Mitochondrial dysfunction and oxidative stress mediate the physiological impairment induced by the disruption of autophagy

Introduction

There is a growing interest in the role of macroautophagy, herein simply termed autophagy, in both normal homeostasis and in a variety of pathological conditions [1,2]. This interest has been sparked in part by observations suggesting that in lower organisms, autophagy is an important determinant of lifespan [3]. For instance, in C elegans, the life extending effects of mutation in the daf-2 pathway requires an intact autophagy program [4,5]. Similarly, in the worm, the increase in lifespan seen with dietary restriction is not evident when autophagy is impaired [6]. Consistent with these observations, genetic manipulations that can increase autophagy in Drosophila result in flies with an extended lifespan and an increase in overall stress resistance [7].

Less is known regarding the role of autophagy in mammalian systems. Prior to the establishment of the molecular and biochemical basis for autophagy, it was well appreciated that aging tissues were often characterized by the accumulation of damaged cellular components. In addition, consistent with a defect in autophagy, it was also evident that animal tissues exhibited an age-dependent decline in the turnover rates of long lived proteins [8]. These and other studies have suggested that autophagic flux declines with age and that the magnitude and timing of this decline is in general concordance with the age-dependent accumulation of damaged proteins and organelles seen within aging tissues. Recent genetic mouse models have strengthened this association. While complete knockouts of essential autophagy genes appear to be lethal in the neonatal stage, various conditional knockout models have been recently described. Among these recent results are the description of tissue specific deletions of essential autophagy genes in liver, brain, pancreas and heart [9-14]. While significant differences exist in these various model systems, most were characterized by the rapid appearance of various pathologies and physiological impairments that can also be observed as a consequence of normal aging.

Relatively little is known about the downstream mediators of the often profound physiological alterations observed following the disruption of autophagic flux. Most likely there is no single pathway across all tissues and organs, and even within a single tissue type, multiple mediators may exist. For instance, while the accumulation of misfolded and aggregated proteins normally cleared in part by autophagy are likely to play a prominent role in models of neurodegeneration, in other tissues, the role of accumulation of damaged protein aggregates is less clear. This tissue specificity was reinforced by recent observations demonstrating that deletion of p62, a ubiquitin and LC3/Atg8 binding protein, rescues the pathological changes observed in autophagy deficient liver but does not appear to alter the phenotypic changes seen following deletion of Atg7 in the brain [15].

One important function of autophagy is the turnover of organelles including mitochondria. While several reports have described the structural changes in mitochondrial appearance evident in electron micrographs taken of autophagy deficient mammalian tissues [9,12-14], the functional alterations, if any, of these mitochondria have not been reported. Here, using a variety of cellular and in vivo models of Atg7 deficiency, we have assessed the magnitude of mitochondrial dysfunction and the contribution of the corresponding oxidative stress in mediating the physiological impairment observed following disruption of autophagic flux.

Results

Discussion

Using a variety of cellular and in vivo models, we demonstrate that impairment of autophagy leads to the accumulation of damaged and dysfunctional mitochondria and a corresponding increase in intracellular ROS levels. In a model of autophagy deficiency occurring within the pancreatic β cell, we further demonstrate that the overall physiological impairment in glucose tolerance and insulin secretion can be significantly ameliorated by the simple addition of an antioxidant. While assessment of autophagic flux by p62 expression suggests that NAC treatment does not directly affect autophagy, the ultimate improvement of glucose tolerance and glucose-stimulated insulin secretion suggests that at least for this in vivo model, continuous oxidative stress plays an important pathophysiological role.

A very recent report has suggested that oxidative stress may also play a role in the alterations in innate immunity observed in autophagy deficient cells [21]. In particular, ROS appeared to mediate the increase in interferon secretion and resistance to viral infection seen in Atg5^-/- cells. The form of autophagy studied in these experiments is quite specialized and involves the delivery of viral nucleic acids to the endosome rather than the standard situation where cargo is delivered to the lysosome. Similarly, it should be noted that in this context, Atg5 deficient cells demonstrate increased interferon protection and increased protection from viral infection rather than the usual situation where autophagy disruption results in a loss of function phenotype. Interestingly, in this recent study, Atg5^-/- MEFs exhibited an approximate 2-fold increase in the number of mitochondrial genomes per cell, while for presently unclear reasons, we did not observe a similar increase in mitochondrial number in our Atg7^-/- MEFs (see Figure 2C).

Based on these recent in vitro observations regarding Atg5 deletion in cells and our in vivo observations regarding Atg7 deletion in the pancreas, it is tempting to speculate that a rise in ROS levels may be a universal downstream mediator of the positive or negative alterations seen in autophagy deficient tissues. Nonetheless, other evidence suggests that such a broad conclusion is unlikely to be always correct. For instance, while the accumulation of ubiquitin positive protein aggregates in inclusion bodies are prominent features in Atg5 and Atg7 deficient neurons and cardiomyocytes, these changes are not observed in T lymphocytes or dendritic cells that lack Atg5 expression [10, 11, 14, 22, 23]. This may reflect fundamental differences in the role of autophagy in rapidly dividing versus postmitotic cells. Similarly, even within a single tissue or organ, the effects of abrogating or inhibiting autophagy cannot always be easily predicted. For instance, deletion of Atg5 in adult mice leads to the spontaneous appearance of contractile dysfunction, while the same deletion performed early in cardiogenesis does not result in any basal myocardial phenotype [14]. Even less straightforward are observations that while cardiac specific deletion of Atg5 results in an animal that is less able to withstand myocardial pressure overload, heterozygous deletion of beclin, another essential autophagy gene, results in mice with the seemingly opposite cardiac phenotype [14, 24]. Thus, the downstream mediators and ultimate consequences of inhibiting autophagy may vary widely depending on the strategy used to disrupt autophagic flux and in what tissue or organ the disruption occurs.

Our data suggests that antioxidant treatment with NAC is particularly beneficial in preventing the glucose intolerance phenotype observed following deletion of Atg7 within β cells. Pancreatic secretion of insulin is well known to be sensitive to changes in the cellular redox state and overall mitochondrial function. Given our results on the metabolic profile of Atg7^-/- MEFs cultured in the presence of NAC, it is tempting to think that in the setting of Atg7 deletion, the in vivo use of antioxidants interrupts the ‘vicious cycle' of mitochondrial generated ROS inducing further mitochondrial damage. These observations suggest that antioxidant targeted therapy might be beneficial for at least a subset of the growing number of conditions where deficiency or impairment of autophagy is thought to contribute.

Methods

Mice and cells. Atg7^F/F mice have been previously described [9] and were crossed with either MCK-Cre mice (Jackson Laboratory) or RIP2-Cre mice (Jackson Laboratory) to generate mice with a conditional deletion of Atg7 within skeletal muscle or pancreatic β cells. For genotype analysis, tissues were digested overnightat 55 °C with Gitschier buffer (67 mM Tris-HCL,pH 8.8, 16.6 mM ammonium sulfate, 6.5 mM MgCl₂, 0.5% TritonX-100, 1% - mercaptoethanol, 100 μg/ml proteinase K). Samples were incubated at 95 °C for 5 min, shaken for 20 min with an Eppendorf thermomixer, and centrifuged at 16,100 x g for 2 min. Three primers 5'- TGGCTGCTACTTCTGCAA TGA TGT-3', 5'- GCAAGCTCACTAGGC TG CAGAACC-3', and 5'-GGTCCA GAGTCCGGTCTC GG-3' were used to detect the flox Atg7 allele in genomic DNA.

Cre-mediated recombination was assessed by PCR analysis of genomic DNA using primers 5'- GGTCTGGCAG TAAAAACTATC-3' and 5'-GTGA AACAGCATTGCTGTCACTT-3', and 5'-GGGTCC CA AAGGCCGCC-3' and 5'-GGATAGTTTTTACT GCCAGAC CGC-3' for Rip2-CRE and MCK-CRE, respectively [25, 26]. Similar to what has been recently described [12, 13], we observed no obvious phenotypic differences between Atg7^+/+, Atg7^+/+: MCK- Cre, Atg7^+/+:Rip2-Cre, Atg7^F/F, Atg7^F/+:MCK-Cre or Atg7^F/+:RIP-Cre mice, and so we predominantly present the latter two genotypes as representative controls in this study.

For chronic NAC treatment, randomized 4- week old mice were treated with 1 g/L of NAC in the drinking water for the indicated duration of the study. All animal experiments were conducted in accordance with the guidelines of the Animal Care and Use Committee, National Heart Lung and Blood Institute, NIH.

Mouse embryonic fibroblasts were prepared from E12-E14 day old embryos using standard methods. MEFs were prepared following a cross between Atg7^F/F mice with a transgenic mouse line carrying the Cre recombinase under the control of the adenoviral promoter EIIa that is known to be broadly expressed in the developing embryo [27]. PCR analyses using primers, 5'- GCTG CTACTTC TGCAATGATGT-3' and 5'GCAAGCTCACTAGG CTGCAGAACC-3', were used to detect the wild-type Atg7 gene and primers, 5'- GCTGCTAC TTCTGCAAT GATGT-3' and 5'- ATG GTAC ATGCTAAGCCTCTGGAC-3', were used to detect the deleted Atg7 gene. MEFs were cultured in growth medium consisting of Dulbecco's Modified Eagle's medium (DMEM; Invitrogen) supplementedwith 15% fetal bovine serum (FBS), 50 units/ml penicillin and 50 μg/ml streptomycin. For in vitro NAC treatment, freshly thawed primary MEFs at passage 2 were placed in growth medium supplemented with 500 μM NAC for 10 days with media being changed every other day.

Glucose tolerance test, insulin measurement and islet isolation. Overnight fasted male mice were given intraperitoneal injections of D-glucose (20% solution; 1 g/kg body weight) and blood glucose was determined using a one-touch Ascensia Elite glucometer (Fisher). Tail vein blood was collected at 0 and 15 min following glucose injection and plasma insulin levels were measured with a rat insulin radioimmunoassay kit according to the manufacturer's recommendations (Millipore).

For pancreatic intracellular insulin measurement, small sections of pancreas were digested with ethanol/acid buffer (25 ml absolute ethanol, 8.3 ml ddH₂O, and 0.5 ml concentrated HCl) overnight at 4°C as previously described [30]. Insulin was measured with a Rat/Mouse Insulin ELISA Kit Insulin (Crystal Chem Inc.). Tissue weight was used to normalize the intracellular insulin levels.

Pancreatic islets were isolated using a standard protocol [31]. Briefly, the pancreas was perfused with a solution of 0.5 mg/ml Collagenase Type V (Sigma) dissolved in Hanks Balanced Salt Solution (HBSS) containing calcium and magnesium (Mediatech Inc). The digestion was performed at 37°C for 15-20 min after which the collagenase was neutralized with HBSS supplemented with 1% FBS. The collagenase treated pancreas was then sequentially filtered through 1.5 mm and 0.8 mm metal mesh filters. Islets were subsequently enriched by centrifugation in Histopaque 1077 (Sigma) and hand-picked under direct light microscopic visualization.

Isolated mitochondrial and intact metabolic studies . Purified mitochondria were isolated from freshly harvested skeletal muscle using standard methods [28]. Briefly, isolated skeletal muscle were rapidly harvested, washed and minced in ice-cold Ionic Medium (100 mM surcrose, 10 mM EDTA, 100 mM Tris-HCL, 46 mM KCl, pH 7.4). The tissues were digested with 5% Proteinase Type XXIV (Sigma) in Ionic Medium for 5 min on ice and the protease was subsequently inactivated by the addition of Ionic Medium supplemented with 0.5% BSA. Samples were then homogenizedwith a glass-Teflon motorized homogenizer and the mitochondrial fraction isolated by differential centrifugation. Mitochondria were subsequently washed twice and then resuspended in Suspension Medium (230 mM mannitol, 70 mM sucrose, 0.02 mM EDTA, 20 mM Tris-HCl, 5 mM K₂HPO₄, pH 7.4) prior to functional assessment. Succinate (5 mM), rotenone (1 μM), Antimycin A (0.25 μM) and ADP (1 mM) were used to assay complex II dependent respiration.

Measurement of intact cellular respiration was performed using the Seahorse XF24 analyzer (Seahorse Bioscience Inc.). Primary MEFs were plated at a density of 40,000 cells/well on XF24 tissue culture plate. Purified pancreatic islets were isolated and cultured overnight in DMEM supplemented with 10% FBS, 50 units/ml penicillin and 50 μg/ml streptomycin. Two-three hours prior to respiration measurements, 50-100 islets were transferred to poly-L-lysine coated XF24 tissue culture plate. Prior to the respiration assay, primary MEFs or islets were rinsed and cultured in DMEM running medium (8.3g/L DMEM (Sigma), 200 mM GlutaMax-1 (Invitrogen), 100mM sodium pyruvate (Sigma), 25 mM D-glucose (Sigma), 63.3 mM NaCl (Sigma), and phenol red (Sigma), adjust pH to 7.4 with NaOH) according to manufacturer's protocol. Oxygen consumption was measured under basal conditions, in the presence of the mitochondrial inhibitors oligomycin (0.5 μM) or antimycin A (0.25 μM), or in the presence of the mitochondrial uncoupler FCCP (0.5 μM or 1 μM) to assess maximal oxidative capacity. Lactate measurements were made by determining the change in extracellular pH over time, as previously described [29]. Oxygen consumption and lactate measurements were normalized to cell number for primary MEFs and protein concentration for pancreatic islets.

Histology and Immunohistochemistry. Histological analysis was performed on 8-10 week old and 16-24 week old mice. Skeletal muscle or pancreatic tissue was isolated from Atg7^+/+:MCK-Cre, Atg7^F/+:MCK-Cre, and Atg7^F/F:MCK-Cre, or Atg7^+/+:RIP2-Cre, Atg7^F/+:RIP2-Cre, and Atg7^F/F:RIP2-Cre mice, respectively. Tissues were fixed in 10% formalin and paraffin embedded tissue sections were used for subsequent hematoxylin and eosin (H&E) staining. Immunohistochemistry analysis with anti-insulin (DAKO), anti-glucagon (Sigma), anti-somatostatin (DAKO), and antipolypeptide (Millipore) antibodies was performed using standard protocols. Cryostat sections were fixed with 4% paraformaldehyde in PBS prior to immunohistochemical analysis with either a nitrotyrosine antibody (Millipore) or p62 antibody (Progen Biotechnik). For quantification purposes, an islet was considered to stain positive for p62 or nitrotyrosine if greater than 5 cells within the islet exhibited positive staining. Rhodamine or Cy2 conjugated secondary antibodies (Jackson Immunological Research Laboratories) were used for visualization. Electron micrographs were performed on ultrathin sections of tissues that were fixed in 2 or 2.5% glutaraldehye plus 1% paraformaldehyde, post-fixed with 1% OsO₄, stained en bloc with 1% uranyl acetate and embedded in Embed 812 (Electron Microscopy Sciences) using standard methods. The sections were stained with lead citrate and uranyl acetate before viewing.

Western Blot Analysis. Primary MEFs or isolated pancreatic islets were lysed with NonidetP-40 lysis buffer (1.0% Nonidet P-40, 50 mM Tris-HCl pH 7.4,150 mM NaCl, 5 mM EDTA) supplemented with protease inhibitor tablet (Roche) and phosphatase inhibitors (1 mM Na₃VO₃, 1 mM β-glycerolphosphate, 10 mM NaF) for 15 min on ice prior to clarificationby centrifugation at 16,100 x g for 15 min at 4 °C. Skeletal muscle was suspended in homogenizing buffer (25 mM Tris. HCl pH 7.4, 150 mM NaCl, 5 mM EDTA, 1% Triton X-100, 0.1% SDS, 10% glycerol, 1 mM DTT, 0.5 % sodium deoxycholic acid) supplemented with protease and phosphatase inhibitors and homogenized using a tissue lyser (Qiagen). Protein concentration was determined with the Pierce BCA assay Kit (Thermo Fisher Scientific). Protein lysates were resolved on precast Tris-Glycine SDS gels (Invitrogen) and transferred onto nitrocellulose. Immunoblot analysis was performed with antibodies directed against Atg7 (ProSci Inc and Sigma), actin (Sigma), the Complex I component NDFS3 (NADH-ubiquinone oxidoreductase, MitoSciences), the Complex II component FP (flavoprotein subunit of complex II, MitoSciences), and the Complex III component Core1 (Ubiquinol-cytochrome-c reductase complex core protein 1, MitoSciences).

For detection of mitochondria complexes on native blue gels, purified mitochondria from skeletal muscle were resolved on Native PAGE Novex Bis-Tris gels according to the manufacturer's instructions (Invitrogen). Briefly, 1 mg of a purified mitochondria pellet was resuspended with 100 μl of 1x NativePAGE buffer and 12.5 μl of 10% maltoside (Sigma). The samples were incubated on ice for 30 min with frequent vortexing before pelleting by centrifugation at 16,100 x g for 10 min at 4°C. After centrifugation, 100 μl of supernatant was transferred to a new tube containing 6.3 μl of Coomassie blue additive and samples (approximately 50 μg) were then resolved by Native PAGE Novex Bis-Tris gel electrophoresis.

ROS measurements. Primary MEFs were cultured on Nunc Lab-Tek two chamber glass slides. As an internal control, each dual chamber slide contained one well of WT MEFs and one well of Atg7^-/- MEFs, or one well of Atg7^-/- MEFs and one well of Atg7^-/- MEFs that have been treated with 500 μM NAC for 4 days. Cells were incubated with HBSS containing 50 μM 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H₂DCFDA; Invitrogen) for 30 min at 37°C. Cells were then rinsed and mounted with mounting medium (Vector Labs) and visualized with a Leica SP1 confocal microscope as previously described [32]. Several random fields were taken of each genotype and mean fluorescence intensity was calculated in three or more separate experiments with data from over 250 cells using at least two independently isolated WT and Atg7^-/- MEF cell isolates. Attached cells were measured rather than MEF cell suspensions since we have previously observed dramatic alterations in ROS levels when attached cells are trypsinized [33].

Mitochondrial number. The ratio of mitochondrial DNA to nuclear DNA was determined as previously described [34]. In brief, quantitative PCR analysis using SYBR green (Applied Biosystems) was performed with 25 μg of isolated DNA (Qiagen). Mitochondrial DNA was assessed using primers, 5'-CTCTTAT CCACGC TTCCGTTACG-3' and 5'-GATGGTGGTACTCCCGCT GTA-3' for the mitochondrial-encoded ND1 gene. Nuclear DNA level was determined by amplifying the genomic H19 locus using primers 5'-GTACCCACCTGTCGTCC-3'and 5'-GTCCACGAGACCAATGA CTG-3'. The relative amount of mitochondrial to nuclear DNA was determined by normalized ND1 to H19 levels.