound molecular alterations. Initially, the Na+/K+ pump, which is the major Na+ extrusion pathway in cardiomyocytes, comes to a halt leading to the accumulation of Na+ in the cytosol. High intracellular Na+ concentration in turn blocks or reverses the function of the plasma membrane Na+/Ca2+ exchanger, causing Ca2+ influx. Simultaneously, the intracellular K+ and Mg2+ concentration drops [1,2]. In addition to the ionic changes, inability to resynthesize ATP by oxidative phosphorylation also leads to acidification due to accumulation of inorganic phosphate and the glycolytic end-product lactate [3]. As a result, ischemia causes extensive biochemical changes in the cytosol, but also affects the cytoskeleton *
Corresponding author. Fax: +353 91 494596. E-mail address: [email protected] (A. Samali).
0006-291X/$ - see front matter 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2006.09.009
and other cellular organelles. For example, the intermediate filaments of the cytoskeleton collapse into large perinuclear aggregates, the actin fibres relocalize around the nucleus, and microtubules break up. Mitochondria swell and lose structure, leading to cessation of the oxidative phosphorylation [4]. To cope with the stress, both cytosolic and mitochondrial heat shock proteins, including Hsp70, 60, and 27, are induced [5]. In the brain, ischemia has been shown to also target the endoplasmic reticulum (ER) and ischemic neurons activate the ER stress response (unfolded protein response, UPR) to protect themselves [6]. Generally, the UPR is triggered by accumulation of protein aggregates in the ER lumen. The ER lumen has a high redox potential to maintain an oxidizing environment, which, together with the high protein and Ca2+ concentration, provides the ideal milieu for protein folding. Physiological or pathophysiological conditions that alter this milieu hinder protein folding and cause aggregation of nascent polypeptide chains [7]. Three ER
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transmembrane receptors, namely PERK, ATF6, and Ire1, monitor the homeostasis of the ER and trigger the UPR. The UPR acts on several levels; it rapidly attenuates general protein synthesis, induces the expression of ER chaperone proteins, and enhances the degradation of malfolded proteins [7]. Although the UPR is primarily an adaptive response, if the stress persists, the ER stress receptors can also trigger pro-apoptotic pathways to initiate cell death [8]. The sarco-endoplasmic reticulum of cardiomyocytes forms an extensive network that interweaves the cell. In addition to its general cellular functions, the sarcoplasmic reticulum is also vital for contractibility of cardiomyocytes. Nevertheless, to date little is known as to how ischemia affects the homeostasis of the sarcoplasmic reticulum, and only indirect evidence is available to suggest that ischemia in the heart induces ER stress. It has been shown that reduction of ER Ca2+ activates ATF6 and induces Grp78 in cardiomyocyte cultures [9]. Also, ER preconditioning with low dose tunicamycin protects H9c2 neonatal cardiomyocyte cells against prolonged ATP depletion or oxidative stress [10]. A DNA microarray analysis of the heart from MPC-1 (monocyte chemotactic protein-1) overexpressing mice, an experimental model of ischemia, identified induction of ER chaperones [11]. The involvement of apoptosis in cardiomyocyte death during ischemia is a relatively new concept and to date most studies have focused on the role of the death receptor and mitochondrial death pathways [12,13]. Here, we examined whether ischemia induces the UPR and more importantly, whether induction of the UPR during ischemia activates the ER stress death pathway. We show for the first time that ischemia induces all three arms of the UPR: PERK, ATF6, Ire1, and their downstream targets. In addition to the UPR, induction of CHOP and cleavage of pro-caspase-12 were also detected. These data demonstrate activation of UPR during ischemia in cardiomyocytes and that ER stress is involved in the onset of ischemic cell death. Materials and methods Cell culture and treatments. Primary cultures of neonatal cardiomyocytes were isolated from 1 to 4-day-old Sprague–Dawley rats, as described previously [14]. Briefly, rats were euthanized and hearts excised. After scalpel homogenization, overnight trypsin digestion at 4 C, and a collagenase treatment for 20 min at 37 C, cardiomyocytes were enriched by Percoll gradient centrifugation (Amersham) and plated at a density of 1 · 105/ml in DMEM/F12 medium supplemented with 10% newborn calf serum, 100 U/ ml penicillin, 100 lg/ml streptomycin, 1 mM sodium pyruvate (GibcoBRL), 5% insulin transferrin selenite (ITS) liquid supplement media, and 100 lM 5-bromo-2-deoxyuridine on culture plates coated with 0.2% gelatin. Cells were cultured at 37 C and 5% CO2. To mimic endogenous ischemia, cultures were exposed to hypoxic conditions (O2/N2/CO2, 0.5:94.5:5), using a hypoxia gas chamber (Russkin) in the absence of glucose and serum, using glucose-free DMEM (Gibco-BRL) supplemented with 10 mM 2-deoxyglucose, 100 U/ml penicillin, 100 lg/ml streptomycin, 1 mM sodium pyruvate, and 5% ITS liquid supplement media. Thapsigargin (2 lM) in normal culture medium was used to induce ER stress. All animal experiments were performed in accordance with the ethical regulations of NUI, Galway. All reagents were from Sigma, unless otherwise stated.
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Western blot analysis. Cells were lysed in whole cell lysis buffer (20 mM Hepes, pH 7.5, 350 mM NaCl, 1 mM MgCl2, 0.5 mM EDTA, 0.5 mM EGTA, 1% Igepal-630, 0.5 mM dithiothreitol (DTT), 100 lM PMSF, and 1 lg/ml pepstatin), except for detection of eIF2a phosphorylation, where cells were directly lysed in Laemmli buffer. Cellular proteins were separated by electrophoresis on 8–12% SDS–polyacrylamide gels and transferred onto nitrocellulose membranes. After blocking (5% non-fat milk, 0.05% Tween 20 in PBS), blots were incubated with antibodies to caspase12 (Sigma, rat monoclonal), PKCd (Santa Cruz, rabbit polyclonal), caspase-3 (Cell Signalling Technology, rabbit polyclonal), sarcomeric a-actinin (Sigma, mouse monoclonal), Grp78 (StressGen, rabbit polyclonal), XBP-1 (Santa Cruz, rabbit polyclonal), phospho-eIF2a, and total eIF2a (both from Cell Signalling Technology, rabbit monoclonal) and CHOP (Santa Cruz, rabbit polyclonal). To verify equal protein loading, blots were probed for actin (Sigma, rabbit polyclonal). All primary antibodies were diluted 1:1000 and the appropriate horseradish peroxidise-conjugated goat secondary antibodies (Pierce) were used at a 1:5000 dilution. Protein bands were detected with SuperSignal Ultra Chemiluminescent Substrate (Pierce) on X-ray film (Agfa). Analysis of caspase activity. Cell lysates (25 ll) and 50 lM carbobenzoxy-Asp-Glu-Val-Asp-7-amino-4-methyl-coumarin (DEVD-AMC) in reaction buffer (100 mM Hepes, 10% sucrose, 5 mM DTT, 0.0001% Igepal-630, and 0.1% 3-[(3-cholamidopropyl) dimethylammonio] propane-1sulphonic acid (CHAPS), pH 7.25) were added in duplicate to a microtiter plate. Liberated AMC was measured using 355 nm excitation and 460 nm emission wavelengths. Fluorescence was measured over time and the data analysed by linear regression to determine enzyme activity. Enzyme activity was expressed as nmoles AMC liberated by 1 mg total cellular protein per minute. Hoechst staining. Following treatment, cells grown on coverslips were fixed in 3.7% formaldehyde for 10 min at room temperature and permeabilized with methanol for an additional 10 min. After a PBS wash, cells were stained with 100 lg/ml Hoechst 33342. The excess stain was removed by a series of PBS washes. The stained nuclei were visualized using an Olympus BX51 fluorescent microscope. Haematoxylin–eosin staining. Cells were seeded onto 35 mm dishes at a density of 5 · 104 cells/ml. After treatment the cells were fixed in 3.7% formaldehyde for 5 min at room temperature. The cytosol was stained with eosin Y for 4 min, rinsed in tap water, and stained in Harris haematoxylin solution for 5 min. Pictures were taken using an Olympus BX51 microscope at a final magnification of 100·. Immunocytochemical staining for a-actinin. Cells were fixed in 3.7% formaldehyde for 5 min at room temperature and permeabilized with 0.2% Triton X-100 for an additional 5 min. After rinsing with PBS, coverslips were blocked in 1% BSA containing 5% goat serum for 1 h at room temperature. Cells were incubated with 1:1000 dilution of sarcomeric antia-actinin antibody in 1% BSA for 1 h at room temperature. The excess antibody solution was removed by 3 series of PBS washes. Cells were then incubated in 1% BSA containing 1:100 dilution of FITC-conjugated antimouse IgG (Sigma) and 100 lg/ml Hoechst 33342 stain in darkness for 45 min at room temperature. The excess antibody solution was removed and cells were mounted using Vectashield mounting medium. The stained cells were visualized using an Olympus BX51 fluorescent microscope. RT-PCR. Total RNA from cells exposed to thapsigargin or ischemic treatment was isolated using the GenElute mammalian total RNA extraction kit (Sigma) according to the manufacturer’s instructions. Reverse transcription was carried out with 2 lg total RNA and oligo(dT) (Invitrogen) using 20 U/25 ll reaction AMV reverse transcriptase (Sigma) according to the manufacturer’s instructions. ATF4-, XBP1-, spliced XBP1 (sXBP1), and GADD34-specific sequences were amplified during 32 cycles of 30 s denaturing at 94 C, 60 s annealing at 56 C, and 60 s extension at 72 C, with the following primers: ATF4 forward 5 0 -C CGAGATGAGCTTCCTGA-3 0 ; ATF4 reverse 5 0 -CTCCTTGCCGGT GTCTGA; XBP1 forward 5 0 -CAGACTACGTGCGCCTCTGC-3 0 ; XBP1 reverse 5 0 -CTTCTGGGTAGACCTCTGGG; sXBP1 forward 5 0 -TCT GCTGAGTCCGCAGCAGG-3 0 ; sXBP1 reverse 5 0 -CTCTAAGACTAG AGGCTTGG; GADD34 forward: TTTCTAGGCCAGACACATGG; G ADD34 reverse: TGTTCCTTTTTCCTCCGTGG. GAPDH was used as
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a loading control, its cDNA was amplified during 26 cycles of 30 s denaturing at 94 C, 60 s annealing at 56 C, and 60 s extension at 72 C, with the following primers: GAPDH forward 5 0 -ACCACAGTCCA TGCCATC-3 0 ; GAPDH reverse 5 0 -TCCACCACCCTGTTGCTG.
Results Ischemia initiates apoptosis in cardiomyocytes All experiments were carried out on primary neonatal rat cardiomyocyte preparations. The purity of the preparations was between 85% and 97% cardiomyocytes, based on sarcoplasmic a-actinin immunostaining (data not shown). To mimic in vivo ischemic conditions, cells were subjected to a combination of serum, glucose, and oxygen deprivation (SGO) as described in Materials and methods. After 24 h ischemia shrunken cells with condensed or fragmented nuclei, characteristic of apoptotic cells, were detectable in cardiomyocyte cultures (Fig. 1A). The apoptotic mode of cell death was confirmed by detecting activation and activity of caspase proteases. Cardiomyocyte cultures subjected to 4 h ischemia expressed a 3-fold increase in DEVDase activity, which was associated with processing of pro-caspase-3 and cleavage of the caspase-3 substrate, PCKd (Fig. 1B). Primary cardiomyocyte cultures may contain a small percentage of fibroblasts or other non-cardiomyocyte cells. Immunostaining for a-actinin, a cardiomyocyte-specific caspase-3 substrate [15], revealed its breakdown and loss of staining following 4 h ischemia (Fig. 1C). Western blot analysis identified 45 and 25 kDa processed fragments, confirming cleavage of a-actinin in ischemic cardiomyocytes (Fig. 1C). Collectively, these data confirmed that exposure of primary neonatal cardiomyocytes to ischemia induces apoptotic cell death. Ischemia triggers the unfolded protein response in primary cardiomyocytes In order to examine the effect of ischemia on the ER, induction of Grp78 (Bip), a marker of ER stress was determined. Ischemia (2–4 h) induced Grp78 both at mRNA and protein levels (Fig. 2A and B). To compare the kinetics and the extent of Grp78 induction, cardiomyocyes were treated with a specific ER stressor, thapsigargin (an inhibitor of the SERCA pump). Thapsigargin caused a similar increase in Grp78 expression as ischemia, although the kinetics of mRNA induction was faster, between 0 and 2 h treatment (Fig. 2A and B). Since induction of Grp78 is indicative of the activation of the UPR, we examined which of the three branches of the UPR are activated by ischemia in cardiomyocytes. The activation of the UPR receptors Ire1, ATF6, and PERK was detected by monitoring their target molecules. In primary cardiomyocytes, induction of a sXBP1 mRNA (frame-shift splice variant of XBP1) occurred during the first 2 h of exposure to ischemia, and the level and the kinetics of induction were similar to those of thapsigar-
Fig. 1. Ischemia induces apoptotic cell death in primary rat cardiomyocytes. To mimic ischemia, cardiomyocytes were exposed to serum, oxygen, and glucose deprivation for the times indicated. (A) Morphological changes were determined in cardiomyocytes subjected to 24 h ischemia by both Haematoxylin–Eosin staining (upper panel) and Hoechst staining (lower panel). (B) DEVDase activity in ischemic cardiomyocytes was measured in whole cell lysates. The graph shows the average of three separate experiments ± SD. (C) Caspase-mediated degradation of sarcomeric a-actinin in ischemic cardiomyocytes detected by immunostaining (upper panel) and by Western blotting of whole cell lysates (lower panel).
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Fig. 2. Ischemia induces expression of Grp78 at both mRNA and protein levels. Cardiomyocyte cultures were subjected to ischemia or thapsigargin (Tg, 2 lM) for the indicated times. (A) Total RNA was isolated from ischemic (upper panel) and thapsigargin-treated (lower panel) cardiomyocytes and the Grp78 message was amplified by RT-PCR. GAPDH was used as a loading control. (B) Induction of Grp78 protein expression in ischemic (upper panel) and thapsigargin-treated (lower panel) cardiomyocytes detected by Western blotting. Actin was used as a loading control. The images are representatives of three separate experiments.
gin-treated cells, indicating that ATF6 and Ire1 are both activated during ischemia (Fig. 3A and B). Activation of ATF6 transcription factor induces expression of XBP1 mRNA [16]. Ire1 has an endoribonuclease domain, by which it splices a 26 nucleotide fragment out of the XBP1 mRNA. The generated frame-shift splice variant (sXBP1) codes for a transcription factor that induces the expression of ER chaperones. Therefore, accumulation of sXBP1 mRNA in the cell is the result of the coordinated action of ATF6 and Ire1.
Phosphorylation of eIF2a was clearly detectable as early as 1 h after induction of ischemia and the level of phosphorylation was comparable to that induced by thapsigargin (Fig. 3C and D). Phosphorylation of eIF2a was followed by accumulation of both ATF4 and GADD34 mRNA, detected by RT-PCR (Fig. 3E and F). These results indicate that in addition to the Ire1 and ATF6 pathways, the PERK pathway is also activated early during ischemia. Activated PERK phosphorylates and inhibits the a-subunit of the eukaryotic initiation factor 2 (eIF2a), which leads to atten-
Fig. 3. Ischemia activates all three arms of the UPR. Cardiomyocyte cultures were subjected to ischemia or treated with thapsigargin (Tg, 2 lM) for the times indicated. Accumulation of spliced XBP1 (sXBP1) mRNA in response to (A) ischemia, and (B) thapsigargin treatment was detected by RT-PCR in total RNA samples. GAPDH mRNA was also amplified and used as a loading control. Phosphorylation of eIF2a in response to (C) ischemia and (D) thapsigargin treatment was detected in whole cell lysates by Western blotting using p-eIF2a antibody. Expression of total eIF2a and actin was detected to serve as loading controls. Induction of the PERK-targets ATF4 and GADD34 mRNAs during (E) ischemia, and (F) thapsigargin treatment was determined by RT-PCR on total RNA samples. GAPDH message was also amplified and used as a loading control. All images are representative of three independent experiments.
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Fig. 4. Prolonged ischemia leads to induction of an ER stress-mediated apoptotic pathway. Cardiomyocytes were exposed to ischemia or thapsigargin (Tg, 2 lM) for 0–24 h and whole cell lysates were prepared. (A) Expression of CHOP in ischemic (upper panel) and thapsigargin-treated (lower panel) cardiomyocytes was detected by Western blotting. (B) Prolonged ischemia (upper panel) and thapsigargin treatment (lower panel) induce pro-caspase-12 processing detected on Western blots of whole cell lysates. Membranes were probed for actin and used as a loading control.
uation of general (cap-dependent) protein translation. Inhibition of the cap-dependent protein synthesis enables the translation of ATF4 transcription factor and GADD34 protein phosphatase 1 (PP1)-interacting protein, through an alternative, cap-independent translation pathway [17]. The ER stress-induced apoptotic pathway is activated during prolonged ischemia During prolonged or severe ER stress the cytoprotective UPR can switch to a pro-apoptotic response to initiate cell death. Therefore, we questioned whether the ER stress-mediated apoptotic pathway is a component of ischemiainduced cell death in primary cardiomyocytes. So far, the ER stress-mediated apoptotic pathway is only partially characterized, although some ER stress-specific components of the pathway have been identified. For instance, the transcription factor CHOP has been shown to play a central role by altering the balance of pro- and anti-apoptotic Bcl-2 proteins to promote apoptosis [18]. Another specific component of the pathway is caspase-12, which is localized to the ER and is processed only in response to ER stress [8]. To confirm that these proteins are actual markers of ER stress-induced apoptosis in cardiomyocytes, both proteins were examined in thapsigargin-treated cardiomyocytes. Induction of CHOP as well as processing of pro-caspase12 was detectable in response to thapsigargin treatment (Fig. 4A and B). Exposure of primary cardiomyocyte cultures to ischemia also led to induction of CHOP, which was detectable following 2 h ischemia (Fig. 4A). Ischemia also triggered processing of pro-caspase-12, which occurred between 4 and 8 h of ischemia (Fig. 4B). These results point out that not only the protective UPR is prompted by ischemic stress, but prolonged ischemia induces severe damage that initiates the ER stress-mediated apoptotic pathway. Discussion While there is mounting evidence emphasizing the role of the UPR and ER stress in cerebral ischemia, there is
only circumstantial evidence available implying involvement of the ER stress response in cardiac ischemia [19]. It was recently demonstrated that Grp78 induction and XBP1 splicing occur in hypoxic cardiomyocytes [20,21]. Zhang et al. also demonstrated that pre-induction of Grp78 by tunicamycin, an ER stressor, protects cardiomyocytes from lethal injury due to prolonged ATP depletion or excessive oxidative stress [10]. There is no information, however, regarding the activation of the UPR, in particular which branches of the UPR are activated, or the kinetics of their activation. Here, we show that ischemia, without reperfusion, is sufficient to activate all three branches of the UPR in primary neonatal rat cardiomyocytes. We detected marked induction of sXBP1 mRNA, a marker for the coordinated action of active ATF6 and Ire1. The early appearance of sXBP1 mRNA, during the first 2 h of ischemia, and the level of induction were comparable to those induced by thapsigargin, the classical ER stressor. Activation of the third branch of the UPR, the PERK pathway also occurred. The PERK substrate eIF2a was phosphorylated during the first hour of ischemia, to a similar extent as it was phosphorylated by thapsigargin. Phosphorylation of eIF2a was followed by the induction of the transcription factor ATF4 and the PP1-interacting protein GADD34, both genes specific targets of the PERK/eIF2a pathway. The UPR is primarily an adaptive response, aiming to restore ER homeostasis and protect the cell from stress. The importance of the UPR in protecting stressed cardiomyocytes was first shown by Vitadello et al., who demonstrated that overexpression of Grp94 reduces ischemic cell death in cardiomyocytes [22]. However, if the stress is prolonged and the adaptive response fails, the UPR receptors can launch a pro-apoptotic response and initiate cell suicide. This apoptotic programme originates from the ER, independent of the mitochondria, and may contribute to cardiomyocyte loss. Although the ER stress-mediated apoptotic pathway is only partially characterized, two specific hallmarks of the process have been identified [23]. These are the induction of the pro-apoptotic transcription
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factor CHOP and activation of the ER-localized caspase, caspase-12. Prolonged ischemia led to induction of CHOP, which occurred between 4 and 8 h of exposure. Processing of pro-caspase-12 was also detectable, and occurred in the same time frame, between 4 and 8 h of ischemic exposure. These results emphasize that ischemia not only induces the UPR, but if prolonged, the UPR receptors launch a proapoptotic response which probably drives the cell towards death. Although, we have shown that cardiomyocytes respond to an ischemic insult by activating all three UPR receptors to counteract the cellular stresses evoked by SGO deprivation, in the setting of prolonged ischemia the UPR fails to restore cellular homeostasis and a proapoptotic ER stress pathway is initiated. The regulation of this effect may be via upregulation of CHOP and cleavage of caspase-12. The activated ER stress death pathway, in concert with the mitochondrial death pathway, causes caspase activation, cytoskeletal breakdown, and nuclear fragmentation, leading to the final demise of the cell. The ER stress pathway is therefore a novel target for prevention of ischemia-mediated cardiomyocyte loss. Acknowledgments This work was supported by grants from Higher Education Authority of Ireland (PRTLI-III), SFI CSET grant to REMEDI and SFI PI award to A.S. References [1] M. Avkiran, Protection of the ischaemic myocardium by Na+/H+ exchange inhibitors: potential mechanisms of action, Basic Res. Cardiol. 96 (2001) 306–311. [2] W.E. Cascio, T.A. Johnson, L.S. Gettes, Electrophysiologic changes in ischemic ventricular myocardium: I. Influence of ionic, metabolic, and energetic changes, J. Cardiovasc. Electrophysiol. 6 (1995) 1039–1062. [3] L.H. Opie, Myocardial metabolism and heart disease, Jpn. Circ. J. 42 (1978) 1223–1247. [4] T. Iwai, K. Tanonaka, R. Inoue, S. Kasahara, N. Kamo, S. Takeo, Mitochondrial damage during ischemia determines post-ischemic contractile dysfunction in perfused rat heart, J. Mol. Cell. Cardiol. 34 (2002) 725–738. [5] A. Samali, S. Orrenius, Heat shock proteins: regulators of stress response and apoptosis, Cell Stress Chaperones 3 (1998) 228–236. [6] T. Hayashi, A. Saito, S. Okuno, M. Ferrand-Drake, R.L. Dodd, P.H. Chan, Oxidative injury to the endoplasmic reticulum in mouse brains after transient focal ischemia, Neurobiol. Dis. 15 (2004) 229–239. [7] D.T. Rutkowski, R.J. Kaufman, A trip to the ER: coping with stress, Trends Cell Biol. 14 (2004) 20–28.
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[8] E. Szegezdi, U. Fitzgerald, A. Samali, Caspase-12 and ER-stressmediated apoptosis: the story so far, Ann. N.Y. Acad. Sci. 1010 (2003) 186–194. [9] D.J. Thuerauf, H. Hoover, J. Meller, J. Hernandez, L. Su, C. Andrews, W.H. Dillmann, P.M. McDonough, C.C. Glembotski, Sarco/endoplasmic reticulum calcium ATPase-2 expression is regulated by ATF6 during the endoplasmic reticulum stress response: intracellular signaling of calcium stress in a cardiac myocyte model system, J. Biol. Chem. 276 (2001) 48309–48317. [10] P.L. Zhang, M. Lun, J. Teng, J. Huang, T.M. Blasick, L. Yin, G.A. Herrera, J.Y. Cheung, Preinduced molecular chaperones in the endoplasmic reticulum protect cardiomyocytes from lethal injury, Ann. Clin. Lab. Sci. 34 (2004) 449–457. [11] A. Azfer, J. Niu, L.M. Rogers, F.M. Adamski, P.E. Kolattukudy, Activation of endoplasmic reticulum stress response during the development of ischemic heart disease, Am. J. Physiol. Heart Circ. Physiol. 291 (2006) H1411–H1420. [12] J.L. Reeve, A.M. Duffy, T. O’Brien, A. Samali, Don’t lose heart–therapeutic value of apoptosis prevention in the treatment of cardiovascular disease, J. Cell. Mol. Med. 9 (2005) 609–622. [13] T.M. Scarabelli, R.A. Gottlieb, Functional and clinical repercussions of myocyte apoptosis in the multifaceted damage by ischemia/ reperfusion injury: old and new concepts after 10 years of contributions, Cell Death Differ. 11 (Suppl. 2) (2004) S144–S152. [14] P. Simpson, S. Savion, Differentiation of rat myocytes in single cell cultures with and without proliferating nonmyocardial cells. Crossstriations, ultrastructure, and chronotropic response to isoproterenol, Circ. Res. 50 (1982) 101–116. [15] C. Communal, M. Sumandea, P. de Tombe, J. Narula, R.J. Solaro, R.J. Hajjar, Functional consequences of caspase activation in cardiac myocytes, Proc. Natl. Acad. Sci. USA 99 (2002) 6252–6256. [16] A.H. Lee, N.N. Iwakoshi, L.H. Glimcher, XBP-1 regulates a subset of endoplasmic reticulum resident chaperone genes in the unfolded protein response, Mol. Cell. Biol. 23 (2003) 7448–7459. [17] J.D. Blais, V. Filipenko, M. Bi, H.P. Harding, D. Ron, C. Koumenis, B.G. Wouters, J.C. Bell, Activating transcription factor 4 is translationally regulated by hypoxic stress, Mol. Cell. Biol. 24 (2004) 7469–7482. [18] K.D. McCullough, J.L. Martindale, L.O. Klotz, T.Y. Aw, N.J. Holbrook, Gadd153 sensitizes cells to endoplasmic reticulum stress by down-regulating Bcl2 and perturbing the cellular redox state, Mol. Cell. Biol. 21 (2001) 1249–1259. [19] D.J. DeGracia, H.L. Montie, Cerebral ischemia and the unfolded protein response, J. Neurochem. 91 (2004) 1–8. [20] K. Terai, Y. Hiramoto, M. Masaki, S. Sugiyama, T. Kuroda, M. Hori, I. Kawase, H. Hirota, AMP-activated protein kinase protects cardiomyocytes against hypoxic injury through attenuation of endoplasmic reticulum stress, Mol. Cell. Biol. 25 (2005) 9554–9575. [21] D.J. Thuerauf, M. Marcinko, N. Gude, M. Rubio, M.A. Sussman, C.C. Glembotski, Activation of the unfolded protein response in infarcted mouse heart and hypoxic cultured cardiac myocytes, Circ. Res. 99 (2006) 275–282. [22] M. Vitadello, D. Penzo, V. Petronilli, G. Michieli, S. Gomirato, R. Menabo, F. Di Lisa, L. Gorza, Overexpression of the stress protein Grp94 reduces cardiomyocyte necrosis due to calcium overload and simulated ischemia, Faseb J. 17 (2003) 923–925. [23] E. Szegezdi, S.E. Logue, A.M. Gorman, E. Samali, Mediators of ER stress-induced apoptosis, EMBO Rep. 7 (2006) 880–885.