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Developing brain is highly susceptible to hypoxic-ischemic (HI) injury leading to severe neurological disabilities in surviving infants and children. Previously, we have reported induction of neuronal pentraxin 1 (NP1), a novel neuronal protein of long-pentraxin family, following HI neuronal injury. Here, we investigated how this specific signal is propagated to cause the HI neuronal death. We used wild-type (WT) and NP1 knockout (NP1-KO) mouse hippocampal cultures, modeled in vitro following exposure to oxygen glucose deprivation (OGD), and in vivo neonatal (P9-10) mouse model of HI brain injury. Our results show induction of NP1 in primary hippocampal neurons following OGD exposure (4–8h) and in the ipsilateral hippocampal CA1 and CA3 regions at 24–48 h post-HI compared to the contralateral side. We also found increased PTEN activity concurrent with OGD time-dependent (4–8h) dephosphorylation of Akt (Ser473) and GSK-3β (Ser9). OGD also caused a time-dependent decrease in the phosphorylation of Bad (Ser136), and Bax protein levels. Immunofluorescence staining and subcellular fractionation analyses revealed increased mitochondrial translocation of Bad and Bax proteins from cytoplasm following OGD (4h) and simultaneously increased release of Cyt C from mitochondria followed by activation of caspase-3. NP1 protein was immunoprecipitated with Bad and Bax proteins; OGD caused increased interactions of NP1 with Bad and Bax, thereby, facilitating their mitochondrial translocation and dissipation of mitochondrial membrane potential (ΔΨm). This NP1 induction preceded the increased mitochondrial release of cytochrome C (Cyt C) into the cytosol, activation of caspase-3 and OGD time-dependent cell death in WT primary hippocampal neurons. In contrast, in NP1-KO neurons there was no translocation of Bad and Bax from cytosol to the mitochondria, and no evidence of ΔΨm loss, increased Cyt C release and caspase-3 activation following OGD; which resulted in significantly reduced neuronal death. Our results indicate a regulatory role of NP1 in Bad/Bax-dependent mitochondrial release of Cyt C and caspase-3 activation. Together our findings demonstrate a novel mechanism by which NP1 regulates mitochondria-driven hippocampal cell death; suggesting NP1 as a potential therapeutic target against HI brain injury in neonates.
Brain injury due to fetal or neonatal asphyxia (Johnston, 1997; Lorenz et al., 1998; Northington et al., 2001a; Northington et al., 2001b;) or neonatal stroke is a leading cause of morbidity and mortality in surviving infants and children worldwide with a high societal cost (Balduini et al., 2004; Gresham et al., 1997; Shiber et al., 2010; Sudlow and Warlow, 1997; Volpe, 2001), and has an incidence as high as 1 in 4000 (Ferriero, 2004; Nelson, 2007). Yet, there is no promising therapy for neonatal brain injury at present; very few trials have documented limited clinical benefit (Blomgren and Hagberg, 2006; Kalenderian et al., 2009; Pezzini and Padovani, 2009; Uchino et al., 2001), in part, owing to incomplete understanding of the pathological secondary injury mechanisms that proceed from the primary insult (Blomgren et al., 2003; Blomgren and Hagberg, 2006; Pezzini and Padovani, 2009; Smaha, 2004). The hippocampus is often injured in neonatal stroke (Osredkar et al., 2010), and that disruption of oxygen and glucose supply resulting from occlusion of a blood vessel ultimately leads to neuronal losses and necrosis (Banasiak et al., 2000). Previously, we demonstrated the induction of a novel neuronal protein neuronal pentraxin 1 (NP1) in in vivo rat model of hypoxic-ischemic (HI) brain injury (Hossain et al., 2004), and that NP1 induction in HI cortical neuronal cell death is regulated via a glycogen synthase kinase (GSK)-3α/β-dependent mechanism (Russell et al., 2011). However, how this specific signal of NP1 induction is propagated to cause the HI neuronal death remains to be elucidated.
The cellular decision as to whether cells will undergo death or survival process is determined by the integration of multiple survival and death signals. The PI3-K/Akt signaling pathway plays a critical role in the survival of neuronal cells (Brunet et al., 2001). Whereas, PTEN (phosphatase and tensin homolog deleted on chromosome 10), which negatively regulates PI3-K/Akt pathway, plays a central role in cell migration, survival and apoptosis (Salmena et al., 2008) For example, PTEN deletion has been reported to prevent ischemic brain injury (Shi et al., 2011), and enhances regenerative ability of adult corticospinal neurons (Liu et al., 2010). In addition, there are many downstream effectors of PI3-K/Akt signaling pathway such as glycogen synthase kinase 3 (GSK-3) and Bcl-2/Bcl-xL-associated death protein (Bad) that play crucial roles in the process of cellular survival or deaths (Cross et al., 1995; Datta et al., 1997; del Peso et al., 1997; Hetman et al., 2000). In particular, GSK-3 is a critical activator of cell death in numerous models of neuronal apoptosis (Hetman et al., 2000; Li et al., 2000; Phiel et al., 2003), including HI (Russell et al., 2011). GSK-3β exerts some of its pro-apoptotic effects in neurons by regulating the mitochondrial localization of Bax, a key component of the intrinsic apoptotic cascade (Linseman et al., 2004).
Neuronal cell death signaling pathways in mitochondria has been demonstrated in the ischemic brain (Chan, 2004; Perez-Pinzon et al., 1999). This often occurs via an intrinsic cell death pathway triggered by the translocation of Bad and Bax, two proapoptotic Bcl-2 family members, to mitochondria (Armstrong and Jones, 2002; Yuan and Yankner, 2000; Zong et al., 2001), reduction in mitochondrial membrane potential (ΔΨm) (Deckwerth and Johnson, 1993), opening of the mitochondrial permeability transition pore (PTP) (Zarotti and Szabo, 1995), release of mitochondrial cytochrome c (Cyt C) (Narita et al., 1998; Wang et al., 1999; Zong et al., 2001) and formation of apoptosome with Apaf-1 and pro-caspase-9 leading to activation of the downstream executioner caspases, including caspase-3 (Li et al., 1997a, 1997b; Zou et al., 1999). These findings suggest potential mitochondrion-targeted strategies for preventing HI neuronal cell death. In this study, we have used primary hippocampal neuronal cultures, modeled in vitro, following oxygen glucose deprivation and in vivo neonatal mouse model of HI to investigate the role of NP1 in causing cell death in the hippocampus, and also the underlying mitochondrial mechanisms of the NP1-mediated death program. Furthermore, we used NP1−/− primary hippocampal neurons to specifically demonstrate the contribution of NP1 in mediating mitochondrial perturbation and cell death.
Primary hippocampal neuronal cultures were prepared from wild-type (WT), NP1 knockout (NP1-KO) mice (C57BL/6 background) and NP triple knockout (NP-TKO; all pentraxin family members-neuronal pentraxin 1, neuronal pentraxin 2 and neuronal pentraxin receptor are knocked down) mice (C57BL/6 and 129 mixed background) at postnatal day 1 or 2 (PND1 or 2) as described previously (Berbari et al., 2007) with modifications. NP1-KO and TKO mice were kindly provided by Dr. Paul Worley, Dept. of Neuroscience, School of medicine, Johns Hopkins University, Baltimore, MD, USA. After plating, at 2 days in vitro (DIV), half of the media were replaced with fresh medium additionally containing cytosine arabinofuranoside (AraC) to a final concentration of 5 μM to prevent non-neuronal proliferation. Thereafter, culture media was changed by half every 3 to 4 days. Cultures were used for experiments at 12–14 DIV. With this protocol, > 95% of cultured cells was MAP-2 (microtubule-associated protein-2) immunoreactive neurons (Chemicon, Temecula, CA) (Hossain et al., 2002).
To induce OGD, primary hippocampal neurons cultured at DIV 12–14 were placed in glucose-free Earl’s balanced salt solution (EBSS) and exposed to humidified 95% N2/5% CO2 using anaerobic modular incubator chambers (Billups-Rothenberg, Del Mar, CA) for various time periods (4–8h) at 37ºC (Russell et al., 2011). Samples were collected immediately after OGD without any reoxygenation. Control cultures were placed in EBSS containing glucose and exposed to humidified 95% air/5% CO2 at 37ºC.
Common carotid artery (CCA) occlusion combined with controlled hypoxia in postnatal day 9–10 mouse pups was performed as described previously (Hossain et al., 2004) to produce hypoxic-ischemic brain injury. Briefly, under deep inhalant anesthesia (isofluorane in an oxygen-nitrous oxide mixture delivered via a Drager vaporizer), a 3–4 mm incision was made in the neck, and the right common carotid artery was isolated, double ligated, and cut between the ligatures. After the surgical procedure, the animals recovered for 2 h in a temperature-controlled incubator, and then animals were placed in an enclosed, vented 500 ml Plexiglas chamber that was partially submerged in 36°C water. Hypoxia was induced by continuous flow of warm, humidified gas (10% oxygen, balanced nitrogen) for 1 h. A subset of control animals were mock-treated with a small incision in their neck without CCA occlusion and placed in the Plexiglas chamber at normal air temperature (sham control). All procedures involving animals were performed in accordance with the NIH guide for the Care and Use of Laboratory Animals and were approved by The Johns Hopkins University Animal Care and Use Committee.
Immediately after the indicated periods of exposure, cell cytotoxicity was determined by lactate dehydrogenase (LDH) assay. LDH released into the media after OGD exposure was measured using the Cytotoxicity Detection Kit (LDH) (Roche Diagnostics Corporation, Indianapolis, IN, USA) as described previously (Hossain et al., 2004). Percent cell death was determined using the formula: % cytotoxicity = OGD LDH release (OD490) / maximum LDH release (OD490) after correcting for baseline absorbance of LDH release at 490 nm.
Mouse pups were sacrificed at the indicated time-points after CCA ligation and hypoxia by perfusion fixation with ice-cold 4% paraformaldehyde in 1XPBS and then cryoprotected in 15% sucrose for overnight and then transferred to 30% sucrose solution for overnight followed by freezing with dry ice. Representative coronal brain sections (20 μm) from control and hypoxic–ischemic animals were immunostained as described previously (Hossain et al., 2004). Mouse monoclonal anti-NP1 (1:200) (BD Transduction Laboratories, Temecula, CA, USA) was used as the primary antibody, while donkey anti-mouse (1:200) Alexa fluor 568 is the secondary antibody (Invitrogen, Carlsbad, CA, USA). Immunofluorescence was visualized using an inverted fluorescence microscope (Olympus IX51fitted with DP2-DSW-V3.2 application software) at 10 X and ZEISS Axioimager M2 (AxioVision SE64 Rel.4.8.1 application software) at 100 X magnification.
Live immunostaining of primary hippocampal cultures (grown on coverslips) with NP1 was done as described previously with minor modifications (Hossain et al., 2004). Briefly, hippocampal neurons were live labeled with NP1 (1:100) for 45 min at 37° C. Neurons were then fixed with 3.7% formaldehyde, and permeabilized cells were stained with Alexa Fluor 568 (red)-conjugated anti-mouse secondary antibody (Invitrogen). For Cyt C-specific immunostaining, neurons were fixed with 3.7% formaldehyde followed by antigen retrieval using 5% urea in 0.1 M Tris/HCl at 95°C for 10 min. Thereafter, neurons were permeabilized and double labeled with Cyt C (1:100; BD Bioscience Pharmingen, San Diego, CA, USA) and VDAC (1:300; Abcam, Cambridge, MA, USA).
Total RNA was extracted using TRIzol reagent (Invitrogen) according to manufacturer’s protocol. cDNA was synthesized from 1 μg of purified total RNA using iScript™ cDNA Synthesis Kit (Bio-Rad laboratories, Richmond, CA, USA), following the manufacturer’s instructions. Quantitative real-time PCR was performed in triplicate by using iQ SYBR Green Supermix on CFX96™ Real-Time System (Bio-Rad). The mRNA level was normalized by housekeeping gene HPRT. The primers set used for NP1 were 5′-GCT GCG AGA GCC AGA GCA CC-3′ (sense) and 5′-TTG CCC GAG TTG GCT GAG CG-3′ (anti-sense), and for HPRT were 5′-CCT GGC GTC GTG ATT AGT GAT G-3′ (sense) and 5′-CAG AGG GCT ACA ATG TGA TGG C-3′ (anti-sense).
Western blotting was performed with standard methods as described previously (Hossain et al., 2011). Cultured neurons were exposed to OGD or mouse hippocampal tissues were collected at the indicated time-points after HI. Whole cell lysates or the proteins extracted from the tissue were subjected to Western blot analysis. Blots were probed with primary antibodies for NP1 (1:1000, BD Transduction Laboratories), PTEN (1:1000, Cell Signaling, Beverly, MA, USA), phospho-Akt (s473) (1:1000, Cell Signaling), phospho-GSK-3β (1:1000, Cell Signaling), GSK-3β (1:1000, Cell Signaling), phospho-Bad (s136) (1:1000, Cell Signaling), Bax (1:1000, Cell Signaling), Cyt C (1:1000; BD Biosciences Pharmingen), procaspase-3 and cleaved caspase-3 (1:1000; Cell Signaling), VDAC (1:1000, Abcam) and β-actin (1:5000, Sigma, St. Louis, MO, USA). Horseradish peroxidase (HRP)-conjugated secondary antibodies (GE Healthcare, Piscataway, NJ, USA) were used at 1:5000 dilutions for 1 h at RT. HRP reaction product was then visualized by enhanced chemiluminescence using an ECL Western blotting detection kit (Pierce, Rockford, IL, USA). Digitized images were quantified using NIH ImageJ software.
Subcellular fractionation was performed as described previously (Russell et al., 2008; Sharma et al., 2011). Briefly, cell pellets were homogenized in isotonic MB (mitochondrial buffer) consisting of 210 mM mannitol, 70 mM sucrose, 1 mM EDTA and 10 mM Hepes, pH 7.5, supplemented with protease and phosphatase inhibitor cocktails (Calbiochem, San Diego, CA, USA) followed by centrifugation at 500 g for 5 min. The pellet is the nuclear fraction. The post-nuclear supernatant was centrifuged at 10,000 g for 30 min at 4°C for the mitochondrial fraction. The pellet containing the mitochondrial fraction was suspended in 50 μl of MB containing 0.1% Triton X-100 to break up the mitochondria; the supernatant was used as the crude cytosolic fraction. To confirm the separation of cytosolic and mitochondrial fractions, blots were stripped and incubated with actin and VDAC, respectively, which also served as a loading control.
After exposure of the cells to the conditions as indicated in the figure legends, total cellular extracts from hippocampal cultures were prepared in immunoprecipitation buffer (10 mM sodium phosphate, 100 mM NaCl, 1% Triton X-100) containing 1X protease inhibitor cocktail set, and phosphatase inhibitor cocktail (Calbiochem) following the method as described previously (Hossain et al., 2004). One hundred micrograms of total proteins were subjected to immunoprecipitation by incubation overnight with Bad (1:100, Cell Signaling) and Bax (1:100, Cell Signaling) at 4 °C with constant shaking. Then, 20 μl of protein A/G-agarose conjugated beads (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) was added to each sample and incubated for 2–4 hr at 4°C. The beads were collected, washed, and boiled for 3 min in 50 μl of 1 X electrophoresis sample buffer followed by Western blot analysis.
Mitochondrial membrane potential, ΔΨm (delta psi), was measured using a JC-1 kit (Molecular Probes, Eugene, OR, USA) following manufacturer’s protocol. Briefly, cells were incubated with 1.5 μM of JC-1 (5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-benzimidazolylcarbocyanine iodide) for 15 min at 37°C. After incubation, cells were washed once with PBS, and images were obtained using an inverted fluorescent microscope (Olympus IX51) fitted with a DP-72 digital color camera.
Statistics were performed using GraphPad Prism software, Version 5.01. For one experimental and one control group, two-tailed Student’s t-test was used to determine if differences exist between means. Comparisons involving multiple groups were done by ANOVA, followed by Bonferroni/Dunn post-hoc test where appropriate. Significance level was assigned at P<0.05.
Hippocampal brain regions are highly susceptible to injury in the event of stroke (Osredkar et al., 2010). First, we asked whether NP1 is contributing to the neuronal death in the hippocampus. We used mature hippocampal cultures (12-14 DIV) to investigate OGD-induced neuronal death, because of the higher degree of sensitivity of mature hippocampal neurons (12-18 DIV) to hypoxic damage (Loreto and Balestrino, 1997). Increased induction of NP1 mRNA and protein levels were observed after exposure of primary hippocampal neurons to OGD. Quantification of NP1 mRNA and protein levels revealed an OGD time-dependent increase with more than two-fold induction observed at 4 h of exposure, which reached the maximum at 8 h (approximately 4 fold), compared to the normoxic control cultures (Fig. 1A & B). We used 4 h time point of OGD exposure in subsequent experiments as we observed clear evidence of injury following 4 h of OGD exposure in WT hippocampal neurons, similar to that we reported previously (Russell et al., 2011). Our immunocytochemical experiments showed enhanced NP1-specific immunostaining of the hippocampal neurons after OGD exposure (4 h) compared to the normoxic control cultures (Fig. 1C), further confirming the induction of NP1 protein in response to OGD exposure. Comparing the time-course of NP1 induction and the appearance of characteristic features of neuronal death, it appears that NP1 induction is an early event, and that the induction occurs before actual cell death. To further confirm NP1 induction in hippocampal neurons, next we immunostained brain sections from WT neonatal mouse HI brains. Fluorescence microscopy showed enhanced NP1-specific immunoreactivity in the ipsilateral hippocampus, which was localized in the pyramidal layer of CA3 and CA1 regions, but not in the dentate gyrus (DG) 48 h after HI onset (Fig. 1D, top). Further analysis of brain sections by fluorescence microscopy of the HI brain sections revealed intense NP1-specific fluorescence associated with individual neurons in the hippocampal regions (inset). Western analyses of the hippocampus tissue homogenate also showed a time-dependent increase in the NP1 protein levels after HI (24 & 48 h) compared to the sham controls (Fig. 1D, bottom); corroborating the induction of NP1 in the hippocampus after HI injury.
To correlate between NP1 induction and cell death in hippocampal cultures, we measured cell cytotoxicity by lactate dehydrogenase (LDH) activity assay following OGD exposure. Quantification of LDH release showed an OGD-induced increase in hippocampal cell death (Fig. 2A), which is consistent with our previous reports showing hypoxia-induced cell death in primary cortical neurons (Hossain et al., 2004; Russell et al., 2011). Cell cytotoxicity was evident by an increase in LDH release (~3-fold at 6h post-OGD) compared to the normoxic control cultures.
To directly demonstrate the link between NP1 induction and cell death, we used hippocampal neurons from both NP1-KO and NP-TKO mice, and exposed them to identical OGD conditions. Our cytotoxicity results showed very negligible change in the release of LDH in NP1-KO and NP-TKO neurons after exposure to OGD compared to the controls (Fig. 2A). Our results suggest that neurons from both NP1-KO and NP-TKO mice are significantly protected from OGD compared to the WT cultures, as evidenced by decreased LDH release. Also, light microscopic analysis showed characteristic changes of dying cells with disintegration of processes and cell bodies in WT hippocampal neurons following OGD (Fig. 2B). In contrast, both NP1-KO and NP-TKO neurons were significantly protected from OGD compared to the neurons from WT mice, which retained morphological features of a healthy neuron, such as larger cell bodies, phase brightness and intact processes.
Exposure of hippocampal neurons to OGD time-dependently decreased the phosphorylation of PTEN with a small increase in the PTEN protein levels, drastically dephosphorylated Akt (Supplemental Fig. 1), which is in agreement with our previous reports (Russell et al., 2011). We also found significant dephosphorylation of GSK-3β at 4 h, and complete dephosphorylation occurred beyond 4 h of OGD exposure (6–8 h) (Fig. 3A), suggesting the activation of GSK-3β under such conditions. Interestingly in NP-TKO (Fig. 3B) and NP1-KO (Fig. 3C) neurons, the pGSK-3β levels remained unchanged up to 4 h OGD compared to respective normoxia control, however, a delayed dephosphorylation (i.e. activation) of the kinase was observed at 6–8 h of exposure. It is worth noting that we found no significant difference in pGSK-3β protein levels between the WT and KO neurons under control normoxia condition. We also observed no change in β-actin protein levels, used as an internal control, across the all experimental time points examined. The blockade of GSK-3 activity by using GSK-3-specific inhibitor SB216763 revealed significant inhibition of NP1 expression (Fig. 3D) and neuronal death (Fig. 3E) compared to that observed in SB216763 untreated OGD group (Fig. 3D-E). Our results clearly show GSK-3β signaling is contributing to NP1-induced hippocampal cell death following OGD exposure.
It is known that Akt phosphorylates Bad and blocks the Bad-induced death of primary neurons by blocking its translocation the mitochondria (Datta et al., 1997). Also, GSK-3β exerts some of its pro-apoptotic effects in neurons by regulating the mitochondrial localization of Bax (Linseman et al., 2004). Here, we examined the Bax protein levels, and phosphorylation of Bad (at Ser136) under similar OGD conditions in WT and NP1-KO cultures. Western blot analysis using phospho-specific Bad primary antibody showed that phosphorylation of Bad (Ser136) drastically reduced at 4 h of OGD insult with essentially complete disappearance after 8 h in WT neurons (Fig. 4A, top); whereas, in NP1-KO cultures, no change in phospho-Bad protein levels was detected, at least, up to 4h of OGD exposure. However, a much delayed and less pronounced dephosphorylation of Bad was observed after 6h (Fig. 4B, top). We also found that the Bax protein levels were decreased in WT hippocampal neurons after OGD exposure (4 to 8h) (Fig. 4A, bottom panels). In contrast, there was no change in the Bax protein levels in NP1-KO cultures up to 4–6h of OGD exposure, although Bax was decreased after 6 h (Fig. 4B, bottom panels). However, the basal levels of phospho-Bad and Bax protein were found to be comparable between WT and NP1-KO groups. These results, therefore, indicate that a differential distribution of the proapoptotic Bad and Bax proteins might have ensued in WT and NP1-KO cultures between the cytosolic and mitochondrial compartments in the event of OGD exposure.
Dephosphorylation of Bad allows it to get translocated from the cytosol to the mitochondria where the protein initiates Cyt C release to promote apoptosis (Shou et al., 2004; Wang et al., 1999). Also, GSK-3β plays a role in the translocation of Bax to the mitochondria (Banasiak et al., 2000; Linseman et al., 2004; Narita et al., 1998). To examine OGD-induced mitochondrial translocation of Bad and Bax and release of Cyt C in WT and NP1-KO hippocampal neurons, we performed subcellular fractionations followed by Western blotting analyses (Russell et al., 2008; Sharma et al., 2011). In cytosolic fractions, significantly decreased levels of Bad (~ 70%; p<0.05) and Bax (~ 58%; p<0.01) were observed with concurrent increase in Cyt C protein level (~ 6-fold; p<0.001) in the OGD exposed WT neurons compared to the normoxia controls. On the other hand, in the mitochondrial fractions Bad (~ 4-fold; p<0.05) and Bax (~2-fold; p<0.01) levels were increased significantly with simultaneous decrease of Cyt C protein level (~ 75%; p<0.01) after OGD exposure compared to the normoxia controls. These results suggest increased mitochondrial translocation of Bad and Bax after OGD exposure (4 h) of WT hippocampal neurons with consequent mitochondrial release of Cyt C into the cytoplasm (Fig. 5A). In contrast, no significant changes of Bad, Bax and Cyt C levels were observed in both the cytoplasmic and mitochondrial fractions from NP1-KO hippocampal neurons following OGD exposure compared to that in control normoxia NP1-KO cultures. Thus, it appears that OGD exposure could not trigger the translocation of Bad and Bax from cytosol to mitochondria in NP1-KO neurons (Fig. 5B). Our results clearly demonstrate a role of NP1 in mitochondrial localization of Bad and Bax following OGD.
To directly demonstrate the association of NP1 with Bad and Bax proteins, and the role of GSK-3 in the interaction between NP1 and Bad/Bax, we performed complementary co-immunoprecipitation experiments with or without SB216763 (10 μM) pretreatment using Bad- and Bax- specific primary antibodies. Our results showed that NP1 was co-precipitated with both Bad (Fig. 5C) and Bax (Fig. 5D) proteins, and increased precipitation of NP1 was observed after OGD exposure, whereas pretreatment with GSK-3 inhibitor blocked the NP1/Bad and NP1/Bax associations. These results clearly show that NP1 forms protein complexes with Bad and Bax proteins which is GSK-3β-dependent, and that OGD enhances NP1 interactions with Bad and Bax to facilitate their mitochondrial localization.
Translocation of Bcl-2 pro-apoptotic proteins, including Bad and Bax, to the mitochondria leads to loss of mitochondrial membrane potential (ΔΨm) (Armstrong and Jones, 2002), which is an early episode in several instances of apoptosis (Green and Reed, 1998). Next, we investigated the change in ΔΨm in both WT and NP1-KO cultures at an earlier time-point of OGD (2h) by using a cationic membrane potential indicator JC-1. In healthy cells, the dye stains the mitochondria bright red. Whereas in apoptotic cells, the JC-1 dye cannot accumulate in the mitochondria and remains in the cytoplasm in a green fluorescent monomeric form (Cossarizza et al., 1993). Exposure of WT neurons to OGD showed drastic decrease in red fluorescence, which is a measure of ΔΨm loss, after 2h of OGD relative to unexposed control cultures (6A, top). The normalized JC-1 fluorescence (red-to-green ratio) showed significant decrease (~65% decrease; p<0.001) following OGD of the hippocampal neurons compared to the normoxic controls. Importantly in NP1-KO cultures, there was no significant change in the JC-1 fluorescence, i.e. red-to-green ratio, between the control and OGD groups (Fig. 6A bottom). Mitochondria in both groups exhibited yellow appearance, suggesting no significant loss of ΔΨm.
Next, we examined the subcellular localization of Cyt C in WT and NP1-KO neurons after OGD by immunofluorescence using Cyt C-specific antibody as described previously (Russell, 2008). WT neurons exposed to OGD exhibited pronounced Cyt C release, as evidenced by the presence of red fluorescence in the cytoplasm (Fig. 6B top right). In contrast, there was no evidence of Cyt C (red) fluorescence in the cytoplasm after exposure of NP1-KO neurons to OGD. Cyt C remained mostly in the mitochondria (Fig. 6B bottom right), which appears as yellow i.e. Cyt C-specific immunofluorescence being overlapped with mitochondrial marker VDAC. Mitochondrial release of Cyt C triggers procaspase-3 proteolysis/activation to mediate the caspase-mediated cell death (Budd et al., 2000). In agreement with this, we found that cleaved (activated) caspase-3 was significantly increased in WT cultures after OGD compared to the control cultures (Fig. 6C left). There was also a decrease in the procaspase-3 level following OGD in WT cultures compared to the normoxic controls. However, activation of caspase-3 has also been reported without change in procaspase-3 protein levels in neuronal death in similar neonatal rodent models of HI (Namura et al., 1998; Nakajima et al, 2000). In contrast, levels of cleaved caspase-3 were comparable between the control and OGD groups in NP1-KO neurons (Fig. 6C right). Our results, thus, clearly demonstrate the involvement of NP1 in mediating mitochondria-driven prodeath caspase pathways. In a separate study we also observed increased NP1 induction in neurons of HI brain sections, which also showed enhanced cleaved caspase-3 (+) immunoreactvity (unpublished data), further supporting the involvement of NP1 in mitochondria-mediated neuronal death.
The present study demonstrates induction of NP1 in primary hippocampal neuronal cultures following OGD and in the ipsilateral hippocampal CA1 and CA3 areas following HI in neonatal mice, and further delineates how induction of NP1 is involved in mediating neuronal death. Firstly, OGD caused activation of PTEN with subsequent dephosphorylation of Akt (i.e. inactivation), and prodeath kinase GSK-3β (i.e. activation) in WT hippocampal cultures. Secondly, OGD induced down-regulation of Bax and decreased phosphorylation of Bad that resulted in enhanced mitochondrial translocation of the Bax and Bad proteins. This was also evident by the dissipation of ΔΨm, increased mitochondrial release of Cyt C with subsequent activation of caspase -3 ultimately leading to neuronal death. Particularly interesting is our identification of the association of NP1 with the prodeath proteins Bad and Bax, which further facilitated Bax and Bad mitochondrial localization and OGD-induced neuronal death. Thirdly, in contrast to the WT cultures, the NP1−/− null hippocampal cultures showed no significant change in mitochondrial translocation of Bad and Bax, ΔΨm, and no apparent increase in mitochondrial release of Cyt C and caspase-3; resulting in significant protection of the NP−/− neurons against the OGD insult. This is the first demonstration of significant neuroprotection in NP1-KO hippocampal neuronal cultures under ischemic stroke condition, further supporting a crucial role of NP1 in neuronal death.
The brain is mostly susceptible to oxidative stress because it consumes a large quantity of oxygen, and hippocampus, in particular, is often vulnerable in hypoxia-ischemia (Osredkar et al., 2010). Previous report from our laboratory showed NP1 induction in hypoxic-ischemic brain injury (Hossain et al., 2004), and GSK-3α/β regulated the expression of the NP1 in cultured cortical neurons (Russell et al., 2011). In this study, we show the specificity of NP1 induction in OGD-induced neuronal death using a homogenous population of mature hippocampal neuronal cultures. We also found HI-time dependent induction of NP1 in the hippocampal (pyramidal layers of CA3 and CA1) regions; the same brain areas that developed infarction (i.e. injury) following HI. In fact, NP1 induction occurred prior (24 h and prior) to the development of the infarction (data not shown), indicating NP1 as an upstream component of the death cascade. This was evidenced by OGD time-dependent expression of NP1 mRNA and protein in cultured hippocampal neurons and the observed OGD-induced neuronal death. Live immunostaining of hippocampal neurons with NP1 antibody further consolidated our findings that NP1 induction occurs before the actual cell death. In addition, the intact morphological characteristics and significantly less cell death observed in NP1-KO and NP-TKO neurons following OGD specifically establish a role of NP1 inhibition in neuroprotection. Taken together our results provide strong evidence that NP1 induction plays a key role in OGD-induced hippocampal neuronal death.
Ample evidence suggests that apoptotic and anti-apoptotic signaling pathways are activated after cerebral HI, and that a shift in the balance between apoptotic and anti-apoptotic cellular factors determines cells fate. The PI3-K/Akt pathway regulates the survival response against pro-apoptotic stimuli (Cross et al., 1995; Hetman et al., 2000), whereas, PTEN, which acts as an antagonist of PI3-K (Oudit et al., 2004) has been reported to play a role in mediating mitochondria-dependent apoptosis (Zhu et al., 2006) and ischemic brain damage (Ning et al., 2004). We found OGD time-dependent decrease in the phosphorylation of PTEN and Akt, abolishing their protective effects. Whereas, decrease in the phosphorylation of the downstream target GSK-3β results in activation of the death kinase, which is consistent with previous reports (Enguita et al., 2005; Russell et al., 2011). Similarly, blockade of GSK-3 activity by pharmacological inhibitor decreased the OGD-induced expression of NP1 and its interaction with Bad and Bax, further supporting the GSK-3-dependent induction of NP1. Interestingly, the activation of GSK-3β appears to be suppressed in NP-KO cultures, at least during the early time-points of OGD. This is possibly because of the fact that knock down of NP1 gene is neuroprotective- a condition where prosurvival PI3-K/Akt pathways are active that may have contributed to the observed phosphorylation of GSK-3β (inactivation), shifting the balance toward the survival path.
The Bcl-2 family of proteins Bad and Bax play a crucial role in intracellular apoptotic signal transduction by regulating the permeability of mitochondrial membrane (Yuan and Yankner, 2000; Zong et al., 2001). Akt phosphorylates Bad (Datta et al., 1997) and phosphorylation of Bad promotes binding to 14-3-3 proteins to be sequestered in the cytosol (Datta et al., 2000; Yang et al., 1995; Zha et al., 1996). Thus, Akt deactivation and decreased phosphorylation of Bad following OGD resulted in enhanced mitochondrial localization of Bad. Similarly, Bax translocation to mitochondria triggers neuronal apoptosis (Zong et al., 2001). Bax is a direct target of GSK-3β that modulates Bax expression and function at the transcriptional and posttranslational levels to promote mitochondrial apoptosis (Linseman et al., 2004; Watcharasit et al., 2003). Our findings suggest that activated GSK-3β following OGD resulted in decreased Bax protein levels in the cytoplasm with concurrent increase in the mitochondrial fraction. Importantly, OGD exposure of NP1−/− null hippocampal neurons did not display any change in Bax protein and phospho-Bad levels. Also, knockdown of NP1 prevented the translocation of Bad and Bax from cytosol to mitochondria; indicating a specific involvement of NP1 in mitochondrial localization of Bad/Bax proteins following OGD.
The cell death signaling pathway in mitochondria has been demonstrated in the ischemic brain and that mitochondrial membrane depolarization and Cyt C release from inner membrane determine the final steps of apoptosis (Merry and Korsmeyer, 1997; Narita et al., 1998). Mitochondrial membrane potential is a very important marker for the function of mitochondria and a decrease in ΔΨm predicts cell injury (Green and Reed, 1998). To specifically demonstrate the role of NP1 in reduction of ΔΨm, we found that NP1-KO cultures resisted the OGD-induced dissipation of ΔΨm and there was significant inhibition of mitochondrial release of Cyt C. While we cannot rule out the contribution of other stressors on OGD-induced mitochondrial outer membrane permeabilization, our results clearly showed stabilization and neuroprotection from mitochondrial cell death pathway as a specific characteristic of NP1-KO hippocampal neurons. Most interestingly, our co-immunoprecipitation experiments revealed that NP1 formed protein complexes with Bad and Bax, and that OGD caused an increase in the interactions of both NP1-Bad and NP1-Bax. However, the increased association of NP1/Bad and NP1/Bax following OGD was reversed by pretreatment with SB216763, which further confirmed the GSK-3 regulation on NP1 expression and ensuing interactions with Bad and Bax proteins. Our current results are also in agreement with the report that shows NP1 facilitates the accumulation of Bax in mitochondria during apoptosis in cerebellar granule neurons in culture (Clayton et al., 2012). Therefore, we suggest that NP1 influences the translocation of these proapoptotic proteins to the mitochondria, thereby contributing to mitochondrial outer membrane permeabilization. We also observed cell death after OGD was associated with robust increase in activated caspase-3 in WT neurons but not in the NP1-KO cultures under identical condition, supporting a role for NP1 in mitochondria-mediated caspase-3-dependent cell death pathway as evident by LDH release in WT vs. NP1-KO neurons. It appears that absence of NP1 desensitizes neurons to the ischemic injury by preventing mitochondria-mediated activation of cell death pathway.
In summary, our results clearly demonstrate a mechanism (Fig. 7) by which NP1 influences mitochondria-mediated cell death by facilitating the translocation of proapoptotic Bad and Bax proteins to the mitochondria, decreasing ΔΨm and releasing Cyt C into the cytosol with subsequent activation of caspase-3. Our results present a novel mechanism of neuronal death involving PTEN/PI3K/Akt/GSK-3-dependent NP1 induction and its contribution to mitochondria-mediated hippocampal neuronal death following HI insult. Taken together, our findings suggest a novel neuroprotective strategy in neonates suffering from HI brain injury.
We gratefully acknowledge Dr. Paul Worley for providing the knockout mice. This work was supported by the National Institutes of Health grant RO1 NS046030 and the Cerebral Palsy International Research Foundation grant R-793-09.
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