Mutations in SURF1 (surfeit locus protein 1) COX (cytochrome c oxidase) assembly protein are associated with Leigh's syndrome, a human mitochondrial disorder that manifests as severe mitochondrial phenotypes and early lethality. In contrast, mice lacking the SURF1 protein (Surf1−/−) are viable and were previously shown to have enhanced longevity and a greater than 50% reduction in COX activity. We measured mitochondrial function in heart and skeletal muscle, and despite the significant reduction in COX activity, we found little or no difference in ROS (reactive oxygen species) generation, membrane potential, ATP production or respiration in isolated mitochondria from Surf1−/− mice compared with wild-type. However, blood lactate levels were elevated and Surf1−/− mice had reduced running endurance, suggesting compromised mitochondrial energy metabolism in vivo. Decreased COX activity in Surf1−/− mice is associated with increased markers of mitochondrial biogenesis [PGC-1α (peroxisome-proliferator-activated receptor γ co-activator 1α) and VDAC (voltage-dependent anion channel)] in both heart and skeletal muscle. Although mitochondrial biogenesis is a common response in the two tissues, skeletal muscle has an up-regulation of the UPRMT (mitochondrial unfolded protein response) and heart exhibits induction of the Nrf2 (nuclear factor-erythroid 2-related factor 2) antioxidant response pathway. These data are the first to show induction of the UPRMT in a mammalian model of decreased COX activity. In addition, the results of the present study suggest that impaired mitochondrial function can lead to induction of mitochondrial stress pathways to confer protective effects on cellular homoeostasis.

INTRODUCTION

SURF1 (surfeit locus protein 1) is a nuclear-encoded small hydrophobic protein localized to the mitochondrial inner membrane that aides in the initial assembly of 13 subunits of the COX (cytochrome c oxidase; complex IV) holoenzyme. Complex IV facilitates the final transfer of electrons in the ETC (electron transport chain) from cytochrome c to molecular oxygen forming water and thus plays a key role in mitochondrial oxidative phosphorylation. In humans, loss-of-function mutations in SURF1 lead to a devastating disease phenotype characterized by severe neurological deficits and early lethality [1]. However, Surf1−/− mice engineered to express a truncated and unstable SURF1 protein do not display a deleterious phenotype, despite a significant reduction in complex IV activity. In fact, the complex IV-deficient Surf1−/− mice have been reported to exhibit enhanced longevity compared with their wild-type littermates [2]. This phenotype linking mitochondrial dysfunction and increased longevity is consistent with studies in yeast, worms [3] and flies [4], in which certain mutations of the ETC can lead to extended lifespan.

We postulated that the loss of SURF1 and the reduction in complex IV activity might initiate compensatory responses to mitochondrial stress that would be beneficial to cellular homoeostasis. Previous studies in the long-lived clk-1, isp-1 and cco-1 Caenorhabditis elegans mitochondrial mutants point to a potential role of the UPRMT (mitochondrial unfolded protein response) [5], mitochondrial biogenesis [68] and Nrf2 (nuclear factor-erythroid 2-related factor 2) activation in mediating longevity in these mutants (see [9] for a review). The UPRMT is an evolutionarily conserved signalling mechanism initiated by mitochondrial stress. In C. elegans, the UPRMT is mediated through the translocation of transcription factors UBL-5/DEV-1 and ATFS-1 (ZC376.7) from the cytosol to the nucleus [1012]. This results in an increase in the expression of mitochondrial-specific chaperones such as Hsp60 (heat-shock protein 60) and proteases such as ClpP (caseinolytic peptidase) [13]. These proteins play an important role in mitochondrial protein quality control [14]. A role for this pathway in the maintenance of mitochondrial homoeostasis has also been reported in mammalian cell culture and is positively regulated by CHOP [C/EBP (CCAAT/enhancer-binding protein)-homologous protein]. Within 20 bp of the CHOP DNA-binding site, upstream and downstream, are two conserved sequences, MURE1 and MURE2 (mitochondrial unfolded response elements 1 and 2 respectively) that positively regulate the expression of UPRMT proteins [1517]. However, the identity of what binds MURE1 and MURE2 remains unknown.

Mitochondrial biogenesis is another mitochondrial homoeostasis mechanism [18]. Mitochondrial biogenesis involves the production of new mitochondria, replication of mtDNA and production of nuclear-encoded mitochondrial proteins. This process is largely under the control of the transcription co-activator PGC-1α (peroxisome-proliferator-activated receptor γ co-activator 1α) [18]. Interestingly, in C. elegans, the lifespan extension mediated by ETC inhibition occurs only when the ETC is inhibited during L3/L4 development [7], and thus coincides with mitochondrial biogenesis [19]. The levels of mtDNA are also increased in the isp-1 and clk-1 mitochondrial mutants, consistent with an increase in mitochondrial biogenesis [6,8]. Thus it has been postulated that mitochondrial biogenesis might be a significant factor underlying lifespan extension in response to ETC deficits in C. elegans [5].

Nrf2 transcription factor is an integral antioxidant signalling mechanism. Under basal conditions Nrf2 is rapidly degraded by the proteasome. However, following oxidative stress, Nrf2 localizes to the nucleus where it binds the conserved ARE (antioxidant response element) DNA sequence. Binding of Nrf2 to the ARE results in the up-regulation of many phase I and phase II detoxifying enzymes as well as antioxidants such as glutathione transferases, Prdxs (peroxiredoxins), Trxs (thioredoxins) and HO-1 (haem oxygenase 1) [20].

To test whether mitochondrial compensatory responses are up-regulated in the Surf1−/− mice, we measured mitochondrial function and changes in mitochondrial stress responses (UPRMT, mitochondrial proliferation and Nrf2) in two highly energetic tissues, heart and skeletal muscle, in wild-type and long-lived Surf1−/− mice. We hypothesized that mechanisms associated with longevity in invertebrates might also be induced in a mammalian model of reduced ETC activity. Furthermore, we investigated in vivo physiological changes that result from loss of functional SURF1.

MATERIALS AND METHODS

Animals

All experiments were performed with the approval by the IACUC (Institutional Animal Care and Use Committee) at the University of Texas Health Science Center at San Antonio. Surf1−/− mice, generated as described previously [2], were bred from Surf1+/− heterozygous crosses in a B6D2F1/J (C57/Bl6J×DBA2) background. All wild-type animals were littermate controls of the Surf1−/− animals. Male mice aged 5–7 months were used for all experiments and killed using CO2 asphyxiation.

Mitochondrial isolation

Heart and hind-limb skeletal muscle mitochondria were isolated using differential centrifugation as described previously [21]. Heart and hind-limb skeletal muscles were removed, rinsed and minced in Chappell–Perry Buffer I [100 mM KCl, 50 mM Tris/HCl, 5 mM MgCl2 and 1 mM EDTA (pH 7.2)] with 1 mM ATP (Grade II, Sigma) and 1.5 mg of protease (Type I: crude, from bovine pancreas, Sigma) per 0.5 g of tissue. The minced tissue was placed on a shaker for 10 min and then homogenized. The homogenate was spun at 600 g for 10 min. The supernatant was filtered through a cheesecloth followed by centrifugation at 14000 g for 10 min. The supernatant was discarded and the pellet was resuspended in Chappell–Perry Buffer II [100 mM KCl, 50 mM Tris/HCl, 1 mM MgCl2 and 0.2 mM EDTA (pH 7.2)] with 0.2 mM ATP and BSA (100 mg/100 ml, Sigma) and spun at 7000 g for 10 min. The pellet was resuspended in Chappell–Perry Buffer II with ATP and spun twice at 3500 g. The final pellet was used for all mitochondrial assays.

Complex activity assays

The final mitochondrial pellet was resuspended in ACA/BT buffer [750 mM 6-aminocaprioic acid and 50 mM BisTris (pH 7.0)] plus 1% n-dodecylmaltoside and 1×protease inhibitor (Cocktail set III, Calbiochem) for 45 min with constant agitation at 4°C. The suspension was then spun at 100000 g for 15 min at 4°C. The protein concentration in the supernatant was measured using the Bradford method and then used for the complex activity assays as described previously [22].

Complex I activity assay

Complex I activity was measured by monitoring the oxidation of NADH with a spectrophotometer at 340 nm with ubiquinone-2 as the electron acceptor in the presence of DCIP (dichlorophenol-indophenol) as described previously [22]. To perform the assay, 1 ml of respiration buffer [250 mM sucrose, 10 mM KH2PO4, 1 mM EGTA and 10 mM Tris (pH 7.4)] was placed into a polystyrene cuvette with 100 μM NADH, 50 μM DCIP, 2 mM KCN and 2 μM antimycin A. After setting up the blank, 50 μM ubiquinone-2 was added followed by 20 μg of mitochondrial protein, and the rate of change in absorbance was measured for 1 min (dA/min). As a negative control, complex I acti-vity was inhibited with the addition of 10 μM rotenone and the rate was subtracted from the rate of NADH oxidation. The final rate (dA/min) was converted into mM/min per mg by dividing dA/min by 6.22 mM−1·cm (molar absorption coefficient for NADH) and then divided by the amount of mitochondrial protein. The final rate was normalized to wild-type control values.

Complex II activity assay

Complex II activity was measured by succinate-dependent reduction of DCIP at 600 nm using ubiquione-2 as an electron acceptor as described previously [22]. To perform the assay, 1 ml of respiration buffer [250 mM sucrose, 10 mM KH2PO4, 1 mM EGTA and 10 mM Tris (pH 7.4)] was placed into a polystyrene cuvette with 20 mM succinate, 50 μM DCIP, 2 mM KCN and 2 μM antimycin A. After setting up the blank, 50 μM ubiquinone-2 (Sigma) was added followed by 40 μg of mitochondrial protein and the rate of change in absorbance was measured for 1 min. As a negative control, activity of complex II was inhibited by the addition of 2 mM malonate and the rate was subtracted from the rate of DCIP reduction from complex II activity. The final rate (dA/min) was converted into mM/min per mg by dividing dA/min by 21 mM−1·cm (molar absorption coefficient for succinate) and then divided by the amount of mitochondrial protein. The final rate was normalized to wild-type control values.

Complex III activity assay

Complex III activity was measured by the reduction of cytochrome c3+ at 550 nm using D-ubiquinol-2 as an electron acceptor as described previously [22]. Reduced ubiquinol was prepared as follows. A 100 μl aliquot of ubiquinone (20 mM in ethanol) was reduced with a pinch of sodium borohydrate (Sigma). After mixing, 100 μl of ethylene glycol was added to stop the reduction of ubiquinone. A 2 μl aliquot of 12 M HCl was added and the mixture was spun to separate the sodium borohydrate from the reduced ubiquinol-2. To perform the assay, 1 ml of respiration buffer [250 mM sucrose, 10 mM KH2PO4, 1 mM EGTA and 10 mM Tris (pH 7.4)] was placed into a polystyrene cuvette with 10 μM rotenone and 2 mM KCN. After setting up the blank, 100 μM D-ubiquinol-2 was added followed by 10 μg of mitochondrial protein and the rate of change in absorbance was measured for 1 min. As a negative control 2 μM antimycin A was added to inhibit the activity of complex III and the rate was subtracted from the rate of cytochrome c oxidation. The final rate (dA/min) was converted into mM/min per mg by dividing dA/min by 21 mM−1·cm (molar absorption coefficient for cytochrome c) and then divided by the amount of mitochondrial protein. The final rate was normalized to wild-type control values.

Complex IV activity assay

Complex IV activity was measured by monitoring the oxidation of cytochrome c2+ at 550 nm using a spectrophotometer as described previously [22]. Reduced cytochrome c2+ was prepared as follows. A 500 μl aliquot of 8 mM cytochrome c3+ (Sigma) was reduced with sodium borohydrate (Sigma). To perform the assay, 1 ml of respiration buffer [250 mM sucrose, 10 mM KH2PO4, 1 mM EGTA and 10 mM Tris (pH 7.4)] was placed into a polystyrene cuvette with 40 μM cytochrome c2+. After setting up the blank, 5 μg of mitochondrial protein was added, and rate of cytochrome c2+oxidation was measured at 550 nm. As a negative control, complex IV activity was inhibited by the addition of 2 mM KCN and the rate of change was then subtracted from the rate of cytochrome c2+ oxidation (dA/min). The rate was converted into mM/min per mg by dividing dA/min by 21 mM−1·cm (molar absorption coefficient for cytochrome c) and then divided by the amount of mitochondrial protein. The final rate was normalized to wild-type control values.

Mitochondrial respiration

Mitochondrial oxygen consumption was measured using a Clark electrode (Oxytherm Oxygen Electrode Control Unit, Hansatech Instruments) as described previously [23,24]. Briefly, mitochondria were resuspended in respiration buffer [250 mM sucrose, 10 mM KH2PO4, 1 mM EGTA and 10 mM Tris (pH 7.4)] with 0.3% BSA; glutamate/malate (5 mM) was used as the respiratory substrate. State 3 respiration was initiated with the addition of 0.3 mM ADP. State 4 respiration was measured as oxygen consumption following the consumption of ADP. RCR (respiratory control ratio) was measured as state 3/state 4 respiration rates.

ATP production

ATP production was measured in freshly isolated heart or skeletal muscle mitochondria using the ATP Bioluminescence Assay Kit CLSII (Roche, catalogue number 1699695) and a fluorescence microplate reader at 562 nm as described previously [22]. Freshly isolated mitochondria were resuspended in ROS (reactive oxygen species) buffer [125 mM KCl, 10 mM Hepes, 5 mM MgCl2 and 2 mM K2HPO4 (pH 7.44)] to a final concentration of 0.04 μg/μl. In a solid white 96-well plate, 100 μl aliquots of ROS buffer with mitochondria per well was added. The complex II-linked substrates used were succinate (5 mM) and rotenone (5 nM). Complex I-linked substrates glutamate (2.5 mM) and malate (2.5 mM) for skeletal muscle mitochondria and pyruvate (2.5 mM) and malate (2.5 mM) for heart mitochondria were used. To each well, 100 μl of luciferase reagent containing 0.3 mM ADP was added and the kinetic luminescence was determined. The results were then calculated from a standard curve generated with an ATP standard provided by the manufacturer (Roche).

Membrane potential

Membrane potential was determined as described previously [25]. Mitochondria were resuspended in ROS buffer [125 mM KCl, 10 mM Hepes, 5 mM MgCl2 and 2 mM K2HPO4 (pH 7.44)] at a concentration of 0.05 μg/μl with 5 μM Safranin O (ICN Biomedical) and 100 μl was aliquoted into a black flat-bottomed 96-well microplate. Plates with complete buffer and mitochondria were incubated at 37°C for 5 min before the addition of substrates. For complex I-linked substrates in skeletal muscle, glutamate (5 mM) and malate (5 mM) were used. For complex I-linked substrates in heart, pyruvate (5 mM) and malate (5 mM) were used. For complex II-linked substrates succinate (10 mM) and rotenone (1 μM) were used. Fluorescence of the quench-dye Safranin O was measured using excitation at 485 nm and emission at 590 nm using a microplate reader, and wells with only ROS buffer and Safranin O were used as negative controls.

H2O2 production assay

Mitochondrial release of H2O2 was measured as described previously [26]. In the presence of horseradish peroxidase, the Amplex Red reagent reacts with H2O2 to produce the red fluorescent oxidation product resorufin. The fluorescence of resorufin was measured with a fluorescence microplate reader using excitation at 544 nm and emission at 590 nm. Isolated mitochondria were diluted in Amplex Red reagent buffer [77.8 μM Amplex Red (Invitrogen, catalogue number a22188), 37.5 units/ml SOD (superoxide dismutase) and 1 unit/ml horseradish peroxidase in ROS buffer] to a final concentration of 0.4 μg/μl. A 100 μl aliquot of diluted mitochondria were used per reaction. Resorufin fluorescence was measured following the addition of succinate (5 mM) and rotenone (5 nM) or glutamate (2.5 mM) and malate (2.5 mM). H2O2 production was determined by comparing the value to an Amplex Red–H2O2 standard curve.

Superoxide production

Extramitochondrial superoxide release was measured by EPR using the spin trap DIPPMPO [5-(di-isopropoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide, Alexis Biochemicals] as described prevously [27]. DIPPMPO forms an adduct with superoxide, resulting in the generation of DIPPMPO-OOH, which decays to the DIPPMPO-OH adduct by the action of glutathione peroxidases. EPR measurements were performed using an X-band MS200 spectrometer (Magnetech). Mitochondria (1 μg/μl) were incubated at 37°C with DIPPMPO (50 mM) for 30 min in 125 mM KCl, 10 mM MOPS, 2 mM diethylene triamine penta-acetic acid, 5 mM MgCl2 and 2 mM K2HPO4 (pH 7.44) in the presence of succinate (24 mM), rotenone (2.4 μM) and antimycin A. These substrates were used to drive respiration starting at complex II. Although it did not affect the DIPPMPO-OH signal, low levels of catalase (10 units/ml) were also added, to prevent the appearance of small additional and unidentified spectrum peaks after extended incubation. For measurements, 40 μl of sample was transferred to 50 μl capillary tubes and these were measured at room temperature with the following settings: receiver gain, 5×105; microwave power, 20 mW; microwave frequency, 9.55 GHz; modulation amplitude, 2 G; scan time, 40 s; and scan width, 100 G, with an accumulation of ten scans. EPR data are expressed as RIU (relative intensity units)/20 μg of mitochondrial protein. As a negative control, measurements were also made in the presence of SOD.

Electron microscopy

Heart and hind-limb skeletal muscle were excised and fixed in phosphate-buffered 4% formaldehyde with 1% glutaraldehyde. The samples were rinsed in 0.1 M phosphate buffer and then post-fixed with 1% Zetterqvist's buffered osmium tetroxide for 30 min. The samples were dehydrated in 70% alcohol for 10 min and then in 95% alcohol for 10 min. The samples were then twice dehydrated in 100% alcohol for 10 min and then twice dehydrated in propylene oxide for 10 min, for each dehydration. The samples were resin-infiltrated first in a 1:1 propylene oxide/resin mix for 30 min and then in 100% resin for 30 min under 25 psi (1 psi=6.9 kPa) vacuum. The samples were then flat-embedded in a BEEM capsule and filled to the top and polymerized at 85°C for 90 min. The embedded samples were then sectioned and stained with a drop of uranyl acetate and microwaved for 30 s. The section was then gently dipped into distilled water and placed on filter paper. A drop of Reynold's lead citrate stain was then placed on the section and microwaved for 20 s and then rinsed in distilled water. The section was then photographed at 20000× using a JEOL 1230 transmission electron microscope with AMT imaging software. Five randomly chosen images per animal (n=3) were quantified for mitochondrial number and area using NIS-Elements BR imaging software.

Western blot analysis

Heart and gastrocnemius skeletal muscle were homogenized in buffer containing 50 mM Hepes, 150 mM NaCl, 2 mM EDTA, 10% glycerol, 1% Igepal, 1 mM MgCl2, 1 mM CaCl2 and 1×protease inhibitor. Homogenized samples were spun at 15000 g for 10 min at 4°C and supernatant was collected. Protein concentrations were measured using the Bradford method and diluted to equal concentrations. Samples were then diluted in a 1:1 ratio in Laemmli sample buffer (Bio-Rad Laboratories) with 50 μl of 2-mercaptoethanol per 950 μl of Laemmli sample buffer and boiled for 10 min. The final samples were then run on a 4–20% gradient Criterion™ XT precast gel (Bio-Rad Laboratories) with 1×Tris/glycine/SDS buffer. Gels were transferred on to PVDF membrane using a Trans-Blot SD Semi-Dry Transfer Cell (Bio-Rad Laboratories) in Tris/glycine/SDS buffer containing 20% methanol. Blots were blocked for 1 h in 1% BSA in TBS-T [Tris-buffered saline buffer (50 mM Tris/HCl, pH 7.5 and 150 mM NaCl) containing 0.05% Tween 20]. Primary antibodies used were: anti-Hsp60 (StressGen), anti-ClpP (Sigma), anti-(Lon protease) (Dr Luke Szweda, Oklahoma Medical Research Foundation, Oklahoma City, OK, U.S.A.), anti-Trx2 (AbCam), anti-CHOP (Cell Signaling Technology), anti-PGC-1α (AbCam), anti-TFAM (mitochondrial transcription factor A, Sigma), anti-(complex II 70 kDa subunit) (Molecular Probes), anti-VDAC (voltage-dependent anion channel, Cell Signaling Technology), anti-(cytochrome c) (Cell Signaling Technology), anti-Nrf2 (Dr Scott Plafker, Oklahoma Medical Research Foundation, Oklahoma City, OK, U.S.A.) and anti-HO-1 (StressGen). Secondary antibodies used were from Vector Laboratories. Western blots were imaged using premium X-ray film from Phenix Research Products.

Quantitative real-time PCR

Total RNA was extracted from 50 mg of flash-frozen hind-limb skeletal muscle and cardiac tissue using the RNeasy Plus kit (Qiagen, catalogue number 73442). SuperScript II reverse transcriptase and random hexamer primers were used for first-strand cDNA synthesis according to the manufacturer's instructions (Life Technologies, catalogue number 18064071). Quantitative real-time PCR was performed with an ABI Prism machine using Power SYBR Green PCR Master Mix (Applied Biosystems, catalogue number 4367659). The calculations were performed using the comparative method (2−ΔΔCT) and the following primers were used: PGC1α, 5′-CGGAAATCATATCCAACCAG-3′ (forward) and 5′-TGAGGACCGCTAGCAAGTTTG-3′ (reverse); and γ-actin, 5′-GCCGCCGCGTCCTT-3′ (forward) and 5′-ATGACGAGTGCGGCGATT-3′ (reverse).

Open-field activity monitoring

Open-field activity monitoring is used to determine total activity as described prevously [28]. Mice are placed in a large clear cage (16 inches×9 inches×5.5 inches) with a grid size (7×15) for 40 h with corncob bedding and free access to food and water. The mice are habituated to the apparatus for 16 h and data are recorded during the subsequent 24 h to assess activity. The number of beam breaks across the X–Y axis is measured to determine total activity (Kinder Scientific and Software from MotoMonitor). These values were summed during the 12-h light and 12-h dark cycle for each mouse.

Endurance capacity

Treadmill running was used to measure endurance capacity as described previously [26] using a mouse treadmill (Columbus Instruments). Following an acclimation period of 10 min without treadmill movement, the treadmill was started at 7 m/min for 5 min. The speed was increased to 12 m/min for no longer than 2.5 h. Animals were removed from the treadmill when they fell on to the encouragement platform and would not engage the treadmill with physical prodding. The experiment was performed by an individual blinded to the genotype of the animals.

Lactate measurements

Blood lactate was measured using the Lactate Plus lactate meter (NOVA Biomedical) with blood collected from the tail as described previously [26].

Grip strength

A chatillion force gauge (Ametek) connected to a wire mesh was used to measure the grip strength of the animals. Each animal was grabbed by the tail and allowed to grip the wire mesh, once grip was established the animal was pulled back by its tail until the grip was lost. This was repeated five times per animal and the maximum grip strength was recorded. The experimenter was blinded to the genotypes of the animal.

2D Echocardiography

Cardiac function was assessed as described previously [29]. Mice were anaesthetized with isofluorane (0.5–2% in a 100% oxygen mix) and placed on to an isothermal heat pad. Echocardiograms were performed using a Vevo 770™ High-Resolution In Vivo Imaging System (Visual Sonics), with heart rate continuously monitored throughout the experiment. Mice were given intraperitoneal injections of dobutamine (3 μg/kg of body weight) and 2D echocardiography was performed before injection of dobutamine, and at 10 and 30 min following injection.

Cardiac glucose uptake

Cardiac glucose uptake was assessed as described in [30]. Mice were anaesthetized under 1.2% isofluorane throughout the measurements. 18FDG (2-[18F]fluoro-2-deoxy-D-glucose) (0.5 mCi) dissolved in 1 ml of physiological saline solution was injected through the tail vein. Before scanning, 40 min of 18FDG uptake was allowed. The animal was then moved to the scanner bed (Focus 220 MicroPET, Siemens) and placed in the prone position. Emission data were acquired for 20 min in a 3D list mode with intrinsic resolution of 1.5 mm. For image reconstruction, 3D PET (positron-emission tomography) data were rebinned into multiple frames of 1 s duration using a Fourier algorithm. After rebinning the data, a 3D image was reconstructed for each frame using a 2D filtered back projection algorithm. Decay and dead-time corrections were applied to the reconstruction process. Because 18FDG is taken up by the whole body of the animal, we chose an ROI (region of interest) for our measurements that encompassed the whole heart, as outlined with a white line in Figure 7, and determined CMRGlc (cardiac metabolic rate of glucose) for whole heart using the mean SUV (standardized uptake value) equation:

 
formula

where A is the activity of the brain, W is the body weight of the mouse and Ainj is the injection dose of 18FDG.

RESULTS

COX activity is decreased in skeletal muscle and heart of Surf1−/− mice

To confirm the effect of loss of the SURF1 protein on ETC complex activities, mitochondria were isolated from heart and hind-limb skeletal muscle (gastrocnemius) and the enzymatic activities of complexes I, II, III and IV were measured. Compared with wild-type control littermates, loss of SURF1 resulted in a 71% and 53% decrease in COX activity in heart (Figure 1A) and skeletal muscle (Figure 1B) respectively. The activities of the other ETC complexes were not affected (Figure 1).

Complex IV activity is specifically affected in Surf1−/− heart and skeletal muscle

Figure 1
Complex IV activity is specifically affected in Surf1−/− heart and skeletal muscle

(A) Enzymatic activity of complexes I, II, III and IV in isolated heart mitochondria from wild-type (white bars) and Surf1−/− (grey bars) mice. Values are normalized to the wild-type control for each assay. (B) Enzymatic activity of complexes I, II, III, and IV in isolated skeletal muscle (SkM) mitochondria from wild-type (white bars) and Surf1−/− (grey bars) mice. n=5 per experimental group, data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05. WT, wild-type.

Figure 1
Complex IV activity is specifically affected in Surf1−/− heart and skeletal muscle

(A) Enzymatic activity of complexes I, II, III and IV in isolated heart mitochondria from wild-type (white bars) and Surf1−/− (grey bars) mice. Values are normalized to the wild-type control for each assay. (B) Enzymatic activity of complexes I, II, III, and IV in isolated skeletal muscle (SkM) mitochondria from wild-type (white bars) and Surf1−/− (grey bars) mice. n=5 per experimental group, data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05. WT, wild-type.

State 3 respiration is decreased in heart, but not skeletal muscle, in mitochondria isolated from Surf1−/− mice

To determine the effect of lessened complex IV activity on mitochondrial respiration, we isolated mitochondria from heart and hind-limb skeletal muscle (gastrocnemius) of Surf1−/− mice and measured mitochondrial state 3 and state 4 respiration using a Clark-type electrode. Although state 3 respiration was modestly decreased in heart mitochondria (−16%) from Surf1−/− mice, it remained unchanged in skeletal muscle mitochondria compared with wild-type controls (Table 1). There was no significant difference in state 4 respiration between Surf1−/− and wild-type mice in either tissue. RCR, the ratio of state 3 to state 4 respiration and a measure of coupling of oxygen consumption to phosphorylation, was not affected between Surf1−/− and wild-type mice in heart or skeletal muscle mitochondria (Table 1). Additionally, the ADP to oxygen ratio (P:O ratio) was not significantly different between Surf1−/− and wild-type (Table 1). Taken together these data suggest that there is no difference in the coupling efficiency of the mitochondria, despite a decrease in state 3 respiration in heart mitochondria.

Table 1
Mitochondrial respiration in heart and skeletal muscle from Surf1−/− and wild-type mice

State 3 respiration, state 4 respiration, RCR and ADP to oxygen ratio (P:O) were measured in isolated heart or hind-limb skeletal muscle mitochondria. n=6–10 per experimental group, data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05.

Tissue Genotype State 3 State 4 RCR (state3/state4) ADP:O 
Heart Wild-type 325±27 58±5 5.3±0.5 1.98±0.2 
 Surf1−/− 264±17 * 59±6 4.8±0.6 2.10±0.2 
Skeletal muscle Wild-type 548±58 53±3 10.5±1.3 3.38±0.4 
 Surf1−/− 527±44 52±3 10.7±1.6 2.93±0.4 
Tissue Genotype State 3 State 4 RCR (state3/state4) ADP:O 
Heart Wild-type 325±27 58±5 5.3±0.5 1.98±0.2 
 Surf1−/− 264±17 * 59±6 4.8±0.6 2.10±0.2 
Skeletal muscle Wild-type 548±58 53±3 10.5±1.3 3.38±0.4 
 Surf1−/− 527±44 52±3 10.7±1.6 2.93±0.4 

ATP production is unaffected by COX deficiency in isolated mitochondria from heart and skeletal muscle in Surf1−/− mice

A major function of the ETC is production of ATP. Because Surf1−/− mice have a deficit in COX activity, we hypothesized this might result in a decrease in ATP production. To measure ATP production we used a luciferase-based assay with complex I- or complex II-linked substrates. Surprisingly, despite the reduction in COX activity, no significant difference in ATP production was found in isolated heart or skeletal muscle mitochondrial preparations from Surf1−/− mice compared with wild-type mice (Figures 2A and 2B). These data are in agreement with our recently published work showing no significant difference in ATP production in isolated brain mitochondria in vitro and no difference in ATP levels in vivo of Surf1−/− brain [30].

Mild reduction of membrane potential in Surf1−/− heart, but not skeletal muscle, mitochondria and there is no effect on ATP production in either tissue

Figure 2
Mild reduction of membrane potential in Surf1−/− heart, but not skeletal muscle, mitochondria and there is no effect on ATP production in either tissue

(A) ATP production in isolated heart mitochondria using either complex I-linked (pyruvate+malate) or complex II-linked (succinate+rotenone) substrates. (B) ATP production in isolated skeletal muscle mitochondria using either complex I-linked (glutamate+malate) or complex II-linked (succinate+rotenone) substrates. (C) Membrane potential was measured by the change in Safranin O fluorescence using either complex I-linked (pyruvate+malate) or complex II-linked (succinate+rotenone) substrates in isolated heart mitochondria. Data are shown as fluorescent units ×1000. (D) Membrane potential was measured by the change in Safranin O fluorescence using either complex I-linked (glutamate+malate) or complex II-linked (succinate+rotenone) substrates in isolated skeletal muscle mitochondria. n=5–10 per experimental group, data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05. FLU, fluorescence units; Glut, glutamate; Mal, malate; Pyr, pyruvate; Rot, rotenone; SkM, skeletal muscle; Suc, succinate.

Figure 2
Mild reduction of membrane potential in Surf1−/− heart, but not skeletal muscle, mitochondria and there is no effect on ATP production in either tissue

(A) ATP production in isolated heart mitochondria using either complex I-linked (pyruvate+malate) or complex II-linked (succinate+rotenone) substrates. (B) ATP production in isolated skeletal muscle mitochondria using either complex I-linked (glutamate+malate) or complex II-linked (succinate+rotenone) substrates. (C) Membrane potential was measured by the change in Safranin O fluorescence using either complex I-linked (pyruvate+malate) or complex II-linked (succinate+rotenone) substrates in isolated heart mitochondria. Data are shown as fluorescent units ×1000. (D) Membrane potential was measured by the change in Safranin O fluorescence using either complex I-linked (glutamate+malate) or complex II-linked (succinate+rotenone) substrates in isolated skeletal muscle mitochondria. n=5–10 per experimental group, data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05. FLU, fluorescence units; Glut, glutamate; Mal, malate; Pyr, pyruvate; Rot, rotenone; SkM, skeletal muscle; Suc, succinate.

Membrane potential is decreased in heart, but not skeletal muscle, mitochondria isolated from Surf1−/− mice

The ETC complexes I, III and IV pump protons from the mitochondrial matrix into the intermembrane space, creating an electrochemical gradient termed the membrane potential. This gradient is important for ATP production, mitochondrial calcium buffering and protein import into the mitochondria. A previous study using primary neuron cultures from Surf1−/− and control mice did not find any difference in the membrane potential [2]. However, the membrane potential was not measured in other tissues or in isolated mitochondria. We measured membrane potential in heart and skeletal muscle mitochondria using the cationic fluorescent probe Safranin O that is accumulated and quenched inside energized mitochondria [31]. Membrane potential was significantly lower (−18%) in isolated heart mitochondria from Surf1−/− mice compared with wild-type mice (Figures 2C and 2D). This difference was observed using both complex I-linked substrates (pyruvate+malate) and complex II-linked substrates (succinate+rotenone) in heart mitochondria. However, no difference in membrane potential was found in skeletal muscle mitochondria isolated from Surf1−/− and wild-type mice in the presence of complex I-linked substrates (glutamate+malate) or complex II-linked substrates (succinate+rotenone).

Mitochondrial ROS generation is not increased in heart or skeletal muscle mitochondria from Surf1−/− mice

Generation of ROS from mitochondrial ETC complexes I and III can damage proteins, lipids and DNA and is thought to contribute to a number of deleterious phenotypes including aging [32]. We measured H2O2 and superoxide release from isolated mitochondria using the Amplex Red assay and EPR with the spin trap DIPPMPO respectively. In heart and skeletal muscle mitochondria, H2O2 production in mitochondria respiring on complex I-linked substrates (pyruvate+malate or glutamate+malate) was not different between Surf1−/− and wild-type mice. However, with complex II-linked substrates and inhibition of reverse electron transfer through complex I using rotenone (succinate+rotenone), both heart and skeletal muscle mitochondria from Surf1−/− mice showed a significant decrease (−23% and −7% respectively) in H2O2 production compared with mitochondria from wild-type mice (Figures 3A and 3B).

ROS production is not increased in mitochondria isolated from Surf1−/− mice

Figure 3
ROS production is not increased in mitochondria isolated from Surf1−/− mice

For all histograms, white bars indicate wild-type and grey bars indicate Surf1−/− mice, n=6–10 for all assays. (A) H2O2 production in isolated heart mitochondria assayed with complex I-linked (pyruvate+malate) or complex II-linked (succinate+rotenone) substrates. (B) H2O2 production in isolated hind-limb skeletal muscle mitochondria with complex I-linked (glutamate+malate) or complex II-linked substrates. (C) Maximal superoxide production in isolated heart and skeletal muscle mitochondria. Data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05. AU, absorbance unit; Glut, glutamate; Mal, malate; Pyr, pyruvate; Rot, rotenone; SkM, skeletal muscle; Suc, succinate.

Figure 3
ROS production is not increased in mitochondria isolated from Surf1−/− mice

For all histograms, white bars indicate wild-type and grey bars indicate Surf1−/− mice, n=6–10 for all assays. (A) H2O2 production in isolated heart mitochondria assayed with complex I-linked (pyruvate+malate) or complex II-linked (succinate+rotenone) substrates. (B) H2O2 production in isolated hind-limb skeletal muscle mitochondria with complex I-linked (glutamate+malate) or complex II-linked substrates. (C) Maximal superoxide production in isolated heart and skeletal muscle mitochondria. Data are means±S.E.M. and significance was determined by two-tailed t test, *P<0.05. AU, absorbance unit; Glut, glutamate; Mal, malate; Pyr, pyruvate; Rot, rotenone; SkM, skeletal muscle; Suc, succinate.

We also measured maximal superoxide production using complex II-linked substrate and inhibitors (succinate+rotenone+antimycin A) using EPR (Figure 3C). This assay allows specific measurement of superoxide anion release from isolated mitochondria. Consistent with the observations from the H2O2 production assays, Surf1−/− heart mitochondria showed a significant decrease (−20%) in extra-mitochondrial superoxide release compared with wild-type mitochondria. However, superoxide release did not differ in skeletal muscle mitochondria from Surf1−/− and wild-type mice.

Surf1−/− mice have an increase in mitochondrial number in heart and skeletal muscle

PGC-1α is a transcriptional co-activator that is an important regulator of mitochondrial biogenesis that responds to a variety of stimuli, including calorie restriction or exercise [18]. We hypothesized that decreased complex IV activity might induce mitochondrial biogenesis to compensate for deficits in mitochondrial function. To test this hypothesis, we determined whether PGC-1α levels were changed in Surf1−/− mice. We have previously shown that adipose tissue from Surf1−/− mice displays significantly increased levels of PGC-1α and increased mitochondrial biogenesis [33]. In the present study we found that PGC-1α protein is significantly up-regulated in heart (66%) and skeletal muscle (2.3-fold) of Surf1−/− mice compared with littermate controls (Figures 4C and 4D). The mRNA level of PGC-1α was significantly elevated in skeletal muscle (1.1-fold), but unchanged in the heart (Figure 4E). The mitochondrial biogenesis marker TFAM was increased in skeletal muscle (1.1-fold) but unchanged in heart. In addition, succinate dehydrogenase (complex II, 70 kDa subunit) in the inner mitochondrial membrane was significantly elevated in heart (43%) and skeletal muscle (70%). VDAC in the outer mitochondrial membrane was increased in heart (2.6-fold) and in skeletal muscle (1.7-fold), further supporting an increase in mitochondrial number. In heart, we used EM to confirm the changes in mitochondrial biogenesis (Figure 4A). We found a 10% elevation in number and a 53% increase in total area of mitochondria from Surf1−/− heart EM sections (Figure 4B).

Mitochondrial number is increased in Surf1−/− mice heart and skeletal muscle

Figure 4
Mitochondrial number is increased in Surf1−/− mice heart and skeletal muscle

(A) Representative images of an electron micrograph from heart and skeletal muscle from wild-type and Surf1−/− mice. n=3, images shown are ×20000 magnification. (B) Quantification of mitochondrial size and number of five randomly chosen images per animal (n=3). (C and D) PGC-1α, TFAM, porin (VDAC), complex II (70 kDa subunit) and cytochrome c protein levels from wild-type and Surf1−/− heart (C) and skeletal muscle (D) tissue homogenates. Data are means±S.E.M. of relative density relative to the β-tubulin loading control. Quantification was done using NIH ImageJ software, n=6. (E) Quantitative real-time PCR of PGC-1α expressed relative to γ-actin, n=7. Significance was determined by two-tailed t test, *P<0.05. CII, complex II; SkM, skeletal muscle; WT, wild-type.

Figure 4
Mitochondrial number is increased in Surf1−/− mice heart and skeletal muscle

(A) Representative images of an electron micrograph from heart and skeletal muscle from wild-type and Surf1−/− mice. n=3, images shown are ×20000 magnification. (B) Quantification of mitochondrial size and number of five randomly chosen images per animal (n=3). (C and D) PGC-1α, TFAM, porin (VDAC), complex II (70 kDa subunit) and cytochrome c protein levels from wild-type and Surf1−/− heart (C) and skeletal muscle (D) tissue homogenates. Data are means±S.E.M. of relative density relative to the β-tubulin loading control. Quantification was done using NIH ImageJ software, n=6. (E) Quantitative real-time PCR of PGC-1α expressed relative to γ-actin, n=7. Significance was determined by two-tailed t test, *P<0.05. CII, complex II; SkM, skeletal muscle; WT, wild-type.

Skeletal muscle from Surf1−/− mice exhibits elevated UPRMT-associated proteins

The UPRMT was shown to be required for mediating the longevity phenotype of the C. elegans mitochondrial mutants [5,34]. We hypothesized that the UPRMT may also be up-regulated in the Surf1−/− mice. To test our hypothesis we measured protein levels of UPRMT markers in heart and hind-limb skeletal muscle (Figures 5A and 5C). The markers used to measure UPRMT induction include Hsp60, ClpP, Lon protease, Trx2 and CHOP. In heart tissue homogenates Lon (77%) and Trx2 (54%) were significantly up-regulated, but Hsp60 or ClpP were not. CHOP trended towards an increase in Surf1−/− heart, but was not statistically significant (54%, P=0.1). However, in skeletal muscle homogenates, Hsp60 (51%,), ClpP (57%), Lon (29%) and CHOP (22%) were all found to be significantly up-regulated, consistent with the activation of UPRMT.

Induction of proteins involved in the UPRMT and Nrf2 pathway in heart and skeletal muscle from Surf1−/− mice

Figure 5
Induction of proteins involved in the UPRMT and Nrf2 pathway in heart and skeletal muscle from Surf1−/− mice

Representative Western blots and quantification of Western blots of UPRMT-associated proteins (A and C) (Hsp60, ClpP, Lon protease, Trx2 and CHOP) or Nrf2 and HO-1 (B and D). White bars indicate values from wild-type mice and grey bars indicate values from Surf1−/− mice for heart (A and B) and skeletal muscle (C and D) tissue homogenates, n=4–6. Data are means±S.E.M. of relative densities of each protein relative to the β-tubulin loading control. Quantification was done using NIH ImageJ software and significance was determined by two-tailed t test, *P<0.05. WT, wild-type.

Figure 5
Induction of proteins involved in the UPRMT and Nrf2 pathway in heart and skeletal muscle from Surf1−/− mice

Representative Western blots and quantification of Western blots of UPRMT-associated proteins (A and C) (Hsp60, ClpP, Lon protease, Trx2 and CHOP) or Nrf2 and HO-1 (B and D). White bars indicate values from wild-type mice and grey bars indicate values from Surf1−/− mice for heart (A and B) and skeletal muscle (C and D) tissue homogenates, n=4–6. Data are means±S.E.M. of relative densities of each protein relative to the β-tubulin loading control. Quantification was done using NIH ImageJ software and significance was determined by two-tailed t test, *P<0.05. WT, wild-type.

Surf1−/− heart does not show an induction in the UPRMT, but does show elevated Nrf2 and HO-1 expression

Nrf2 nuclear localization and binding to the ARE results in the up-regulation of genes, including antioxidants and HO-1. Induction of HO-1 has previously been shown to be cardioprotective and anti-apoptotic following heart failure [35] and ischaemia/reperfusion injury [36]. In heart tissue homogenates from Surf1−/− mice there was a 68% increase in Nrf2 protein expression compared with wild-type. Consistent with increased Nrf2 expression, HO-1 (82%) was significantly increased in Surf1−/− heart (Figure 5B). However, in skeletal muscle homogenates, the expression of Nrf2 and HO-1 did not differ between wild-type and Surf1−/− mice (Figure 5D).

In Surf1−/− mice, basal spontaneous activity is not affected; however, endurance capacity is significantly decreased and associated with increased blood lactate in response to moderate exercise

In order to determine whether the decrease in COX activity resulted in physiological deficits, we measured both basal activity level and endurance capacity in wild-type and Surf1−/−mice. To measure basal activity level, open-field activity with laser break counts was used. The number of laser breaks was summed during the 12-h light and 12-h dark cycle separately (Figure 6A). No significant differences in cage activity were observed between Surf1−/− and wild-type mice in either the light or the dark cycle, suggesting that the Surf1 deletion does not affect the basal activity level.

Surf1−/− mice show no difference in basal activity but detriments in grip strength and endurance capacity associated with elevated blood lactate

Figure 6
Surf1−/− mice show no difference in basal activity but detriments in grip strength and endurance capacity associated with elevated blood lactate

(A) Activity of wild-type (white bars) and Surf1−/− (grey bars) mice measured during the light and dark cycle over a 24-h period. Activity was recorded as the number of laser breaks and summed for each animal, n=8. Data are means±S.E.M. (B) Endurance capacity in wild-type (●) and Surf1−/− (■) mice. Each point represents a single animal and the time removed due to exhaustion, up to 150 min. Data are significant (P=0.0007) as determined by the log-rank mantel-cox test; n=15–20. (C) Blood lactate levels following moderate exercise. Wild-type and Surf1−/− mice had blood lactate readings measured before treadmill running at 12 m/min and then measured immediately following 15 and 35 min of treadmill running; n=6–9. (D) Grip strength for wild-type and Surf1−/− animals in GF (grams force). For each animal, the value for grip strength was the best of five attempts, n=17 animals. Data are means±S.E.M. and are significant at each time point as measured by two-tailed t test, *P<0.05. WT, wild-type.

Figure 6
Surf1−/− mice show no difference in basal activity but detriments in grip strength and endurance capacity associated with elevated blood lactate

(A) Activity of wild-type (white bars) and Surf1−/− (grey bars) mice measured during the light and dark cycle over a 24-h period. Activity was recorded as the number of laser breaks and summed for each animal, n=8. Data are means±S.E.M. (B) Endurance capacity in wild-type (●) and Surf1−/− (■) mice. Each point represents a single animal and the time removed due to exhaustion, up to 150 min. Data are significant (P=0.0007) as determined by the log-rank mantel-cox test; n=15–20. (C) Blood lactate levels following moderate exercise. Wild-type and Surf1−/− mice had blood lactate readings measured before treadmill running at 12 m/min and then measured immediately following 15 and 35 min of treadmill running; n=6–9. (D) Grip strength for wild-type and Surf1−/− animals in GF (grams force). For each animal, the value for grip strength was the best of five attempts, n=17 animals. Data are means±S.E.M. and are significant at each time point as measured by two-tailed t test, *P<0.05. WT, wild-type.

To determine whether the decrease in COX activity would result in decreased exercise performance, we subjected wild-type and Surf1−/− mice to endurance exercise on a mouse treadmill (Figure 6B). Surf1−/− mice performed significantly worse compared with their wild-type littermates (P<0.001). This suggests that whereas lower COX activity does not affect basal activity of the animal, endurance capacity is significantly limited.

Surf1−/− mice were previously shown to have a significant increase in blood lactate under basal conditions [2]. Consistent with these findings, we found that Surf1−/− mice had a significant increase in blood lactate under basal conditions (28%). Following treadmill running for 15 and 35 min, the Surf1−/− mice had a significantly greater increase in blood lactate (15 min: 55%; 35 min: 72%) compared with wild-type mice (Figure 6C). In addition, Surf1−/− mice also showed a significant decrease in grip strength (−13%) (Figure 6D). Taken together these data support a limiting effect in vivo of mitochondrial function in the Surf1−/− mice in response to the reduction in COX activity.

Elevated glucose uptake and mitochondrial biogenesis may compensate for mitochondrial dysfunction and support normal cardiac function in Surf1−/− mice

Mitochondrial dysfunction in cardiac muscle has previously been shown to adversely affect cardiac function, namely endurance capacity. To determine whether the mild mitochondrial dysfunction in the Surf1−/− mice would adversely affect cardiac function under conditions of stress, we performed 2D echocardiography before and 10 and 30 min after intraperitoneal injection of dobutamine, a β-adrenergic receptor agonist that stresses the heart by elevating heart rate. Heart rate, end-diastolic dimension and fractional shortening, were measured in wild-type and Surf1−/− mice (Figures 7A–7C). We found no significant differences in any of these measurements between wild-type and Surf1−/− mice and all animals displayed a normal response to dobutamine stress. In addition, we found no significant differences in end-systolic dimension, septal wall thickness or posterior wall thickness (results not shown). This suggests that Surf1−/− mice have normal cardiac function despite in vitro decreases in COX activity, mitochondrial respiration and membrane potential in isolated mitochondria. To address this discrepancy, we measured in vivo glucose uptake in the heart of Surf1−/− and wild-type animals. Surf1−/− mice have a significant increase in glucose uptake as determined by PET scan (33%), suggesting a potential mechanism by which Surf1−/− mice hearts compensate for sub-optimal mitochondrial function (Figures 7D and 7E).

No difference in cardiac function between wild-type and Surf1−/− mice but enhanced glucose uptake in heart of Surf1−/− mice in vivo

Figure 7
No difference in cardiac function between wild-type and Surf1−/− mice but enhanced glucose uptake in heart of Surf1−/− mice in vivo

For all histograms, white bars indicate wild-type and grey bars indicate Surf1−/− mice. (A) Heart rate in beats/min before and 10 and 30 min following treatment with dobutamine, n=4–5. (B) End-diastolic dimension as measured by 2D echocardiography before and 10 and 30 min following dobutamine stress treatment. (C) Fractional shortening as measured by 2D echocardiography before and 10 and 30 min following treatment with dobutamine. (D) In vivo cardiac metabolic rate of glucose (CMRGlc) maps of representative wild-type and Surf1−/− mice obtained by PET. Circle indicates the location of the heart. Colour indicates rate of 18FDG uptake, n=6. (E) Quantification of in vivo metabolic glucose uptake. Data are means±S.E.M. Significance was determined by two-tailed t test, *P<0.05. SUV, standardized uptake value; WT, wild-type.

Figure 7
No difference in cardiac function between wild-type and Surf1−/− mice but enhanced glucose uptake in heart of Surf1−/− mice in vivo

For all histograms, white bars indicate wild-type and grey bars indicate Surf1−/− mice. (A) Heart rate in beats/min before and 10 and 30 min following treatment with dobutamine, n=4–5. (B) End-diastolic dimension as measured by 2D echocardiography before and 10 and 30 min following dobutamine stress treatment. (C) Fractional shortening as measured by 2D echocardiography before and 10 and 30 min following treatment with dobutamine. (D) In vivo cardiac metabolic rate of glucose (CMRGlc) maps of representative wild-type and Surf1−/− mice obtained by PET. Circle indicates the location of the heart. Colour indicates rate of 18FDG uptake, n=6. (E) Quantification of in vivo metabolic glucose uptake. Data are means±S.E.M. Significance was determined by two-tailed t test, *P<0.05. SUV, standardized uptake value; WT, wild-type.

DISCUSSION

The loss of the SURF1 complex IV assembly protein in mice was previously shown to lower complex IV assembly, lower complex IV content and cytochrome oxidase activity and was unexpectedly associated with an increased median lifespan (~20% in both males and females) [2]. Because it is well established in C. elegans that ETC inhibition, including inhibition of complex IV, can result in increased lifespan, we reasoned that Surf1−/− mice would provide an excellent opportunity to investigate potential mechanisms by which ETC inhibition might initiate protective mechanisms in a mammalian model. In the present paper, using mitochondria from two highly energetic tissues, heart and skeletal muscle, we report that loss of Surf1 leads to induction of mitochondrial stress response pathways, including mitochondrial biogenesis, the UPRMT and Nrf2 activation.

Mice with a truncated SURF1 protein were originally generated to study Leigh's syndrome, a human mitochondrial disorder that stems from loss-of-function mutations in the SURF1 gene and is characterized by neurological deficits and early lethality. Surprisingly, mice lacking the SURF1 protein lack any debilitating phenotype and paradoxically have an increase in lifespan. A key difference between human Leigh's syndrome and Surf1−/− mice is the relative decline in COX activity. Leigh's syndrome patients can have a greater than 90% loss of COX activity, whereas Surf1−/− mice have a less severe decline of COX activity ranging from 50 to 75% of wild-type values across a number of tissues. Although the Surf1−/− mice do show mild changes in mitochondrial function, we propose that a less severe reduction in complex IV activity might actually lead to induction of compensatory pathways that prevent the deleterious phenotypes present in Leigh's syndrome.

In light of the dramatic deleterious phenotypes of mitochondrial mutations in humans, we were surprised to find that the indices of mitochondrial function we measured in isolated mitochondria from Surf1−/− mice (e.g. generation of ROS, membrane potential, ATP production and respiration) failed to show dramatic changes, despite a substantial decline in complex IV activity. The fact that there was no reduction in ATP production is consistent with previous observations in a complex I mitochondrial mutant mouse model (Ndufs4−/− mice) that reported no change in ATP levels despite undetectable complex I activity in isolated liver mitochondria measured spectrophotometrically. Unlike Surf1−/− mice, the Ndufs4−/− mice exhibit a severe phenotype resulting in greatly reduced survival [37]. Although we did not find comprehensive mitochondrial dysfunction in the Surf1−/− mice, our results do show some mild alterations in mitochondria isolated from the heart in Surf1−/− mice, including a 16% decrease in state 3 respiration and a 19% decrease in membrane potential. Because of the critical role of complex IV in respiration, we hypothesized that a decrease in COX assembly and complex IV activity would result in a decline in oxygen consumption. The lower state 3 respiration in heart is consistent with our hypothesis and with a previous study in isolated brain mitochondria from Surf1−/− mice [30]. However, the in vitro decline in state 3 respiration may not be indicative of respiration levels in vivo, highlighted by the fact that there is no difference in whole-animal oxygen respiration of the Surf1−/− mice compared with wild-type [33]. Surprisingly, in skeletal muscle, indices of mitochondrial function from Surf1−/− mice were virtually indistinguishable from wild-type mice. The mechanism behind the differential responses in heart and skeletal muscle is still unclear, but could be the result of a threshold effect, as complex IV activity was reduced to a greater extent (>70% reduction) in heart than in skeletal muscle (~50%).

In skeletal muscle of Surf1−/− mice, decreased complex IV activity was associated with a significant decrease in grip strength but no change in muscle mass (results not shown), suggesting the muscles may be weaker. This phenotype is consistent with muscle weakness that occurs in humans with complex IV deficiency due to Surf1 mutations [1]. The Surf1−/− mice also show a pronounced reduction in endurance capacity that is associated with elevated levels of blood lactate before and after moderate exercise. Blood lactate during exercise is predominately produced from skeletal muscle and indicates a greater reliance on anaerobic glycolysis [38]. Surprisingly, despite a decrease in grip strength and endurance capacity in the Surf1−/− mice, there is no significant change in the overall basal activity level of the Surf1−/− mice, suggesting that the mitochondrial limitations are evident only under conditions of physiological challenge. These results and the data in heart mitochondrial function in vivo and in vitro in the Surf1−/− mice highlight the discrepancy between in vitro measures in isolated mitochondria and physiological parameters measured in vivo. Despite only mild alterations in Surf1−/− skeletal muscle mitochondrial function in vitro, exercise tolerance was clearly limited in vivo.

Our previous data suggest that the reduction in complex IV activity results in up-regulation of compensatory responses that are actually beneficial [30,33]. For example, in a recent study we reported that, in brain, loss of SURF1 leads to decreased respiration and increased ROS generation in isolated brain mitochondria. However, these mild deficits were associated with elevated brain glucose metabolism, increased cerebral blood flow and surprisingly enhanced memory [30]. We have also reported that loss of SURF1 leads to key metabolic changes resulting in reduced adiposity, increased insulin sensitivity and induction of mitochondrial biogenesis in adipose tissue [33]. These findings suggest that mild mitochondrial dysfunction from loss of SURF1 may in fact have positive effects overall, and is consistent with the mitohormesis hypothesis that suggests mild mitochondrial dysfunction could be beneficial because it results in compensatory responses that overcome the deleterious effects [39].

Consistent with this idea, the findings of the present study show that a number of stress response pathways, including mitochondrial biogenesis, the UPRMT and the Nrf2 pathway, pathways known to respond to mitochondrial stress, are elevated in Surf1−/− mice. For example, mitochondrial biogenesis has previously been shown to be elevated in response to energy stress conditions such as calorie restriction and endurance exercise in mice [18]. In C. elegans mitochondrial mutants, lifespan extension is limited to inhibition of the ETC during the L3/L4 stage of development [7] during which mitochondrial biogenesis occurs [19], supporting a potential link between mitochondrial biogenesis and lifespan extension. This is consistent with elevated levels of PGC-1α in response to caloric restriction [40] which is linked to increased lifespan in a number of species. Mitochondrial biogenesis in mice overexpressing PGC-1α specifically in muscle is also associated with changes in insulin sensitivity and increased lifespan, further supporting a role for mitochondrial biogenesis longevity [41,42]. In the Surf1−/− mice, markers of mitochondrial biogenesis are elevated in heart and skeletal muscle compared with wild-type mice. These data suggest that increasing the number of mitochondria is an important compensatory response to energy stress caused by SURF1 deficiency.

Nrf2 is a transcription factor that co-ordinates expression of the ~200 genes involved in drug detoxification, endogenous antioxidant production, NADPH regeneration and metabolism. Under basal conditions, Nrf2 is constitutively targeted for degradation by the proteasome. Following stress, however, Nrf2 binds the ARE resulting in the up-regulation of cytoprotective genes. Among the genes up-regulated by Nrf2 is HO-1, which catalyses the degradation of haem to ferrous iron, carbon monoxide and biliverdin. These products are important in mediating the antioxidant, anti-inflammatory and anti-apoptotic effects of HO-1 up-regulation. Overexpression of HO-1, specifically in heart, has been shown to be cardioprotective in response to ischaemia/reperfusion injury [36]. Because of the role of Nrf2 and HO-1 in antioxidant defence and cardioprotection, we asked whether Nrf2 and HO-1 were altered in Surf1−/− mice. Interestingly, protein levels of Nrf2 and HO-1 are increased in heart. The combined increase in mitochondrial number, HO-1 and 33% enhanced glucose uptake we found in heart may compensate for the decline in function of the individual mitochondrion and allows for normal heart physiological function.

Initiation of the UPRMT results in the up-regulation of mitochondrial-specific chaperones and proteases that aim to maintain mitochondrial proteostasis by refolding or degrading unfolded proteins. The UPRMT was shown to be required for the lifespan extension of the cco-1 complex IV mitochondrial mutant in C. elegans [5]. Furthermore, disruption of mitochondrial ribosomal protein S5 in C. elegans resulted in decreased mitochondrial respiration and induced the UPRMT [34]. This compensatory response was attributed to a mitonuclear protein imbalance and also resulted in enhanced longevity of C. elegans, and showed similar increases in mammalian cell culture. These data, along with the data from Surf1−/− animals, add further evidence to the UPRMT being an evolutionarily conserved longevity response and merits further investigation. We find that some, but not all, proteins involved in the UPRMT are elevated in the heart. The most commonly used markers for the UPRMT induction are Hsp60 and ClpP; however, neither were up-regulated in the heart of the Surf1−/− mice. At this point it is unclear whether the increase in the UPRMT in response to COX deficiency is beneficial to skeletal muscle function. This tissue specificity of the UPRMT induction has important implications and merits further research into this pathway. Although it may be important in maintaining the mitochondrial function under conditions of mitochondrial dysfunction, it is unclear what the physiological outcomes of this response are or what affect the UPRMT has on the lifespan of mammalian mitochondrial mutants. These results are significant because they provide evidence for an evolutionarily conserved stress response between invertebrates and mammals (UPRMT). Furthermore, the present paper is the first report of the UPRMT in a mammalian in vivo model of mitochondrial ETC insufficiency. Studies are ongoing to define the potential role of the UPRMT in the longevity of the Surf1−/− mouse.

Abbreviations

     
  • ARE

    antioxidant response element

  •  
  • CHOP

    C/EBP (CCAAT/enhancer-binding protein)-homologous protein

  •  
  • ClpP

    caseinolytic peptidase

  •  
  • COX

    cytochrome c oxidase

  •  
  • DCIP

    dichlorophenol-indophenol

  •  
  • DIPPMPO

    5-(di-isopropoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide

  •  
  • ETC

    electron transport chain

  •  
  • 18FDG

    2-[18F]fluoro-2-deoxy-D-glucose

  •  
  • HO-1

    haem oxygenase 1

  •  
  • Hsp60

    heat-shock protein 60

  •  
  • MURE

    mitochondrial unfolded response element

  •  
  • Nrf2

    nuclear factor-erythroid 2-related factor 2

  •  
  • PET

    positron-emission tomography

  •  
  • PGC-1α

    peroxisome-proliferator-activated receptor γ co-activator 1α

  •  
  • RCR

    respiratory control ratio

  •  
  • ROS

    reactive oxygen species

  •  
  • SOD

    superoxide dismutase

  •  
  • SURF1

    surfeit locus protein 1

  •  
  • SUV

    standardized uptake value

  •  
  • TFAM

    mitochondrial transcription factor A

  •  
  • Trx

    thioredoxin

  •  
  • UPRMT

    mitochondrial unfolded protein response

  •  
  • VDAC

    voltage-dependent anion channel

AUTHOR CONTRIBUTION

Daniel Pulliam performed experiments, and was involved in experimental design, data analysis and interpretation, and writing the paper. Sathyaseelan Deepa performed experiments. Yuhong Liu performed mitochondria experiments. Shauna Hill helped perform animal experiments and aided in writing/editing the paper. Ai-Ling Lin performed in vivo glucose uptake experiments and data analysis. Arunabh Bhattacharya and Yun Shi were involved in experimental design and data interpretation. Lauren Sloane designed and performed the animal activity assay. Carlo Viscomi and Massimo Zeviani generated the Surf1−/− mouse and edited the paper. Holly Van Remmen was involved in experimental design, data analysis and interpretation, and writing the paper.

We received assistance from the Cardiac Proteomics Core (Merry Lindsey Laboratory) and Barbara Hunter from the Electron Microscopy Core at UTHSCSA (University of Texas Health Science Center at San Antonio). We thank Dr Luke Szweda and Dr Scott Plafker (Oklahoma Medical Research Foundation) for providing the Lon protease and Nrf2 antibodies respectively.

FUNDING

This work was supported by an Ellison Medical Foundation Senior Scholar award to H.V.R. and the Biology of Aging Training Grant [grant number T32 AG021890] (to D.P.) In vivo imaging studies were supported by the National Institutes of Health [grant number K01AG040164] and the American Federation for Aging Research [grant number A12474] (to A.-L.L.).

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