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Mitochondria are organelles centrally important for bioenergetics as well as regulation of apoptotic death in eukaryotic cells. High mobility group box 1 (HMGB1), an evolutionarily conserved chromatin-associated protein which maintains nuclear homeostasis, is also a critical regulator of mitochondrial function and morphology. We show that heat shock protein beta-1 (HSPB1/ HSP27) is the downstream mediator of this effect. Disruption of the HSPB1 gene in embryonic fibroblasts with wild-type HMGB1 recapitulates the mitochondrial fragmentation, deficits in mitochondrial respiration, and adenosine triphosphate (ATP) synthesis observed with targeted deletion of HMGB1. Forced expression of HSPB1 reverses this phenotype in HMGB1 knockout cells. Mitochondrial effects mediated by HMGB1 regulation of HSPB1 expression, serves as a defense against mitochondrial abnormality, enabling clearance and autophagy in the setting of cellular stress. Our findings reveal a novel role for HMGB1 in autophagic surveillance with important effects on mitochondrial quality control.
Mitochondria play a primary role in bioenergetics in most eukaryotic cells (Chan, 2006a). Gene products derived from both the mitochondrial and nuclear genomes (Poyton and McEwen, 1996) are necessary for the assembly and function of respiratory-competent mitochondria. High mobility group box 1 (HMGB1) is an evolutionarily conserved protein, found in abundance within the nucleus. HMGB1 knockout mice die shortly after birth with severe hypoglycemia, pointing to the essential nature of the protein (Calogero et al., 1999). Within the nucleus, HMGB1 binds and bends DNA, facilitating numerous nuclear functions including maintenance of genome stability, transcription, replication, recombination and repair. HMGB1 plays a role in neurodegeneration, aging and cancer (Kang et al., 2010b; Lotze and Tracey, 2005; Qi et al., 2007; Tang et al., 2010c, d), which are often accompanied by mitochondrial abnormalities (Chan, 2006b; Poyton and McEwen, 1996). It can be rapidly mobilized to other sites in the cell, as well as actively released from cells. This study was initiated to determine whether HMGB1 affects mitochondrial function.
To address this possibility, we first assessed real-time oxidative phosphorylation (OXPHOS) by determining the oxygen consumption rate (OCR) and glycolysis by measuring the extracellular acidification rate (ECR) using an extracellular flux analyzer (Qian and Van Houten, 2010; Wu et al., 2007) in Hmgb1+/+ wild type and Hmgb1−/− knockout immortalized mouse embryonic fibroblasts (MEFs) (Fig. 1A). Analysis of OCR and ECR was performed in the presence of four individual inhibitors: 1) oligomycin (Olig) which inhibits mitochondrial adenosine triphosphate (ATP) synthesis 2) p-trifluoromethoxy carbonyl cyanide phenyl hydrazone (FCCP) which uncouples OXPHOS 3) 2-deoxyglucose (2DG) which inhibits hexokinase in the glycolytic pathway and 4) rotenone (Rote) which inhibits Complex I in the respiratory-chain. There was a significant decrease in basal OXPHOS and glycolysis with HMGB1 deficiency (“+/+” versus “−/−“, p < 0.001) (Fig. 1B), as well as substantial decreases in reserve capacity (FCCP stimulated OCR). To determine whether alterations in mitochondrial function are directly due to the loss of HMGB1, we transfected a plasmid expressing HMGB1 and a downstream green fluorescent protein (GFP) cDNA (to assess transfection efficiency) into Hmgb1−/− MEFs. Expression of HMGB1 restored glycolysis, mitochondrial respiration, and ATP production (Fig. 1A–C). To further rule out the possibility that decreased mitochondrial respiration in the Hmgb1−/− MEFs resulted from chronic metabolic or other unrelated genetic changes that may have occurred during clonal selection, we examined OCR and ECR after reducing HMGB1 expression by short hairpin RNA (shRNA). Decreased HMGB1 expression in parental MEFs or NIH/3T3 mouse fibroblasts or Panc02 mouse or Panc2.03 human pancreatic cancer cells or human HCT116 colon cancer cells resulted in decreased OXPHOS, phenocopying HMGB1-deficient cells (Fig. 2 and Fig. 3). Furthermore, steady-state levels of ATP are decreased by 20–40% in Hmgb1−/− or HMGB1 knockdown cells (Fig. 1C, ,2C2C and and3C),3C), which is accompanied by diminished cell growth and proliferation (Fig. 1D). To determine whether the deficiency in mitochondrial respiration in Hmgb1−/− cells is due to a decrease in mitochondrial mass, we quantified levels of representative proteins from complex I-IV (listed in Fig. S1A) in Hmgb1+/+ and Hmgb1−/− MEFs by immunoblotting. All of the measured proteins were expressed at similar levels in both cell types. In contrast, we observed an increase in mitochondrial mass in Hmgb1−/− MEFs by MitoTracker Green staining (Fig. S1B) or coomassie brilliant blue staining (Fig. S1C).
Mitochondria are dynamic organelles, and their morphologic changes are tightly associated with their function (Chan, 2006a, b). To assess the effect of HMGB1 on mitochondrial morphology, we visualized mitochondrial architecture using immunohistochemistry with antibodies against Complex I subunit GRIM-19. Morphological analysis revealed that the mitochondria, which exhibit long and tubular morphology in Hmgb1+/+ cells, became dramatically shorter and rounder in Hmgb1−/− cells (Fig. 1E). Moreover, expression of Hmgb1 cDNA in Hmgb1−/− MEFs restored mitochondrial morphology. HMGB1 deficient cells exhibit mitochondrial dysfunction with respiratory deficits, a fragmented morphology and loss in the mitochondrial membrane potential (Fig. 1E). In addition, a balance between fission and fusion is critical to maintain proper mitochondrial morphology and function. The key mediators of mitochondrial fusion and fission include dynamin-related protein 1 (DRP1), fission 1 (FIS1), optic atrophy 1 (OPA1), mitofusin 1 (MEN1) and mitofilin (Chan, 2006a, b). All of these mediators are present in similar amounts in whole cell lysate of both Hmgb1+/+ and Hmgb1−/− cells (Fig. S1A). However, the short and long forms (S and L) of OPA1 (Suen et al., 2008) are decreased in isolated mitochondria of Hmgb1−/− cells (Fig. S1A).
Heat shock proteins (HSPs) and mitophagy have also been proposed to regulate mitochondrial dynamics and quality (Kim et al., 2007; Tatsuta and Langer, 2008; Youle and Narendra, 2011). The principle HSPs that have chaperone activity belong to several families: HSP100 (e.g. HSP110), HSP90, HSP70, HSP60, HSP40 and the small heat-shock proteins [e.g. heat shock protein beta-1 (HSPB1), αB-crystallin, HSP27] (Craig et al., 1994; Sorger, 1991). In erythroid cells, NIX (also known as BNIP3L), a member of the Bcl-2 gene family, is required for mitophagy (Schweers et al., 2007). The protein expression of HSPB1 (also known as HSP25 in mice, HSP27 in humans), but not other HSPs or NIX, is significantly inhibited in Hmgb1−/− cells (Fig. 4A). Like HMGB1 (Calogero et al., 1999), HSPB1 is expressed in various cell types and tissues, and the failure to obtain knockout mice suggests that HSPB1 is essential for growth and development (Arrigo, 2007; Garrido, 2002). Expression of HMGB1 restored HSPB1 mRNA and protein expression that were reduced in Hmgb1−/− cells (Fig. 4B). Furthermore, we confirmed a similar HMGB1-dependent regulation of HSPB1 protein expression following heat shock, which was not observed in expression of HSP70 (Fig. 4C). In both Hmgb1+/+ and Hmgb1−/− cells, there are similar levels of heat shock factor 1 (HSF1), the major heat shock transcription factor that regulates stress inducible synthesis of HSPs (Sorger, 1991), and heat shock element (HSE) activity (Fig. 4A and 4D). These results reveal that HSPB1 levels are regulated by HMGB1.
To determine whether loss of HSPB1 can mimic HMGB1-deficiency, we reduced HSPB1 expression levels by about 80% using two individual shRNA in Hmgb1+/+ MEFs when compared with control shRNA. Decreased HSPB1 expression resulted in reduced OCR, ECR (Fig. 3B and and4E),4E), and ATP production (Fig. 3C and and4F),4F), and increased mitochondrial fragmentation (Fig. 4G), which was remarkably similar to the phenotype of HMGB1-deficient cells (Fig. 1). To also determine whether HMGB1 regulates mitochondrial quality directly through HSPB1, we expressed mouse HSPB1 cDNA in Hmgb1−/− MEFs. Expression of HSPB1 in Hmgb1−/− cells at the physiological level observed in the Hmgb1+/+ cells (Fig. 5A) corrected the deficit in mitochondrial respiration (Fig. 5B), ATP production (Fig. 5C) and mitochondrial fragmentation (Fig. 5D) observed in HMGB1-deficient cells. This suggests that HSPB1 expression can rescue the mitochondrial phenotype observed in HMGB1-deficient cells.
The increased number of dysfunctional mitochondria in cells deficient in HMGB1 thus appears to be due to the loss of one of HMGB1’s transcriptional targets, HSPB1. Autophagy (which includes macro-, micro-, and chaperone-mediated autophagy) is an important biological mechanism for the elimination of damaged/obsolete macromolecules and organelles (Kroemer et al., 2010; Yang and Klionsky, 2010). Mitochondrial autophagy (mitophagy) is responsible for elimination of dysfunctional and impaired mitochondria (Kim et al., 2007; Youle and Narendra, 2011). Mitophagy involves at least a three-step process: 1] autophagosome formation to engulf the dysfunctional mitochondria, 2] lysosome-autophagosome fusion and 3] degradation of dysfunctional mitochondria by autolysosomes. Blocking the formation of autophagosomes by 3-methyladenine (3-MA, class III phosphoinositide 3-kinase inhibitor) (Stroikin et al., 2004), blocking the fusion of autophagosomes with lysosomes by bafilomycin A1 (vacuolar ATPase inhibitor) (Bjorkoy et al., 2005), or blocking lysosomal degradation by pepstatin and E64D (lysosomal protease inhibitors) (Mizushima and Yoshimori, 2007) increased rotenone-mediated ATP depletion and mitochondrial fragmentation in MEF cells (Fig. S2A–B). In addition, autophagy inhibitors (e.g. 3-MA and bafilomycin A1) decreased OCR and ECR (Fig. S2C). Our findings support the notion that autophagy/mitophagy is involved in sustaining mitochondrial respiration and morphology following cellular stress and mitochondrial injury.
Inhibition of the mitochondrial respiratory chain complex I by rotenone induces autophagy or apoptosis in a variety of cells by mediating ATP depletion, ROS generation, or causing mitochondrial depolarization (Chen et al., 2007; Clark et al., 2006; Moon et al., 2005) (Fig. 6A). We focused on the early stages of mitochondrial injury induced by rotenone since autophagy preceded mitochondria-mediated apoptosis in MEFs (Fig. S2D–E). There was a significant increase in mitochondrial fragmentation following rotenone treatment in HMGB1 and HSPB1 deficient cells (Fig. S2F–G). The microtubule-associated protein light chain-3 (LC3), a mammalian homolog of Atg8, is thought to recruit mitochondria into autophagosomes. Loss of HMGB1 or HSPB1 inhibited amino acid- and ATP depletion-induced autophagy as assessed by the expression of LC3-II (Fig. 6C), LC3 punctae (Fig. S3A and B) and lysosomal-associated membrane protein 2 (LAMP2)/LC3 colocalization (Fig. S3D). Loss of HMGB1 and HSPB1 also decrease the mitochondrial membrane potential and increase apoptosis (Fig. S3B). Moreover, autophagy inhibitors (e.g. 3-MA) increased rotenone-induced apoptosis (Fig. S3C), suggesting that autophagy is a cell survival mechanism under stress. To determine whether HMGB1 and HSPB1 specifically regulate mitophagy, we assessed the process of mitophagy in the absence of HMGB1 or HSPB1. Indeed, loss of HMGB1 and HSPB1 decreased colocalization of mitochondria not only with LC3, but also with LAMP2 (Fig.6A and S4), suggesting that the loss of HMGB1 or HSPB1 results in defective autophagy with secondary consequences on mitochondria.
Ultrastructural EM analysis reveals that wild type cells exhibit normal mitochondrial morphology, with fragments of mitochondria present in some autophagosomes or lysosomes following mitochondrial injury induced by rotenone (Fig. 6B). As expected, polyubiquitin-binding protein, sequestosome-1 (p62) bodies and expression (Bjorkoy et al., 2005) are increased in HMGB1 and HSPB1 deficient cells when rotenone-induced autophagy is impaired (Fig.6C and Fig. S5A–B). In addition, the degradation of exogenously introduced p62 following starvation is also impaired in HMGB1 and HSPB1 deficient cells (Fig. S5C).
A well characterized function of HSPB1 is to interact with the actin cytoskeleton (Lavoie et al., 1993a; Lavoie et al., 1993b), a dynamic structure that maintains cell shape, enables cellular locomotion and plays important roles in the transport and morphology of intracellular vesicles and organelles including mitochondria (Boldogh and Pon, 2006). Stress fibers, which are bundles of easily visualized actin filaments, appear in response to rotenone-induced mitochondrial injury. The colocalization of mitochondria and actin is also increased following rotenone treatment in wild type cells (Fig. 6A), indicating that such interactions are an important regulatory mechanism following mitochondrial injury. Similar to HMGB1 deficiency, treatment of wild type cells with the cytoskeleton inhibitor, cytochalasin D (“CytD”) decreases stress fibers and colocalization of mitochondria/actin, mitochondria/autophagosome (e.g. LC3) and mitochondria/autolysosomes (e.g. LAMP2) with mitochondrial injury (Fig. 6A). Moreover, CytD increases rotenone-induced apoptosis (Fig. 6A). In contrast, expression of HSPB1 in HMGB1 deficient cells undergoing mitochondrial injury restored the interaction between mitochondria and the cytoskeleton, and increased the dynamic stages of autophagy (Fig. S6).
In response to stress, phosphorylation and increased expression of HSPB1 modulates actin polymerization and reorganization (Lavoie et al., 1993b; Rousseau et al., 1997). We next explored whether alterations in the phosphorylation of HSPB1 (Ser15 and Ser86) influences autophagy. ATP depletion induced by rotenone treatment increased the expression of pHSPB1 (both Ser15 and Ser86) in Hmgb1+/+ cells but not Hmgb1−/− cells (Fig. 6C). Previous studies have demonstrated that these phosphorylated HSPB1 mutations decreased F-actin stabilization and had no influence on the chaperone activity of HSPB1 (Benndorf et al., 1994; Rogalla et al., 1999). Expression of phosphorylation-deficient mutants of HSPB1 (S15A and S86A), however, did not restore the deficit in autophagy, ATP production or morphology, whereas wild type HSPB1 cDNA did (Fig. 6C and S6). These findings suggest that both Ser15 and Ser86 are necessary for HSPB1-mediated autophagy.
Upon mitochondrial membrane depolarization, PTEN induced putative kinase 1 (Pink1), a kinase of the outer mitochondrial membrane, induces Parkin translocation to stressed mitochondria (Geisler et al., 2010; Narendra et al., 2008). Subsequently, Parkin mediates the formation of ubiquitin chains on voltage-dependent anion channel 1 (VDAC1). This leads to the recruitment of the autophagy receptor p62, which in turn, by binding to LC3, directs damaged mitochondria to the autophagosome (Geisler et al., 2010). There has, however, been a contradictory report showing that VDAC1 may not be required for Parkin-mediated mitophagy (Narendra et al., 2010). We found that loss of HMGB1 or HSPB1 inhibited rotenone-induced Parkin translocation, VDAC1 ubiquitylation (Fig. 6D, 6E and S7A), but not Parkin expression (Fig.1A, ,2A,2A, ,3A3A and and5A).5A). Moreover, knockdown of Pink1 or Parkin abolish the HSPB1-restored ATP production and reduced mitochondrial fragmentation in Hmgb1−/− cells (Fig. 6F). These findings suggest that both Pink1 and Parkin are required for HSPB1/HMGB1-mediated mitophagy.
Although mitophagy is a well established mechanism necessary for elimination of dysfunctional mitochondria and regulation of mitochondrial quality in yeast or mammalian cells by mediators such as NIX (Sandoval et al., 2008), autophagy-related gene 32 (Okamoto et al., 2009), OPA1 (Twig et al., 2008) and DRP1 (Twig et al., 2008), it is unclear whether or how mitophagy triggered by dysfunctional mitochondria is regulated by nuclear mediators. Our findings indicate that the nuclear protein HMGB1 modulates mitochondrial respiration and morphology by helping to sustain autophagy in mitochondria maintenance through regulation of HSPB1 gene expression (Fig 7). Phosphorylation of HSPB1 is necessary to regulate the actin cytoskeleton, which affects the dynamics of autophagy in response to mitochondrial injury. Since loss of either HMGB1 or HSPB1 results in autophagy deficiency, increase in the number of impaired mitochondria results in an apparent increase in mitochondrial fragmentation. Moreover, loss of HMGB1 or HSPB1 results in decreased aerobic respiration and subsequent ATP production. Indeed, deletion of HMGB1 (Calogero et al., 1999) or HSPB1 (Garrido, 2002) in mice results in early postembryonic lethality. HMGB1 is localized within the nucleus, but in several cell types and environments, HMGB1 can be detected within the cytoplasm and readily within the extracellular space (Lotze and Tracey, 2005). Our recent studies demonstrated that both extracellular (exogenous) and endogenous HMGB1 promote autophagy and cell survival following application of cancer chemotherapeutic agents and nutritional depletion (Liu et al., 2010; Tang et al., 2010a; Tang et al., 2010b). Cytoplasmic HMGB1 is a Beclin 1-binding protein active in autophagy (Kang et al., 2010a; Tang et al., 2010b). Interestingly, rotenone-mediated mitochondrial injury increased the expression of HMGB1 and HSPB1 in mitochondria (Fig. S7B), although this change and its significance in the regulation of autophagy/mitophagy is unknown.
The cellular cytoskeleton plays multiple roles in regulation of both selective and non-selective autophagy (Monastyrska et al., 2009). For example, histone deacetylase-6 (HDAC6) promotes autophagy and mitophagy by assembling an F-actin network that stimulates autophagosome-lysosome fusion and substrate degradation (Lee et al., 2010). Interestingly, mitochondria degradation in yeast involves both a selective and nonselective process of autophagy (Kissova et al., 2007). Indeed, some specific regulators of mitophagy also play roles in nonselective autophagy. For example, Parkin-mediated K63-linked polyubiquitination is required for autophagic clearance of misfolded proteins (Olzmann and Chin, 2008). Nix is important for reactive oxygen species (ROS)-mediated autophagy induction (Ding et al., 2010). More recently, the mammalian Atg1 homolog UNC-51-like kinase 1 (ULK1), has been shown to be required for both autophagy and mitophagy following starvation (Egan et al., 2011). Our findings suggest that HMGB1 and HSPB1 coordinately regulate both macroautophagy and mitophagy following starvation and mitochondrial injury. However, it is still unclear as to the actual mechanisms by which selective and non-selective autophagy are switched. Delineation of discrete autophagic pathways will improve our understanding of both mitophagy and autophagy, but also enable development therapeutic strategies to target diseases such as Parkinson’s and cancer.
The antibodies to heat shock proteins 90 (HSP90), HSP70, HSP60, HSP40, and heat shock factor 1 (HSF1), green fluorescent protein (GFP), B-cell leukemia/lymphoma 2 (Bcl-2) and His-tag were obtained from Cell Signaling Technology (Danvers, MA, USA). The antibody to HSP110 was obtained from Stressgen (Ann Arbor, MI, USA). The antibodies to actin, Nix and HSPB1 (HSP25/27) were obtained from Sigma (St. Louis, MO, USA). The antibodies to pHSPB1 (Ser15), pHSPB1 (Ser86), PTEN induced putative kinase 1 (Pink1), Parkin, voltage-dependent anion channel 1 (VDAC1) and ubiquitin were obtained from Abcam (Cambridge, MA, USA). The antibodies to high-mobility group box 1 (HMGB1), microtubule-associated protein light chain 3 (LC3)-I/II, dynamin-related protein 1 (DRP1), mitofusin 1 (MEN1), optic atrophy 1 (OPA1) and lysosomal-associated membrane protein 2 (LAMP2) were obtained from Novus (Littleton, CO, USA). The antibodies to αB-crystallin, p62 and fission 1 (FIS1) were from Santa Cruz Technology (Santa Cruz, CA, USA). The antibodies to the Complex I subunits NDUFA9 and GRIM-19, Complex II subunit 70 kDa Fp, Complex III subunit Core 2, Complex IV subunit 1, Complex V subunit α, cytchrome C, ATP synthase β, and mitofilin were obtained from Mitosciences (Eugene, OR, USA). Hoechst33342 and phalloidin were obtained from Invitrogen (San Diego, California). All other chemical reagents were obtained from Sigma (St. Louis, MO, USA).
pEGFPN1-HMGB1 plasmid was a kind gift from Dr. George Hoppe (Cole Eye Institute, USA) (Hoppe et al., 2006). The pUNO1-HMGB1 plasmid was obtained from InvivoGen. The pcDNA4-HisMaxC-HSPB1, pcDNA4-HisMaxC-S15A and pcDNA4-HisMaxC-S86A were kind gifts from Dr. Yoon-Jin Lee (Korea Institute of Radiological and Medical Sciences)(Lee et al., 2005). pEGFPC1-p62 plasmid was kind gift from Dr. Eileen White (The Cancer Institute of New Jersey, USA) (Mathew et al., 2009). Mouse or human HSPB1 shRNA, HMGB1 shRNA, Pink1 shRNA and Parkin shRNA were obtained from Sigma. Nuclear and cytoplasmic extraction kit, and Mitochondria and Membrane Isolation kit were obtained from Pierce (Rockford, IL, USA).
NIH/3T3 fibroblasts, Panc02, Panc2.03 and HCT116 cells were derived from the American Type Culture Collection (Manassas, VA, USA) or National Institutes of Health (Bethesda, MD, USA). Hmgb1+/+ and Hmgb1−/− immortalized mouse embryonic fibroblasts were a kind gift from Dr. Marco E. Bianchi (San Raffaele Institute, Italy) (Calogero et al., 1999). All cell lines were cultured in medium supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM glutamine and penicillin-streptomycin mix (Invitrogen, final concentration 50µg/ml each) in a humidified incubator with 5% CO2 and 95% air.
Expression vector transfection or shRNA transfection using FuGENE® HD Transfection Reagent (Roche Applied Science, Stockholm, Sweden) or Lipofectamine 2000 reagent (Life Technologies) were performed according to the manufacturer’s instructions. siRNA was transfected into cells using X-tremeGENE siRNA reagent (Roche Applied Science) according to the manufacturer’s instructions. In general, the transfection efficiency is > 80%.
Proteins in cell lysates were resolved on 4–12% Criterion XT Bis-Tris gels (Bio-Rad, USA) and transferred to a nitrocellulose membrane as previously described (Tang et al., 2007a; Tang et al., 2007b). After blocking, the membrane was incubated for overnight at 4°C with various primary antibodies. After incubation with peroxidase-conjugated secondary antibodies for 1 h at 25°C, signals were visualized by enhanced chemiluminescence detection (Pierce) according to the manufacturer's instruction. The relative band intensities were quantified using the Gel-pro Analyzer® software (Media Cybernetics, Bethesda, MD, USA).
Cells were cultured on glass cover-slips and fixed in 3% formaldehyde for 30 min at room temperature prior to detergent extraction with 0.1% Triton X-100 for 10 min at 25°C. Cover slips were saturated with 2% bovine serum albumin (BSA) in phosphate buffered saline (PBS) for 1 h at room temperature and processed for immunofluorescence with primary antibodies followed by Alexa Fluor or Cy3-conjugated secondary antibodies respectively. Nuclear morphology was analyzed with the fluorescent dye Hoechst 33342. Actin was stained with phalloidin. Mitochondria were stained using anti-Complex I subunit GRIM-19 or anti-ATP synthase β. Between all incubation steps, cells were washed three times for 3 min with 0.5% BSA in PBS. Images were taken with an Olympus Fluoview 1000 confocal microscope (Olympus Corp, Tokyo, Japan). Fluorescent intensities were measured by Image-Pro Plus platform (Media Cybernetics, Bethesda, MD, USA). The mitochondria of untreated cells were filamentous and showed a threadlike tubular structure, while mitochondria in stressed cells were fragmented and appeared shortened and punctate. Quantification of mitochondrial fragmentation was performed as described previously (Brooks et al., 2009).
Cells were lysed at 4°C in ice-cold modified radioimmunoprecipitation (RIPA) lysis buffer (Millipore, Billerica, MA, USA), and cell lysates were cleared by centrifugation (12000 g, 10 min). Concentrations of proteins in the supernatant were determined by bicinchoninic acid (BCA) assay. Prior to immunoprecipitation, samples containing equal amount of proteins were pre-cleared with Protein A or protein G agarose/sepharose (Millipore) (4°C, 3 h) and subsequently incubated with various irrelevant IgG or specific antibodies (2–5 µg/ml) in the presence of protein A or G agarose/sepharose beads for 2 h or overnight at 4°C with gentle shaking. Following incubation, agarose/sepharose beads were washed extensively with PBS, and proteins were eluted by boiling in 2× sodium dodecyl sulfate (SDS) sample buffer before SDS-PAGE electrophoresis. To detect ubiquitylation of VDAC1, cell extracts were immunoprecipitated with anti-VDAC1 antibody, and analyzed with an anti-ubiquitin antibody by western blotting.
The ATP content in whole cell extracts was determined with a luminescent ATP detection kit (ATPlite; PerkinElmer Life Sciences, Boston, MA) according to the manufacturer's instructions. The luminescent intensity was measured using a microplate reader (Synergy 2, BioTek instruments, Winooski, VT). In parallel, the cell number in whole cell samples were counted by trypan blue exclusion assay. The results were expressed as relative ATP level compared with controls after normalizing for cell number.
The percentage of cells with LC3 punctae was determined by quantifying the number of positively staining cells from 50–100 randomly chosen cells from 15 to 20 random fields. Autophagic flux assays were performed by western blotting for LC3-I/II and p62, by imaging for the percentage of cells with LC3 and p62 punctae and colocalization of LC3/mitochondria, actin/mitochondria and LAMP2/mitochondria. In brief, images were collected using a laser-scanning confocal microscope (Fluoview FV-1000; Olympus) using a 60× Plan Apo/1.45 oil immersion objective at 25°C and Fluoview software (FV10-ASW 1.6; Olympus). Images were subsequently analyzed for fluorescent intensity levels and colocalization of various stains by Image-Pro Plus 5.1 software (Media Cybernetics).
Transmission electron microscopic (TEM) assessment of autophagosomes and autolysosomes was performed as previously described (Tang et al., 2010b). In brief, cells were fixed with 2% paraformaldehyde and 2% glutaraldehyde in 0.1 mol/L phosphate buffer (pH 7.4), followed by incubation for 6 hours in 1% OsO4. After dehydration with graded alcohols, the samples were embedded in epoxy resin (Epon, Momentive Specialty Chemicals, Houston). Following embedding, thin sections (70 nm) were cut using a microtome (Leica Ultracut R), mounted on copper grids and post-stained with 2% uranyl acetate and 1% lead citrate, dried, and analyzed using a transmission electron microscope at 25°C (JEOL 100CX, Peabody, MA, USA). Thick sections were cut (300 nm) and stained with 1% toluidine blue. Images were acquired digitally from a randomly selected pool of 10 to 15 fields under each condition.
Apoptosis in cells was assessed using the Annexin V-FITC Apoptosis Detection Kit (BD Pharmingen, San Jose, CA, USA) by flow cytometric analysis (Kang et al., 2010c; Tang et al., 2010a) or a TUNEL (Terminal deoxynucleotidyl transferase dUTP nick end labeling) kit from Roche Applied Science. Mitochondrial membrane potential depolarization was measured by flow cytometry using a fluorescent cationic dye, 1,1’3,3’- tetraethylbenzamidazolocarbocyanin iodide (JC-1, Molecular Probes, San Diego, CA, USA). JC-1 dye exhibits potential-dependent accumulation in mitochondria, indicated by a fluorescence emission shift from green (~529 nm) to red (~590 nm). Consequently, mitochondrial depolarization is assessed by a decrease in the red/green fluorescence intensity ratio.
Cells were transiently transfected in a 12-well plate with an HSE luciferase reporter plasmid and control empty plasmid using Lipofectamine 2000 reagent (Invitrogen) according to the manufacturer’s instructions. After 24–48h, the cells were exposed to various treatments. Luciferase activity was determined using an assay system including the reporter lysis buffer obtained from Promega (Madison, WI, USA) as described previously (Wang et al., 2002). The results were expressed as relative HSE activity after normalizing for the control empty plasmid.
Cellular OXPHOS and glycolysis were monitored using the Seahorse Bioscience Extracellular Flux Analyzer (XF24, Seahorse Bioscience Inc., North Billerica, MA, USA) by measuring the Oxygen Consumption Rate (OCR, indicative of respiration) and Extracellular Acidification Rate (ECR, indicative of glycolysis) in real time as previously described (Qian and Van Houten, 2010; Wu et al., 2007). Briefly, 30,000–50,000 cells were seeded in 24-well plates designed for XF24 in 150 µl of appropriate growth media and incubated overnight. Prior to measurements, cells were washed with unbuffered media once, then immersed in 675 µl unbuffered media and incubated in the absence of CO2 for 1 hour. The OCR and ECR were then measured in a typical 8-minute cycle of mix (2–4 minutes), dwell (2 minutes) and measure (2–4 minutes) as recommended by Seahorse Bioscience. The basal levels of OCR and ECR were recorded first, followed by the OCR and ECR levels following injection of compounds that inhibit the respiratory mitochondrial electron transport chain [METC], ATP synthesis, or glycolysis.
Subcellular fractionation of cells was carried out with a mitochondria, membrane, or nuclear isolation kit obtained from Pierce (Rockford, IL, USA) according to the manufacturer’s instructions.
Data are expressed as means ± SD of two or three independent experiments performed in triplicate. One-way ANOVA was used for comparison among the different groups. When the ANOVA was significant, post hoc testing of differences between groups was performed using an LSD test. A p-value < 0.05 was considered significant.
This project was funded by a grant from the NIH (M. T. L), start-up funding from the University of Pittsburgh (D.T.) and PA C.U.R.E. (B. V. H) and. We thank Dr. George Hoppe (Cole Eye Institute, USA) for pEGFPN1-HMGB1; Dr. Yoon-Jin Lee (Korea Institute of Radiological and Medical Sciences) for pcDNA4-HisMaxC-HSPB1, pcDNA4-HisMaxC-S15A and pcDNA4-HisMaxC-S86A; Dr. Marco E. Bianchi (San Raffaele Institute, Italy) for Hmgb1+/+ and Hmgb1−/− MEFs; Dr. Eileen White (The Cancer Institute of New Jersey, USA) for pEGFPC1-p62; Drs. Patricia Loughran, Donna B Stolz, and Simon Watkins (Center for Biologic Imaging, UPMC) for EM sample preparation and image analysis.
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