Free Radical Biology and Medicine 75 (2014) 241–251
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Original Contribution
Defects in mitochondrial DNA replication and oxidative damage in muscle of mtDNA mutator mice Jill E. Kolesar a,1, Adeel Safdar b,c,d,1, Arkan Abadi c, Lauren G. MacNeil c, Justin D. Crane c,d, Mark A. Tarnopolsky c,d,n, Brett A. Kaufman a,nn a
Department of Animal Biology, University of Pennsylvania School of Veterinary Medicine, Philadelphia, PA 19104, USA Department of Kinesiology, McMaster University, Hamilton, ON L8N 3Z5, Canada c Department of Pediatrics, McMaster University, Hamilton, ON L8N 3Z5, Canada d Department of Medicine, McMaster University, Hamilton, ON L8N 3Z5, Canada b
art ic l e i nf o
a b s t r a c t
Article history: Received 30 April 2014 Received in revised form 24 July 2014 Accepted 28 July 2014 Available online 12 August 2014
A causal role for mitochondrial dysfunction in mammalian aging is supported by recent studies of the mtDNA mutator mouse (“PolG” mouse), which harbors a defect in the proofreading-exonuclease activity of mitochondrial DNA polymerase gamma. These mice exhibit accelerated aging phenotypes characteristic of human aging, including systemic mitochondrial dysfunction, exercise intolerance, alopecia and graying of hair, curvature of the spine, and premature mortality. While mitochondrial dysfunction has been shown to cause increased oxidative stress in many systems, several groups have suggested that PolG mutator mice show no markers of oxidative damage. These mice have been presented as proof that mitochondrial dysfunction is sufficient to accelerate aging without oxidative stress. In this study, by normalizing to mitochondrial content in enriched fractions we detected increased oxidative modification of protein and DNA in PolG skeletal muscle mitochondria. We separately developed novel methods that allow simultaneous direct measurement of mtDNA replication defects and oxidative damage. Using this approach, we find evidence that suggests PolG muscle mtDNA is indeed oxidatively damaged. We also observed a significant decrease in antioxidants and expression of mitochondrial biogenesis pathway components and DNA repair enzymes in these mice, indicating an association of maladaptive gene expression with the phenotypes observed in PolG mice. Together, these findings demonstrate the presence of oxidative damage associated with the premature aging-like phenotypes induced by mitochondrial dysfunction. & 2014 Elsevier Inc. All rights reserved.
Introduction Mitochondria are host to numerous biosynthetic, bioenergetic, and signaling processes that ultimately couple cellular metabolism to homeostasic regulatory mechanisms. As the dominant site of reducing equivalent (electrons) and oxygen consumption, the
Abbreviations: cat, catenanes; CCC, covalently closed circles; ddC, dideoxycytosine; ETC, electron transport chain; MEFs, mouse embryonic fibroblasts; mtDNA, mitochondrial DNA; NT, nitrotyrosine; PC, protein carbonyls; NC, nicked circles; rc, relaxed circles; ROS, reactive oxygen species; WT, wild-type; 1D-IMAGE, onedimensional intact mtDNA agarose gel electrophoresis; 2D-IMAGE, twodimensional intact mtDNA agarose gel electrophoresis; 4-HNE, 4-hydroxy-2nonenal; 8-OHdG, 8-oxodeoxyguanisine. n Corresponding author at: Departments of Pediatrics and Medicine, HSC-2H26, McMaster University Medical Center, 1200 Main St. W., Hamilton, ON L8N 3Z5, Canada. fax: 1–905-577–8380. nn Corresponding author. fax: 1 215 573 5188. E-mail addresses: [email protected] (M.A. Tarnopolsky), [email protected] (B.A. Kaufman). 1 Co-first authors. http://dx.doi.org/10.1016/j.freeradbiomed.2014.07.038 0891-5849/& 2014 Elsevier Inc. All rights reserved.
mitochondrial electron transport chain (ETC) is a major source of reactive oxygen species (ROS). Alterations in ETC function, in particular in complexes (Co) I and III, can cause redox imbalances and oxidative damage to nearby proteins, lipids, and nucleic acids. Mitochondrial DNA (mtDNA), which is present in thousands of copies per cell, encodes components of ETC complexes I, III, and IV. mtDNA is intrinsically more sensitive than nuclear DNA to oxidative damage and is more slowly repaired than nuclear DNA [1]. Thus, mtDNA may be damaged even under conditions where nuclear DNA is not. In the context of mitochondrial dysfunction, this effect is exacerbated by the close proximity of mtDNA to the source of mitochondrial ROS. mtDNA therefore serves as a sensitive marker for the detection of overall mitochondrial oxidative stress A multitude of diseases and conditions involve both mitochondrial dysfunction and elevated oxidative stress (ex. [2–4]). For example, myocardial infarction has been shown to induce mitochondrial dysfunction and oxidative stress resulting in mtDNA damage [5]. Age-related neuropathologies such as Parkinson's
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disease and Alzheimer's disease show a strong association between mitochondrial dysfunction and oxidative stress (reviewed in [2]). Evidence also exists for elevated oxidative stress in cases where the primary cause of mitochondrial dysfunction is known, such as in myoclonic epilepsy and ragged-red fibers (MERFF) syndrome caused by the mtDNA mutation A8344G [6]. More recently, mice engineered to harbor a mt-ND6 mutation mimicking a prevalent Leber’s hereditary optic neuropathy (LHON) mutation show clear complex I defects and elevated ROS generation [7]. It is therefore generally accepted that mitochondrial dysfunction, in particular involving Co I and III, can cause elevated ROS that potentially alter physiology through damage or homeostatic signaling mechanisms. Mitochondrial respiratory function and mtDNA sequence stability both decline during normal aging [4,8]. A causal role for somatic (and possibly stem cell) mtDNA mutations in mammalian aging is supported by recent studies of the mtDNA mutator mouse (“PolG” mouse), which harbors a defect in the proofreading-exonuclease activity of mitochondrial DNA polymerase gamma [9,10]. These mice exhibit accelerated aging (reduced life span) and phenotypes characteristic of human aging including muscle atrophy, cardiomyopathy, anemia, skin and fur aging (thin dermis and gray fur), and systemic mitochondrial dysfunction [9–12]. Somewhat surprisingly, several groups have examined markers of oxidative stress in these mice and found no change [10,12], while others have provided support for oxidative stress in the PolG mice [13,14]. Resolving this discrepancy is crucial to solidifying the causal links among mitochondrial dysfunction, ROS, and aging, which have been the subject of debate [15]. In an effort to definitively determine whether PolG mice are under mitochondrial oxidative stress, we examined multiple aspects of mtDNA integrity as a primary readout of oxidative damage occurring in PolG mitochondria. mtDNA damage is frequently detected by a long-template PCR assay [16] that relies on base modification and breaks in the template to serve as an indication of oxidative stress. To our knowledge, this PCR technique has not been reported in PolG mice, likely because the replication pausing known to occur in these mice [17] could produce truncated templates that may lead to misinterpretation of reduced amplification efficiency as mtDNA damage. In the following study, we developed novel methods that detect both replication pausing and oxidative damage directly in intact mtDNA. The application of these methods to the PolG mutator mouse muscle, along with standard methods, strongly indicates that the mitochondrial compartment is specifically under oxidative stress in these mice. We show that mitochondrial biogenesis and
antioxidant defense pathways are highly dysregulated, similar to what is observed in normal human aging, suggesting that responses to mitochondrial dysfunction may be exacerbated by maladaptive gene expression.
Results and discussion Reduced tissue mass and mitochondrial dysmorphy are associated with mitochondrial dysfunction in PolG mice PolG mice suffer from numerous muscle pathologies [9–12]. For example, we and others have observed a significant reduction in skeletal muscle mass (Fig. 1A) and lower Complex IV activity (Fig. 1B) in quadriceps muscle (mainly fast-twitch) from independent cohorts of PolG mice compared to WT [9–12]. A similar reduction in respiratory chain activity was found in saponinpermeabilized soleus (mainly slow-twitch) muscle (Supplementary Fig. 1; [18]). The mitochondria in PolG quadriceps also show aberrant ultrastructure by electron microscopy, consistent with the quantitative decrease in mitochondrial number and increase in mitochondrial volume we previously described [19]; Supplementary Fig. 2). These results confirm that the mice used in this study have the same phenotypes as those previously reported, which is critical for interpreting further tests of the existence of mitochondrial oxidative stress in PolG mice. MtDNA structural defects, lower mtDNA levels, and DNA breaks in PolG mutator mice Multiple alterations to mtDNA molecules have been reported in PolG mouse tissues, including control region multimers (CRM) [20], elevated levels of linear mtDNA content [9], and replication pausing [17]. Because these alterations would manifest across entire genomes, we surveyed PolG mtDNA for these and other alterations using the high-resolution approach of two-dimensional intact mtDNA agarose gel electrophoresis (2D-IMAGE) [21]. This approach allows the separation and direct detection of up to 24 mtDNA structural variants, in particular structures with singlestranded DNA content thought to be DNA replication intermediates. 2D-IMAGE and associated mtDNA analyses of PolG muscle revealed the presence of a discrete mtDNA variant (labeled var.), whose migration characteristics (between 1n and 2n nicked circles, NC) are consistent with replication intermediates detected in cells during recovery from mtDNA depletion [21] (Fig. 2). The
Fig. 1. Decreased muscle weight and mitochondrial activity in quadriceps of PolG mutator mouse. (A) Mass determination of quadriceps (quad.), gastrocnemius (gastroc.), soleus, and extensor digitorum longus (EDL) at 8 months of age (n¼ 10 WT, n¼ 10 PolG (mt/mt)). (B) Protein-content normalized cytochrome c oxidase (COX) activity in quadriceps expressed as percentage of WT (n¼ 3 WT, n¼ 3 PolG (mt/mt)). All values are mean þ/– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05; nn o 0.01, nnn o 0.001.
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Fig. 2. Mitochondrial DNA alterations and damage in PolG mutator mice quadriceps by 2D-IMAGE. (A) Typical 2D-IMAGE profile of DNA preparations probed for mtDNA in WT and PolG quadriceps. Major structures are indicated, including a known variant (indicated var.) and shorter form (indicated #) in PolG samples. (B) Relative abundance of major mtDNA structural and topological isomers in PolG and WT quadriceps (n¼ 3 WT, n¼3 PolG (mt/mt)). (C) Relative abundance of circular and linear mtDNA, and the circular to linear (C:L) mtDNA ratio in PolG and WT quadriceps (n¼ 3 WT, n¼ 3 PolG (mt/mt)). (D). Linearization of mtDNA by XhoI digestion shows full-length mtDNA, mtDNA structure previously assigned “deletion” in the PolG mouse, and control region multimers (CRM). (E) Relative mtDNA content of PolG and WT quadriceps by qPCR and Southern blot after SacI digestion (n ¼3 WT, n ¼3 PolG (mt/mt)). All values are mean þ /– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05; nn o0.01, nnn o 0.001. Abbreviations: catenanes, cat; nicked circles, nc; linear, lin; covalently closed circles, ccc; variant, var.; linear degradation products, lin. deg; and n.a., not applicable.
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2D-IMAGE characteristics of the mtDNA variant are identical to the replication intermediates described in PolG mouse liver (Fig. 2A; [17]), and this variant migrates as an apparent “deletion” by standard Southern blot protocols after XhoI digestion (Fig. 2D; [9]). In addition, we observed an accumulation of a heterogeneous DNA population that extends upward from the 1n linear on the linear diagonal toward larger genome lengths (Fig. 2A, labeled #). The diminished abundance of catenanes and CCC molecules, previously shown to be produced during replication [21], coupled with increased replication intermediates is consistent with the reported deficiency in replication processivity of the PolG mutator protein [17]. Southern blotting also revealed hybridizing species near the well, which have been described for CRM tandem repeats (Fig. 2D; [20]). 2D-IMAGE successfully detects all reported aspects of altered replication described for the PolG mutator mice, but also provides new information regarding the status of mtDNA integrity.
Quantification of the prominent mtDNA structural and topological conformers in PolG muscle using 2D-IMAGE revealed a significant shift in the relative abundance of circular and linear mtDNA. Comparison of summed circular DNA (1n NC, catenanes, and CCC) abundance relative to linear DNA generated a circular to linear (C:L) ratio of 10:1 for WT muscle and 2:1 for PolG muscle (Fig. 2C). In contrast to other reports [9,22], we found that mtDNA levels were lower in skeletal muscle of the PolG mouse both by qPCR for total mtDNA and by Southern blot for full-length mtDNA detection after SacI digestion (Fig. 2E, [23]). There is strong evidence for linear mtDNA instability causing rapid genome depletion [24], suggesting that the mechanisms that underlie the elevated levels of doublestrand breaks in mtDNA (manifested as linear molecules) could contribute to the observed genome depletion. We hypothesized that either replicative pausing or oxidative damage could contribute to the increased linear mtDNA content in
Fig. 3. Quantitation of circular and linear mtDNA by 1D-IMAGE analysis. (A,C,E) Quantitation of major topoisomers by 1D-IMAGE and phosphorimage analysis. (B,D,F) Determination of relative abundance of circular mtDNA by ratio to linear mtDNA (circular:linear; C:L). Samples analyzed: (A,B) 24-h treatment of MEFs with ddC; (C,D) C2C12 myotubes exposed to 200 or 400 mM H2O2 for 15 min; (E,F) Quadriceps from WT vs PolG (mt/mt) mice. P values in panels C and D (n ¼3 each) were determined by one-way ANOVA with Bonferroni post hoc analysis against the untreated sample indicated (n o 0.05; nn o 0.01, nnn o 0.001). P values in panels E and F (n¼ 3 each) were determined by Student’s t test (n o 0.05; nn o0.01, nnn o0.001).
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the PolG muscle. To test the effects of replication arrest on levels of double strand breaks, we exposed cultured MEFs to dideoxycytosine (ddC), a deoxynucleotide chain terminator that preferentially affects mtDNA replication [25]. Single dimension intact mtDNA agarose gel electrophoresis (1D-IMAGE) showed that ddC exposure causes dose-dependent linearization, culminating in the depletion of circular mtDNA (Fig. 3A and B). To test whether oxidation caused linearization of mtDNA, we exposed C2C12 myotubes to acute oxidative stress for 1D-IMAGE analysis (Fig. 3C and D). Short hydrogen peroxide (H2O2) exposures were selected to prevent complication of data interpretation due to adaptive gene expression. The formation of double-strand breaks (linears) occurred in a dose-dependent manner in myotubes in short exposures (15 min). Longer exposures (30 min) showed a plateau in linearization (Supplementary Fig. 3), possibly due to limited availability of iron for Fenton chemistry to produce hydroxyl radicals [26]. The effects of oxidative damage on the formation of linear mtDNA were confirmed by 2D-IMAGE of myotubes (Supplementary Fig. 4), MEFs (Supplementary Fig. 5), and neonatal cardiomyocytes (Supplementary Fig. 6). Cells were adherent in all cases, with MEFs and neonatal cardiomyocytes being subconfluent. These results suggest that acute oxidant exposure linearizes circular mtDNA in a dose-responsive manner. To distinguish between replicative pausing and oxidative damage to mtDNA in the PolG mouse, we devised a method to describe the relative abundance of single-strand breaks in mtDNA (Fig. 4). We have previously shown that S1 nuclease will cut across from single-strand breaks in circular double-stranded mtDNA genomes to produce linear
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molecules detectable in 1D and 2D-IMAGE profiles [21]. First, we tested whether hydrogen peroxide and ddC treatments resulted in differential S1 resistance. As expected, circular DNA (2n catenanes and relaxed circles) in H2O2 samples showed increased S1 sensitivity and formed more linear mtDNA relative to the untreated control (Fig. 4A and B). In contrast, the ddC-treated samples showed a different diagnostic pattern upon S1 treatment, with linear molecules decreasing during replication pausing (Fig. 4C and D). These results suggested that S1 nuclease treatment may be used as a tool to detect the presence of oxidative damage in mtDNA. When comparing the relative S1 sensitivity of WT and PolG mouse muscle, we observed only a slight increase in full-length 1n linear molecules upon S1 digestion of WT mtDNA (Fig. 4E), suggesting limited oxidative stress. In contrast, circular mtDNA in PolG muscle was more sensitive to S1 nuclease treatment and produced more linear mtDNA degradation products (Fig. 4F), consistent with oxidative damage. We also noted that the expressions of base excision repair enzymes Nth1, UNG1, and OGG1 were significantly downregulated in the PolG samples (Fig. 4G). Taken together, the single-strand breaks in PolG mtDNA structures are consistent with oxidative damage and may persist due to insufficient expression of DNA repair enzymes.
Mitochondrial dysfunction in PolG mice is associated with increased oxidative damage to mitochondrial protein and mtDNA Initial examination by others of PolG tissue homogenates for the contribution of mitochondrial dysfunction to oxidative stress failed to
Fig. 4. PolG mtDNAs demonstrate increased sensitivity to S1 nuclease digestion, consistent with oxidative damage. Identical amounts of total DNA were treated in parallel with endonuclease S1 and resolved by 1D-IMAGE (A,C,E) and the fraction of DNA resistant to digestion relative to control was determined (B,D,F). (A) Representative 1DIMAGE of H2O2-treated myotubes reveals increased S1 sensitivity of catenanes and relaxed circles, producing linear mtDNA. (B) Graphical representation of S1 resistance of H2O2-treated myotubes (n¼ 3). (C) 1D-IMAGE of MEFs cultured overnight in the presence of varying concentrations of the chain terminator dideoxycytosine (ddC) reveals a distinctive S1 pattern. (D) Graphical representation of ddC-exposed samples for S1 resistance of topoisomers normalized to untreated control. (E) Representative 1D-IMAGE of PolG (mt/mt) muscle reveals elevated S1 sensitivity of all major topoisomers relative to WT. (F) Graphical representation of PolG (mt/mt) S1 resistance for topoisomers normalized to WT (n¼ 2 WT, n¼2 PolG (mt/mt)). (G) Relative gene expression of base excision repair enzymes Nth1, UNG1, and OGG1 (n¼ 3 per genotype). All values are mean þ /– SEM; P values determined by one-way ANOVA with Bonferroni post hoc analysis (B) or unpaired Student’s t test (F,G); P values: n o 0.05; nn o 0.01, nnn o 0.001.
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muscle by Western blot analysis for VDAC, ANT1, and CS (Supplementary Fig. 8). By comparing oxidative modifications normalized to mitochondrial content in these fractions, we observed a significant increase in PC, 4-HNE, and nitrotyrosine (NT) in PolG mitochondrial fractions vs WT (Fig. 5A–C). To further support the occurrence of oxidative modifications in mitochondria, we examined levels of 8-OHdG DNA modification within circular mtDNA (Fig. 5E) and found that the PolG muscle mtDNA contained significantly elevated levels of 8-OHdG relative to controls. The amount of mtDNA analyzed was confirmed by qPCR (data not shown). These data clearly show the presence of increased oxidative protein and base modification in muscle mitochondria of this strain of PolG mice, and support our observations of single-strand breaks in mtDNA originating from oxidation. Additional studies will be necessary to determine whether strain background, notably variation in the NNT gene, contributes to the manifestation of oxidative damage [28,29]. Differential effects on nonenzymatic and enzymatic redox capacity in PolG tissues
Fig. 5. Mitochondrial markers of oxidative stress. (A) Relative levels of protein carbonylation (PC) in mitochondrial preparations of PolG (mt/mt) and WT quadriceps normalized to VDAC abundance (n¼ 6 WT, n ¼6 PolG (mt/mt)). (B) Relative levels of 4-HNE lipid modification of proteins from quadriceps mitochondrial preparations of PolG (mt/mt) and WT muscle normalized to VDAC abundance (n¼6 WT, n ¼5 PolG (mt/mt)). (C) Tyrosine nitrosylation in mitochondrial preparations of PolG (mt/mt) and WT muscle normalized to VDAC abundance (n ¼6 WT, n¼ 4 PolG (mt/mt)). (D) 8-OHdG DNA base modification in circular DNA isolated from PolG (mt/mt) and WT quadriceps muscle normalized to mt-Co2 as determined by qPCR (n¼ 6 WT, n¼ 4 PolG (mt/mt)). All values are mean þ/– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05; nn o0.01, nnn o0.001.
reveal elevated ROS or increased oxidative damage despite significant accumulation of mtDNA point mutations [10,27]. In agreement with aspects of these studies, we also found no difference in markers of oxidative damage in whole muscle homogenates of PolG and WT mice as measured by 4-hydroxy-2-nonenal (4-HNE; marker of lipid oxidation) and protein carbonyls (PC; marker of protein oxidation) (Supplementary Fig. 7). However, we did find a consistently lower protein content for several well-known protein markers of total mitochondrial abundance VDAC (outer mitochondrial membrane), CS (mitochondrial matrix), and ANT1 (inner mitochondrial membrane) (data not shown). The lower mitochondrial protein levels are consistent with the decreased mitochondrial numbers observed by EM in PolG muscle [19]. We interpret these findings as evidence for limited total cellular oxidative damage, which agrees with previous reports [9,10,12] in the context of reduced mitochondrial content. To determine whether ROS-mediated damage occurs specifically in muscle mitochondria of PolG mice, we analyzed mitochondriaenriched fractions for several oxidative damage markers (including those applied to whole tissue lysates in Supplementary Fig. 7). As expected from the whole tissue lysates and electron microscopy, we found that the relative recovery of mitochondria was lower in PolG
To establish whether conditions in the PolG mouse were conducive to oxidative stress, we examined total nonenzymatic redox capacity and indicators of enzymatic redox capacity in WT and PolG animals. Nonenzymatic antioxidant capacity was measured by Trolox equivalent antioxidant capacity (TEAC) assays in brain, heart, kidney, and blood from WT and PolG mice. We found increased capacity in whole tissue extracts from quadriceps and brain (Fig. 6A and B) but decreased capacity in heart, kidney, and plasma (Fig. 6C– E). These data show that there are tissue-specific modulations of nonenzymatic antioxidant capacity in all of the PolG tissues tested. Importantly, despite the higher capacity in muscle, there is still evidence for mitochondrial-specific oxidative stress in this tissue. Because the TEAC assay is not specific for a particular antioxidant, it is not feasible to speculate as to the causes of this upregulation. Enzymatic antioxidant defenses use catalytic activities to remove ROS, including superoxide anion radicals, hydroxyl radicals, and hydrogen peroxide. To approximate the ability of the PolG mouse to remove these molecules, we measured levels of total tissue glutathione (a major reducing cofactor), and the enzymatic antioxidant activities of catalase (which removes hydrogen peroxide), and superoxide dismutase (SOD, which converts superoxide anions to hydrogen peroxide). We found that total quadricep glutathione levels were significantly reduced in PolG mice (Fig. 7A), as were catalase and total SOD activities (SOD1 and SOD2) (Fig. 7B and C). Analysis of MnSOD and Cu/Zn SOD showed similarly low activities (Supplementary Figs. 9A and B) despite their transcript levels being unaffected (Supplementary Figs. 9C and D; [12]). SOD activities have been shown to be inactivated by exposure to superoxide and nitric oxide radicals [30]. In addition, we have previously shown an age-related increase in SOD2 nitration that consequently impairs its activity in aged muscle [31]. The decreases in catalase activity, SOD activity, and glutathione levels in PolG muscle would be expected to limit intracellular enzymatic antioxidant responses. Aberrant response to mitochondrial dysfunction The decrease in total mitochondrial content suggested a potential dysregulation of the mitochondrial biogenesis expression axis in PolG muscle, so we next examined changes in expression of peroxisome proliferator-activated receptor gamma coactivator 1 alpha (PGC-1α/ Ppargc1a), a major transcriptional coactivator associated with mitochondrial biogenesis (Fig. 8 A). Studies in primary muscle cells from patients with mitochondrial disorders and in murine skeletal muscle from models of mitochondrial myopathy indicate aberrant expression of PGC-1α [32,33]. Consistent with this observation, we detected a
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Fig. 6. Trolox equivalent antioxidant capacity assays show altered response in PolG mutator mice. Trolox standards were used to calibrate reducing capacity in WT vs PolG (mt/mt) tissues: (A) quadriceps; (B) brain; (C) heart; (D) kidney; and (E) plasma. (A–D) Tissues were normalized to grams of protein. All values are mean þ/– SEM (n¼ 7 WT, n¼ 7 PolG (mt/mt)); P values determined by unpaired Student’s t test; P values: n o 0.05; nn o 0.01, nnn o 0.001.
Fig. 7. Reduced antioxidant capacity in PolG mutator mouse quadriceps. (A) Total glutathione content (n¼ 5 WT, n¼ 5 PolG (mt/mt)), (B) catalase activity (n¼ 10 WT, n ¼10 PolG (mt/mt)), and (C) total superoxide dismutase (SOD) activity (n¼ 10 WT, n¼ 10 PolG (mt/mt)) in PolG (mt/mt) and WT quadriceps.
strong repression of PGC-1α transcription. Downstream nuclear transcription factors that regulate mitochondrial content, such as nuclear respiratory factor 1 (Nrf1) and one subunit of nuclear respiratory factor 2 (Gabpa, Gabpb), are likewise significantly downregulated. As would be expected from the downregulation of these transcription factors, transcript levels of nuclear-encoded (CoxIV, Cs, and Tfam) and mitochondrial-encoded genes (mt-Nd1, mt-CoI, and mt-Atp6) were also lower. The impact of decreased transcript levels is demonstrated in the reduced protein content described above. These findings show a coordinated downregulation of mitochondrial oxidative metabolic networks in skeletal muscle of PolG mice, supporting the data of others [12], and our earlier data in an independent cohort of PolG mice [11].
We postulated that the changes in glutathione levels could reflect diminished expression of not only glutathione biosynthesis enzymes but also upstream antioxidant regulatory pathways. To test this idea, we examined the transcript levels of Gclc, which encodes γ-GCLC, the catalytic subunit of the rate-limiting step in glutathione synthesis [34]. As expected, the reduced total muscle glutathione content in PolG mice was reflected in Gclc expression (Fig. 8B). A major upstream regulator of Gclc expression is the nuclear factor-erythroid 2 p45related factor 2 (NFE2L2/Nrf2), which governs an important signaling pathway involved in mediating enzymatic antioxidant responses. Under oxidative stress, Nrf2 is released from its cytosolic repressor, Kelch-like ECH-associated protein 1 (KEAP1), to translocate into the nucleus and activate expression of phase II antioxidants that mount a
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Fig. 8. Alteration in Pgc1α and Nrf2 expression axes. (A) Ppargc1a expression and downstream targets in PolG (mt/mt) and WT quadriceps muscle (n ¼6 WT, n¼ 6 PolG (mt/ mt)). (B) Nrf2-mediated antioxidant gene expression analysis in quadriceps of WT and PolG mutator mouse (n¼ 6 WT, n¼ 6 PolG (mt/mt)). All values are average þ/– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05, nnn o0.001.
pleiotropic cellular defense program. Compared to WT mice, we found that the skeletal muscle of PolG animals demonstrated a significant reduction in Nrf2 expression and its downstream phase II antioxidants including catalase, HMOX1, and enzymes involved in regulating cellular glutathione metabolism and redox homeostasis such as γ-GCLC, glutathione peroxidases, and glutathione S-transferases (Fig. 8B). These findings are highly relevant to aging and mitochondrial dysfunction in humans, as our earlier work had demonstrated that Nrf2 and KEAP1 levels were dysregulated in sedentary aged human individuals, who also showed significant mitochondrial dysfunction, higher oxidative stress markers, and alterations in antioxidant enzyme compensation [35]. Together, our data suggest that dysregulation of antioxidant defenses occurs in PolG mice, which is consistent with our observation of increased hallmarks of mitochondrial oxidative damage in older adults [31,35].
Materials and methods Breeding of mtDNA mutator and littermate wild-type mice Mice heterozygous for the mitochondrial polymerase gamma knock-in mutation (C57Bl/6 J, PolgA þ /D257A) were a kind gift of Dr. Tomas A. Prolla, University of Wisconsin–Madison (USA) [10].
We generated homozygous knock-in mtDNA mutator mice (PolG; PolgAD257A/D257A) and littermate wild-type mice (WT; PolgA þ / þ ) from the heterozygous mice colony maintained at the McMaster University Central Animal Facility as previously described [11]. During breeding, all animals were housed three to five per cage in a 12-h light/dark cycle and were fed ad libitum (Harlan-Teklad 8640 22/5 rodent diet) after weaning. The presence of the polymerase gamma homozygous knock-in mutation was confirmed as previously described [10]. Briefly, genotyping was performed using an Extract-N-Amp Tissue PCR Kit (Sigma-Aldrich, St. Louis, MO) with genotyping primers (Supplemental Table S1). Study design Tissue harvesting was carried out as previously described using an independent cohort of mice [11]. Briefly, at 3 months of age, mice were housed individually in microisolator cages in a temperature- and humidity-controlled room and maintained on a 12h light-dark cycle with food and water ad libitum [36]. Equal numbers of age- and sex- matched PolG and littermate wild-type (WT) female and male mice were utilized in this study (n ¼10/ group; ♀¼♂). At 8 months of age (previously described [11] end point for mutator mice) animals were euthanized, and tissues collected for molecular analyses. This study was approved by the
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McMaster University Animal Research and Ethics Board under the global Animal Utilization Protocol 05-03-11, which strictly followed guidelines of the Canadian Council of Animal Care. Skeletal muscle mitochondrial isolation Four hundred milligrams of fresh skeletal muscle tissue (quadriceps) was homogenized in 2 ml of Isolation Buffer 1 (IB1: 67 mM sucrose, 50 mM Tris, 50 mM KCl, 10 mM EDTA, 0.2% fatty acid free BSA, pH 7.4, w/KOH) using a Polytron mechanical homogenizer at medium setting for 30 s. The homogenate was transferred to a chilled glass-on-glass dounce-type homogenizer and further homogenized with 40 strokes using a tight fitting pestle. The samples were centrifuged at 700g for 15 min at 4 1C. The resulting supernatant was collected and the pellet was resuspended in 1 ml of IB1, before recentrifugation as above. The pooled supernatant was then centrifuged at 12,000g for 20 min to pellet mitochondria. This crude mitochondrial fraction was snap-frozen in liquid nitrogen and stored at -80 1C. Protein analysis, and markers of protein/lipid oxidative damage Mitochondrial pellets were resuspended in 150 μl of NP40 lysis buffer (50 mM Tris, 150 mM NaCl, 1% NP40, pH 7.4) supplemented with protease inhibitor cocktail (Roche Diagnostics, Indianapolis IN) and phosphatase inhibitor (PhosSTOP, Roche Diagnostics). Protein concentrations were determined using the Pierce BCA Protein Assay Kit (Thermo Scientific, Waltham MA). Denatured proteins were separated by SDS-PAGE and the expression of specific proteins was analyzed by Western blotting with antiVDAC (Cell Signaling Technology, Danvers MA), anti-4-hydroxy-2nonenal antibody (4-HNE) (Abcam, Cambridge MA), and antinitrotyrosine antibody (Abcam). For determination of protein carbonyls (PC) samples were first processed using the OxyBlot Protein Oxidation Detection Kit (Millipore, Billerica MA) and then separated by SDS-PAGE. Quantification of the immunoreactive signal from specific bands (VDAC) or whole-lane staining (4HNE, NT, and PC) was performed using ImageJ software (http:// rsbweb.nih.gov/ij/docs/index.html, NIH). Whole tissue lysate for enzyme assays Total protein was extracted from tissue samples as previously described [11]. Briefly, 30 mg of skeletal muscle (quadriceps femoris) was homogenized on ice in a 2 ml Wheaton glass homogenizer (Fisher Scientific, Waltham MA) with 25 vol of phosphate homogenization buffer (50 mM KPi, 5 mM EDTA, 0.5 mM DTT, 1.15% KCl supplemented with a Complete Mini ETDA-free protease inhibitor cocktail tablet and a PhosSTOP phosphatase inhibitor cocktail tablet [Roche Diagnostics] per 10 ml buffer). The lysate was centrifuged at 600g for 15 min at 4 1C to pellet cellular debris. The supernatant was aliquoted, snap-frozen in liquid nitrogen, and stored at -80 1C until further analysis. Superoxide dismutase and catalase enzyme activity Muscle total superoxide dismutase (Mn-SOD and Cu/Zn-SOD) activity was determined in muscle lysate by measuring the kinetic consumption of superoxide radical (O-2) by SOD in a competitive reaction with cytochrome c, as previously described [31]. Absorption was recorded at 550 nm every 15 s for 2 min at 37 1C. One unit (U) of SOD activity was defined as the amount of enzyme that caused a 50% inhibition of the reduction of cytochrome c. Total SOD activity was expressed in units per milligram of protein. In a separate cuvette, the same sample was analyzed under identical conditions in the presence of 0.2 M KCN (pH 8.5–9.5), a potent
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inhibitor of cytosolic Cu/Zn-SOD [37] for determination of mitochondrial Mn-SOD activity. Cu/Zn-SOD activity was approximated by subtracting Mn-SOD activity from total SOD activity. Both MnSOD and Cu/Zn-SOD activities were expressed in units per milligram of protein. Catalase activity was determined by measuring the kinetic decomposition of H2O2 as previously described [38]. Catalase activity was measured by combining 960 μl of K2HPO4 buffer (50 mM with 50 mM EDTA and 0.01% Triton X-100, pH 7.2– 7.4) with 150 μg of muscle homogenate. Then 10 μl of H2O2 (1 M) was added to the cuvette and mixed by inversion to initiate the reaction. Absorbance was measured at 240 nm every 15 s for 2 min. Catalase activity was calculated and reported in micromoles per minute per milligram protein. All samples were analyzed in duplicate on a Cary UV-Vis spectrophotometer (Varian, Inc.). Measurement of mtDNA oxidative damage by 8-OHdG immunoblotting Total genomic/mitochondrial DNA was isolated from skeletal muscle using the QIAamp DNA Mini Kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. DNA samples were treated with RNase (Fermentas, Ottawa, ON) to remove RNA contamination. To prevent auto/air oxidation of DNA, ethanethiol (4% v/v, Sigma-Aldrich) was added to RNase-DNase-free water to elute total genomic/mitochondrial DNA from the column. This step is critical in assessing mtDNA oxidation in response to experimental conditions and prevents artifactual oxidation of DNA bases by prooxidant environmental sources that could serve to negate differences between groups [39]. DNA concentration and quality were assessed using Nanodrop 2000 (Thermo Scientific). DNA preparations were digested with KpnI and DraIII endonucleases (New England Biolabs, Ipswich, MA) followed by treatment with Exonuclease III (New England Biolabs) to degrade any contaminating nuclear DNA. One microgram of resulting mtDNA from each sample was dot-blotted on a nitrocellulose membrane (Amersham, Piscataway, NJ). Immunoblotting was carried out using mouse monoclonal 8-OHdG (N45.1) antibody (Japan Institute for the Control of Aging, Baltimore, MD). Membranes were then incubated with anti-mouse HRP-linked secondary antibody (Bio-Rad Laboratories, Hercules, CA) and visualized by enhanced chemiluminescence (Amersham, Pittsburgh, PA). Relative intensities of the circular dots were digitally quantified by using ImageJ analysis software (version 1.37, Scion Image). Real-time PCR of mtDNAencoded COXII was carried out to further ensure equal dotting of mtDNA from each sample. Total antioxidant capacity Plasma samples from WT and PolG mice were analyzed for TEAC using an assay based on the decolorizing of a solution of 2,20 azinobis-3-ethylbenzothiazoline-6-sulfonic acid (ABTS) radical cations (ABTS. þ ) by antioxidant solutions. After preparing stock solutions of ABTS (5.00 10-4 M) and sodium persulfate solution (6.89 10 3 M) (Sigma-Aldrich) in PBS (pH 7.4), 1 ml of sodium persulfate solution was added to 99 ml of ABTS solution and allowed to equilibrate in the dark for 16 h. All measurements were taken in duplicate by adding 40 μl of each sample or standard to 160 μl of ABTS. þ , incubating at 37 1C for 30 min, and measuring absorbance at 734 nm using a Benchmark Plus 96-well microplate reader (Bio-Rad Laboratories). Serial dilutions of the vitamin E analog Trolox (Sigma-Aldrich) were prepared in ethanol and used to generate a standard curve. Plasma samples were diluted 80-fold to ensure that they were within the linear portion of the standard curve. All results from tissues are expressed as millimolar Trolox equivalent per gram protein and results from serum expressed millimolar Trolox equivalent.
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Glutathione content
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Cell culture treatments C2C12 myoblasts were grown to confluence in Dulbecco’s modified Eagle medium (DMEM) plus 10% serum (1:1 fetal calf serum:fetal bovine serum). Myoblasts were differentiated in DMEM plus 2% horse serum for 5 days and visually monitored for myotube formation. For each treatment, myotubes were washed with PBS and incubated in serum-free DMEM containing 0, 200, or 400 mM hydrogen peroxide (Sigma-Aldrich) for 15 or 30 min as indicated. Stock H2O2 concentration was calculated using absorption at 240 nm [40]. 1D- and 2D-IMAGE analyses Extensive methodology explaining Southern blotting of digested DNA, single and double dimension separation of intact mtDNA, and S1-nuclease treatment has recently been described [21]. Additional methods Blood and tissue collection, total RNA isolation from skeletal muscle, real-time quantitative PCR, and electron microscopy are all described in Supplementary Materials and Methods. Statistical analyses All aforementioned molecular indices between the groups (PolG vs WT mice) were analyzed using two-tailed unpaired Student’s t test or one-way ANOVA with Bonferroni post hoc comparison analysis using Excel (Microsoft, Redmond, WA) or Statistica 5.0 software (Statsoft, Tulsa, OK). Statistical significance was established at a minimum P value r0.05. Data are presented as mean 7standard error of the mean (SEM).
Acknowledgments This work was supported by Muscular Dystrophy Association [MDA-69064] and Children’s Hospital of Philadelphia pilot award to B.A.K.; Canadian Institute of Health Research (CIHR) and a kind donation from Mr. Warren Lammert and family to M.A.T. A.S. was funded by Banting Postdoctoral Fellowship (CIHR) and American Federation for Aging Research and Ellison Medical Foundation Postdoctoral Fellowship. Funding for open access charge: Departmental funds. The authors thank Dr. Joseph Baur for his critical reading of this manuscript.
Appendix A.
Supplementary Information
Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.freeradbiomed. 2014.07.038.
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Original Contribution
Defects in mitochondrial DNA replication and oxidative damage in muscle of mtDNA mutator mice Jill E. Kolesar a,1, Adeel Safdar b,c,d,1, Arkan Abadi c, Lauren G. MacNeil c, Justin D. Crane c,d, Mark A. Tarnopolsky c,d,n, Brett A. Kaufman a,nn a
Department of Animal Biology, University of Pennsylvania School of Veterinary Medicine, Philadelphia, PA 19104, USA Department of Kinesiology, McMaster University, Hamilton, ON L8N 3Z5, Canada c Department of Pediatrics, McMaster University, Hamilton, ON L8N 3Z5, Canada d Department of Medicine, McMaster University, Hamilton, ON L8N 3Z5, Canada b
art ic l e i nf o
a b s t r a c t
Article history: Received 30 April 2014 Received in revised form 24 July 2014 Accepted 28 July 2014 Available online 12 August 2014
A causal role for mitochondrial dysfunction in mammalian aging is supported by recent studies of the mtDNA mutator mouse (“PolG” mouse), which harbors a defect in the proofreading-exonuclease activity of mitochondrial DNA polymerase gamma. These mice exhibit accelerated aging phenotypes characteristic of human aging, including systemic mitochondrial dysfunction, exercise intolerance, alopecia and graying of hair, curvature of the spine, and premature mortality. While mitochondrial dysfunction has been shown to cause increased oxidative stress in many systems, several groups have suggested that PolG mutator mice show no markers of oxidative damage. These mice have been presented as proof that mitochondrial dysfunction is sufficient to accelerate aging without oxidative stress. In this study, by normalizing to mitochondrial content in enriched fractions we detected increased oxidative modification of protein and DNA in PolG skeletal muscle mitochondria. We separately developed novel methods that allow simultaneous direct measurement of mtDNA replication defects and oxidative damage. Using this approach, we find evidence that suggests PolG muscle mtDNA is indeed oxidatively damaged. We also observed a significant decrease in antioxidants and expression of mitochondrial biogenesis pathway components and DNA repair enzymes in these mice, indicating an association of maladaptive gene expression with the phenotypes observed in PolG mice. Together, these findings demonstrate the presence of oxidative damage associated with the premature aging-like phenotypes induced by mitochondrial dysfunction. & 2014 Elsevier Inc. All rights reserved.
Introduction Mitochondria are host to numerous biosynthetic, bioenergetic, and signaling processes that ultimately couple cellular metabolism to homeostasic regulatory mechanisms. As the dominant site of reducing equivalent (electrons) and oxygen consumption, the
Abbreviations: cat, catenanes; CCC, covalently closed circles; ddC, dideoxycytosine; ETC, electron transport chain; MEFs, mouse embryonic fibroblasts; mtDNA, mitochondrial DNA; NT, nitrotyrosine; PC, protein carbonyls; NC, nicked circles; rc, relaxed circles; ROS, reactive oxygen species; WT, wild-type; 1D-IMAGE, onedimensional intact mtDNA agarose gel electrophoresis; 2D-IMAGE, twodimensional intact mtDNA agarose gel electrophoresis; 4-HNE, 4-hydroxy-2nonenal; 8-OHdG, 8-oxodeoxyguanisine. n Corresponding author at: Departments of Pediatrics and Medicine, HSC-2H26, McMaster University Medical Center, 1200 Main St. W., Hamilton, ON L8N 3Z5, Canada. fax: 1–905-577–8380. nn Corresponding author. fax: 1 215 573 5188. E-mail addresses: [email protected] (M.A. Tarnopolsky), [email protected] (B.A. Kaufman). 1 Co-first authors. http://dx.doi.org/10.1016/j.freeradbiomed.2014.07.038 0891-5849/& 2014 Elsevier Inc. All rights reserved.
mitochondrial electron transport chain (ETC) is a major source of reactive oxygen species (ROS). Alterations in ETC function, in particular in complexes (Co) I and III, can cause redox imbalances and oxidative damage to nearby proteins, lipids, and nucleic acids. Mitochondrial DNA (mtDNA), which is present in thousands of copies per cell, encodes components of ETC complexes I, III, and IV. mtDNA is intrinsically more sensitive than nuclear DNA to oxidative damage and is more slowly repaired than nuclear DNA [1]. Thus, mtDNA may be damaged even under conditions where nuclear DNA is not. In the context of mitochondrial dysfunction, this effect is exacerbated by the close proximity of mtDNA to the source of mitochondrial ROS. mtDNA therefore serves as a sensitive marker for the detection of overall mitochondrial oxidative stress A multitude of diseases and conditions involve both mitochondrial dysfunction and elevated oxidative stress (ex. [2–4]). For example, myocardial infarction has been shown to induce mitochondrial dysfunction and oxidative stress resulting in mtDNA damage [5]. Age-related neuropathologies such as Parkinson's
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disease and Alzheimer's disease show a strong association between mitochondrial dysfunction and oxidative stress (reviewed in [2]). Evidence also exists for elevated oxidative stress in cases where the primary cause of mitochondrial dysfunction is known, such as in myoclonic epilepsy and ragged-red fibers (MERFF) syndrome caused by the mtDNA mutation A8344G [6]. More recently, mice engineered to harbor a mt-ND6 mutation mimicking a prevalent Leber’s hereditary optic neuropathy (LHON) mutation show clear complex I defects and elevated ROS generation [7]. It is therefore generally accepted that mitochondrial dysfunction, in particular involving Co I and III, can cause elevated ROS that potentially alter physiology through damage or homeostatic signaling mechanisms. Mitochondrial respiratory function and mtDNA sequence stability both decline during normal aging [4,8]. A causal role for somatic (and possibly stem cell) mtDNA mutations in mammalian aging is supported by recent studies of the mtDNA mutator mouse (“PolG” mouse), which harbors a defect in the proofreading-exonuclease activity of mitochondrial DNA polymerase gamma [9,10]. These mice exhibit accelerated aging (reduced life span) and phenotypes characteristic of human aging including muscle atrophy, cardiomyopathy, anemia, skin and fur aging (thin dermis and gray fur), and systemic mitochondrial dysfunction [9–12]. Somewhat surprisingly, several groups have examined markers of oxidative stress in these mice and found no change [10,12], while others have provided support for oxidative stress in the PolG mice [13,14]. Resolving this discrepancy is crucial to solidifying the causal links among mitochondrial dysfunction, ROS, and aging, which have been the subject of debate [15]. In an effort to definitively determine whether PolG mice are under mitochondrial oxidative stress, we examined multiple aspects of mtDNA integrity as a primary readout of oxidative damage occurring in PolG mitochondria. mtDNA damage is frequently detected by a long-template PCR assay [16] that relies on base modification and breaks in the template to serve as an indication of oxidative stress. To our knowledge, this PCR technique has not been reported in PolG mice, likely because the replication pausing known to occur in these mice [17] could produce truncated templates that may lead to misinterpretation of reduced amplification efficiency as mtDNA damage. In the following study, we developed novel methods that detect both replication pausing and oxidative damage directly in intact mtDNA. The application of these methods to the PolG mutator mouse muscle, along with standard methods, strongly indicates that the mitochondrial compartment is specifically under oxidative stress in these mice. We show that mitochondrial biogenesis and
antioxidant defense pathways are highly dysregulated, similar to what is observed in normal human aging, suggesting that responses to mitochondrial dysfunction may be exacerbated by maladaptive gene expression.
Results and discussion Reduced tissue mass and mitochondrial dysmorphy are associated with mitochondrial dysfunction in PolG mice PolG mice suffer from numerous muscle pathologies [9–12]. For example, we and others have observed a significant reduction in skeletal muscle mass (Fig. 1A) and lower Complex IV activity (Fig. 1B) in quadriceps muscle (mainly fast-twitch) from independent cohorts of PolG mice compared to WT [9–12]. A similar reduction in respiratory chain activity was found in saponinpermeabilized soleus (mainly slow-twitch) muscle (Supplementary Fig. 1; [18]). The mitochondria in PolG quadriceps also show aberrant ultrastructure by electron microscopy, consistent with the quantitative decrease in mitochondrial number and increase in mitochondrial volume we previously described [19]; Supplementary Fig. 2). These results confirm that the mice used in this study have the same phenotypes as those previously reported, which is critical for interpreting further tests of the existence of mitochondrial oxidative stress in PolG mice. MtDNA structural defects, lower mtDNA levels, and DNA breaks in PolG mutator mice Multiple alterations to mtDNA molecules have been reported in PolG mouse tissues, including control region multimers (CRM) [20], elevated levels of linear mtDNA content [9], and replication pausing [17]. Because these alterations would manifest across entire genomes, we surveyed PolG mtDNA for these and other alterations using the high-resolution approach of two-dimensional intact mtDNA agarose gel electrophoresis (2D-IMAGE) [21]. This approach allows the separation and direct detection of up to 24 mtDNA structural variants, in particular structures with singlestranded DNA content thought to be DNA replication intermediates. 2D-IMAGE and associated mtDNA analyses of PolG muscle revealed the presence of a discrete mtDNA variant (labeled var.), whose migration characteristics (between 1n and 2n nicked circles, NC) are consistent with replication intermediates detected in cells during recovery from mtDNA depletion [21] (Fig. 2). The
Fig. 1. Decreased muscle weight and mitochondrial activity in quadriceps of PolG mutator mouse. (A) Mass determination of quadriceps (quad.), gastrocnemius (gastroc.), soleus, and extensor digitorum longus (EDL) at 8 months of age (n¼ 10 WT, n¼ 10 PolG (mt/mt)). (B) Protein-content normalized cytochrome c oxidase (COX) activity in quadriceps expressed as percentage of WT (n¼ 3 WT, n¼ 3 PolG (mt/mt)). All values are mean þ/– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05; nn o 0.01, nnn o 0.001.
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Fig. 2. Mitochondrial DNA alterations and damage in PolG mutator mice quadriceps by 2D-IMAGE. (A) Typical 2D-IMAGE profile of DNA preparations probed for mtDNA in WT and PolG quadriceps. Major structures are indicated, including a known variant (indicated var.) and shorter form (indicated #) in PolG samples. (B) Relative abundance of major mtDNA structural and topological isomers in PolG and WT quadriceps (n¼ 3 WT, n¼3 PolG (mt/mt)). (C) Relative abundance of circular and linear mtDNA, and the circular to linear (C:L) mtDNA ratio in PolG and WT quadriceps (n¼ 3 WT, n¼ 3 PolG (mt/mt)). (D). Linearization of mtDNA by XhoI digestion shows full-length mtDNA, mtDNA structure previously assigned “deletion” in the PolG mouse, and control region multimers (CRM). (E) Relative mtDNA content of PolG and WT quadriceps by qPCR and Southern blot after SacI digestion (n ¼3 WT, n ¼3 PolG (mt/mt)). All values are mean þ /– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05; nn o0.01, nnn o 0.001. Abbreviations: catenanes, cat; nicked circles, nc; linear, lin; covalently closed circles, ccc; variant, var.; linear degradation products, lin. deg; and n.a., not applicable.
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2D-IMAGE characteristics of the mtDNA variant are identical to the replication intermediates described in PolG mouse liver (Fig. 2A; [17]), and this variant migrates as an apparent “deletion” by standard Southern blot protocols after XhoI digestion (Fig. 2D; [9]). In addition, we observed an accumulation of a heterogeneous DNA population that extends upward from the 1n linear on the linear diagonal toward larger genome lengths (Fig. 2A, labeled #). The diminished abundance of catenanes and CCC molecules, previously shown to be produced during replication [21], coupled with increased replication intermediates is consistent with the reported deficiency in replication processivity of the PolG mutator protein [17]. Southern blotting also revealed hybridizing species near the well, which have been described for CRM tandem repeats (Fig. 2D; [20]). 2D-IMAGE successfully detects all reported aspects of altered replication described for the PolG mutator mice, but also provides new information regarding the status of mtDNA integrity.
Quantification of the prominent mtDNA structural and topological conformers in PolG muscle using 2D-IMAGE revealed a significant shift in the relative abundance of circular and linear mtDNA. Comparison of summed circular DNA (1n NC, catenanes, and CCC) abundance relative to linear DNA generated a circular to linear (C:L) ratio of 10:1 for WT muscle and 2:1 for PolG muscle (Fig. 2C). In contrast to other reports [9,22], we found that mtDNA levels were lower in skeletal muscle of the PolG mouse both by qPCR for total mtDNA and by Southern blot for full-length mtDNA detection after SacI digestion (Fig. 2E, [23]). There is strong evidence for linear mtDNA instability causing rapid genome depletion [24], suggesting that the mechanisms that underlie the elevated levels of doublestrand breaks in mtDNA (manifested as linear molecules) could contribute to the observed genome depletion. We hypothesized that either replicative pausing or oxidative damage could contribute to the increased linear mtDNA content in
Fig. 3. Quantitation of circular and linear mtDNA by 1D-IMAGE analysis. (A,C,E) Quantitation of major topoisomers by 1D-IMAGE and phosphorimage analysis. (B,D,F) Determination of relative abundance of circular mtDNA by ratio to linear mtDNA (circular:linear; C:L). Samples analyzed: (A,B) 24-h treatment of MEFs with ddC; (C,D) C2C12 myotubes exposed to 200 or 400 mM H2O2 for 15 min; (E,F) Quadriceps from WT vs PolG (mt/mt) mice. P values in panels C and D (n ¼3 each) were determined by one-way ANOVA with Bonferroni post hoc analysis against the untreated sample indicated (n o 0.05; nn o 0.01, nnn o 0.001). P values in panels E and F (n¼ 3 each) were determined by Student’s t test (n o 0.05; nn o0.01, nnn o0.001).
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the PolG muscle. To test the effects of replication arrest on levels of double strand breaks, we exposed cultured MEFs to dideoxycytosine (ddC), a deoxynucleotide chain terminator that preferentially affects mtDNA replication [25]. Single dimension intact mtDNA agarose gel electrophoresis (1D-IMAGE) showed that ddC exposure causes dose-dependent linearization, culminating in the depletion of circular mtDNA (Fig. 3A and B). To test whether oxidation caused linearization of mtDNA, we exposed C2C12 myotubes to acute oxidative stress for 1D-IMAGE analysis (Fig. 3C and D). Short hydrogen peroxide (H2O2) exposures were selected to prevent complication of data interpretation due to adaptive gene expression. The formation of double-strand breaks (linears) occurred in a dose-dependent manner in myotubes in short exposures (15 min). Longer exposures (30 min) showed a plateau in linearization (Supplementary Fig. 3), possibly due to limited availability of iron for Fenton chemistry to produce hydroxyl radicals [26]. The effects of oxidative damage on the formation of linear mtDNA were confirmed by 2D-IMAGE of myotubes (Supplementary Fig. 4), MEFs (Supplementary Fig. 5), and neonatal cardiomyocytes (Supplementary Fig. 6). Cells were adherent in all cases, with MEFs and neonatal cardiomyocytes being subconfluent. These results suggest that acute oxidant exposure linearizes circular mtDNA in a dose-responsive manner. To distinguish between replicative pausing and oxidative damage to mtDNA in the PolG mouse, we devised a method to describe the relative abundance of single-strand breaks in mtDNA (Fig. 4). We have previously shown that S1 nuclease will cut across from single-strand breaks in circular double-stranded mtDNA genomes to produce linear
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molecules detectable in 1D and 2D-IMAGE profiles [21]. First, we tested whether hydrogen peroxide and ddC treatments resulted in differential S1 resistance. As expected, circular DNA (2n catenanes and relaxed circles) in H2O2 samples showed increased S1 sensitivity and formed more linear mtDNA relative to the untreated control (Fig. 4A and B). In contrast, the ddC-treated samples showed a different diagnostic pattern upon S1 treatment, with linear molecules decreasing during replication pausing (Fig. 4C and D). These results suggested that S1 nuclease treatment may be used as a tool to detect the presence of oxidative damage in mtDNA. When comparing the relative S1 sensitivity of WT and PolG mouse muscle, we observed only a slight increase in full-length 1n linear molecules upon S1 digestion of WT mtDNA (Fig. 4E), suggesting limited oxidative stress. In contrast, circular mtDNA in PolG muscle was more sensitive to S1 nuclease treatment and produced more linear mtDNA degradation products (Fig. 4F), consistent with oxidative damage. We also noted that the expressions of base excision repair enzymes Nth1, UNG1, and OGG1 were significantly downregulated in the PolG samples (Fig. 4G). Taken together, the single-strand breaks in PolG mtDNA structures are consistent with oxidative damage and may persist due to insufficient expression of DNA repair enzymes.
Mitochondrial dysfunction in PolG mice is associated with increased oxidative damage to mitochondrial protein and mtDNA Initial examination by others of PolG tissue homogenates for the contribution of mitochondrial dysfunction to oxidative stress failed to
Fig. 4. PolG mtDNAs demonstrate increased sensitivity to S1 nuclease digestion, consistent with oxidative damage. Identical amounts of total DNA were treated in parallel with endonuclease S1 and resolved by 1D-IMAGE (A,C,E) and the fraction of DNA resistant to digestion relative to control was determined (B,D,F). (A) Representative 1DIMAGE of H2O2-treated myotubes reveals increased S1 sensitivity of catenanes and relaxed circles, producing linear mtDNA. (B) Graphical representation of S1 resistance of H2O2-treated myotubes (n¼ 3). (C) 1D-IMAGE of MEFs cultured overnight in the presence of varying concentrations of the chain terminator dideoxycytosine (ddC) reveals a distinctive S1 pattern. (D) Graphical representation of ddC-exposed samples for S1 resistance of topoisomers normalized to untreated control. (E) Representative 1D-IMAGE of PolG (mt/mt) muscle reveals elevated S1 sensitivity of all major topoisomers relative to WT. (F) Graphical representation of PolG (mt/mt) S1 resistance for topoisomers normalized to WT (n¼ 2 WT, n¼2 PolG (mt/mt)). (G) Relative gene expression of base excision repair enzymes Nth1, UNG1, and OGG1 (n¼ 3 per genotype). All values are mean þ /– SEM; P values determined by one-way ANOVA with Bonferroni post hoc analysis (B) or unpaired Student’s t test (F,G); P values: n o 0.05; nn o 0.01, nnn o 0.001.
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muscle by Western blot analysis for VDAC, ANT1, and CS (Supplementary Fig. 8). By comparing oxidative modifications normalized to mitochondrial content in these fractions, we observed a significant increase in PC, 4-HNE, and nitrotyrosine (NT) in PolG mitochondrial fractions vs WT (Fig. 5A–C). To further support the occurrence of oxidative modifications in mitochondria, we examined levels of 8-OHdG DNA modification within circular mtDNA (Fig. 5E) and found that the PolG muscle mtDNA contained significantly elevated levels of 8-OHdG relative to controls. The amount of mtDNA analyzed was confirmed by qPCR (data not shown). These data clearly show the presence of increased oxidative protein and base modification in muscle mitochondria of this strain of PolG mice, and support our observations of single-strand breaks in mtDNA originating from oxidation. Additional studies will be necessary to determine whether strain background, notably variation in the NNT gene, contributes to the manifestation of oxidative damage [28,29]. Differential effects on nonenzymatic and enzymatic redox capacity in PolG tissues
Fig. 5. Mitochondrial markers of oxidative stress. (A) Relative levels of protein carbonylation (PC) in mitochondrial preparations of PolG (mt/mt) and WT quadriceps normalized to VDAC abundance (n¼ 6 WT, n ¼6 PolG (mt/mt)). (B) Relative levels of 4-HNE lipid modification of proteins from quadriceps mitochondrial preparations of PolG (mt/mt) and WT muscle normalized to VDAC abundance (n¼6 WT, n ¼5 PolG (mt/mt)). (C) Tyrosine nitrosylation in mitochondrial preparations of PolG (mt/mt) and WT muscle normalized to VDAC abundance (n ¼6 WT, n¼ 4 PolG (mt/mt)). (D) 8-OHdG DNA base modification in circular DNA isolated from PolG (mt/mt) and WT quadriceps muscle normalized to mt-Co2 as determined by qPCR (n¼ 6 WT, n¼ 4 PolG (mt/mt)). All values are mean þ/– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05; nn o0.01, nnn o0.001.
reveal elevated ROS or increased oxidative damage despite significant accumulation of mtDNA point mutations [10,27]. In agreement with aspects of these studies, we also found no difference in markers of oxidative damage in whole muscle homogenates of PolG and WT mice as measured by 4-hydroxy-2-nonenal (4-HNE; marker of lipid oxidation) and protein carbonyls (PC; marker of protein oxidation) (Supplementary Fig. 7). However, we did find a consistently lower protein content for several well-known protein markers of total mitochondrial abundance VDAC (outer mitochondrial membrane), CS (mitochondrial matrix), and ANT1 (inner mitochondrial membrane) (data not shown). The lower mitochondrial protein levels are consistent with the decreased mitochondrial numbers observed by EM in PolG muscle [19]. We interpret these findings as evidence for limited total cellular oxidative damage, which agrees with previous reports [9,10,12] in the context of reduced mitochondrial content. To determine whether ROS-mediated damage occurs specifically in muscle mitochondria of PolG mice, we analyzed mitochondriaenriched fractions for several oxidative damage markers (including those applied to whole tissue lysates in Supplementary Fig. 7). As expected from the whole tissue lysates and electron microscopy, we found that the relative recovery of mitochondria was lower in PolG
To establish whether conditions in the PolG mouse were conducive to oxidative stress, we examined total nonenzymatic redox capacity and indicators of enzymatic redox capacity in WT and PolG animals. Nonenzymatic antioxidant capacity was measured by Trolox equivalent antioxidant capacity (TEAC) assays in brain, heart, kidney, and blood from WT and PolG mice. We found increased capacity in whole tissue extracts from quadriceps and brain (Fig. 6A and B) but decreased capacity in heart, kidney, and plasma (Fig. 6C– E). These data show that there are tissue-specific modulations of nonenzymatic antioxidant capacity in all of the PolG tissues tested. Importantly, despite the higher capacity in muscle, there is still evidence for mitochondrial-specific oxidative stress in this tissue. Because the TEAC assay is not specific for a particular antioxidant, it is not feasible to speculate as to the causes of this upregulation. Enzymatic antioxidant defenses use catalytic activities to remove ROS, including superoxide anion radicals, hydroxyl radicals, and hydrogen peroxide. To approximate the ability of the PolG mouse to remove these molecules, we measured levels of total tissue glutathione (a major reducing cofactor), and the enzymatic antioxidant activities of catalase (which removes hydrogen peroxide), and superoxide dismutase (SOD, which converts superoxide anions to hydrogen peroxide). We found that total quadricep glutathione levels were significantly reduced in PolG mice (Fig. 7A), as were catalase and total SOD activities (SOD1 and SOD2) (Fig. 7B and C). Analysis of MnSOD and Cu/Zn SOD showed similarly low activities (Supplementary Figs. 9A and B) despite their transcript levels being unaffected (Supplementary Figs. 9C and D; [12]). SOD activities have been shown to be inactivated by exposure to superoxide and nitric oxide radicals [30]. In addition, we have previously shown an age-related increase in SOD2 nitration that consequently impairs its activity in aged muscle [31]. The decreases in catalase activity, SOD activity, and glutathione levels in PolG muscle would be expected to limit intracellular enzymatic antioxidant responses. Aberrant response to mitochondrial dysfunction The decrease in total mitochondrial content suggested a potential dysregulation of the mitochondrial biogenesis expression axis in PolG muscle, so we next examined changes in expression of peroxisome proliferator-activated receptor gamma coactivator 1 alpha (PGC-1α/ Ppargc1a), a major transcriptional coactivator associated with mitochondrial biogenesis (Fig. 8 A). Studies in primary muscle cells from patients with mitochondrial disorders and in murine skeletal muscle from models of mitochondrial myopathy indicate aberrant expression of PGC-1α [32,33]. Consistent with this observation, we detected a
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Fig. 6. Trolox equivalent antioxidant capacity assays show altered response in PolG mutator mice. Trolox standards were used to calibrate reducing capacity in WT vs PolG (mt/mt) tissues: (A) quadriceps; (B) brain; (C) heart; (D) kidney; and (E) plasma. (A–D) Tissues were normalized to grams of protein. All values are mean þ/– SEM (n¼ 7 WT, n¼ 7 PolG (mt/mt)); P values determined by unpaired Student’s t test; P values: n o 0.05; nn o 0.01, nnn o 0.001.
Fig. 7. Reduced antioxidant capacity in PolG mutator mouse quadriceps. (A) Total glutathione content (n¼ 5 WT, n¼ 5 PolG (mt/mt)), (B) catalase activity (n¼ 10 WT, n ¼10 PolG (mt/mt)), and (C) total superoxide dismutase (SOD) activity (n¼ 10 WT, n¼ 10 PolG (mt/mt)) in PolG (mt/mt) and WT quadriceps.
strong repression of PGC-1α transcription. Downstream nuclear transcription factors that regulate mitochondrial content, such as nuclear respiratory factor 1 (Nrf1) and one subunit of nuclear respiratory factor 2 (Gabpa, Gabpb), are likewise significantly downregulated. As would be expected from the downregulation of these transcription factors, transcript levels of nuclear-encoded (CoxIV, Cs, and Tfam) and mitochondrial-encoded genes (mt-Nd1, mt-CoI, and mt-Atp6) were also lower. The impact of decreased transcript levels is demonstrated in the reduced protein content described above. These findings show a coordinated downregulation of mitochondrial oxidative metabolic networks in skeletal muscle of PolG mice, supporting the data of others [12], and our earlier data in an independent cohort of PolG mice [11].
We postulated that the changes in glutathione levels could reflect diminished expression of not only glutathione biosynthesis enzymes but also upstream antioxidant regulatory pathways. To test this idea, we examined the transcript levels of Gclc, which encodes γ-GCLC, the catalytic subunit of the rate-limiting step in glutathione synthesis [34]. As expected, the reduced total muscle glutathione content in PolG mice was reflected in Gclc expression (Fig. 8B). A major upstream regulator of Gclc expression is the nuclear factor-erythroid 2 p45related factor 2 (NFE2L2/Nrf2), which governs an important signaling pathway involved in mediating enzymatic antioxidant responses. Under oxidative stress, Nrf2 is released from its cytosolic repressor, Kelch-like ECH-associated protein 1 (KEAP1), to translocate into the nucleus and activate expression of phase II antioxidants that mount a
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Fig. 8. Alteration in Pgc1α and Nrf2 expression axes. (A) Ppargc1a expression and downstream targets in PolG (mt/mt) and WT quadriceps muscle (n ¼6 WT, n¼ 6 PolG (mt/ mt)). (B) Nrf2-mediated antioxidant gene expression analysis in quadriceps of WT and PolG mutator mouse (n¼ 6 WT, n¼ 6 PolG (mt/mt)). All values are average þ/– SEM; P values determined by unpaired Student’s t test; P values: n o 0.05, nnn o0.001.
pleiotropic cellular defense program. Compared to WT mice, we found that the skeletal muscle of PolG animals demonstrated a significant reduction in Nrf2 expression and its downstream phase II antioxidants including catalase, HMOX1, and enzymes involved in regulating cellular glutathione metabolism and redox homeostasis such as γ-GCLC, glutathione peroxidases, and glutathione S-transferases (Fig. 8B). These findings are highly relevant to aging and mitochondrial dysfunction in humans, as our earlier work had demonstrated that Nrf2 and KEAP1 levels were dysregulated in sedentary aged human individuals, who also showed significant mitochondrial dysfunction, higher oxidative stress markers, and alterations in antioxidant enzyme compensation [35]. Together, our data suggest that dysregulation of antioxidant defenses occurs in PolG mice, which is consistent with our observation of increased hallmarks of mitochondrial oxidative damage in older adults [31,35].
Materials and methods Breeding of mtDNA mutator and littermate wild-type mice Mice heterozygous for the mitochondrial polymerase gamma knock-in mutation (C57Bl/6 J, PolgA þ /D257A) were a kind gift of Dr. Tomas A. Prolla, University of Wisconsin–Madison (USA) [10].
We generated homozygous knock-in mtDNA mutator mice (PolG; PolgAD257A/D257A) and littermate wild-type mice (WT; PolgA þ / þ ) from the heterozygous mice colony maintained at the McMaster University Central Animal Facility as previously described [11]. During breeding, all animals were housed three to five per cage in a 12-h light/dark cycle and were fed ad libitum (Harlan-Teklad 8640 22/5 rodent diet) after weaning. The presence of the polymerase gamma homozygous knock-in mutation was confirmed as previously described [10]. Briefly, genotyping was performed using an Extract-N-Amp Tissue PCR Kit (Sigma-Aldrich, St. Louis, MO) with genotyping primers (Supplemental Table S1). Study design Tissue harvesting was carried out as previously described using an independent cohort of mice [11]. Briefly, at 3 months of age, mice were housed individually in microisolator cages in a temperature- and humidity-controlled room and maintained on a 12h light-dark cycle with food and water ad libitum [36]. Equal numbers of age- and sex- matched PolG and littermate wild-type (WT) female and male mice were utilized in this study (n ¼10/ group; ♀¼♂). At 8 months of age (previously described [11] end point for mutator mice) animals were euthanized, and tissues collected for molecular analyses. This study was approved by the
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McMaster University Animal Research and Ethics Board under the global Animal Utilization Protocol 05-03-11, which strictly followed guidelines of the Canadian Council of Animal Care. Skeletal muscle mitochondrial isolation Four hundred milligrams of fresh skeletal muscle tissue (quadriceps) was homogenized in 2 ml of Isolation Buffer 1 (IB1: 67 mM sucrose, 50 mM Tris, 50 mM KCl, 10 mM EDTA, 0.2% fatty acid free BSA, pH 7.4, w/KOH) using a Polytron mechanical homogenizer at medium setting for 30 s. The homogenate was transferred to a chilled glass-on-glass dounce-type homogenizer and further homogenized with 40 strokes using a tight fitting pestle. The samples were centrifuged at 700g for 15 min at 4 1C. The resulting supernatant was collected and the pellet was resuspended in 1 ml of IB1, before recentrifugation as above. The pooled supernatant was then centrifuged at 12,000g for 20 min to pellet mitochondria. This crude mitochondrial fraction was snap-frozen in liquid nitrogen and stored at -80 1C. Protein analysis, and markers of protein/lipid oxidative damage Mitochondrial pellets were resuspended in 150 μl of NP40 lysis buffer (50 mM Tris, 150 mM NaCl, 1% NP40, pH 7.4) supplemented with protease inhibitor cocktail (Roche Diagnostics, Indianapolis IN) and phosphatase inhibitor (PhosSTOP, Roche Diagnostics). Protein concentrations were determined using the Pierce BCA Protein Assay Kit (Thermo Scientific, Waltham MA). Denatured proteins were separated by SDS-PAGE and the expression of specific proteins was analyzed by Western blotting with antiVDAC (Cell Signaling Technology, Danvers MA), anti-4-hydroxy-2nonenal antibody (4-HNE) (Abcam, Cambridge MA), and antinitrotyrosine antibody (Abcam). For determination of protein carbonyls (PC) samples were first processed using the OxyBlot Protein Oxidation Detection Kit (Millipore, Billerica MA) and then separated by SDS-PAGE. Quantification of the immunoreactive signal from specific bands (VDAC) or whole-lane staining (4HNE, NT, and PC) was performed using ImageJ software (http:// rsbweb.nih.gov/ij/docs/index.html, NIH). Whole tissue lysate for enzyme assays Total protein was extracted from tissue samples as previously described [11]. Briefly, 30 mg of skeletal muscle (quadriceps femoris) was homogenized on ice in a 2 ml Wheaton glass homogenizer (Fisher Scientific, Waltham MA) with 25 vol of phosphate homogenization buffer (50 mM KPi, 5 mM EDTA, 0.5 mM DTT, 1.15% KCl supplemented with a Complete Mini ETDA-free protease inhibitor cocktail tablet and a PhosSTOP phosphatase inhibitor cocktail tablet [Roche Diagnostics] per 10 ml buffer). The lysate was centrifuged at 600g for 15 min at 4 1C to pellet cellular debris. The supernatant was aliquoted, snap-frozen in liquid nitrogen, and stored at -80 1C until further analysis. Superoxide dismutase and catalase enzyme activity Muscle total superoxide dismutase (Mn-SOD and Cu/Zn-SOD) activity was determined in muscle lysate by measuring the kinetic consumption of superoxide radical (O-2) by SOD in a competitive reaction with cytochrome c, as previously described [31]. Absorption was recorded at 550 nm every 15 s for 2 min at 37 1C. One unit (U) of SOD activity was defined as the amount of enzyme that caused a 50% inhibition of the reduction of cytochrome c. Total SOD activity was expressed in units per milligram of protein. In a separate cuvette, the same sample was analyzed under identical conditions in the presence of 0.2 M KCN (pH 8.5–9.5), a potent
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inhibitor of cytosolic Cu/Zn-SOD [37] for determination of mitochondrial Mn-SOD activity. Cu/Zn-SOD activity was approximated by subtracting Mn-SOD activity from total SOD activity. Both MnSOD and Cu/Zn-SOD activities were expressed in units per milligram of protein. Catalase activity was determined by measuring the kinetic decomposition of H2O2 as previously described [38]. Catalase activity was measured by combining 960 μl of K2HPO4 buffer (50 mM with 50 mM EDTA and 0.01% Triton X-100, pH 7.2– 7.4) with 150 μg of muscle homogenate. Then 10 μl of H2O2 (1 M) was added to the cuvette and mixed by inversion to initiate the reaction. Absorbance was measured at 240 nm every 15 s for 2 min. Catalase activity was calculated and reported in micromoles per minute per milligram protein. All samples were analyzed in duplicate on a Cary UV-Vis spectrophotometer (Varian, Inc.). Measurement of mtDNA oxidative damage by 8-OHdG immunoblotting Total genomic/mitochondrial DNA was isolated from skeletal muscle using the QIAamp DNA Mini Kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. DNA samples were treated with RNase (Fermentas, Ottawa, ON) to remove RNA contamination. To prevent auto/air oxidation of DNA, ethanethiol (4% v/v, Sigma-Aldrich) was added to RNase-DNase-free water to elute total genomic/mitochondrial DNA from the column. This step is critical in assessing mtDNA oxidation in response to experimental conditions and prevents artifactual oxidation of DNA bases by prooxidant environmental sources that could serve to negate differences between groups [39]. DNA concentration and quality were assessed using Nanodrop 2000 (Thermo Scientific). DNA preparations were digested with KpnI and DraIII endonucleases (New England Biolabs, Ipswich, MA) followed by treatment with Exonuclease III (New England Biolabs) to degrade any contaminating nuclear DNA. One microgram of resulting mtDNA from each sample was dot-blotted on a nitrocellulose membrane (Amersham, Piscataway, NJ). Immunoblotting was carried out using mouse monoclonal 8-OHdG (N45.1) antibody (Japan Institute for the Control of Aging, Baltimore, MD). Membranes were then incubated with anti-mouse HRP-linked secondary antibody (Bio-Rad Laboratories, Hercules, CA) and visualized by enhanced chemiluminescence (Amersham, Pittsburgh, PA). Relative intensities of the circular dots were digitally quantified by using ImageJ analysis software (version 1.37, Scion Image). Real-time PCR of mtDNAencoded COXII was carried out to further ensure equal dotting of mtDNA from each sample. Total antioxidant capacity Plasma samples from WT and PolG mice were analyzed for TEAC using an assay based on the decolorizing of a solution of 2,20 azinobis-3-ethylbenzothiazoline-6-sulfonic acid (ABTS) radical cations (ABTS. þ ) by antioxidant solutions. After preparing stock solutions of ABTS (5.00 10-4 M) and sodium persulfate solution (6.89 10 3 M) (Sigma-Aldrich) in PBS (pH 7.4), 1 ml of sodium persulfate solution was added to 99 ml of ABTS solution and allowed to equilibrate in the dark for 16 h. All measurements were taken in duplicate by adding 40 μl of each sample or standard to 160 μl of ABTS. þ , incubating at 37 1C for 30 min, and measuring absorbance at 734 nm using a Benchmark Plus 96-well microplate reader (Bio-Rad Laboratories). Serial dilutions of the vitamin E analog Trolox (Sigma-Aldrich) were prepared in ethanol and used to generate a standard curve. Plasma samples were diluted 80-fold to ensure that they were within the linear portion of the standard curve. All results from tissues are expressed as millimolar Trolox equivalent per gram protein and results from serum expressed millimolar Trolox equivalent.
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Glutathione content
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Cell culture treatments C2C12 myoblasts were grown to confluence in Dulbecco’s modified Eagle medium (DMEM) plus 10% serum (1:1 fetal calf serum:fetal bovine serum). Myoblasts were differentiated in DMEM plus 2% horse serum for 5 days and visually monitored for myotube formation. For each treatment, myotubes were washed with PBS and incubated in serum-free DMEM containing 0, 200, or 400 mM hydrogen peroxide (Sigma-Aldrich) for 15 or 30 min as indicated. Stock H2O2 concentration was calculated using absorption at 240 nm [40]. 1D- and 2D-IMAGE analyses Extensive methodology explaining Southern blotting of digested DNA, single and double dimension separation of intact mtDNA, and S1-nuclease treatment has recently been described [21]. Additional methods Blood and tissue collection, total RNA isolation from skeletal muscle, real-time quantitative PCR, and electron microscopy are all described in Supplementary Materials and Methods. Statistical analyses All aforementioned molecular indices between the groups (PolG vs WT mice) were analyzed using two-tailed unpaired Student’s t test or one-way ANOVA with Bonferroni post hoc comparison analysis using Excel (Microsoft, Redmond, WA) or Statistica 5.0 software (Statsoft, Tulsa, OK). Statistical significance was established at a minimum P value r0.05. Data are presented as mean 7standard error of the mean (SEM).
Acknowledgments This work was supported by Muscular Dystrophy Association [MDA-69064] and Children’s Hospital of Philadelphia pilot award to B.A.K.; Canadian Institute of Health Research (CIHR) and a kind donation from Mr. Warren Lammert and family to M.A.T. A.S. was funded by Banting Postdoctoral Fellowship (CIHR) and American Federation for Aging Research and Ellison Medical Foundation Postdoctoral Fellowship. Funding for open access charge: Departmental funds. The authors thank Dr. Joseph Baur for his critical reading of this manuscript.
Appendix A.
Supplementary Information
Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.freeradbiomed. 2014.07.038.
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