HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) All Versions of this Article: 291/6/C1172    most recent 00195.2006v1 Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Benard, G. Articles by Rossignol, R. Search for Related Content PubMed PubMed Citation Articles by Benard, G. Articles by Rossignol, R.

MITOCHONDRIAL MODELING AND FUNCTION

Physiological diversity of mitochondrial oxidative phosphorylation

¹INSERM, U688 Physiopathologie Mitochondriale, Université Victor Segalen-Bordeaux 2, Bordeaux, France; and ²IFR 31, Institut Louis Bugnard, BP 84225, UMR 5018 CNRS UPS, Toulouse, France

Submitted 20 April 2006 ; accepted in final form 7 June 2006

	ABSTRACT

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
To investigate the physiological diversity in the regulation and control of mitochondrial oxidative phosphorylation, we determined the composition and functional features of the respiratory chain in muscle, heart, liver, kidney, and brain. First, we observed important variations in mitochondrial content and infrastructure via electron micrographs of the different tissue sections. Analyses of respiratory chain enzyme content by Western blot also showed large differences between tissues, in good correlation with the expression level of mitochondrial transcription factor A and the activity of citrate synthase. On the isolated mitochondria, we observed a conserved molar ratio between the respiratory chain complexes and a variable stoichiometry for coenzyme Q and cytochrome c, with typical values of [1–1.5]:[30–135]:[3]:[9–35]:[6.5–7.5] for complex II:coenzyme Q:complex III:cytochrome c:complex IV in the different tissues. The functional analysis revealed important differences in maximal velocities of respiratory chain complexes, with higher values in heart. However, calculation of the catalytic constants showed that brain contained the more active enzyme complexes. Hence, our study demonstrates that, in tissues, oxidative phosphorylation capacity is highly variable and diverse, as determined by different combinations of 1) the mitochondrial content, 2) the amount of respiratory chain complexes, and 3) their intrinsic activity. In all tissues, there was a large excess of enzyme capacity and intermediate substrate concentration, compared with what is required for state 3 respiration. To conclude, we submitted our data to a principal component analysis that revealed three groups of tissues: muscle and heart, brain, and liver and kidney.

respiratory chain; tissues; stoichiometry; cytochrome c; coenzyme Q

Here, we performed a global comparative study of oxidative phosphorylation on tissue lysates and mitochondria isolated from rat skeletal muscle, heart, liver, kidney, and brain under similar experimental conditions. For this, we used a wide array of biochemical techniques, including polarography, enzymology, Western blot, cytochrome spectroscopy, and HPLC coupled to electrochemical measurements (HPLC-EC). In this manner, we measured the relative and absolute content of respiratory chain complexes II, III, and IV, their kinetics, and stoichiometry with respective substrate. We also analyzed the global functioning of the respiratory chain by measuring the redox status of cytochrome c and CoQ₉ at state 3, and the respiratory control ratio (RCR). Mitochondrial structure and content were also determined in the different tissues. Our results show that, in tissues, oxidative phosphorylation capacity is highly variable and diverse, as determined by different combinations of 1) the mitochondrial content, 2) the amount of respiratory chain complexes, and 3) their intrinsic activity. We discuss these differences in regard to the variability of OXPHOS control and the dramatic tissue specificity of mitochondrial diseases (43, 50).

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Chemicals. All chemicals used for mitochondrial preparations and enzymological and polarographic studies were purchased from Sigma. Chemicals for Western blot experiments were purchased from Bio-Rad.

Animals. Male Wistar rats weighing 180–200 g, having free access to water and standard laboratory diet, were used for this study. The animals were killed by cervical shock and decapitation. All of the procedures were performed in accordance with institutional guidelines for the care and use of laboratory animals (Declaration of Helsinki and Title 45, U.S. Code of Federal Regulations, Part 46, Protection of Human Subjects). Our experimental protocol was performed in accordance with the guidelines of the French National Institute for Science and Medical Research (INSERM).

Preparation of rat muscle mitochondria. Rat muscle mitochondria were isolated by differential centrifugation. Muscle from two hind legs were collected in isolation medium I (in mM: 210 mannitol, 70 sucrose, 50 Tris·HCl, pH 7.4, and 10 K⁺ EDTA) and digested by trypsin (0.5 mg/g of muscle) for 30 min. The reaction was stopped by addition of trypsin inhibitor (soybean 3:1 inhibitor to trypsin). The homogenate was centrifuged at 1,000 g for 5 min. The supernatant was strained through gauze and centrifuged at 7,000 g for 10 min. The resulting pellet was resuspended in ice-cold isolation medium II (in mM: 225 mannitol, 75 sucrose, 10 Tris·HCl, pH 7.4, and 0.1 K⁺ EDTA), and a new series of centrifugations (1,000 and 7,000 g) was performed. The last mitochondrial pellet was resuspended into a minimum volume of isolation medium II to obtain a mitochondrial concentration between 50 and 80 mg/ml. Protein concentration was measured by the Biuret method using BSA as standard.

Preparation of rat liver and kidney mitochondria. Liver was collected in isolation medium A (in mM: 250 sucrose, 10 Tris·HCl, pH 7.6, and 1 K⁺ EGTA) and homogenized. The homogenate was centrifuged at 1,000 g for 5 min. The supernatant was strained through gauze and centrifuged at 7,000 g for 10 min. The resulting pellet was resuspended in ice-cold isolation medium B (in mM: 250 sucrose, 10 Tris·HCl, pH 7.6, and 0.1 K⁺ EGTA), and a new series of centrifugations (1,000 and 7,000 g) was performed. The last mitochondrial pellet was resuspended in a minimum volume of isolation medium B to obtain a mitochondrial concentration of between 50 and 70 mg/ml.

Preparation of rat brain mitochondria. Brain mitochondria were isolated from whole brain. Rats were killed by decapitation without stunning, and the brains were removed and homogenized in isolation buffer (in mM: 250 sucrose, 10 Tris·HCl, pH 7.4, and 0.5 K⁺ EDTA). The homogenate was centrifuged at 1,000 g for 5 min. The supernatant was strained through gauze and recentrifuged at 7,000 g for 10 min. The resulting pellet was resuspended in ice-cold isolation buffer, and a new series of centrifugations (1,000 and 7,000 g) was performed. The crude mitochondrial pellet was resuspended in a final volume of 10 ml in 3% Ficoll medium (3% Ficoll, 250 mM sucrose, 10 mM Tris·HCl, pH 7.4, and 0.5 mM K⁺ EDTA). This suspension was carefully layered onto 20 ml of 6% Ficoll medium (6% Ficoll, 250 mM sucrose, 10 mM Tris·HCl, pH 7.4, and 0.5 mM K⁺ EDTA) and centrifuged for 30 min at 11,500 g. The mitochondrial pellet was resuspended in isolation medium and centrifuged for 10 min at 12,500 g. The mitochondria were made up to a concentration of 50 mg protein/ml in the isolation buffer.

Transmission electron microscopy. To allow comparisons of the mitochondrial organization in muscle, heart, liver, kidney, and brain, we prepared sections from tissues obtained from the same animal. These were also submitted to the same procedure of fixation and coloration. First, the tissues were dissected immediately after death and fixed by immersion in solution A (2.5% glutaraldehyde, 4% paraformaldehyde, 4% saccharose, and 2% polyvinylpyrolidone in 0.1 M cacodylate buffer). These were sectioned in 40-mm³ blocks, immersed for 1 h in fixative solution A, and rinsed with water. The blocks were recut into smaller samples of 12 mm³ and immersed in a second fixative (solution B: 2% osmic acid in 0.1 M cacodylate buffer) for 1 h. After dehydration, the blocks were embedded in epoxy (epon) resin and cut into longitudinal or transversal sections 0.1 µm thick. The different sections were stained with a solution of uranyl acetate and lead citrate. Sections were observed on a Philips CM10 microscope. Morphometric analysis was performed by randomly analyzing selected tissue sections of the different tissues (n 5) obtained from three different animals.

Western blotting. Samples were diluted into SDS-PAGE tricine sample buffer (Bio-Rad) containing 2% -mercaptoethanol by incubation for 30 min at 37°C, and separated on a 10–22% SDS polyacrylamide gradient mini-gel (Bio-Rad) at 150 V. Proteins were transferred electrophoretically to 0.45-µm polyvinylidine difluoride membranes for 2 h at 100 mA in CAPS buffer (3.3 g CAPS, 1.5 liters of 10% methanol, pH 11) on ice. Membranes were blocked overnight in 5% milk-PBS + 0.02% azide and incubated for 3 h with primary antibodies purchased from Mitosciences. The antiporin antibody was purchased from Calbiochem. After three washes with PBS-0.05% Tween 20, the membranes were incubated for 2 h with horseradish peroxidase-conjugated goat anti-mouse antibody (Bio-Rad) diluted in 5% milk-PBS. This secondary antibody was detected using enhanced chemiluminescent Plus reagent (Amersham). The signal was quantified by densitometric analysis with the use of Image J (National Institutes of Health) sofware.

Respiration measurements. Mitochondrial oxygen consumption was monitored at 30°C in a 1-ml thermostatically controlled chamber equipped with a Clark oxygen electrode (Oxy 1, Hansatech) in respiration buffer [75 mM mannitol, 25 mM sucrose, 100 mM KCl, 10 mM Tris phosphate, 10 mM Tris·HCl, pH 7.4, 50 µM EDTA, plus respiratory substrates (10 mM pyruvate in presence of 10 mM malate)]. The mitochondrial concentration used for respiration measurements was 1 mg/ml, and state 3 was obtained by the addition of 2 mM ADP. Respiration rates were expressed in nanograms atom O per minute per milligrams of protein. The RCR is defined as the ratio of state 3 (in the presence of ADP) to state 4 (in the absence of ADP) respiratory rate. The uncoupling ratio was defined as the ratio of the uncoupled respiratory rate (measured in presence of 1–10 µM carbonyl cyanide 3-chlorophenyl hydrazone) to state 3.

Enzymatic determination. Assays of all respiratory chain enzyme activities were carried out spectrophotometrically at 30°C using a double-wavelength Xenius spectrophotometer from SAFAS (Monaco) and standardized reproducible methods as described previously (1). All activities were expressed in nanomoles per minute per milligram. The kinetic parameters (V_max and K_m) of complexes III and IV were obtained by fitting the experimental curve V = f[S], with the Michaelis-Henri equation, V = (V_max x [S])/(K_m + [S]), where [S] is substrate concentration and V is velocity, using Kaleida Graph 3.0.2 (Abelbeck software). Activity measurements were performed for complex III using 10 µg/ml of mitochondrial protein (muscle and liver) with a ubiquinone concentration ranging from 0 to 200 µM. For complex IV, we used 10 µg/ml of mitochondrial protein (muscle and liver) and a reduced cytochrome c concentration that also ranged from 0 to 200 µM.

Complex II (succinate dehydrogenase). The assay was performed by following the decrease in absorbance at 600 nm resulting from the reduction of 2,6-dichlorophenolindo-phenol in 1 ml of medium containing 60 mM KH₂PO₄ (pH 7.4), 3 mM KCN, 20 µg/ml rotenone, 20 mM succinate, and 10 µg mitochondrial protein. The reaction was initiated by the addition of 1.3 mM phenazine methasulfate and 0.18 mM 2,6-dichloroindophenol sodium salt hydrate. The extinction coefficient used for DCIP was 21 mM^–1·cm^–1.

Complex III (ubiquinol cytochrome c reductase). The oxidation of 6.5 mM decylubiquinol by complex III was determined by using cytochrome c (III) as an electron acceptor. The assay was carried out in basic medium supplemented with 2.5 mg/ml BSA, 15 µM cytochrome c (III), and 5 µg/ml rotenone. The reaction was started with 10 µg of mitochondrial protein, and the enzyme activity was measured at 550 nm. The extinction coefficient used for cytochrome c was 18.5 mM^–1·cm^–1.

Complex IV (cytochrome c-oxidase). Two methods were used for determining cytochrome c-oxidase activity. Initially, cytochrome c-oxidase activity was determined spectrophotometrically with cytochrome c (II) as substrate. The oxidation of cytochrome c was monitored at 550 nm at 30°C. The extinction coefficient used for cytochrome c was 18.5 mM^–1·cm^–1. In the second method, we monitored cytochrome c-oxidase activity by inhibiting the rest of the respiratory chain with rotenone and antimycin, using 3 mM ascorbate and 0.5 mM N,N,N',N'-tetramethyl-p-phenylenediamine, as an electron donor system.

Citrate synthase. The reduction of 5,5-dithiobis(2-nitrobenzoic acid) by citrate synthase at 412 nm (extinction coefficient of 13.6 mM^–1·cm^–1) was followed in a coupled reaction with coenzyme A and oxaloacetate. A reaction mixture of 0.2 M Tris·HCl, pH 8.0, 0.1 mM acetyl-coenzyme A, 0.1 mM 5,5-dithiobis(2-nitrobenzoic acid), and 5–20 µg of muscle or brain mitochondrial protein was incubated at 30°C for 5 min. The reaction was initiated by the addition of 0.5 mM oxaloacetate, and the absorbance change was monitored for 5 min.

Determination of cytochrome b_II, b_H, b_L, c, c₁, and aa₃ absolute content by spectrophotometry. Mitochondria isolated from muscle and liver were used at 1 mg/ml, in 1 ml of the respiratory buffer described above. Individual fully reduced (with excess sodium dithionite) or fully oxidized (with excess ferricyanide) absorbance spectra were recorded between 500 and 650 nm on a SAFAS Genius double-wavelength spectrophotometer (SAFAS, Monaco). The concentration of cytochrome aa₃ ([aa₃]) contained in the heme of cytochrome c-oxidase (complex IV) was determined from the difference spectrum [reduced – oxidized (red-ox)] at the maximum absorption value of 605 nm (A_{red-ox 605}), normalized by the absorbance of the isobestic point at 630 nm (A_{red-ox 630}). Values were calculated by the Beer-Lambert law (see Eq. 1), with an extinction coefficient _{red-ox aa3} of 24,000 M^–1·cm^–1 (40) and a cuvette length of 1 cm

(1)

The absolute content of c-type cytochromes (c + c₁) was determined from the same difference spectrum (reduced – oxidized) at the maximum absorption value of 550 nm (A_{red-ox 550}), normalized by the absorbance of the isobestic point at 535 nm (A_{red-ox 535}). The concentration of cytochrome c + c₁ was determined according to the Beer-Lambert law (see Eq. 2), with an extinction coefficient _{red-ox c+c1} of 18,500 M^–1·cm^–1 (26) and a cuvette length of 1 cm

(2)

The total content of cytochrome c was then calculated from this total content in cytochrome c + c₁ by subtracting the contribution of c₁, which was determined by considering its molar equivalence of 1:1, with cytochrome b_H (22). This content in cytochrome b_H was determined experimentally, using a detergent-based partial solubilization of mitochondrial proteins as described in Ref. 40, which leads to the physical separation of complex II and complex III. In this manner, it is possible to determine the part of the absorbance of the different b-type cytochromes from the total (b_H + b_L + b_II), calculated from the maximum absorption peak at 562 nm (A_{red-ox 562}), normalized by the absorbance of the isobestic point at 575 nm (A_{red-ox 575}). The total extinction coefficient of b_H + b_L (_{red-ox bH+L}) was equal to 56,100 M^–1·cm^–1 and that of b_II (_{red-ox bII}) was 26,000 M^–1·cm^–1 (40). The cuvette length was 1 cm. In this manner, the concentration of cytochrome c₁ was calculated from that of b_H according to Eq. 3:

(3)

(4)

with R = b_II/b_total (ratio of the absorbance of cytochrome b_II to the total absorbance of the b-type cytochromes at 562 nm). The total content of cytochrome c was then obtained from c + c₁ (see Eq. 4) by subtracting the amount of c₁ (see Eq. 3), determined in the same volume of 1 ml. This content of cytochrome c was expressed in picomoles per milligram protein.

Determination of steady-state cytochrome c redox status by spectrophotometry. To determine the concentration of reduced cytochrome c during state 3, we performed the above detailed analysis of cytochrome spectra on mitochondria taken from muscle or liver mitochondria respiring as described above in Respiration measurements. After the addition of substrates pyruvate and malate, state 3 respiration was initiated by the addition of 2 mM ADP. Absorbance spectra were recorded individually every 40 s in the presence of different concentrations of myxothiazol or KCN. Steady state was typically reached within 1 min after the addition of ADP, as observed by polarography on the same mitochondrial samples.

Determination of CoQ absolute content and steady-state redox status by HPLC-EC measurement. The absolute CoQ content (oxidized + reduced form) was measured by HPLC-EC as described in Ref. 12. Determination of the concentration of reduced CoQ during state 3 was performed on mitochondria respiring at steady state as described for cytochrome c in Respiration measurements above. During each steady-state assay, respiration samples were withdrawn from the cuvette and immediately stored at –80°C.

Statistical analysis. All of the data presented in this study correspond to the mean value of n experiments ± SD, with n 3. Comparison of the data obtained from isolated muscle and liver mitochondria were performed with Student's t-test, using Excel software (Microsoft). Two sets of data were considered as statistically different when P < 0.05.

Principal component analysis. The principal component analysis (PCA) method is concerned with interpreting the variance-covariance structure through a few linear combinations of the original variables. Its general objectives are dimensionality reduction and ease of interpretation. PCA is performed according to the usual method with Statbox version 3.2 (Grimmersoft) as described in Ref. 8. If we restrict ourselves to the first two principal components, the results may be represented graphically by a biplot.

	RESULTS

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mitochondrial content in the different tissues. We evaluated the global mitochondrial content in tissues by measuring the activity of citrate synthase in tissue lysates. We obtained values of 11,155 ± 576 nmol·min^–1·mg^–1 in heart, compared with 7,415 ± 1,230 in muscle, 3,798 ± 521 in brain, 1,516 ± 71 in kidney, and 736 ± 66 in liver. As another method to evaluate the mitochondrial content in tissues, we determined the amount of respiratory chain complexes I and III per milligram of tissue proteins by Western blot. We also measured the expression level of the mitochondrial transcription factor A (mtTFA). An example of Western blot is shown in Fig. 1A, inset. The corresponding results of the densitometric analysis are given in the histogram of Fig. 1A. It can be seen that heart contains the highest content of complex III core 2 per microgram of tissue proteins, compared with muscle, brain, liver, and kidney, respectively. A very similar hierarchy between tissues can be observed for the expression level of the complex I 20-kDa subunit, as well as for mtTFA (Fig. 1A). Comparison of the tissue respiratory chain content determined by Western blot (complex III core 2) with the activity of citrate synthase (Fig. 1C) revealed a strong correlation (R² = 0.974). Likewise, there was a good correlation between the expression level of mtTFA and that of the complex I 20-kDa subunit (R² = 0.90) or complex III core 2 (R² = 0.98) in tissues.

View larger version (23K): [in this window] [in a new window]   Fig. 1. Respiratory chain content in tissue lysates and correlation with citrate synthase activity or mitochondrial transcription factor A (mtTFA) expression level. A: contents in complex III (CIII) core 2, 20-kDa subunit of complex I (CI 20), and mtTFA were determined by Western blot on 5 µg of protein of the different tissues lysates. Each band density was measured by densitometry and represented in the histogram: CIII (solid bars), mtTFA (dark gray bars), and CI 20 (light gray bars). M, muscle; H, heart; L, liver; K, kidney; B, brain. B: maximal activity of citrate synthase was measured on these different tissue lysates (expressed in nmol·min–1·mg protein–1). C: mtTFA (), CI 20 (), and CIII core 2 () band densities were expressed as a function of citrate synthase activity in the different tissues. Correlation factors (R2) are given at the end of each regression line. Data represent mean values ± SD, as determined on 3 different experiments.

View larger version (141K): [in this window] [in a new window]   Fig. 2. Electron micrographs of mitochondria from different tissues. Mitochondrial and cellular organizations from thin tissue sections was investigated on electron micrograph. A: skeletal muscle (slow type; soleus, longitudinal). B: skeletal muscle (rapid type; peroneus digiti quarti, longitudinal). C: heart (longitudinal). D: liver. E: kidney (the basal corpus and part of the microvilosity of the tubule are represented). F: brain (the basal corpus and the beginning of the axonal region are represented; they correspond to pyramidal region of the cerebral hemispheres). These micrographs were compared at the same magnification and correspond to tissue sections taken from the same animal, prepared according to the same method. Bars = 2 µm.

View larger version (30K): [in this window] [in a new window]   Fig. 3. Comparison of the respiratory chain content in mitochondria isolated from different tissues. Content in respiratory chain complexes I–V were determined in mitochondria isolated from different tissues by performing Western blots on 5 µg of mitochondrial proteins. First, we verified that the signal of each band was not saturated when the amount of mitochondrial proteins was increased from 5 to 10 µg (inset). Each band intensity was determined by densitometry and normalized to that of the porin. In this manner, the histogram can show the relative expression levels of CI 20, 70 kDa subunit of complex II (CII 70), CIII core 2, 5a subunit of complex IV (CIV 5a), and -subunit of complex V (CV) in mitochondria isolated from skeletal muscle (solid bars), heart (dark gray bars), liver (gray bars), kidney (light gray bars), and brain (open bars). Data represent mean values ± SD, as determined on 3 different experiments.

View larger version (16K): [in this window] [in a new window]   Fig. 4. Absolute content in respiratory chain b-type cytochrome in mitochondria isolated from different tissues. The content in the different cytochromes present in the respiratory chain was calculated from the difference absorbance spectra (inset), as explained in EXPERIMENTAL PROCEDURES. This required the physical separation of CII and CIII according to the method developed by Schägger et al. (40). This allowed us to calculate the individual content in cytochrome bL + bH (dark gray sections) and bII (light gray sections), expressed in pmol/mg of mitochondrial proteins.

Respiratory chain complex-specific activity. V_max and K_m of respiratory chain complexes II, III, and IV were measured on mitochondria isolated from the different tissues. Hence, they must be referred to as the apparent kinetic parameters (V_max-app and K_m-app) because they were not measured on the pure enzyme. The maximal activity of citrate synthase was also determined on these mitochondrial preparations. All of the values obtained are listed in Table 2. Results show important differences between enzyme complexes, with complex IV presenting the highest rate of substrate oxidation, compared with complexes III, II, and I. Comparisons of these activities in the different tissues show that the heart presents with the highest velocities, compared with muscle, brain, liver, and kidney, respectively. The same hierarchy between tissues was also observed for the other complexes (Table 2). These differences in V_max-app between tissues could be attributed to variations in the amount of respiratory chain complexes and/or in their catalytic constant. Thus, to evaluate the catalytic constant (k_cat) of respiratory chain complexes II, III, and IV in tissues, we divided their maximal activity (Table 2) by their absolute content (see Table 1), determined on the same isolated mitochondria. In this manner, we obtained important differences in the k_cat values of complex IV, which presented with a higher value in brain (17,232 ± 2,184 min^–1) than in liver (13,043 ± 1,497 min^–1), heart (10,900 ± 1,562 min^–1), muscle (8,339 ± 974 min^–1), and kidney (6,436 ± 748 min^–1). For complex III, we obtained the highest k_cat value in brain (13,377 ± 1,544 min^–1) compared with muscle (6,371 ± 732 min^–1), heart (5,201 ± 497 min^–1), liver (6,337 ± 541 min^–1), and kidney (5,431 ± 630 min^–1).

View larger version (12K): [in this window] [in a new window]   Fig. 5. Principal component analysis of the main mitochondrial features in different tissues: biplot representation of the PCA results for muscle, heart, liver, and kidney. The variables correspond to the mean value of the following parameters obtained in the different tissues: CIII content, CIV content, CIII apparent (app) Km, CIV apparent Km, coenzyme Q9 (CoQ9) content, cytochrome c (cyt c) content, state 3 reduction level of CoQ9, state 3 reduction level of cytochrome c, biochemical threshold value of CIII (taken from Ref. 37), and biochemical threshold value of CIV (taken from Ref. 37). CS, citrate synthase.

	DISCUSSION

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

First, we looked at the mitochondrial content in the different tissues. This simple question raised numerous issues concerning the accuracy of the different methods of quantification generally employed. For instance, the classic measurement of mitochondrial section area on electron micrographs is limited by the very complex three-dimensional organization of mitochondria as a global network (16, 31, 51), the morphometric parameters of which also vary widely on energy status (17–19, 23, 35). Accordingly, in our study, there was an important variability of mitochondrial morphology between tissues: in heart and skeletal muscle, they presented essentially with a regular quasi-crystalline organization (46), whereas in liver and kidney they looked more irregular and less compacted by tissue architecture. In brain, the mitochondria looked more like a collection of small and numerous ovoid sections. Moreover, we observed a strong heterogeneity of mitochondrial intracellular distribution within tissues, so that the number of organellar profiles varies dramatically with the localization of the section. For instance, in brain, there are few mitochondria around the nucleus, in the basal corpus, whereas they are omnipresent along the axone, reaching a centimeter range in length. Likewise, in kidney, there are more mitochondria in the tubule (microvilosities) than in the basal corpus. In the other tissues, mitochondria present with a less heterogeneous organization. The internal organization of mitochondria, i.e., the form of the cristae, is also irregular and variable on physiological conditions (28). Accordingly, in our study, in tissue sections viewed by transmission electron microscopy, we observed important differences in the internal organization of the mitochondria. For instance, there was a variable number of cristae per organellar section, along with differences in their arrangement. The heart presented with the highest number of cristae per surface of mitochondrial section vs. the other tissues (factor of 1.3–4.8). Moreover, we observed variations in the density of the mitochondrial matrix, compared with the cytosol, with again a high value in heart vs. that shown in the other tissues (factor of 1.4–1.9). A high number of cristae, as observed in heart, could indicate a higher mitochondrial content in respiratory chain complexes, in agreement with studies showing a preferential localization of these proteins in the crystal membranes (11, 14). Accordingly, heart mitochondria also present with the darker matrix, which could indicate a higher state of respiration in vivo (17–19). Another method of mitochondrial quantification frequently used is the determination of mitochondrial-to-tissue protein ratio. However, in our conditions, it could not be used to compare accurately the mitochondrial content of the different tissues, given the important variability in the yield of the different methods of organellar isolation from the various tissues.

To avoid these errors, we determined the content of mitochondria on total lysates from the different tissues, by measuring 1) the citrate synthase activity and 2) the expression level of respiratory chain complexes I and III, as well as mtTFA. The results show that heart contains more mitochondria than muscle, liver, brain, and kidney. This is in good correlation with the expression level of mtTFA, suggesting the importance of that protein in determining the tissue OXPHOS capacity. We also observed a good correlation between citrate synthase activity and complex III or complex I content in tissues, confirming its utilization as a marker of the respiratory chain content in tissues. This also suggests that the citrate synthase expression level could be coordinated with that of the respiratory chain complexes.

In the second part of our study, we questioned the possible qualitative differences in mitochondria isolated from the different tissues, regardless of their variable tissue content, as discussed above. The analysis of the relative expression level of seven proteins of the respiratory chain by Western blot indicates a higher content of complex I, II, III, IV, and V in heart- and muscle-isolated mitochondria than in liver, kidney and brain. For instance, the complex III core 2 subunit was more expressed in heart-isolated mitochondria (set as 100%) than in muscle (86 ± 17%), kidney (39 ± 09%), liver (34 ± 06%), and brain (25 ± 07%). This is also true for the other proteins analyzed, suggesting a diversity in intramitochondrial respiratory chain content between tissues. We also determined the absolute content in cytochromes b_H, b_L, b_II, c, c₁, and aa₃, as well as the content in CoQ₉ on the isolated mitochondria. The results show that the highest absolute contents in respiratory chain complexes are in muscle and heart mitochondria, compared with liver, kidney, and brain (factor of 3). However, the molar ratio between complex II, III, and IV was conserved, with boundary values of [0.89–1.45]:[3]:[6.5–7.1]. We chose to set the complex III content at 3 to allow comparison with values in the literature. Indeed, other authors have obtained similar values for the molar ratio between respiratory chain complexes determined on bovine heart using a variety of techniques (4, 20, 40). Our results also support the general idea that fixed values between the different complexes could constitute a strong argument for the "supercomplexes hypothesis" (41). In our study, we precise this organization by determining the actual stoichiometry between respiratory chain complexes and respective substrates. We observe a large variability between tissues, with a higher coenzyme Q₉-to-complex III ratio in liver, than in kidney, muscle, brain, and heart. This is in agreement with values (ranging between 11 and 32) obtained previously in bovine heart (4). The cytochrome c-to-complex IV ratio is higher in brain than in kidney, muscle, heart, and liver. Again, this is in agreement with values obtained in bovine heart (4). Hence, the total content in these two intermediate substrates is not correlated with the expression levels of the different respiratory chain complexes determined in tissues. These different stoichiometries could reflect the adaptability of the tissues to a variable energy demand. However, the values that we obtained might not correspond to the actual stoichiometries, as recent studies also suggest the existence of different physical pools of cytochrome c (32, 42, 48) and CoQ (2) that could play different functions in the cell.

The functional analysis revealed a large diversity in the V_max-app values obtained for complexes II, III, and IV, with muscle and heart generally presenting with the highest values. Such differences in V_max could be explained by a variable content in respiratory chain complexes per milligram of mitochondrial proteins and/or a difference in the k_cat. To take into account the content in respiratory chain complexes, we normalized these V_max-app values to those of citrate synthase. The results show that, after normalization by citrate synthase activity, the highest velocities of complexes II, III, IV are observed in the brain, kidney, and liver, compared with muscle and heart (factor of 2 between brain and heart). To look more closely at the differences between respiratory chain complexes in tissues, we calculated the k_cat values by dividing V_max-app by the molar content in enzyme complex. Again, we observe a higher k_cat for brain than for the other tissues (factor of 1.4–2.6 for complex IV and 2.1–2.6 for complex III). The k_cat values that we obtained are similar to those obtained on pure complex IV (47) and present with comparable variations between the type of tissue.

In the last part of our analysis, we looked at the differences in the integrated function of the respiratory chain between the different tissues. First, we verified the integrity of our mitochondrial preparations, as a prerequisite for the analysis of the respiratory rate. We considered the following indexes: state 3 value of respiration and RCR, as RCR alone is not a good indicator, given its important variability on several parameters, even on intact mitochondria (44). The analysis of the respiratory flux values measured at state 3 revealed an apparent discrepancy. Indeed, although skeletal muscle- and heart-isolated mitochondria present with higher respiratory chain enzyme contents, along with higher V_max values, the state 3 respiratory rate is not different. This could be explained by the fact that most of the control of mitochondrial respiration is supported, at state 3, by the pyruvate carrier (36). However, the situation must be very different in vivo as the various tissues consume different physiological energy substrates, with important preferences in their utilization (27). The steady-state analysis of mitochondrial respiration also reveals large differences between the maximal respiratory chain complexes activity and the flux value, i.e., their velocity at state 3. This indicates again that, in our conditions, the rate of mitochondrial respiration is not tightly controlled by the sole content in OXPHOS complexes, nor their maximal velocities. For instance, the maximal activity of complex IV exceeds by a factor of 15 the activity measured during state 3 respiration in heart-isolated mitochondria. This factor is equal to 12.6, 12.2, 8.1, and 4.9 in muscle, brain, liver, and kidney, respectively. For complex III, this excess capacity ranges from 1.8 to 4.2 in the different tissues. The observation that complex IV does not function at the maximal velocity during state 3 respiration led to the definition of the "excess capacity" (15). This excess could be utilized, at least in part, to accommodate the flux to an increase in energy demand, so that tissues with a high excess capacity could adapt more easily to large-scale variations in energy demand. Interestingly, our observations allow us to extend this notion of excess capacity to the intermediary substrates of the respiratory chain. Indeed, the comparison of the amount of reduced CoQ₉ determined during state 3, compared with the total content, reveals that only 2% is reduced during steady-state 3 mitochondrial respiration in heart, muscle, and brain. Conversely, this proportion is equal to 60% in liver and kidney, using the same substrate combination (pyruvate-malate). For cytochrome c, the fraction used at state 3 is 65% of the total in heart and muscle, compared with 40% in liver, kidney, and brain. Therefore, our results indicate that not all the substrate is engaged (reduced) during mitochondrial respiration. This raises again the problem of a possible existence of different physical and functional pools of mitochondrial CoQ and cytochrome c, as well as their possible compartmentation. This could also be explained by the equilibrium constant or bypass reactions, such as superoxide production from the reduced cytochrome c. Hence, our work generalizes the existence of an excess of both enzyme capacities and substrates concentrations in mitochondria from different tissues. This could be of particular importance for the control of mitochondrial energy production, as well as the physiological adaptation to a sudden change in energy demand or substrate delivery. Such differences could also participate to the observed tissue specificity of mitochondrial diseases. In particular, the excess capacity could be utilized for the compensation of pathological defects in respiratory chain activity and determine in part the biochemical threshold value (37).

To conclude, we aimed to compare the different tissues, while taking into account the numerous data obtained in our study. For this, we performed a PCA that revealed three groups of tissues: 1) heart and muscle (slow type), 2) liver and kidney, and 3) brain. They present with important differences at the level of respiratory chain content, composition, activity, and flux response. Interestingly, each group represents organs of the same embryonic origin, which could suggest that OXPHOS features could be set up early in the tissue development period, or closely related to the type of tissue function. The first group, skeletal muscle and heart, presents with the highest OXPHOS capacity and a low resistance against the occurrence of respiratory chain perturbation, as illustrated by lower threshold values and high control coefficients. Conversely, the second group, liver and kidney, is characterized by a lower OXPHOS capacity and a lower sensitivity to OXPHOS defects. The third group, which contains solely the brain, is between the first and second group regarding the OXPHOS capacity and flux response. More generally, our study demonstrates that oxidative phosphorylation capacity is highly variable and diverse in tissues. It appears to be determined by different combinations of the mitochondrial content, the amount of respiratory chain complexes, and their intrinsic activity (see Fig. 6). This underlines the complexity of the regulation of mitochondrial energy production, which can occur at the level of 1) organellar biogenesis, 2) mitochondrial or nuclear DNA transcription, and 3) enzyme regulation. Different signaling pathways have been described for the modulation of each of those determinants, but their overall control and orchestration are presently not understood. The present observations of a different dosage of mitochondrial content, respiratory chain expression levels, and enzyme complex intrinsic activity in the different tissues also suggest that mitochondrial metabolism is tailored to meet organ-specific features. For instance, a larger mitochondrial compartment per surface of tissue could allow better access to oxygen more efficiently and could allow delivery of ATP throughout the cytosolic compartment. The mitochondrial content and architecture could also be related to some tissue-specific adaptative needs, such as mechanosensing in skeletal muscle (33). Moreover, the different options of a higher content in enzyme complexes (i.e., heart) than more active ones (i.e., brain) could determine differences in tissue sensitivity to changes in energy demand, controlled by the concentration of available substrates. Accordingly, determination of K_m for cytochrome c for complex IV revealed a twice lower value in brain (11.3 ± 0.9 µM) than in heart (28.2 ± 2.7 µM). Hence, the role of energy demand and the type of substrate utilized for energy production could play an important role in defining the mitochondrial compositional features and steady-state functioning characteristics. Accordingly, our group (35) demonstrated previously on living human cells that a change in the type of energy substrate was accompanied by a rapid modulation of the expression of mitochondrial proteins, as well as organellar structural parameters. Our group (7) also showed that physical exercise could modify the content in mitochondria and the amount of respiratory chain complexes in skeletal muscle. Thus, to further detail the diversity in the OXPHOS system between tissues, it will be necessary to consider the numerous regulations that can occur at the level of the respiratory chain or at the level of the mitochondrial network morphofunctional characteristics. For instance, recent evidences suggest that changes in mitochondrial activity can trigger morphological adaptations of the mitochondrial network (23, 35), and clinical studies further indicate that molecular defects affecting its dynamics lead to pathology (38). This suggests that a link between energy status and organellar network configuration must exist. Indeed, mitochondrial overall configuration is controlled by a balance between fusion and fission events, mediated by specific proteins (5, 25) that could participate in the modulation of energy production. Accordingly, the different tissues present with variable expression levels of the fusion and fission proteins, possibly related to specific energy needs. Also, the in-depth study of the other mitochondrial functions related to the OXPHOS system will contribute to understand what regulates mitochondrial structural and functional features in tissues. Our results have implications for the understanding of mitochondrial physiopathology. Indeed, the clinical manifestation of respiratory chain disorders typically present with a tissue specificity, characterized by the fact that a given pathological mutation can affect the different tissues to a variable extent (37). Our data suggest that mutations affecting either the amount of active complexes (i.e., Surf 1) or their catalytic activity (i.e., point mutations) will lead to different degrees of energy defect according to the tissue considered.

View larger version (19K): [in this window] [in a new window]   Fig. 6. Representation of the respiratory chain in different tissues: For each tissue, we show the respiratory chain system, taking into account the content in mitochondria, the amount of respiratory chain complexes II, III, and IV, their intrinsic activity, and the total content in intermediate substrate. We also figured the excess capacity of these enzyme complexes and the excess concentration of reduceable substrate. The amount of mitochondria was represented in the background by a yellow rectangle, the surface of which is proportional to their tissue amounts (as determined by the CS activity). The content in enzyme complexes (gray squares) and substrates (blue squares) is proportional to the surface of the rectangles. Moreover, their color intensity is proportional to the catalytic constant (kcat) (for the enzymes). For the substrates, we figured the excess that is not utilized at state 3 (light blue) compared with the content utilized at state 3 (dark blue). To illustrate the diversity in the determinism of the OXPHOS capacity in tissues, we can compare the heart and the brain, looking more closely at CIII. The heart contains more mitochondria than the brain (factor of 3) and more complex III (factor of 9), but a less active enzyme (factor of 0.4). Thus the overall capacity of CIII in heart is 11 times higher (per mg of tissue proteins) than in brain. However, in both tissues, the excess capacity of CIII is comparable (35% in heart and 25% in brain).

	ACKNOWLEDGMENTS

 
The authors thank INSERM, Université Victor Segalen Bordeaux 2, Association Française contre les Myopathies (AFM), Association contre les Maladies Mitochondriales (Ammi), Ligue contre le Cancer, and Région Aquitaine for financial support. We are also grateful to Jean-Pierre Mazat for discussion. We also thank Devin Oglesbee for text corrections.

	FOOTNOTES

 

Address for reprint requests and other correspondence: R. Rossignol, INSERM U688, Physiopathologie mitochondriale, Université Victor Segalen-Bordeaux 2, 146 rue Léo-Saignat, F-33076 Bordeaux Cedex, France (e-mail: rossig{at}u-bordeaux2.fr)

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 EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
1. Aleardi AM, Benard G, Augereau O, Malgat M, Talbot JC, Mazat JP, Letellier T, Dachary-Prigent J, Solaini GC, and Rossignol R. Gradual alteration of mitochondrial structure and function by -amyloids: importance of membrane viscosity changes, energy deprivation, reactive oxygen species production, and cytochrome c release. J Bioenerg Biomembr 37: 207–225, 2005.[CrossRef][Web of Science][Medline]

2. Bianchi C, Genova ML, Parenti Castelli G, and Lenaz G. The mitochondrial respiratory chain is partially organized in a supercomplex assembly: kinetic evidence using flux control analysis. J Biol Chem 279: 36562–36569, 2004.[Abstract/Free Full Text]

3. Cairns CB, Walther J, Harken AH, and Banerjee A. Mitochondrial oxidative phosphorylation thermodynamic efficiencies reflect physiological organ roles. Am J Physiol Regul Integr Comp Physiol 274: R1376–R1383, 1998.[Abstract/Free Full Text]

4. Capaldi RA, Halphen DG, Zhang YZ, and Yanamura W. Complexity and tissue specificity of the mitochondrial respiratory chain. J Bioenergetics and Biomembranes 20: 291–311, 1988.[CrossRef][Web of Science][Medline]

5. Chan DC. Mitochondrial fusion and fission in mammals. Annu Rev Cell Dev Biol First published May 11, 2006; doi:10.1146/annurev.cellbio.22.010305.104638.

6. Collins TJ, Berridge MJ, Lipp P, and Bootman MD. Mitochondria are morphologically and functionally heterogeneous within cells. EMBO J 21: 1616–1627, 2002.[CrossRef][Web of Science][Medline]

7. Duclos M, Gouarne C, Martin C, Rocher C, Mormede P, and Letellier T. Effects of corticosterone on muscle mitochondria identifying different sensitivity to glucocorticoids in Lewis and Fischer rats. Am J Physiol Endocrinol Metab 286: E159–E167, 2004.[Abstract/Free Full Text]

8. Durrieu G, Letellier T, Antoch J, Deshouillers J, Malgat M, and Mazat J. Identification of mitochondrial deficiency using principal component analysis. Mol Cell Biochem 174: 149–156, 1997.[CrossRef][Web of Science][Medline]

9. Fell D. Metabolic control analysis: a survey of its theoretical and experimental development. Biochem J 286: 313–330, 1992.[Web of Science][Medline]

10. Forner F, Foster LJ, Campanaro S, Valle G, and Mann M. Quantitative proteomic comparison of rat mitochondria from muscle, heart and liver. Mol Cell Proteomics 5: 608–619, 2006.[Abstract/Free Full Text]

11. Frey TG, Renken CW, and Perkins GA. Insight into mitochondrial structure and function from electron tomography. Biochim Biophys Acta 1555: 196–203, 2002.[Medline]

12. Galinier A, Carriere A, Fernandez Y, Bessac AM, Caspar-Bauguil S, Periquet B, Comtat M, Thouvenot JP, and Casteilla L. Biological validation of coenzyme Q redox state by HPLC-EC measurement: relationship between coenzyme Q redox state and coenzyme Q content in rat tissues. FEBS Lett 578: 53–57, 2004.[CrossRef][Web of Science][Medline]

13. Ghadially F. Ultrastructural Pathology of the Cell and Matrix. Boston, MA: Butterworth-Heinemann, 1997.

14. Gilkerson RW, Selker JM, and Capaldi RA. The cristal membrane of mitochondria is the principal site of oxidative phosphorylation. FEBS Lett 546: 355–358, 2003.[CrossRef][Web of Science][Medline]

15. Gnaiger E, Lassnig B, Kuznetsov A, Rieger G, and Margreiter R. Mitochondrial oxygen affinity, respiratory flux control and excess capacity of cytochrome c oxidase. J Exp Biol 201: 1129–1139, 1998.[Abstract]

16. Griparic L and van der Bliek AM. The many shapes of mitochondrial membranes. Traffic 2: 235–244., 2001.[CrossRef][Web of Science][Medline]

17. Hackenbrock CR. Ultrastructural bases for metabolically linked mechanical activity in mitochondria. I. Reversible ultrastructural changes with change in metabolic steady state in isolated liver mitochondria. J Cell Biol 30: 269–297, 1966.[Abstract/Free Full Text]

18. Hackenbrock CR. Ultrastructural bases for metabolically linked mechanical activity in mitochondria. II. Electron transport-linked ultrastructural transformations in mitochondria. J Cell Biol 37: 345–369, 1968.[Abstract/Free Full Text]

19. Hackenbrock CR, Rehn TG, Weinbach EC, and Lemasters JJ. Oxidative phosphorylation and ultrastructural transformation in mitochondria in the intact ascites tumor cell. J Cell Biol 51: 123–137, 1971.[Abstract/Free Full Text]

20. Hatefi Y. The mitochondrial electron transport and oxidative phophorylation system. Annu Rev Biochem 54: 1015–1069, 1985.[CrossRef][Web of Science][Medline]

21. Heinrich R and Rapoport TA. A linear steady-state treatment of enzymatic chains. General properties, control and effector strength. Eur J Biochem 42: 89–95, 1974.[Web of Science][Medline]

22. Iwata S, Lee JW, Okada K, Lee JK, Iwata M, Rasmussen B, Link TA, Ramaswamy S, and Jap BK. Complete structure of the 11-subunit bovine mitochondrial cytochrome bc1 complex. Science 281: 64–71, 1998.[Abstract/Free Full Text]

23. Jakobs S, Martini N, Schauss AC, Egner A, Westermann B, and Hell SW. Spatial and temporal dynamics of budding yeast mitochondria lacking the division component Fis1p. J Cell Sci 116: 2005–2014, 2003.[Abstract/Free Full Text]

24. Kacser H and Burns JA. The control of flux. In: Rate Control of Biological Processes, edited by Davies DD. Cambridge Univ. Press, 1973, p. 65–104.

25. Karbowski M and Youle RJ. Dynamics of mitochondrial morphology in healthy cells and during apoptosis. Cell Death Differ 10: 870–880, 2003.[CrossRef][Web of Science][Medline]

26. Kuhn-Nentwig L and Kadenbach B. Isolation and properties of cytochrome c oxidase from rat liver and quantification of immunological differences between isozymes from various rat tissues with subunit-specific antisera. Eur J Biochem 149: 147–158, 1985.[Web of Science][Medline]

27. Leverve XM and Fontaine E. Role of substrates in the regulation of mitochondrial function in situ. IUBMB Life 52: 221–229, 2001.[Web of Science][Medline]

28. Mannella CA. The relevance of mitochondrial membrane topology to mitochondrial function. Biochim Biophys Acta 1762: 140–147, 2006.[Medline]

29. Mitchell P. Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191: 144–148, 1961.[CrossRef][Medline]

30. Mootha VK, Bunkenborg J, Olsen JV, Hjerrild M, Wisniewski JR, Stahl E, Bolouri MS, Ray HN, Sihag S, Kamal M, Patterson N, Lander ES, and Mann M. Integrated analysis of protein composition, tissue diversity, and gene regulation in mouse mitochondria. Cell 115: 629–640, 2003.[CrossRef][Web of Science][Medline]

31. Ogata T and Yamasaki Y. Ultra-high-resolution scanning electron microscopy of mitochondria and sarcoplasmic reticulum arrangement in human red, white, and intermediate muscle fibers. Anat Rec 248: 214–223, 1997.[CrossRef][Medline]

32. Ott M, Robertson JD, Gogvadze V, Zhivotovsky B, and Orrenius S. Cytochrome c release from mitochondria proceeds by a two-step process. Proc Natl Acad Sci USA 99: 1259–1263, 2002.[Abstract/Free Full Text]

33. Passerieux E, Rossignol R, Chopard A, Carnino A, Marini JF, Letellier T, and Delage JP. Structural organization of the perimysium in bovine skeletal muscle: junctional plates and associated intracellular subdomains. J Struct Biol 154: 206–216, 2006.[CrossRef][Web of Science][Medline]

34. Pfeiffer T, Schuster S, and Bonhoeffer S. Cooperation and competition in the evolution of ATP-producing pathways. Science 292: 504–507, 2001.[Abstract/Free Full Text]

35. Rossignol R, Gilkerson R, Aggeler R, Yamagata K, Remington SJ, and Capaldi RA. Energy substrate modulates mitochondrial structure and oxidative capacity in cancer cells. Cancer Res 64: 985–993, 2004.[Abstract/Free Full Text]

36. Rossignol R, Letellier T, Malgat M, Rocher C, and Mazat JP. Tissular variation in the control of oxidative phosphorylations, implication for mitochondrial diseases. Biochem J 347: 45–53, 2000.[CrossRef][Web of Science][Medline]

37. Rossignol R, Malgat M, Mazat JP, and Letellier T. Threshold effect and tissue specificity. J Biol Chem 274: 33426–33432, 1999.[Abstract/Free Full Text]

38. Santel A. Get the balance right: mitofusins roles in health and disease. Biochim Biophys Acta 1763: 490–499, 2006.[Medline]

39. Saraste M. Oxidative phosphorylation at the fin de sciècle. Science 283: 1488–1493, 1999.[Abstract/Free Full Text]

40. Schägger H and Pfeiffer K. The ratio of oxidative phosphorylation complexes i-v in bovine heart mitochondria and the composition of respiratory chain supercomplexes. J Biol Chem 276: 37861–37867, 2001.[Abstract/Free Full Text]

41. Schägger H and Pfeiffer K. Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J 19: 1777–1783, 2000.[CrossRef][Web of Science][Medline]

42. Scorrano L, Ashiya M, Buttle K, Weiler S, Oakes SA, Mannella CA, and Korsmeyer SJ. A distinct pathway remodels mitochondrial cristae and mobilizes cytochrome c during apoptosis. Dev Cell 2: 55–67, 2002.[CrossRef][Web of Science][Medline]

43. Smeitink JA, Zeviani M, Turnbull DM, and Jacobs HT. Mitochondrial medicine: a metabolic perspective on the pathology of oxidative phosphorylation disorders. Cell Metab 3: 9–13, 2006.[CrossRef][Web of Science][Medline]

44. Tarjan EM and Von Korff RW. Factors affecting the respiratory control ratio of rabbit heart mitochondria. J Biol Chem 242: 318–324, 1967.[Abstract/Free Full Text]

45. Taylor SW, Fahy E, Zhang B, Glenn GM, Warnock DE, Wiley S, Murphy AN, Gaucher SP, Capaldi RA, Gibson BW, and Ghosh SS. Characterization of the human heart mitochondrial proteome. Nat Biotechnol 21: 281–286, 2003.[CrossRef][Web of Science][Medline]

46. Vendelin M, Beraud N, Guerrero K, Andrienko T, Kuznetsov AV, Olivares J, Kay L, and Saks VA. Mitochondrial regular arrangement in muscle cells: a "crystal-like" pattern. Am J Physiol Cell Physiol 288: C757–C767, 2005.[Abstract/Free Full Text]

47. Vijayasarathy C, Biunno I, Lenka N, Yang M, Basu A, Hall IP, and Avadhani NG. Variations in the subunit content and catalytic activity of the cytochrome c oxidase complex from different tissues and different cardiac compartments. Biochim Biophys Acta 1371: 71–82, 1998.[Medline]

48. Vyssokikh M, Zorova L, Zorov D, Heimlich G, Jurgensmeier J, Schreiner D, and Brdiczka D. The intra-mitochondrial cytochrome c distribution varies correlated to the formation of a complex between VDAC and the adenine nucleotide translocase: this affects Bax-dependent cytochrome c release. Biochim Biophys Acta 1644: 27–36, 2004.[Medline]

49. Waino W. The Mamalian Mitochondrial Respiratory Chain. London and New York: Academic, 1970.

50. Wallace D. Mitochondrial diseases in man and mouse. Science 283: 1482–1488, 1999.[Abstract/Free Full Text]

51. Yaffe MP. Dynamic mitochondria. Nat Cell Biol 1: E149–E150, 1999.[CrossRef][Web of Science][Medline]

This article has been cited by other articles:

				A.M. Hall, R.J. Unwin, M.G. Hanna, and M.R. Duchen Renal function and mitochondrial cytopathy (MC): more questions than answers? QJM, October 1, 2008; 101(10): 755 - 766. [Abstract] [Full Text] [PDF]

				J. A. Mayr, D. Meierhofer, F. Zimmermann, R. Feichtinger, C. Kogler, M. Ratschek, N. Schmeller, W. Sperl, and B. Kofler Loss of Complex I due to Mitochondrial DNA Mutations in Renal Oncocytoma Clin. Cancer Res., April 15, 2008; 14(8): 2270 - 2275. [Abstract] [Full Text] [PDF]

				T. Trian, G. Benard, H. Begueret, R. Rossignol, P.-O. Girodet, D. Ghosh, O. Ousova, J.-M. Vernejoux, R. Marthan, J.-M. Tunon-de-Lara, et al. Bronchial smooth muscle remodeling involves calcium-dependent enhanced mitochondrial biogenesis in asthma J. Exp. Med., December 24, 2007; 204(13): 3173 - 3181. [Abstract] [Full Text] [PDF]

				G. Benard, N. Bellance, D. James, P. Parrone, H. Fernandez, T. Letellier, and R. Rossignol Mitochondrial bioenergetics and structural network organization J. Cell Sci., March 1, 2007; 120(5): 838 - 848. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) All Versions of this Article: 291/6/C1172    most recent 00195.2006v1 Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Benard, G. Articles by Rossignol, R. Search for Related Content PubMed PubMed Citation Articles by Benard, G. Articles by Rossignol, R.