A new activating role for CO in cardiac mitochondrial biogenesis

Pleiotropic effects of carbon monoxide (CO), attributed to heme protein binding, produce cellular hypoxia and interfere with enzymatic function (Coburn and Forman, 1987; Piantadosi, 2002), but also allow for adaptation (Maines, 1988; Thom et al., 2000). Endogenous CO produced by heme catabolism has physiological roles in eukaryotic cells (Verma et al., 1993), and both endogenous and exogenous CO ameliorate experimental cardiac, lung and vascular injuries (Otterbein et al., 2003; Sato et al., 2001; Song et al., 2003), and protect against certain inflammatory states (Fujita et al., 2001). Thus, CO can exhibit anti-inflammatory (Wagener et al., 2003), anti-proliferative (Taille et al., 2003) and anti-apoptotic effects (Zhang et al., 2003) by largely undetermined physiological mechanisms.

The cellular physiology of CO is complex because the gas binds to multiple heme proteins, including cytochrome P₄₅₀, guanylate cyclase and cytochrome c oxidase (Coburn and Forman, 1987). The latter inhibits mitochondrial electron transport, which can be deleterious to aerobic contractile function (Liao et al., 1996). However, we considered that the physiological activities of CO might be linked to oxidative metabolism (especially with respect to adaptation) by optimizing mitochondrial biogenesis. This requires nuclear and mitochondrial genomic orchestration by regulatory factors, including nitric oxide (NO), which activates guanylate cyclase and yields transcriptional response of, for example, peroxisome proliferator-activated receptor gamma coactivator 1 alpha (PPARGC1A, hereafter referred to as PGC-1α) (Nisoli and Carruba, 2006; Nisoli et al., 2003).

CO specifically binds to reduced cytochrome a₃ but has a weak affinity for guanylate cyclase – much less than NO – and, therefore, may be poised to play a complementary role in mitochondrial biogenesis. This idea derives from observations that cytochrome c oxidase naturally metabolizes CO to carbon dioxide (CO₂) (Young and Caughey, 1986), binding of CO to cytochrome oxidase a₃ heme increases the mitochondrial H₂O₂ leak rate (Zhang and Piantadosi, 1992), and the pro-survival phosphatidylinositide 3-kinase (PI3-K)/Akt pathway (Cantley, 2002) activates replication of mitochondrial DNA (mtDNA) by oxidant-dependent regulation of phosphorylation of nuclear respiratory factor 1 (NRF1) and expression of Tfam (Piantadosi and Suliman, 2006). Our hypothesis was that mitochondrial H₂O₂ production deriving from CO binding to cytochrome c oxidase is an activating factor in mitochondrial biogenesis.

We empirically identified by light microscopy brief CO exposures in mice that produced no mortality or evidence of histological damage. Exposures of 1 hour were chosen, which transiently depressed the resting oxygen consumption (VO₂) and the CO₂ production rates (VCO₂), but then allowed resting-energy expenditure to recover to normal or above (Fig. 1A). The molecular work focused on the heart because of its high metabolic rate and rapid response.

In mice, cardiac mitochondrial protein content increased for all five respiratory complexes by 2D gel electrophoresis within 24 hours of CO exposure (Fig. 1B). Mitochondrial state–state-3 respiration and coupling were preserved, although succinate-linked state-3 increased after CO (Fig. 1C). Low temperature spectra (77°K) of succinate-stimulated mitochondria before and after CO incubations demonstrated the formation of a CO–cytochrome-a₃ complex and early selective reduction of the cytochrome bc₁ region of the chain (Fig. 1D). As detected by electron microscopy (EM), the cross-sectional area and, hence, volume density of interfibrillar mitochondria increased at 24 hours by ∼30%, and was accompanied by profuse budding and ultrastructual heterogeneity characteristic of mitochondrial biogenesis (Fig. 1E). Overall, Fig. 1 illustrates that CO activates cardiac mitochondrial biogenesis in vivo.

In mouse heart tissue, the physiological CO concentration was on average 9 picomoles/mg and increased five- to 20-fold after 1 hour of incubation with CO (Fig. 2A, left panel). The intracellular CO content rose rapidly with the formation of carboxyhemoglobin (COHb), which increases the tissue capillary diffusion barrier and causes retention of endogenous CO (Cronje et al., 2004). At 6 hours, cellular CO concentration had declined to baseline (not shown) but expression of PGC-1α mRNA had increased (Fig. 2A, middle) in relation to the measured cardiac CO concentration (Fig. 2A, right).

A comparison of wild-type (Wt) mice and endothelial nitric oxide synthase (eNOS)-deficient (eNOS^–/–) mice was performed to evaluate the role of NO in CO-induced mitochondrial biogenesis (Clementi and Nisoli, 2005; Nisoli and Carruba, 2006; Nisoli et al., 2003; Nisoli et al., 2004). In Wt and eNOS^–/– strains, mRNA expression for the mitochondrial transcription factor A (TFAM) and DNA polymerase γ (Polγ) was increased after treatment with CO, and cardiac TFAM and Polγ protein had tripled at 24 hours (Fig. 2B, left and middle panels for western blots). Cardiac mtDNA content increased in both strains (Fig. 2B, right). The initial mtDNA content was lower in eNOS^–/– than Wt hearts, but increased 2.5- to 3-fold in both strains after treatment with CO, demonstrating eNOS-independence. The possibility that inducible NOS (iNOS) caused the CO response in eNOS^–/– mice was excluded by inhibiting iNOS with 1400 W (Cayman Chemical, Ann Arbor, MI), which did not alter the responses (data not shown).

The levels of mRNA for NRF1, NRF2 and PGC-1α, the transcription factors and the coactivator that regulate mitochondrial biogenesis, responded to the presence of CO in Wt and eNOS^–/– mouse hearts followed by Tfam mRNA expression (Fig. 3A-D). In Wt mice, mRNA of PGC-1α peaked at 2 hours (Fig. 3A, fourfold; P<0.05), of NRF1 and NRF2 at 2-6 hours (Fig. 3B,C, four- to sixfold; P<0.05) and of Tfam at 24 hours (Fig. 3D, fivefold; P<0.01). In eNOS^–/– mice, levels of these transcripts were lower than in Wt mice but the temporal profiles were comparable.

To establish whether these responses were produced by CO or by hypoxia, Wt mice were exposed to hypoxia for 1 hour (a simulated altitude of 24,000 ft) to control for the hypoxic effect of COHb by lowering cardiac pressure of oxygen (PO₂) levels. Hypoxia had no effect on the expression of cardiac PGC-1α or NRF1 (Fig. 3E,F). To confirm the requirement for binding of CO to heme protein in vivo, CO treatment was given in conjunction with hyperbaric oxygen to prevent binding of CO to reduced cytochrome a₃ by raising mitochondrial PO₂ levels (Piantadosi, 2002). Hyperbaric oxygen abrogated the effect of CO on mRNA levels of both proteins.

Mitogen-activated protein (MAP) kinases, most notably p38, are activated by reactive oxygen species (ROS) (Sugden and Clerk, 1998) and by CO (Zhang et al., 2003). We used the p38 inhibitor SB203580 in vivo to prevent CO-induced phosphorylation of cardiac p38 (Fig. 4A). SB203580 did not affect mRNA levels of PGC-1α, NRF1 or Tfam, or the increase in mtDNA in Wt or eNOS^–/– mice after treatment with CO (Fig. 4A).

The pro-survival, anti-apoptotic serine/threonine kinase Akt is cardioprotective, in part by phosphorylating and activating eNOS (Dimmeler et al., 1999). CO activates cardiac Akt in Wt and eNOS^–/– mice (Fig. 4B, top, P<0.05, densitometry not shown). Akt activation had the expected anti-apoptotic effect, e.g. phosphorylation of Bad (Fig. 4B, middle) (Matsui et al., 2001). In Wt mice, hypoxia (or treatment with hyperbaric oxygen and CO as negative control) did not yield phosphorylation of Akt; moreover, inhibition of p38 did not prevent Akt activation (Fig. 4B, bottom panel). Akt activation by CO, however, was attenuated more than 50% by inhibiting heme oxygenase (data not shown), supporting the hypothesis that cardiac Akt is also activated by endogenous CO.

Rat heart H9c2 cells were treated with the CO-generating molecule dichloromethane (DCM) to increase cellular CO levels via cytochrome P₄₅₀ activity (Kubic and Anders, 1975). CO dose and CO time responses are shown in Fig. 5A (left panel) with cGMP production (middle panel). Mitochondrial viability and function, assessed by MTT assay, was initially unaffected (at 12-24 hours) but increased significantly 36 and 48 hours after treatment with DCM and CO generation, commensurate with mitochondrial biogenesis (Fig. 5A, right panel).

In H9c2 cells, like in the heart, an oxidative stress response to CO was demonstrated by the production of the proteins SOD2 (Fig. 5B), UCP2 (not shown) and HO-1 (not shown). SOD2 protein increased most dramatically (approximately tenfold at 6 hours). ROS generation by CO in H9c2 cells was proximate to mitochondria using the oxidant-sensitive probe Redox Sensor Red CC-1 dye (CC-1) and the mitochondrial-selective dye MitoTracker Green FM (MitoTracker) (Fig. 5C). Control cells showed little CC-1 oxidation and no CC-1 localization to mitochondria (Fig. 5CA-C). After DCM, CC-1 localized early to mitochondria (D-F) and dissipated by 24 hours (G-I). This ROS production depended on CO because the CC-1 fluorescence was eliminated by cytochrome P₄₅₀ blockade (J-M).

In H9c2 cells, CO led to Akt and p38 phosphorylation, but we focused on Akt because the in vivo data did not implicate p38. Moreover, CO-activated Akt phosphorylation in H9c2 cells was unaffected by p38 inhibition, partially abrogated by guanylate cyclase inhibition, and blocked by PI3-K inhibition (Fig. 5D). Full Akt expression was also reduced by heme oxygenase inhibition (data not shown). Akt activation by CO was disrupted by the insertion of catalase into the mitochondria (Fig. 5D), thus demonstrating a role for mitochondrial H₂O₂ in Akt regulation.

Activation of mitochondrial biogenesis in H9c2 cells by CO was indicated by three- to fourfold increases in NRF1, NRF2α, PGC-1α and TFAM protein expression by western blot analysis (Fig. 5E). Guanylate cyclase blockade halved the CO response, but p38 blockade had no effect. By contrast, inhibition of PI3-K/Akt blocked expression of NRF1 and TFAM and attenuated that of PGC-1α and NRF2 (Fig. 5E). CO also increased the copy number of mtDNA, which was prevented by inhibition of guanylate cyclase or PI3-K but not p38 (Fig. 5F).

NRF1 and PGC-1α are binding partners in Tfam expression; therefore NRF1 binding to PGC-1α was checked by co-immunoprecipitation (Fig. 6). NRF1–PGC-1α binding was enhanced in H9c2 cells after treatment with CO, and was sensitive to PI3-K/Akt inhibition. Tfam promoter activation by NRF1 and NRF2 by CO was demonstrated by chromatin immunoprecipitation assay (Fig. 6B). NRF1-Tfam promoter binding after CO in H9c2 cells was abrogated by mitochondrial-targeted catalase (Fig. 6B).

MtDNA synthesis and mitochondrial biogenesis in H9c2 cells was evaluated using bromodeoxyuridine (BrdU) and anti-BrdU antibody (Ab) (Fig. 6C). In control cells, association of BrdU (red) with mitochondria (green) was minimal (Fig. 6CA-C). After CO treatment, in situ BrdU incorporation increased in conjunction with mtDNA replication, commensurate with an increase in mitochondrial mass (green fluorescence intensity, D-F). This finding was confirmed quantitatively by using MTT (not shown). BrdU incorporation into mtDNA was blocked by cytochrome P₄₅₀ inhibition, confirming the requirement of CO (G-I).

Overall, the work demonstrates that mitochondrial production of ROS by CO binding acts as a retrograde activating-factor for mitochondrial biogenesis in the heart. In mice, CO in the physiological range acts on cardiac mitochondrial biogenesis demonstrated by a combination of molecular, biochemical and morphological evidence. This new finding has importance for both exogenous CO and that generated as an endogenous gas.

Mitochondrial biogenesis is regulated by the coordinated expression of specialized nuclear transcription factors that activate genes for mitochondrial proteins (Hood, 2001; Hood et al., 2003). These transcriptional elements were stimulated by CO in the mouse heart by a hypoxia-independent effect. The transcriptional activation mechanisms are not delineated here, but the responses produced significant increases in the copy number of mtDNA, content of OXPHOS protein and mitochondrial density in vivo. The response to CO was confirmed in cardiomyocytes, which in association with cGMP expression and mitochondrial ROS production also showed apposite increases mitochondrial DNA synthesis and functional mitochondria.

The increases in cardiac mtDNA content in Wt and eNOS^–/– mice were the same, unaffected by iNOS inhibition and, in view of the low nNOS content of the heart, support NO-independent effects of CO on mitochondrial biogenesis. The molecular data imply an overlap in the mechanism of action of the two gases: NO relies on guanylate cyclase (Nisoli et al., 2003), whereas CO has dual guanylate-cyclase-dependent and guanylate-cyclase-independent effects – the latter requires mitochondrial production of H₂O₂ and activation of Akt.

CO requires classical O₂-dependent heme-protein binding demonstrated by the effect of hyperbaric oxygen, which keeps oxylabile transition metal centers in the oxidized state. This test is definitive because CO binds only reduced cytochrome oxidase a₃ heme in intact mitochondria (Coburn and Forman, 1987; Piantadosi, 2002). CO elicits other features of mitochondrial redox signaling: inhibition of electron transport (Coburn and Forman, 1987), H₂O₂ generation (Zhang and Piantadosi, 1992) and reversibility by oxygenation (Young and Caughey, 1986).

The binding of CO to cytochrome a₃ promotes reduction of the respiratory carriers in the cytochrome bc₁ region, raising PO₂ levels and increasing mitochondrial H₂O₂ production (Zhang and Piantadosi, 1992), shown here by low-temperature heart-muscle spectra and CC-1 cell labeling studies suggesting an accelerated mitochondrial H₂O₂ leak rate. CO also upregulates SOD2, which scavenges superoxide, augments mitochondrial H₂O₂ release, and may modulate redox signaling (Zhang et al., 2002). In cardiomyocytes, mitochondrial CC-1 uptake in the presence of CO was blocked by mitochondrial-targeted catalase. The loss of mitochondrial H₂O₂ disrupted Akt activation, NRF1 phosphorylation and its binding to the Tfam promoter, events that can be regulated by H₂O₂ (Piantadosi and Suliman, 2006). This role of mitochondrial H₂O₂ is also consistent with the ability of exogenous peroxides to increase cell mitochondrial mass (Lee et al., 2002; Piantadosi and Suliman, 2006).

Mitochondrial biogenesis entails mtDNA replication and transcription, which depend on nuclear-encoded transcription of Tfam (Moraes, 2001) regulated by NRF1 and NRF2 in association with PGC-1α (Virbasius and Scarpulla, 1994; Wu et al., 1999). NRF1 and NRF2 also partner with PGC-1α to activate other components of mitochondrial biogenesis (Kelly and Scarpulla, 2004; Schreiber et al., 2003) including genes encoding components of the respiratory subunit, e.g. cytochrome c and ATP synthase (Kelly and Scarpulla, 2004; Schreiber et al., 2003). PGC-1α is activated by cGMP (Nisoli et al., 2003) and transcriptional activity of GA-repeat-binding protein (mouse NRF2 homologue) forms a positive feedback loop that drives nuclear gene expression for mitochondrial proteins (Mootha et al., 2004).

CO activates two requisite processes in the current paradigm for mitochondrial biogenesis. First, like NO, CO activates guanylate cyclase which promotes mitochondrial biogenesis. Second, CO activates Akt through an established oxidant mechanism involving NRF1, which in concert with NRF2 activates Tfam expression. cGMP may also influence PI3-K-Akt signaling (Li et al., 2000), but H₂O₂ activates Akt by oxidizing cysteine in counter-regulatory phosphatases, such as PTEN, that oppose PI3-K (Leslie et al., 2003). In H9c2 cells, Akt activation by CO was eliminated by two structurally dissimilar PI3-K inhibitors. Likewise, Tfam expression was inhibited, confirming the importance of this pathway in Tfam regulation (Piantadosi and Suliman, 2006). Akt activation was abrogated by mitochondrial-targeted catalase, establishing mitochondrial H₂O₂ as a signal. By contrast, guanylate cyclase inhibition interfered little with Akt activation by CO, thereby implicating H₂O₂ as the primary PI3-K/Akt activation mechanism in this case.

CO also activates p38 MAP kinase (Zhang et al., 2003) but we could not implicate it in early activation of mitochondrial biogenesis. We did not explore the disruption of PGC-1α binding to p160 myb – a negative regulator of its transcriptional activity and of respiration (Fan et al., 2004) – by p38 or the cross-talk postulated between p38 and PI3-K/Akt (Iwasa et al., 2003), because p38 signaling in the heart is complex. Some studies have found p38 protective (Baines et al., 1999), whereas others find it pro-apoptotic (Petrich and Wang, 2004; Ren et al., 2005).

An interpretation of picomolar CO as a respiratory inhibitor must be placed in the context of micromolar cytochrome oxidase in the heart. The CO to cytochrome a₃ stoichiometry was too low to compromise aerobic energy production to the point of damage, and indicated a high gain of mitochondrial biogenesis to mitochondrial oxidation-reduction state. For instance, the spectral data suggest that the CO:O₂ ratio adjusts electron transport in conjunction with the Q cycle to regulate superoxide production and the H₂O₂ leak rate via SOD2 (Trumpower, 1990; Zhang et al., 2002) (Fig. 7). CO is advantaged by binding only the reduced cytochrome oxidase a₃ heme, whereas other terminal inhibitors, such as CN and NO, bind both ferrous (Fe²⁺) and ferric (Fe³⁺) heme. Work on those gases will be of interest but, like CO, requires care to avoid ATP depletion and additional stimulation of biogenesis. Because the Michaelis-Menten constant (K_m) of cytochrome c oxidase for O₂ is low, the reduced cytochrome oxidase a₃ heme available to react with CO is also low, but increases under hypoxic and State 3 conditions (Chance et al., 1970).

Two observations regarding CO and hypoxia are relevant from a physiological perspective. First, the induction of hypoxia by COHb might directly activate Akt, eNOS and certain MAP kinases. This possibility was excluded by hypoxia-controls in mice and in H9c2 cells; picomolar CO stimulated mitochondrial biogenesis in aerobic conditions, mitigating hypoxia as the direct cause. Second, COHb delays endogenous CO clearance both by decreasing the PO₂ level and by increasing the gas back-pressure in tissue (Cronje et al., 2004). This is a simple explanation for the rapid cytoprotection by CO that may recapitulate HO-1 induction (Akamatsu et al., 2004; Fujita et al., 2001). CO and hypoxia also induce HO-1 and mitochondrial heme release (Cronje et al., 2004; Piantadosi et al., 2006), and heme oxygenase inhibition attenuates Akt activation, thus also implicating endogenous CO in mitochondrial biogenesis. The effect is not simple, however, and was set aside for the mitochondrial H₂O₂ mechanism which gives purview to endogenous CO.

With respect to adaptation and cell survival, our evidence is preliminary: CO influences the mitochondrial phenotype by upregulating TFAM, SOD2, the anti-apoptotic pBAD and the resting energy expenditure. We noticed relative activation of complex II after CO despite uniform increases in respiratory complex proteins, which could, in principle, reflect differential assembly, regulation or other adaptive strategies that require energy (Fujita et al., 2001; Wagener et al., 2003). These speculations require confirmation and future investigation.

In summary, modest increases in cellular CO concentration activate mitochondrial biogenesis by a set of molecular responses that includes mitochondrial H₂O₂ production, activation of guanylate cyclase and Akt, and induction of HO-1, independently of eNOS, iNOS and hypoxia. We impute from the physiological behavior of CO that exogenous and endogenous CO interact, and conjecture that endogenous CO participates in accordance with heme turnover and CO clearance rates. These findings have both adaptive and pathogenic implications for conditions that substantially raise the tissue CO content and produce oxidative stress, such as smoking, air pollution, and CO poisoning, and hemolytic or inflammatory states that accelerate heme turnover.

Male 8-week- to 12-week-old mice (C57BL/6, Wt) and eNOS-null mice (eNOS^–/– bred onto C57BL/6, Jackson) were housed in a barrier facility with ad libitum access to standard chow and water. The procedures were approved by the University IACUC. The resting steady-state O₂ consumption and CO₂ production rates (VO₂ and VCO₂, respectively) were measured at the same time of day at constant temperature in a metabolic chamber by timed collections of expired gas and by gas chromatography (Varian model 3800). VO₂ and VCO₂ were computed with standard formulae corrected to standard temperature and pressure dry.

Pre-mixed CO was directed into exposure chambers at 6 l/minute (Cronje et al., 2004) and levels were monitored with a CO analyzer (Interscan, Chatsworth, CA). To control for changes in PO₂, some mice were exposed to conditions mimicking higher altitude (24,000 ft) or hyperbaric conditions (final pressure 5ATA±CO). Others received intraperitoneal (i.p.) injections of 50 μg of the heme oxygenase inhibitor Sn-protoporphyrin (SnPP), of vehicle (DMSO) with or without CO or of p38 inhibitor at 4 hours and 1 hour before treatment with CO (i.p. SB2035801, TOCRIS, Ellisville, MO); control mice received vehicle (0.2 ml 5% DMSO). After euthanasia, hearts were removed for immediate use, were flash-frozen or fixed by perfusion for light microscopy and transmission electron microscopy (TEM) (Suliman et al., 2004). For EM, hearts underwent retro-aortic perfusion first with cold PBS, then with 2% paraformaldehyde and 1% glutaraldehyde in 0.1 M PBS (pH 7.4). Multiple 1-2-mm cubes of the left ventricle were cut and randomly selected for EM. After embedding, ultrathin sections (60 nm) were cut, contrasted with uranyl acetate followed by lead citrate and examined by TEM (Phillips CM12).

Tissue CO analysis was done by calibrated reduction gas chromatography (GC; RGA5; Trace Analytical, Menlo Park, CA, detection limit 1 pM) (Cronje et al., 2004). Duplicate blood-free heart or 1 × 10⁷ cardiomyocyte samples were homogenized in cold PBS (pH 7.0) then homogenized by ultrasound for 1 minute on ice. Homogenates were spun at 2000 g for 20 seconds and 20-60 μl supernatant was placed in 2-ml vials containing 0.5% sulfosalicylic acid, used to release CO into the headspace.

The mtDNA copy number was quantified by competitive PCR using co-amplification of a target template with a known amount of competitor (Suliman et al., 2003a). Mouse cytochrome-b-specific primers 14335 5′-CATCAGTAACACACATTTGT-3′ and 14906 5′-GATTAGCTGGTATGTAGTTG-3′ amplify 571 bp of the target mtDNA and 332 bp of the competitor DNA when cut with SimI. Triplicate samples of 2 × 10³ copies of competitor and 50 ng of target mtDNA were co-amplified in a thermal cycler (PE Applied Biosystems, Foster City, CA) using product optimization on ethidium bromide (EB)-stained gels in the exponential phase. PCR products were separated by electrophoresis in 2% agarose. EB-stained band intensities were measured by densitometry and mtDNA-specific products plotted against competitor density on a log-scale. Copy number was determined from the x intercept or the point at which the ratio of the 571-bp and 332-bp products was 1. A factor of 1.7 was used to correct for molecular weight.

Fresh cardiac interfibrillar mitochondria were isolated, their yield was determined and respiratory complexes were separated by Blue native polyacrylamide gel electrophoresis (BN-PAGE) (Suliman et al., 2004). Samples (2 mg) were mixed in 6-aminocaproic acid (200 μl 0.75 M) Bis-Tris (50 mM, pH 7.0) and 37.5 μl DDM (10%), centrifuged (100,000 g) and Serva Blue was added to the supernatant before 4-12% gradient electrophoresis. Complexes were identified by elution order and in-gel activity stains. Peptides were resolved by 2D Tricine-SDS-PAGE electrophoresis; BN-PAGE bands were excised, incubated in 1% SDS/1% mercaptoethanol for 2 hours and loaded onto 16% separating gels. After electrophoresis, gels were stained with Coomassie Blue followed by silver. Multiple sets of gels, each with tissue from one control and one CO-exposed mouse were carried simultaneously through separation, staining and densitometry. Low-temperature difference spectra of rat heart mitochondria, frozen in 50% glycerol after reduction with sodium dithionite or 1% CO gas, using oxygenated mitochondria as a reference, were used to demonstrate binding of CO to cytochrome-a₃ and reduction of the cytochrome-bc₁ region (Piantadosi, 1989). Mitochondrial function was assessed by polarography in 2-ml thermostatically controlled glass chambers or by conversion of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium) bromide (MTT) to formazan. H9c2 cells were plated at a density of 5 × 10⁴ cells/well in 96-well plates and exposed to DCM at 37°C and 5% CO₂ and stained with MTT. The formazan was dissolved with DMSO and absorbance measured at 540 nm on a plate reader. MTT reduction was defined by untreated controls [100× (absorbance of treated cells) ÷ (absorbance of control cells)].

Proteins were separated by SDS-PAGE (Suliman et al., 2003b). Membranes were incubated with validated polyclonal rabbit Abs against mouse PGC-1α, NRF1, TFAM, and Polγ, or Akt, pAkt, p38, p-p38 and SOD2 (Santa Cruz Biotechnology, Santa Cruz, CA), or anti-tubulin or β-actin (1:1000; Sigma). After five washes in TBST, membranes were incubated at a 1:10,000 dilution of HRP-conjugated goat anti-rabbit or anti-mouse IgG (Amersham). Blots were developed with ECL, proteins quantified on digitized images from the mid-dynamic range and expressed relative to tubulin or β-actin.

Total RNA was prepared from heart or H9c2 cells using Trizol reagent (Invitrogen Corp., Carlsbad, CA). One μg of RNA was reverse-transcribed using Random Hexomer primers and a Superscript enzyme (Invitrogen). Real-time reverse transcriptase (RT)-PCR was performed using the ABI Prism 7000 and SYBR Green master mix (Applied Biosystems, Foster City, CA). After PCR, samples were subjected to melting curve analysis. 18S RNA or β-actin were used as internal controls. ABI Prism 7000 SDS Software was used to quantify differences in gene expression. The threshold cycle (C_t) was determined in the exponential amplification phase. The amount of transcript was normalized to 18S RNA or β-actin by subtracting the mean C_t values for each condition. Because of the exponential PCR reaction, a difference of n in C_t values represents a twofold difference in transcript levels. PCR was performed in triplicate.

H9c2 cells (embryonic rat cardiomyocytes, ATCC, Rockville, MD) were maintained in Dulbecco's modified Eagle's medium (DMEM; Invitrogen) supplemented with 10% fetal bovine serum (Invitrogen) and 2 mmol/l L-glutamine. Cells were incubated with dichloromethane (DCM, 10-150 μM) diluted in 10% DMSO using 10% DMSO in controls. SKF-525A, ODQ, SB203580, Wortmannin and LY294002 were added 30 minutes before treatment at final concentrations of 20 μM, 20 μM, 10 μM, 20 nM, and 25 μM, respectively. H9c2 cells grown to confluence on 96-well plates were assayed with the BioTrak kit (Amersham Pharmacia). Cells were pre-treated for 10 minutes with 0.2 mM IBMX to prevent cGMP degradation followed by DCM/CO. Cells were lysed, centrifuged and supernatants mixed with rabbit anti-cGMP Ab. Aliquots were added to 96-well plates pre-coated with anti-rabbit Ab cGMP-peroxidase conjugate, washed, and peroxidase substrate added. The reaction was terminated with H₂SO₄ and OD₄₅₀ was read against standards.

Redox Sensor Red CC-1 dye (CC-1; Molecular Probes, Eugene, OR), which localizes upon oxidation to mitochondria or lysozomes was used with mitochondrial-selective dye MitoTracker Green FM (MitoTracker; Molecular Probes). H9c2 cells at near confluence were exposed to CO (100 μM DCM) for 8 and 24 hours, and CC-1 (5 μM) and MitoTracker (1 μM) were added for 10 minutes at 37°C. Cells were washed twice in PBS. Localization of MitoTracker (488 nm) and CC-1 (568 nm) was examined using an LSCM (Model 410; Carl Zeiss MicroImaging Inc.). Control and treated samples were scanned at identical parameters and magnification. Confocal images were collected in fluorescence mode followed by electronic image merging.

Mitochondrial-targeted catalase (mtCAT)-expression vectors were constructed as a chimeric cDNA of mouse MnSOD mitochondrial leader sequence and mouse CAT. The MnSOD leader sequence was amplified using a sense primer flanked by a NheI restriction site (5′-GCTAGCGTGTAAACCTCAATAATGTTGTGTCGG-3′) and an antisense primer flanked by an Hpy188I restriction site (5′-GTCCGACATTGCCGGGAGCCCGCGGCCA-3′). The mouse CAT-coding sequence (GenBank accession number X52108) was equipped with an Hpy188I site at the 5′ end with oligonucleotide linkers, using the primers 5′-ATGTCGGACAGTCGGGACCCAGC-3′ and 5′-CAGGTTAGCTTTTCCCTTCGCAGCCAT-3′. PCR fragments of the leader sequence and CAT-coding region were cloned into pGEM-T System I (Promega, Madison, WI). The pGEM-T vector containing the mitochondrial leader sequence was linearized with Hpy188I; the catalase region was removed, treated with Hpy188I and ligated into the pGEM-T-MnSOD linearized vector. The MnSOD-CAT construct was subcloned in-frame into pcDNA3 expression vector. H9c2 cells were transfected with the DNA vector using FuGENE-HD (Roche) and expression verified by lack of H₂O₂ production using CC-1 dye. Cells transfected with the pcDNA3 vector lacking the insert were the controls.

H9c2 cells were lysed and centrifuged at 16,000 g for 30 minutes. Supernatant proteins were cleared with mouse or rabbit IgG and immunoprecipitated with anti-PGC-1. Immunoprecipitates were resolved by SDS-10% PAGE and transferred to nitrocellulose membranes. NRF1 was detected by western blot analysis using a validated Ab.

H9c2 cells (∼4.0 × 10⁶) cultured in 15 cm plates were treated with DCM (100 μM) for indicated times and then cross-linked with 1% formaldehyde for 7.5 minutes, harvested, and sonicated to ∼500bp fragments, incubated for 30 min at 37°C and quenched with 0.125 M glycine. Cells were washed with PBS, harvested and processed for ChIP It assay with anti-NRF1 or anti-NRF2. After ethanol precipitation, DNA was re-suspended in 200 μl/10⁷ cells and 2-5 μl as PCR template. Input samples representing 1% of total DNA were diluted 1:5 and IP fractions 1:2. PCR was carried out on 1 μl samples using sense, 5′-GGCAGTTTGCTGCTGGGT-3′, and antisense, 5′-GGCACTGTGGGAGGCCCA-3′ primers that amplified a 331-bp segment of Tfam from –359 to –28 relative to the transcription start site (Piantadosi and Suliman, 2006).

H9c2 cells were grown in chamber slides and treated with DCM (100 μM) for 48 hours. Twelve hours before fixation, medium was changed to growth medium without pyruvate, uridine or antibiotics supplemented with 1 μM 5-bromo-2′-deoxy-uridine (BrdU) (Roche). After 12 hours, the medium was removed and the cultures were washed with pre-warmed HBSS (37°C). Cells were fixed at room temperature in 4% phosphate-buffered paraformaldehyde and 4% sucrose and washed in PBS. Fixed cells were incubated in methanol (–20°C) for 10 minutes and washed again with PBS. Cultures were incubated for 25 minutes at 37°C in 200 nM MitoTracker (Molecular Probes), washed in PBS and incubated 100 minutes at 37°C with mouse anti-BrdU Ab (Roche) in buffer, or buffer only. Cells were washed and incubated for 60 minutes at 37°C in goat anti-mouse FluoroLink Cy3-labeled Ab (Molecular Probes), rinsed, coverslipped and observed by fluorescence confocal microscopy (Zeiss 410 LSCM, 620x).

Mouse and cell data were expressed as the mean + standard error (s.e.) for n=3-8 for each time and condition. Experimental and control group comparisons were carried out by Student's t-test. Time series data were analyzed by two-way ANOVA, with repeated measure corrections as indicated, and by Fisher's post-hoc test. P values of P<0.05 were considered significant.

This work was supported by NHLBI PO1 42444 (CAP). Parts of this research were presented at the HO Conference 2005 Fourth International Congress, Boston, MA and the 12th Annual Meeting of the Society for Free Radical Biology Medicine, Austin, TX, 2005.