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Am J Physiol Heart Circ Physiol 285: H2132-H2141, 2003. First published July 24, 2003; doi:10.1152/ajpheart.00147.2003
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Hypoxia-induced left ventricular dysfunction in myoglobin-deficient mice

Pradeep P. A. Mammen,1 Shane B. Kanatous,1 Ivan S. Yuhanna,2 Philip W. Shaul,2 Mary G. Garry,1 Robert S. Balaban,4 and Daniel J. Garry1,3

Departments of 1Internal Medicine, 2Pediatrics, and 3Molecular Biology, University of Texas Southwestern Medical Center, Dallas, Texas 75390; and 4Laboratory of Cardiac Energetics, National Heart, Lung, and Blood Institute, Bethesda, Maryland 20892

Submitted 19 February 2003 ; accepted in final form 8 July 2003


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
Myoglobin-deficient mice are viable and have preserved cardiac function due to their ability to mount a complex compensatory response involving increased vascularization and the induction of the hypoxia gene program (hypoxia-inducible factor-1{alpha}, endothelial PAS, heat shock protein27, etc.). To further define and explore functional roles for myoglobin, we challenged age- and gender-matched wild-type and myoglobin-null mice to chronic hypoxia (10% oxygen for 1 day to 3 wk). We observed a 30% reduction in cardiac systolic function in the myoglobin mutant mice exposed to chronic hypoxia with no changes observed in the wild-type control hearts. The cardiac dysfunction observed in the hypoxic myoglobin-null mice was reversible with reexposure to normoxic conditions and could be prevented with treatment of an inhibitor of nitric oxide (NO) synthases. These results support the conclusion that hypoxia-induced cardiac dysfunction in myoglobin-null mice occurs via a NO-mediated mechanism. Utilizing enzymatic assays for NO synthases and immunohistochemical analyses, we observed a marked induction of inducible NO synthase in the hypoxic myoglobin mutant ventricle compared with the wild-type hypoxic control ventricle. These new data establish that myoglobin is an important cytoplasmic cardiac hemoprotein that functions in regulating NO homeostasis within cardiomyocytes.

nitric oxide; knockout mice; cardiac systolic function

MYOGLOBIN (Mb) is a monomeric cytoplasmic hemoprotein that is tissue restricted to cardiomyocytes and oxidative skeletal myofibers. In general, studies support the hypothesis that Mb is an essential protein that facilitates oxygen delivery from the intracapillary erythrocyte to the mitochondria to maintain oxidative phosphorylation for myocardial contractility (16, 27, 57, 58, 63). Although several studies challenge the role of Mb in intracellular oxygen transport (21, 62), a number of studies utilizing pharmacological inhibitors of Mb support a functional role of Mb in facilitating oxygen transport in striated muscle (16, 27, 57, 58). As an alternative, we pursued a gene disruption strategy to generate mice that are viable and fertile despite a complete absence of Mb (20). Mice lacking Mb have preserved cardiac function and exercise capacity compared with their wild-type (WT) littermates under normoxic conditions. Additionally, Mb-deficient (Mb–/–) mice survive by mounting a complex cellular and molecular compensatory response that includes increased capillary density and an induction of the hypoxia gene program (hypoxia-inducible factor-1{alpha}, VEGF, heat shock protein27, etc.) (22, 23, 36). These findings have prompted our laboratory and others to reexamine the functional role of Mb in the heart.

Additional functions have been proposed for Mb, including the role(s) as a reservoir for oxygen, as a cytoprotective protein against reactive oxygen species (i.e., H2O2, , and ONOO–), and as a modulator of the signaling molecule nitric oxide (NO) (19, 28, 30, 59). NO is an important inter- and intracellular physiological messenger capable of evoking a number of cellular responses that are beneficial or potentially toxic. NO avidly binds to heme-containing proteins such as soluble guanylate cyclase (sGC), Mb, hemoglobin, and cytochrome c oxidase, and these hemoproteins are essential in the modulation of NO bioactivity and responsiveness to the concentration of NO (13, 15, 26, 35).

In the present study, we challenged the Mb-null mice with chronic hypoxia in an attempt to further define and explore alternative functional roles for Mb. We hypothesized that the cellular and molecular adaptations in Mb mutant mice would be insufficient to maintain cardiac function in response to chronic hypoxic conditions. Here, we present evidence to support our hypothesis and establish Mb as an important regulator of NO homeostasis in the heart.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
Animals. Homozygous Mb–/– mice were generated utilizing gene disruption technology and genotyped as previously described (20). The handling and use of these mice were in accordance with National Institutes of Health and University of Texas Southwestern Medical Center's institutional guidelines. All the mice used in this study were adult males (2–4 mo old) and were from a C57BL/6J strain background.

Hypoxic exposure. A Plexiglas chamber (87 x 42 x 45 cm) was engineered to maintain a constant hypoxic environment of 10% O2 by infusing a gas mixture of 10% O2 and 90% N2 into the chamber at a rate of 1 l/min. It was continuously monitored for oxygen and carbon dioxide concentrations, temperature, and humidity. Within the hypoxic chamber, mice were individually housed in 15 x 32-cm cages and were maintained in a 12:12-h light-dark cycle. To ensure the mice were exposed to a significant hypoxic stimulus, we measured the hematocrit levels of these mice using conventional methods. Echocardiographic M-mode images (Hewlett-Packard Sonos 5500) were obtained from a parasternal short-axis view at the level of the left ventricular (LV) papillary muscle in lightly anesthetized mice (0.015 ml Avertin/g body wt) as previously described (36). As demonstrated by Roth el al. (47), Avertin has minimal adverse cardiodepressant effects after an equilibrium state has been reached. Thus, as recommended by Roth el al. (47), echocardiographic measurements were performed 15 min after the mice were anesthetized. The heart rate was also monitored during echocardiography.

In selected studies, WT and Mb–/– mice were treated with the NO synthase (NOS) inhibitor N{omega}-nitro-L-arginine methyl ester (L-NAME; Sigma; St. Louis, MO). Mice were pretreated for a 1-wk period under normoxic conditions with L-NAME (0.07 mg · g body wt–1 · day–1), which was dissolved in the drinking water (33, 34, 39, 42). Water consumption was monitored to ensure both the WT and Mb–/– mice received similar doses of L-NAME. The mice were then exposed to chronic 10% hypoxia, and treatment with L-NAME was continued during this hypoxic period. Echocardiography was used to evaluate the LV systolic function in mice before L-NAME treatment and 1 wk after L-NAME treatment under normoxic conditions and after exposure to hypoxic conditions.

Histological and immunohistochemical analyses. Tissues for histological and immunohistochemical studies were perfused via the LV with PBS, followed by 4% paraformaldehyde-PBS (pH 7.4), excised, postfixed for 14–16 h at 4°C, and embedded in paraffin. Selected tissue sections were stained with hematoxylin and eosin or Masson's trichrome for morphological analyses. Immunohistochemical analyses were undertaken using a primary rabbit anti-inducible NOS (iNOS) serum (1:200, Santa Cruz Biotechnology; San Diego, CA), which was detected by a secondary antiserum (1:100 goat anti-rabbit biotinylated serum; Vector Laboratory, Burlingame, CA) and horseradish peroxidase (HRP)-dimethyl-aminoazobenzene (DAB). Substitution controls revealed an absence of signal (data not shown). To limit the loss of immunohistochemical signal, no counterstain was utilized.

Vascular quantitation. Capillary density was quantified in WT and Mb–/– ventricles by staining paraffin-embedded methyl-Carnoy-fixed sections with biotinylated Bandiera simplicifolia lectin B4 (20 mg/ml, Vector Laboratory) and detected with DAB (36). Digital photographs were taken of the LV, and capillary density was quantified at four different levels (50 µm apart) for each heart. With the use of a computerized digital analysis program (Scion Image 1.62c), images were captured, and vascularity was quantitated from four different (random) regions in the lateral LV.

Western blot analysis and biochemical assays. Western blot analysis was performed according to a previously published protocol (20). Rabbit anti-Mb serum (1:3,000, DAKO; Carpenteria, CA) was used as the primary antiserum, which was detected using a HRP-conjugated secondary antiserum. Mouse monoclonal {alpha}-tubulin antibody (1:3,000, Sigma) was used as a standard to ensure equal protein loading of the lanes. Band intensity was quantitated using a computerized digital analysis program (Scion Image 1.62c). NOS enzymatic activity was determined in the supernatant of homogenized hearts from normoxic and hypoxic WT and Mb–/– mice by measuring the conversion of L-[3H]arginine to L-[3H]citrulline as previously described (43, 53). NOS activity was linear with time (up to 60 min) and with protein concentration (up to 60 µg/reaction). Furthermore, NOS activity was fully inhibited by 2 mM L-NAME. Calcium-dependent and -independent NOS activity were partitioned by the addition of 5 mM EGTA to the incubation mixture. Selective inhibition of neuronal NOS (nNOS) and endothelial NOS (eNOS) activities by Ca2+ chelation and the lack of effect on iNOS activity were confirmed in lysates of COS-7 cells singularly transfected with cDNA for each isoform.

cGMP assay. Normoxic and hypoxic mice were euthanized by cervical dislocation, and the hearts were excised within 15 s. The great vessels and atria were removed, and the heart was snap frozen in liquid N2 and stored at –80°C. The frozen ventricles were weighed, homogenized in 1 ml of ice-cold 0.5 N perchloric acid, and then centrifuged at 14,000 rpm at 4°C for 5 min. The supernatant contained cGMP, which was measured using a standard radioimmunoassay as previously described (9, 17).

Statistical analysis. Data analysis between WT and Mb–/– mice at any one hypoxic time period was performed using two-tailed paired Student's t-test. Comparison within a genotype to varying durations of hypoxia was performed utilizing ANOVA (single factor), followed by a Bonferroni post hoc test. Data values are reported as means ± SE.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 DISCLOSURES
 REFERENCES
 
Survival in the absence of Mb is dependent on cellular and molecular adaptations resulting in preserved cardiac function. To further define the role for Mb, we challenged Mb–/– and WT age- and gender-matched mice to 10% O2 for various time periods (1 day to 3 wk). At baseline (normoxic conditions), there was no difference in the hematocrit levels between WT and Mb–/– mice (51.31 ± 0.66% vs. 51.30 ± 0.72%, respectively, n = 21 for each group). We observed an increase in the hematocrit levels in both hypoxic WT and Mb–/– mice, and at any single hypoxic time period there was no difference in the hematocrit levels between WT and Mb–/– mice (Fig. 1A). For example, after 3 wk of hypoxic (10% O2) exposure, we observed a similar increase in the hematocrit levels in both WT and Mb–/– mice compared with their respective baseline levels (73.19 ± 2.55% and 74.75 ± 1.31%, respectively, P < 0.001). This polycythemic response in WT control mice was comparable to previously published results and served as a further confirmation that the mice were exposed to hypoxic conditions (64).



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  		Fig. 1. Increase in hematocrit levels, development of left ventricular (LV) systolic dysfunction, and preserved cardiac structural anatomy in myoglobin (Mb) mutant mice exposed to chronic hypoxia. A: hematocrit levels of wild-type (WT) and Mb-deficient (Mb–/–) mice exposed to chronic hypoxia (10% oxygen). *P < 0.001, hypoxic WT vs. normoxic WT (n = 21 at baseline, n = 6 for each hypoxic group); **P < 0.001, hypoxic Mb–/– vs. normoxic Mb–/– (n = 21 at baseline, n = 6 for each hypoxic group). B: with the use of transthoracic echocardiography, LV systolic function (i.e., fractional shortening) was measured in normoxic and hypoxic WT and Mb–/– mice. *P < 0.001, hypoxic Mb–/– vs. hypoxic WT (n = 6 for each hypoxic group). C: representative M-mode images obtained in WT and Mb–/– mice exposed to 1 wk of chronic 10% hypoxia. LVEDD, LV end-diastolic dimension; LVESD, LV end-systolic dimension. D: histological (hematoxylin-eosin) analysis of hearts obtained from WT and Mb–/– mice exposed to 1 wk of hypoxic (10% oxygen) conditions.

 

Echocardiography was utilized to evaluate the LV systolic function of WT and Mb–/– mice in response to chronic hypoxia by measuring the fractional shortening of the LV. As illustrated in Fig. 1, B and C, Mb–/– mice exposed to chronic 10% O2 for a 1-wk period developed an ~30% reduction in systolic function compared with normoxic Mb–/– mice (0.43 ± 0.02 and 0.59 ± 0.02, respectively, P < 0.05, n = 6 for each group) and hypoxic WT mice (0.43 ± 0.02 and 0.59 ± 0.03, respectively, P < 0.001, n = 6 for each group). This degree of hypoxia-induced LV systolic dysfunction in the Mb–/– mice was constant and without further deterioration, regardless of the period of hypoxic exposure (1–3 wk). In contrast, hypoxic WT mice did not develop myocardial depression at any time period after exposure to hypoxia. Under normoxic conditions, the heart rate was similar between WT and Mb–/– anesthetized mice [615 ± 36 and 585 ± 70 beats/min, respectively, not significant (NS)]. Furthermore, no evidence of LV dysfunction was observed in normoxic WT and Mb–/– hearts using the anesthetic agent. Finally, under hypoxic conditions, no difference was observed in the heart rate between WT and Mb–/– anesthetized mice, indicating that the anesthetic agent did not have a differential effect on cardiac function between the two groups.

As illustrated in Fig. 1D, histological evaluation of hypoxic WT and Mb–/– hearts showed similar structural differences. This was confirmed by echocardiographic measurements of ventricular dimensions (see Table 1). After 1 wk of hypoxic exposure, the anterior wall thickness was similar between WT and Mb–/– hearts (0.99 ± 0.04 vs. 1.00 ± 0.01 mm, respectively, NS, n = 6 in each group). Likewise, there was no difference in the posterior wall thickness between hypoxic WT and Mb–/– hearts (0.95 ± 0.02 vs. 0.98 ± 0.01 mm, respectively, NS, n = 6 in each group). Figure 1D and Table 1 also indicate the development of right ventricular and LV hypertrophy in both hypoxic WT and Mb–/– hearts. In addition, we did not observe any morphological evidence of either ischemic or necrotic myocardial injury in Mb–/– or WT hearts under hypoxic conditions, as assessed by Masson's trichrome staining (data not shown). Furthermore, we did not observe any differential expression of Mn-SOD and p38 in the WT and Mb-null hearts exposed to hypoxic conditions, suggesting that oxidative stress did not contribute to the LV dysfunction in the Mb-null heart (data not shown).


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  		Table 1. Echocardiographic measurements in WT and Mb-/- mice exposed to normoxic and hypoxic conditions

 

Hypoxia has been shown to be associated with an increase in capillary density (29, 60, 61). Using an immunohistochemical protocol, as described in previous studies, we quantitated the capillary density in the LV (23, 36). As demonstrated by our laboratory and others (22, 23, 36), under normoxic conditions there is a significant increase in capillary density in the LV of Mb–/– mice compared with the LV of WT mice (Fig. 2A; 4,729 ± 87 vs. 4,116 ± 88 capillaries/mm2, respectively, P < 0.001, n = 18 for each group). Upon exposure to chronic hypoxia, we observed an ~35% increase in capillary density of the LV in both WT and Mb–/– mice (Fig. 2A; P < 0.001, n = 20 for each group). This increase in capillary density is finite in both hypoxic WT and Mb–/– ventricles, and no further increase occurs with increased exposure to hypoxic conditions.



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  		Fig. 2. Increase in LV capillary density in hypoxic WT and Mb–/– mice. A: LV capillary density in hearts isolated from WT and Mb–/– mice exposed to normoxic (n = 18) and hypoxic (n = 20 for each hypoxic group) conditions. *P < 0.001, normoxic Mb–/– vs. normoxic WT; **P < 0.001, hypoxic WT vs. normoxic WT; ***P < 0.001, hypoxic Mb–/– vs. normoxic Mb–/–. B: representative sections stained with lectin B4 to identify capillaries in the LV under normoxic and hypoxic (1 wk) conditions. Bar = 40 µm.

 

Mb expression is markedly induced in response to hypoxic conditions in the WT ventricle, as observed by Western blot analysis (Fig. 3, A and B). After 2 wk of hypoxic exposure, the Mb content in WT hearts increased by 31% from normoxic levels (1.22 ± 0.08 vs. 0.97 ± 0.01 mm, respectively, P < 0.05, n = 3).



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  		Fig. 3. Ventricular Mb content increases with exposure to hypoxia. A: representative Western blot illustrating an increase in ventricular Mb content with chronic hypoxic exposure (10% oxygen). B: quantification of ventricular Mb content done in triplicate. Data are normalized to {alpha}-tubulin content. *P < 0.05, hypoxic WT vs. normoxic WT.

 

To determine whether the hypoxia-induced LV systolic dysfunction in hypoxic Mb–/– mice was a reversible process, we exposed age- and gender-matched mice to 2 wk of chronic hypoxia and then returned them to normoxic conditions (21% O2). We evaluated the LV function of these mice by echocardiography immediately after hypoxic exposure versus 1 and 7 days after reexposure to normoxic conditions. As previously shown, we observed a 30% reduction in systolic function in Mb–/– mice compared with WT mice after 2 wk of hypoxia (Fig. 4A; P < 0.001, n = 3 for each group). However, this hypoxia-induced LV systolic dysfunction was reversible in Mb–/– mice after they were returned to normoxic conditions for 1 and 7 days (Fig. 4A). The reversibility of the LV dysfunction of the hypoxic Mb-null mice was consistent with a nonstructural etiology.



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  		Fig. 4. Hypoxia-induced LV systolic dysfunction in Mb–/– mice is a reversible process and is mediated by nitric oxide (NO). A: with the use of echocardiography, LV systolic function (i.e., fractional shortening) was measured in WT and Mb–/– mice exposed to 2 wk of chronic hypoxia. These mice were then reexamined using echocardiography after reexposure to normoxic conditions. *P < 0.001, 2-wk hypoxic Mb–/– vs. 2-wk hypoxic WT (n = 3 for each group). B: WT and Mb–/– mice were treated with N{omega}-nitro-L-arginine methyl ester (L-NAME) and exposed to chronic hypoxia, and LV function was evaluated using echocardiography (n = 3 in each group treated with L-NAME). For comparison, LV systolic function in WT and Mb–/– mice not treated with L-NAME and exposed to 1 wk of chronic hypoxia is illustrated (n = 6 for each hypoxic group not treated with L-NAME). Note that L-NAME treatment prevents LV dysfunction in the hypoxic Mb-null heart. *P < 0.001, hypoxic Mb–/– without L-NAME vs. hypoxic WT without L-NAME.

 

Having determined that hypoxia-induced LV dysfunction in Mb mutant mice was a reversible process, we examined whether the LV dysfunction was a NO-mediated mechanism. Therefore, we treated WT and Mb–/– mice with L-NAME, a potent inhibitor of NOS, to determine whether we could prevent the cardiac dysfunction observed in hypoxic Mb–/– mice. As illustrated in Fig. 4B, L-NAME prevented the development of LV systolic dysfunction in hypoxic Mb–/– mice. In addition, L-NAME did not have any adverse effect on the systolic function of normoxic or hypoxic WT mice (Fig. 4B). To determine that both WT and Mb–/– mice were receiving similar amounts of L-NAME, we measured daily water consumption and observed no difference between the two groups (2.44 ± 0.36 vs. 2.51 ± 0.47 ml, respectively, NS, n = 6 in each group).

The above experiment demonstrates that the hypoxia-induced LV systolic dysfunction we observed in hypoxic Mb–/– mice is a NO-mediated process. Because cardiac dysfunction first developed in the Mb mutant mice after 1 wk of hypoxic exposure, we used a conventional assay to measure the ventricular NOS activity level in these mice exposed to 1 wk of chronic hypoxia. Under normoxic conditions, the total NOS activity was similar in WT and Mb–/– hearts (0.50 ± 0.11 vs. 0.42 ± 0.08 pmol · mg protein–1 · min–1, respectively, NS, n = 10 for each group). As shown in Fig. 5A, we observed a more than a twofold increase in total ventricular NOS activity in hypoxic WT mice compared with normoxic WT controls (n = 3 for the hypoxic group). In the hypoxic WT heart, this increase in NOS activity was attributable solely to an increase in Ca2+-dependent NOS activity (eNOS and nNOS), and there was a complete absence of ventricular Ca2+-independent NOS activity (iNOS) in both normoxic and hypoxic WT mice (Fig. 5, B and C). In Mb–/– mice, there was a 1.5-fold increase in the total ventricular NOS activity under hypoxic conditions compared with under normoxic conditions (n = 3 for the hypoxic group). We observed a simultaneous decrease in the Ca2+-dependent NOS activity (Fig. 5B). In addition, there was an approximately sixfold increase in ventricular Ca2+-independent NOS activity (iNOS) in hypoxic Mb–/– mice compared with normoxic Mb–/– mice (Fig. 5C). These results were further corroborated as we observed a marked increase in iNOS expression in the hypoxic Mb-null LV (Fig. 5D) by immunohistochemistry compared with the controls.



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  		Fig. 5. Marked induction of inducible NO synthase (iNOS) activity in the ventricles of hypoxic Mb–/– mice compared with hypoxic WT mice. A–C: NOS activity was measured from the ventricles of WT and Mb–/– mice exposed to 1 wk of hypoxic conditions and normalized to its respective normoxic activity levels. Ca2+-dependent NOS activity represents the combined activity of neuronal NOS and endothelial NOS. Ca2+-independent NOS activity represents the activity of iNOS (n = 3 for each group). *P < 0.05, Mb–/– vs. WT. D: iNOS immunohistochemical analysis in representative sections of normoxic and hypoxic WT and Mb–/– ventricles. Note the absence of iNOS in normoxic and hypoxic WT ventricles but a marked induction of iNOS in Mb-null cardiomyocytes exposed to chronic hypoxia (arrows mark immunopositive cardiomyocytes). Bar = 100 µm.

 

To determine whether this NO-mediated LV dysfunction in the Mb–/– heart was a cGMP-dependent mechanism, we measured cGMP levels in ventricles from hypoxic WT and Mb–/– mice (Fig. 6). We observed no significant difference in cGMP levels in ventricles from hypoxic Mb–/– mice compared with hypoxic WT mice (Fig. 6). These results suggest that the LV dysfunction in the Mb-null mice exposed to chronic hypoxia is due to a NO-dependent, cGMP-independent pathway.



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  		Fig. 6. cGMP levels are unchanged in WT and Mb–/– hypoxic hearts. cGMP levels were measured by a radioimmunoassay, and no significant difference was found between hypoxic WT and Mb–/– ventricles (n = 3 for each group).

 


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
DISCLOSURES
 REFERENCES
 
We have previously observed that adult mice lacking Mb are viable, fertile, and have preserved cardiac function. In the present study, a distinguishable cardiac phenotype was identified only in Mb mutant mice challenged with chronic hypoxic exposure and not in WT control mice. LV systolic dysfunction was observed in Mb-deficient mice after exposure to chronic hypoxic conditions, suggesting that the molecular and cellular adaptations are insufficient to maintain cardiac function in the absence of Mb. The LV dysfunction observed in Mb-null hypoxic mice was reversible with reexposure to normoxic conditions and could be completely abolished when mice were treated with L-NAME, a pharmacological inhibitor of NOSs. These pharmacological studies support the conclusion that the LV systolic dysfunction in hypoxic Mb–/– mice is via a NO-mediated mechanism.

Although our data suggest a NO-mediated mechanism for the hypoxia-induced LV dysfunction in Mb-null mice, it is possible that other genetic and physiological factors may also contribute to our experimental results. In the present study, we measured serum hemocrit levels as an indirect measure of the oxygen-carrying capacity in hypoxic WT and Mb–/– mice. We observed a similar rise in serum hematocrit levels in both hypoxic WT and Mb–/– mice compared with their respective controls. Therefore, the serum oxygen-carrying capacity in both groups of mice was the same.

We also analyzed the structural integrity of hypoxic WT and Mb–/– hearts utilizing both histological and echocardiographic techniques. We observed a similar degree of right ventricular and LV hypertrophy in the respective mouse models after the exposure to chronic hypoxia using morphological techniques. These observations were further extended and supported utilizing echocardiographic techniques to determine the LV wall thickness. Masson's trichrome staining did not reveal any myocardial necrosis with hypoxic exposure in either group. Therefore, alterations in the structural integrity of the hypoxic heart are unlikely to explain our data.

In the present study, we also measured the LV capillary density in response to hypoxia. Our data reveal a significant increase in LV capillary density in both genotype groups, and this increase was finite after 1 wk of hypoxic exposure. Although there was a significantly greater LV capillary density in the 1-wk hypoxic WT heart compared with the 1-wk hypoxic Mb–/– heart, no significant differences were observed between groups after 2 wk of hypoxic exposure. This finite increase in the capillary density in response to chronic hypoxia was consistent with the notion that the LV dysfunction observed in the Mb-null mouse exposed to chronic hypoxia was a result of limited oxygen delivery. The results of the L-NAME experiments argue against such a mechanism and further support a NO-mediated mechanism for the LV dysfunction observed in the Mb-null mouse exposed to chronic hypoxic conditions.

Our data support the hypothesis that Mb has an additional function as an inactivator or scavenger of NO, which is distinct from its traditional role in oxygen flux. NO is an important inter- and intracellular physiological messenger capable of evoking a number of cellular responses that are either beneficial or potentially toxic to the cell (8, 14, 18, 45, 48, 51, 55). NO is generated by the oxidation of L-arginine and is catalyzed by one of three isoforms of NOS (3). Both nNOS and eNOS have Ca2+-dependent NOS activity and are constitutively expressed in the heart. In contrast, iNOS has Ca2+-independent NOS activity and is usually biosynthesized in a cell after it has been stimulated by a stressful event (i.e., ischemia or hypoxia).

Recent studies indicate that the type of NOS that generates NO determines whether NO will have a beneficial or toxic effect on the cell. For example, eNOS, which localizes to the caveolae, has been shown to generate NO that inhibits {beta}-adrenergic-induced inotropy, whereas nNOS, which targets the cardiac sarcoplasmic reticulum, has been shown to facilitate cardiac contractility (2). In contrast, the induction of iNOS activity in hearts exposed to a stress stimulus (i.e., ischemia or hypoxia) is generally believed to result in detrimental effects (i.e., decreased contractile function, sudden death, etc.) (41, 50). However, recent studies have challenged the notion that iNOS expression has detrimental effects in the cardiomyopathic heart (7, 8, 14, 31, 45).

Many of the intracellular effects of NO are mediated by the ability of NO to bind to various heme-containing proteins (i.e., NO binding to sGC). Therefore, hemoproteins (Mb, hemoglobin, sGC, lipoxygenases, or free radicals such as superoxide) are essential in the modulation of NO bioactivity and responsiveness to the concentration of NO (13, 19, 26, 35). We have previously shown that NO generation is altered in skeletal muscle of Mb-null mice after a 30-Hz electrical stimulation (23). These results support the presence of NO-mediated mechanism(s) in Mb mutant skeletal muscle groups to presumably promote oxygen delivery through vasodilation and/or to mediate NO signaling pathways through adaptations that have not been completely defined. Additionally, Flögel et al. (19) demonstrated that Mb not only has a functional role in oxygen metabolism but also avidly binds NO within myocytes.

NO has a number of beneficial effects in various systems and importantly functions as a physiological vasodilator in the coronary artery vascular bed, a regulator of mitochondrial respiration, a modulator of myocardial contractility, and a key metabolite in blood pressure regulation (1, 30, 32, 59). Most of the functions for NO in the cardiovascular system are attributable to the enzyme eNOS, which is expressed in the endothelium as well as the cardiomyocyte. In contrast, iNOS is expressed in the cardiac myocyte primarily under stressful or pathological conditions. Recent data have challenged this notion that iNOS-derived NO is detrimental to the myocyte and may in fact have a beneficial effect on myocardial oxygen consumption in congestive heart failure (8).

The NOS activity in the WT heart, exposed to either normoxic or hypoxic conditions, is attributed solely to Ca2+-dependent NOS isoforms (nNOS and eNOS) and not to the Ca2+-independent NOS isoform (iNOS) (Fig. 5, A–C). In response to chronic hypoxia, Mb content is markedly increased in the WT heart and presumably scavenges excess NO, resulting in preserved systolic function (Fig. 3). However, in Mb-deficient mice, iNOS is the predominant isoform in the heart under hypoxic conditions (Fig. 5C). Low levels of iNOS activity were observed in the hearts of Mb-null mice exposed to normoxic conditions, indicating a low level of baseline stress; this finding is further supported as we have previously reported an induction of the hypoxia-inducible gene program in these mice (Fig. 5, C and D) (36). Under normoxic conditions, the Mb mutant mice are able to maintain cardiac function as a result of cellular and molecular adaptations that steepen the PO2 gradient within the myocyte as well as decrease the distance of O2 diffusion from the capillary to cardiac mitochondria (22, 23, 36). However, these adaptations are insufficient to maintain cardiac systolic function in Mb-null mice exposed to chronic hypoxia as a result of altered NO homeostasis. This conclusion is based on the rescue of cardiac function in hypoxic Mb–/– mice treated with L-NAME. Thus, in the absence of Mb, perturbed NO homeostasis results in cardiac systolic dysfunction. Similar mechanisms may be associated with clinical heart failure (15, 24, 25, 49, 50). The cardiomyopathic heart is characterized by decreased Mb content, altered NO homeostasis, and ventricular dysfunction (24, 25, 44). In this clinical setting, a component of cardiac dysfunction may be a result of altered NO homeostasis due to impaired inactivation of NO by decreased amounts of Mb.

The cytotoxic effects of NO can be mediated either via a cGMP-dependent or -independent pathway. The cGMP-dependent deleterious effects of NO are mediated by either the stimulation of protein kinase G proteins resulting in decreased myofilament Ca2+ sensitivity or by the inhibition of L-type Ca2+ channels (37, 38, 40, 52). Alternatively, the cytotoxic effects of NO that are cGMP independent are mediated either by the nitrosylation of the sarcomeric proteins or by the direct effect of NO on mitochondrial respiration (4–6, 11, 12, 46, 51, 54–56, 65). Because the hypoxia-induced LV systolic dysfunction observed in the Mb–/– mouse is reversible within 24 h upon exposure to normoxic conditions, we hypothesize that this is the result of a metabolic disturbance rather than a structural alteration. Therefore, one possibility is that the deleterious effects of NO in our murine model is acting via a interaction of NO with the electron transport chain.

A previous study by Chung et al. (10) has demonstrated that NO regulation of mitochondrial respiration was observed only in response to pharmacological doses of nitrite (>10 mM) in the isolated, perfused heart preparation. In contrast, in the present study, we hypothesize that hypoxia-induced LV dysfunction in Mb-null mice is due to an interaction of NO with the electron transport chain. The rationale for this hypothesis is supported by a number of published studies. Taylor et al. (58) demonstrated that under hypoxic conditions, ATP and phosphocreatine are depleted significantly more in rat hearts with decreased concentration of functional Mb compared with hearts with normal levels of Mb. Several additional studies (1, 32, 59) have demonstrated that NO modulates oxygen mitochondrial respiration in the heart. NO has been shown to directly inhibit mitochondrial oxidative phosphorylation through reversible binding to cytochrome c oxidase (complex IV), where NO competes with dioxygen to bind at the binuclear site of cytochrome c oxidase (4–6, 11, 12, 46, 54, 56, 65). As reviewed by Trochu et al. (59) and recently demonstrated by both Clement et al. (12) and Shiva et al. (54), this inhibition of cytochrome c oxidase by NO is enhanced under low oxygen tension.

In our present study, we did not observe any difference in cGMP levels in hypoxic WT and Mb–/– hearts. While we cannot exclude the possibility of a direct nitrosylation-mediated process or a NO-guanylate cyclase-cGMP-mediated mechanism, the current data support the hypothesis that the depressed LV systolic function observed in hypoxic Mb–/– mice is a NO-dependent, cGMP-independent mechanism.

In summary, we demonstrated that Mb is induced in the WT heart in response to a hypoxic challenge. In the absence of Mb, altered NO homeostasis is observed under chronic hypoxic conditions, resulting in reversible LV dysfunction via a cGMP-independent mechanism presumably involving the reversible inhibition of cytochrome c oxidase by NO. These results collectively support the paradigm that Mb functions to facilitate oxygen flux and maintain NO concentration gradients in the cardiomyocyte.


   DISCLOSURES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
REFERENCES
 
This study was supported by National Heart, Lung, and Blood Institute Grant HL-63788. P. P. A. Mammen is a Pfizer Postdoctoral Fellow in Cardiovascular Medicine.


   ACKNOWLEDGMENTS
 
We acknowledge the assistance of Drs. David Graber and Ted Chrisman with the cGMP measurements presented in this study.


   FOOTNOTES
 

Address for reprint requests and other correspondence: D. J. Garry, NB11.118A, 5323 Harry Hines Blvd., Univ. of Texas Southwestern Medical Center, Dallas, TX 75390-8573 (E-mail: daniel.garry{at}utsouthwestern.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 

  1. Adler S, Huang H, Loke KE, Xu X, Tada H, Laumas A, and Hintze TH. Endothelial nitric oxide synthase plays an essential role in regulation of renal oxygen consumption by NO. Am J Physiol Renal Physiol 280: F838–F843, 2001.[Abstract/Free Full Text]
  2. Barouch LA, Harrison RW, Skaf MW, Rosas GO, Cappola TP, Kobeissi ZA, Hobai IA, Lemmon CA, Burnett AL, O'Rourke B, Rodriguez ER, Huang PL, Lima JAC, Berkowitz DE, and Hare JM. Nitric oxide regulates the heart by spatial confinement of nitric oxide synthase isoforms. Nature 416: 337–340, 2002.[Medline]
  3. Boucher JL, Moali C, and Tenu JP. Nitric oxide biosynthesis, nitric oxide synthase inhibitors and arginase competition for L-arginine utilization. Cell Mol Life Sci 55: 1015–1028, 1999.[Web of Science][Medline]
  4. Brown GC. Nitric oxide and mitochondrial respiration. Biochim Biophys Acta 1411: 351–369, 1999.[Medline]
  5. Brown GC and Cooper CE. Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett 356: 295–298, 1994.[Web of Science][Medline]
  6. Brunori M, Giuffre A, Sarti P, Stubauer G, and Wilson MT. Nitric oxide and cellular respiration. Cell Mol Life Sci 56: 549–557, 1999.[Web of Science][Medline]
  7. Buchwalow IB, Schulze W, Karczewski P, Kostic MM, Wallukat G, Morwinski R, Krause EG, Müller J, Paul M, Slezak J, Luft FC, and Haller H. Inducible nitric oxide synthase in the myocardium. Mol Cell Biochem 217: 73–82, 2001.[Web of Science][Medline]
  8. Chen YJ, Traverse JH, Du R, Hou MX, and Bache R. Nitric oxide modulates myocardial oxygen consumption in the failing heart. Circulation 106: 273–279, 2002.[Abstract/Free Full Text]
  9. Chrisman TD and Garbers DL. Reciprocal antagonism coordinates C-type natriuretic peptide and mitogen-signaling pathways in fibroblasts. J Biol Chem 247: 4293–4299, 1999.
  10. Chung Y, Xu D, and Jue T. Nitrite oxidation of myoglobin in perfused myocardium: implication for energy coupling in respiration. Am J Physiol Heart Circ Physiol 271: H1166–H1173, 1996.[Abstract/Free Full Text]
  11. Cleeter MWJ, Cooper JM, Darley-Usmar VM, Moncada S, and Schapira AHV. Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. FEBS Lett 345: 50–54, 1994.[Web of Science][Medline]
  12. Clement E, Brown GC, Foxwell N, and Moncada S. On the mechanism by which vascular endothelial cells regulate their oxygen consumption. Proc Natl Acad Sci USA 96: 1559–1562, 1999.[Abstract/Free Full Text]
  13. Cooper CE. Nitric oxide and iron proteins. Biochim Biophys Acta 1411: 290–309, 1999.[Medline]
  14. Cotton JM, Kearney MT, MacCarthy PA, Grocott-Mason RM, McClean DR, Heymes C, Richardson PJ, and Shah AM. Effects of nitric oxide synthase inhibition on basal function and the force-frequency relationship in the normal and failing human heart in vivo. Circulation 104: 2318–2323, 2001.[Abstract/Free Full Text]
  15. DeBelder AJ, Radomski MW, Why HJF, Richardson PJ, Bucknall CA, Salas E, Martin JF, and Moncada S. Nitric oxide synthase activities in human myocardium. Lancet 341: 84–85, 1993.[Web of Science][Medline]
  16. Doeller JE and Wittenberg BA. Myoglobin function and energy metabolism of isolated cardiac myocytes: effect of sodium nitrite. Am J Physiol Heart Circ Physiol 261: H53–H62, 1991.[Abstract/Free Full Text]
  17. Domino SE, Tubb DL, and Garbers DL. Assay of guanylyl cyclase catalytic activity. Methods 195: 345–355, 1991.
  18. Finkel MS, Oddis CV, Jacob TD, Watkins SC, Hattler BG, and Simmons RL. Negative intropic effects of cytokines on the heart mediated by nitric oxide. Science 257: 387–389, 1992.[Abstract/Free Full Text]
  19. Flögel U, Merx M, Gödecke A, Decking KM, and Schrader J. Myoglobin: a scavenger of bioactive NO. Proc Natl Acad Sci USA 98: 735–740, 2001.[Abstract/Free Full Text]
  20. Garry DJ, Ordway GA, Lorenz JN, Radford NB, Chin ER, Grange RW, Bassel-Duby R, and Williams RS. Mice without myoglobin. Nature 395: 905–908, 1998.[Medline]
  21. Glabe A, Chung Y, Xu D, and Jue T. Carbon monoxide inhibition of regulatory pathways in myocardium. Am J Physiol Heart Circ Physiol 274: H2143–H2151, 1998.[Abstract/Free Full Text]
  22. Gödecke A, Flögel U, Zanger K, Ding Z, Hirchenhan J, Decking UKM, and Schrader J. Disruption of myoglobin in mice induces multiple compensatory mechanisms. Proc Natl Acad Sci USA 96: 10495–10500, 1999.[Abstract/Free Full Text]
  23. Grange RW, Meeson A, Chin E, Lau KS, Stull JT, Shelton JM, Williams RS, and Garry DJ. Functional and molecular adaptations in skeletal muscle of myoglobin-mutant mice. Am J Physiol Cell Physiol 281: C1487–C1494, 2001.[Abstract/Free Full Text]
  24. Habib FM, Springall DR, Davies GJ, Oakley CM, Yacoub MH, and Polak JM. Tumor necrosis factor and inducible nitric oxide synthase in dilated cardiomyopathy. Lancet 347: 1151–1155, 1996.[Web of Science][Medline]
  25. Haywood GA, Tsao PS, von der Leyen HE, Mann MJ, Keeling PJ, Trindade PT, Lewis NP, Byrne CD, Rickenbacher PR, Bishopric NH, Cooke JP, McKenna WJ, and Fowler MB. Expression of inducible nitric oxide synthase in human heart failure. Circulation 93: 1087–1094, 1996.[Abstract/Free Full Text]
  26. Henry Y and Guissani A. Interactions of nitric oxide with hemoproteins: roles of nitric oxide in mitochondria. Cell Mol Life Sci 55: 1003–1014, 1999.[Web of Science][Medline]
  27. Hochachka PW. The metabolic implications of intracellular circulation. Proc Natl Acad Sci USA 22: 12233–12239, 1999.
  28. Ishibashi T, Hamaguchi M, Kato K, Kawada T, Ohta H, Sasage H, and Imai S. Relationship between myoglobin contents and increases in cyclic GMP produced by glyceryl trinitrate and nitric oxide in rabbit aorta, right atrium and papillary muscle. Naunyn Schmiedebergs Arch Pharmacol 347: 553–561, 1993.[Web of Science][Medline]
  29. Kayar SR and Banchero N. Myocardial capillarity in acclimation to hypoxia. Pflügers Arch 404: 319–325, 1985.[Web of Science][Medline]
  30. Kojda G and Kottenberg K. Regulation of basal myocardial function by NO. Cardiovasc Res 41: 514–523, 1999.[Free Full Text]
  31. Li Q, Guo Y, Xuan YT, Lowenstein CJ, Stevenson SC, Prabhu SD, Wu WJ, Zhu Y, and Bolli R. Gene therapy with inducible nitric oxide synthase protects against myocardial infarction via a cyclooxygenase-2-dependent mechanism. Circ Res 92: 741–748, 2003.[Abstract/Free Full Text]
  32. Loke KE, Messina EJ, Shesely EG, Kaley G, and Hintze TH. Potential role of eNOS in the therapeutic control of myocardial oxygen consumption by ACE inhibitors and amlodipine. Cardiovasc Res 49: 86–93, 2001.[Abstract/Free Full Text]
  33. Maeso R, Navarro-Cid J, Rodrigo E, Ruilope LM, Cachofeiro V, and Lahera V. Effects of antihypertensive therapy on factors mediating endothelium-dependent relaxation in rats treated chronically with L-NAME. J Hypertens 17: 221–227, 1999.[Web of Science][Medline]
  34. Mattson DL. Comparison of arterial blood pressure in different strains of mice. Am J Hypertens 14: 405–408, 2001.[Web of Science][Medline]
  35. McMahon TJ, Moon RE, Luschinger BP, Carraway MS, Stone AE, Stolp BW, Gow AJ, Pawloski JR, Watke P, Singel DJ, Piantadosi CA, and Stamler JS. Nitric oxide in the human respiratory cycle. Nat Med 8: 711–717, 2002.[Web of Science][Medline]
  36. Meeson MP, Radford N, Shelton JM, Mammen PPA, DiMaio JM, Hutcheson K, Kong Y, Elterman J, Williams RS, and Garry DJ. Adaptive mechanisms that preserve cardiac function in mice without myoglobin. Circ Res 88: 713–720, 2001.[Abstract/Free Full Text]
  37. Mery PF, Lohmann SM, Walter U, and Fischmeister R. Ca2+ current is regulated by cyclic GMP-dependent protein kinase in mammalian cardiac myocytes. Proc Natl Acad Sci USA 88: 1197–1201, 1991.[Abstract/Free Full Text]
  38. Mery PF, Pavoine C, Belhassen L, Pecker F, and Fischmeister R. Nitric oxide regulates cardiac Ca2+ current. J Biol Chem 268: 26286–26295, 1993.[Abstract/Free Full Text]
  39. Michel JB, Xu Y, Blot S, Philippe M, and Chatellier G. Improved survival in rats administered NG-nitro L-arginine methyl ester due to converting enzyme inhibition. J Cardiovasc Pharmacol 28: 142–148, 1996.[Web of Science][Medline]
  40. Mohan P, Brutsaert DL, Paulus WJ, and Sys SU. Myocardial contractile response to nitric oxide and cGMP. Circulation 93: 1223–1229, 1996.[Abstract/Free Full Text]
  41. Mungrue IN, Gros R, You X, Pirani A, Azad A, Csont T, Schulz R, Butany J, Stewart DJ, and Husain MJ. Cardiomyocyte overexpression of iNOS in mice results in peroxynitrite generation, heart block, and sudden death. J Clin Invest 109: 735–743, 2002.[Web of Science][Medline]
  42. Navarro-Cid J, Maeso R, Rodrigo E, Munoz-Garcia R, Ruilope LM, Lahera V, and Cachofeiro V. Renal and vascular consequences of the chronic nitric oxide synthase inhibition. Am J Hypertens 9: 1077–1083, 1996.[Web of Science][Medline]
  43. North AJ, Lau KS, Brannon TS, Wu LC, Wells LB, German Z, and Shaul PW. Oxygen upregulates nitric oxidase synthase gene expression in ovine fetal pulmonary artery endothelial cells. Am J Physiol Lung Cell Mol Physiol 270: L643–L649, 1996.[Abstract/Free Full Text]
  44. O'Brien PJ, O'Grady M, McCutcheon IJ, Shen H, Nowack L, Horne RD, Mirsalimi SM, Julian RT, Grima EA, and Moe GW. Myocardial myoglobin deficiency in various animal models of congestive heart failure. J Mol Cell Cardiol 24: 721–730, 1992.[Web of Science][Medline]
  45. Paulus WJ, Frantz S, and Kelly RA. Nitric oxide and cardiac contractility in human heart failure. Circulation 104: 2260–2262, 2001.[Free Full Text]
  46. Pearce LL, Kanai AJ, Birder LA, Pitt BR, and Peterson J. The catabolic fate of nitric oxide. J Biol Chem 277: 13556–13562, 2002.[Abstract/Free Full Text]
  47. Roth DM, Swaney JS, Dalton ND, Gilpin EA, and Ross J. Impact of anesthesia on cardiac function during echocardiography in mice. Am J Physiol Heart Circ Physiol 282: H2134–H2140, 2002.[Abstract/Free Full Text]
  48. Rouet-Benzineb P, Eddahibi A, Raffestin B, Laplace M, Depond S, Adnot S, and Crozatier B. Induction of cardiac nitric oxide synthase 2 in rats exposed to chronic hypoxia. J Mol Cell Cardiol 31: 1697–1708, 1999.[Web of Science][Medline]
  49. Saito T, Hu F, Tayara L, Fahas L, Shennib H, and Giaid A. Inhibition of NOS II prevents myocardial infarction and congestive heart failure. Am J Physiol Heart Circ Physiol 283: H339–H345, 2002.[Abstract/Free Full Text]
  50. Sam F, Sawyer DB, Xie Z, Chang DLF, Ngoy S, Brenner DA, Siwik DA, Singh K, Apstein CS, and Colucci WS. Mice lacking inducible nitric oxide synthase have improved left ventricular contractile function and reduced apoptotic cell death late after myocardial infarction. Circ Res 89: 351–356, 2002.
  51. Schmidt HHH and Walter U. NO at work. Cell 78: 919–925, 1994.[Web of Science][Medline]
  52. Shah AM, Spurgeon HA, Sollott SJ, Talo A, and Lakatta EG. 8-Bromo-cGMP reduces the myofilament response to Ca2+ in intact cardiac myocytes. Circ Res 74: 970–978, 1994.[Abstract/Free Full Text]
  53. Shaul PW, Yuhanna IS, German Z, Chen Z, Steinhorn RH, and Morin FC III. Pulmonary endothelial NO synthase gene expression is decreased in fetal lambs with pulmonary hypertension. Am J Physiol Lung Cell Mol Physiol 272: L1005–L1012, 1997.[Abstract/Free Full Text]
  54. Shiva S, Brookes PS, Patel RP, Anderson PG, and DarleyUsmar VM. Nitric oxide partitioning into mitochondrial membranes and the control of respiration at cytochrome c oxidase. Proc Natl Acad Sci USA 98: 7212–7217, 2001.[Abstract/Free Full Text]
  55. Stamler JS. Redox signaling: nitrosylation and related target interactions of nitric oxide. Cell 78: 931–936, 1994.[Web of Science][Medline]
  56. Stumpe T, Deckling UKM, and Schrader J. Nitric oxide reduces energy supply by direct action on the respiratory chain in isolated cardiomyocytes. Am J Physiol Heart Circ Physiol 280: H2350–H2356, 2001.[Abstract/Free Full Text]
  57. Takahashi E, Endoh H, and Doi K. Visualization of myoglobin-facilitated mitochondrial O2 delivery in a single isolated cardiomyocyte. Biophys J 78: 3252–3259, 2000.[Web of Science][Medline]
  58. Taylor DJ, Matthews PM, and Radda GK. Myoglobin-dependent oxidative metabolism in the hypoxic rat heart. Respir Physiol 63: 275–283, 1986.[Web of Science][Medline]
  59. Trochu JN, Bouhour JB, Kaley G, and Hintze TH. Role of endothelium-derived nitric oxide in the regulation of cardiac oxygen metabolism. Circ Res 87: 1108–1117, 2000.[Abstract/Free Full Text]
  60. Vetterlein F, Hemeling H, Sammler J, Petho A, and Schmidt G. Hypoxia-induced acute changes in capillary and fiber density and capillary red cell distribution in the rat heart. Circ Res 64: 742–752, 1989.[Abstract/Free Full Text]
  61. Vetterlein F, Prange M, Lubrich D, Pedina J, Neckel M, and Schmidt G. Capillary perfusion pattern and microvascular geometry in heterogeneous hypoxic areas of hypoperfused rat myocardium. Am J Physiol Heart Circ Physiol 268: H2183–H2194, 1995.[Abstract/Free Full Text]
  62. Wittenberg BA and Wittenberg JB. Myoglobin-mediated oxygen delivery to mitochondria of isolated cardiac myocytes. Proc Natl Acad Sci USA 84: 7503–7507, 1987.[Abstract/Free Full Text]
  63. Wittenberg BA and Wittenberg JB. Transport of oxygen in muscle. Annu Rev Physiol 51: 857–878, 1989.[Web of Science][Medline]
  64. Yu AY, Shimoda LA, Iyer N, Huso DL, Sun X, McWilliams R, Beaty T, Sham JSK, Wiener CM, and Semenza GL. Impaired physiological responses to chronic hypoxia in mice partially deficient for hypoxia-inducible factor 1{alpha}. J Clin Invest 103: 691–696, 1999.[Web of Science][Medline]
  65. Zhao X, Sampath V, and Caughey WS. Infrared characterization of nitric oxide bonding to bovine heart cytochrome c oxidase and myoglobin. Biochem Biophys Res Commun 204: 537–543, 1994.[Web of Science][Medline]



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