| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
CELLULAR AND MITOCHONDRIAL METABOLISM
1Department of Biochemistry, Nijmegen Centre for Molecular Life Sciences, 2Microscopical Imaging Centre, Nijmegen Centre for Molecular Life Sciences, 3Department of Paediatrics, Nijmegen Center for Mitochondrial Disorders, and 4Department of Cell Biology, Nijmegen Centre for Molecular Life Sciences, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands; 5Department of General Pediatrics, Heinrich-Heine-University, Düsseldorf, Germany; and 6MicroSpectroscopy Centre, Laboratory of Biochemistry, Wageningen University, Wageningen, The Netherlands
Submitted 13 February 2008 ; accepted in final form 17 March 2008
 |
ABSTRACT |
|---|
 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
NADH:ubiquinone oxidoreductase deficiency; human skin fibroblasts; fluorescence correlation spectroscopy; mitochondrial motility
). This potential is maintained by the electron transport chain (complexes I to IV), which, together with the Fo/F1-ATPase (complex V), constitutes the oxidative phosphorylation (OXPHOS) system. The electron transport chain system expels protons (at complexes I, III, and IV) out of the mitochondrial matrix via controlled electron transfer from NADH (complex I) and succinate (complex II) to oxygen (complex IV) and uses the resulting electrochemical proton gradient to drive the synthesis of ATP (complex V) (6, 12). Alterations in mitochondrial functional state are associated with a large family of metabolic disorders of which defects of the OXPHOS system are the most common (48). Among these disorders, NADH:ubiquinone oxidoreductase or complex I (CI; EC 1.6.5.3) deficiency (OMIM 252010 [OMIM] ) is most frequently observed (49). CI consists of 45 different subunits, together having a weight of close to 1 MDa (10). The catalytic core of the human complex consists of 14 evolutionary conserved proteins, two flavoproteins (encoded by the nuclear NDUFV1 and NDUFV2 genes), five iron-sulfur proteins (encoded by the nuclear NDUFS1, NDUFS2, NDUFS3, NDUFS7, and NDUFS8 genes), and seven hydrophobic proteins (encoded by the mitochondrial ND1 to ND6 and ND4L genes) (8).
In most children, CI deficiency is caused by autosomal recessive mutations in one of the nuclear-encoded subunits of the complex, leading to a reduction in its amount and/or catalytic activity (52). Currently, such mutations have been identified in all seven core subunits (3, 4, 5, 33, 34, 45, 51); in the so-called accessory or supernumerary subunits NDUFS4 (53), NDUFS6 (25), and NDUFA1 (15); in the complex I assembly factors B17.2L (38), C6ORF66 (43), and CIA30 [encoded by the NDUFAF1 gene; (13)]; and in the mitochondrial elongation factor G1 (11). The most common clinical presentations associated with CI deficiency are Leigh syndrome, Leigh syndrome with cardiomyopathy, and cerebral atrophy/leukodystropy (50, 40).
It is well established that changes in mitochondrial metabolism influence mitochondrial shape and matrix configuration (2, 18, 19, 32, 41, 44) and thus the exchange of metabolites across the inner mitochondrial membrane and diffusion of matrix constituents (32, 44, 56). Using confocal microscopy and fluorescence correlation spectroscopy (FCS) of human skin fibroblasts expressing enhanced yellow fluorescent protein (EYFP) in the mitochondrial matrix (mitoEYFP), we recently showed that chronic CI inhibition by rotenone increased mitochondrial length and degree of branching and decreased the fraction of moving mitochondria and their velocity (27, 30). We furthermore showed that chronic rotenone treatment increased matrix EYFP diffusion in both moving and stationary mitochondria. It was proposed that the observed increases in mitochondrial length and degree of branching and matrix protein diffusion may constitute part of an adaptive response to counterbalance the detrimental effects of the decrease in CI activity. In agreement with this idea, analysis of a cohort of 16 fibroblast lines from children with isolated CI deficiency caused by mutations in nuclear-encoded CI subunits showed that mitochondria were fragmented and/or less branched in fibroblasts with a severely reduced CI amount and activity ("class I"), whereas patient cells in which these latter parameters were only moderately reduced displayed a normal mitochondrial morphology ("class II"; 31). Class I cells (8 different patients) carried mutations in the NDUFS1, NDUFS2, NDUFS4, and NDUFS8 subunit of complex I, whereas class II cells (8 different patients) carried mutations in the NDUFS4, NDUFS7, NDUFS8, and NDUFV1 subunit of complex I.
In the present study, we aimed to establish the relationship between residual CI activity (maximal activity of CI determined in a homogenate of a mitochondrial enriched fraction), as a measure of mitochondrial OXPHOS capacity, and mitochondrial shape, mobility, and matrix protein diffusion in inherited CI deficiency. To this end, we applied confocal microscopy and FCS of mitoEYFP-expressing skin fibroblasts of a healthy subject and two genetically characterized CI-deficient patients, one with a R228Q mutation in the NDUFS2 subunit and the other with a R59X/T423M mutation in the NDUFV1 subunit. These patients displayed either a large (61%; NDUFS2) or modest (27%; NDUFV1) decrease in residual CI activity, paralleled by a decrease and increase in mitochondrial degree of branching, respectively (28).
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Transient expression of mitochondria matrix-targeted EYFP. MitoEYFP was expressed using a modified baculovirus vector as described previously (30, 57). Measurements were performed three days after the onset of baculovirus transduction, when virtually all cells expressed the construct. The virus remained present in the culture medium during the entire incubation period.
Quantitative analysis of mitochondrial morphology.
Quantitative analysis of mitochondrial morphology was carried out as described in detail elsewhere (27, 28, 29). Briefly, after correction for background fluorescence, images were subjected to linear contrast stretching followed by top-hat filtering, median filtering, and thresholding. This procedure yields binary images showing white mitochondria against a black background. These images were used to quantify the number of mitochondria per cell (Nc) and their form factor (F) using image analysis software (see below). F is calculated for each individual mitochondrion and equals perimeter2/4
·area. The latter is a combined measure of mitochondrial length and degree of branching.
FCS. Combined FCS and confocal microscopy was performed on a ConfoCor II system (Carl Zeiss, Sliedrecht, The Netherlands) as described previously (30). Briefly, mitoEYFP was excited using the 514-nm line of an Ar-ion laser focused via a dichroic mirror and a Zeiss C-Apochromat objective (x40, 1.2 numerical aperture, water immersion) onto the sample. Laser output power was 20 kW/cm2, and fluorescence emission light was guided via the dichroic mirror and a 545DF35 band-pass filter onto an avalanche photodiode. After selection of mitochondrial regions of interest in the confocal mode, autocorrelation curves were acquired during 10 s at 20°C.
The positions of the pinhole and the correction ring of the objective lens were optimized using an aqueous EYFP solution. Optimal settings were considered to be reached when the highest molecular brightness was observed. Fitting the autocorrelation curves of EYFP, of which the diffusion coefficient in water is known, to Eq. 1 revealed that the FCS detection volume had radii of 0.253 µm and 1.37 µm for the equatorial (rxy) and axial radius (rz), respectively. Intensity signals were software-correlated and individual autocorrelations were displayed on-line. The autocorrelation function (ACF) describing j independent molecular species diffusing freely in a three-dimensional Gaussian-shaped observation volume (Veff = 3/2·rxy2·rz) is given by:
 | (1) |
 |
kin, respectively. The lateral diffusion time dif,j describes the residence time of a particle in the observation volume, which is related to the translational diffusion coefficient:
 | (2) |
The amplitude of the ACF, G(0), represents the average number of molecules 
N
found in the observation volume: G(0) – 1 = 1/N. During fitting, a single-component model (j = 1) was evaluated first. This model was considered valid when the residuals did not exceed 0.05 (9, 21, 22, 26, 47). When the latter was not the case, the single-component model was rejected and a two-component model (j = 2) was evaluated using similar criteria. Recordings displaying excessive bleaching (>5% over the full duration of the recording) or high-amplitude spikes were omitted from the analysis.
To allow selection of the most appropriate model for fitting the ACF (17), we first assessed the mitochondrial diameter by calculating the width at half-maximal height of a 1 pixel wide intensity profile perpendicular to the long axis of the mitochondrial filament (as described in the supplement of Ref. 7). This analysis revealed no difference between control (0.85 ± 0.05 µm, n = 35 mitochondria) and NDUFS2 cells (0.80 ± 0.03 µm, n = 33 mitochondria), and the values obtained agreed well with those determined by electron microscopy (EM; Ref. 1 and Fig. 3). Conversely, mitochondrial diameter was somewhat smaller in NDUFV1 cells (0.60 ± 0.03 µm, n = 24 mitochondria; P > 0.05). In analogy to our previous study (30), Eq. 1 was appropriate to fit all experimental ACFs. To determine diffusion times (dif), ACF curves were fitted between = 0.01 ms and = 100 ms, a range compatible with translational diffusion (14). In aqueous solution, EYFP displayed a single diffusion time () of 0.165 ± 0.04 ms (n = 4), which was equivalent to a translational diffusion constant (D) of 92.3 ± 0.22 µm2/s (30). In control and patient cells, no autocorrelation was detectable in the nucleoplasm and cytosol (30). This rules out interference of autofluorescence and demonstrates that mitoEYFP was exclusively present within the mitochondrial matrix.
|
Image analysis and statistical analysis. Confocal images were processed and analyzed using Image Pro Plus 5.1 software (Media Cybernetics, Silver Spring, MD). Numerical results were visualized using Origin Pro 7.5 (Originlabs, Northampton, MA) and are presented as means ± SE. Statistical differences were determined using Student's t-test (Bonferroni corrected) and were considered significant when P < 0.05.
Chemicals. Culture materials were obtained from Invitrogen (Breda, The Netherlands). All other reagents were from Sigma (St. Louis, MO).
|
RESULTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Quantification of mitochondrial shape in healthy and patient fibroblasts.
Confocal imaging of mitoEYFP-expressing healthy control (CT; Fig. 1A) and CI-deficient (NDUFS2, Fig. 1B, and NDUFV1, Fig. 1C) patient fibroblasts revealed numerous threadlike structures, previously identified as mitochondria (27–31, 57). The shape of these mitochondrial structures was quantified by calculating the mitochondrial form factor (F), a combined measure of mitochondrial length and degree of branching, using a previously described protocol (27–29). This analysis revealed that, compared with healthy control cells, F was lower in NDUFS2 cells and higher in NDUFV1 cells (Fig. 1D and Table 1). In both patient cell lines, the number of mitochondria per cell (Nc) was identical to control. To describe mitochondrial morphology, we calculated the operational parameter "mitochondrial complexity," given by the ratio of F to Nc (28, 31). Relative to control cells, mitochondrial complexity was significantly smaller in NDUFS2 cells and significantly larger in NDUFV1 cells (Fig. 1D). The change in mitochondrial shape as determined in the present study with mitoEYFP was quantitatively identical to that reported previously (28, 31) using mitoEYFP (NDUFV1: F = 138 ± 8%, Nc = 103 ± 2%, n = 30; NDUFS2: F = 64 ± 2%, Nc = 103 ± 2%, n = 27) and the mitochondria-specific cation rhodamine 123 (NDUFV1: F = 133 ± 6%, Nc = 107 ± 5%, n = 79; NDUFS2 F = 65 ± 2%, Nc = 95 ± 5%, n = 336).
|
|
To determine the translational diffusion coefficient of EYFP within the mitochondrial matrix, mitoEYFP-expressing fibroblasts were analyzed by FCS, as recently described in detail (30). Cells were visualized by confocal microscopy for random selection of mitochondrial regions of interest (Fig. 1; crosses). Next, fluorescence intensity fluctuations were acquired during 10 s, after which the experimental ACF of the photodetector output was calculated for each individual region of interest. Finally, translational diffusion times (dif) were computed from the ACF curve (see MATERIALS AND METHODS). In both healthy and patient fibroblasts, 
20% of the mitochondria displayed one single (fast1) diffusion time, and the remainder displayed two separate (fast2 and slow2) diffusion times (Fig. 2A). Importantly, both "types" of mitochondria were present within one and the same cell. When cells were treated with nocodazole (Noc; 10 µg/ml; 2 h), an established inhibitor of mitochondrial movement (36, 37, 42, 63), slow2 was abolished and only a single fast translational diffusion time remained (Noc; Fig. 2A). These results were in accordance with previous findings (30), which demonstrated that the value of slow2 reflects mitochondrial velocity and fast1 and fast2 reflect matrix protein diffusion in stationary and moving mitochondria, respectively.
|
2.5-fold higher in patient cells. In healthy fibroblasts treated with Noc, the translational diffusion coefficient (DNoc) was identical to Dfast1 and Dfast2 in untreated fibroblasts (Fig. 2E and Ref. 30), indicating that, in these cells, blocking mitochondrial movement with Noc did not affect matrix protein diffusion. DNoc was not different between the two patient cell lines (Fig. 2E; NDUFS2: DNoc = 30 ± 2 µm2/s, n = 30 mitochondria; NDUFV1: DNoc = 28 ± 2 µm2/s, n = 45 mitochondria). Calculation of the weighed average of Dfast1 and Dfast2 in untreated patient fibroblasts yielded a value similar to DNoc, suggesting that Noc immobilized moving mitochondria without altering matrix protein diffusion.
Mitochondrial ultrastructure in healthy and patient fibroblasts.
Evidence in the literature demonstrated that alterations in mitochondrial metabolism can be paralleled by changes in mitochondrial ultrastructure (18, 19, 32, 35, 41, 44). As a consequence, protein diffusion in the mitochondrial matrix may be affected (39, 56). These observations suggest that the difference in matrix protein diffusion between control and patient cells may result from alterations in mitochondrial ultrastructure. To investigate this possibility, EM analysis was performed. Obviously, the EM technique cannot discriminate between stationary and moving mitochondria. However, our FCS analysis (Fig. 2A) demonstrated that the large majority is moving (80%) in both control and patient fibroblasts. Visual inspection of a large number of EM images (n > 30 cells) revealed no differences in mitochondrial cristae morphology and matrix density between control and patient fibroblasts (Fig. 3). EM analysis of two additional control cell lines [nos. 4996 and 5118 (28, 31, 54, 55, 58, 63)] also revealed no differences (data not shown). These findings suggest that alterations in matrix protein diffusion observed by FCS are not caused by gross alterations in mitochondrial ultrastructure.
|
DISCUSSION |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Using FCS, we found here that a similar fraction of mitochondria (80%) was moving in both healthy and patient fibroblasts. However, patient mitochondria moved 2.5-times faster. Intramatrix protein diffusion was similar for moving and stationary mitochondria in healthy fibroblasts. In contrast, patient mitochondria displayed either a 20% slower (stationary organelles) or 2.5-times increased intramatrix diffusion (moving organelles). These results clearly demonstrate that the major difference between healthy and patient fibroblasts involves the velocity and matrix protein diffusion rate of moving mitochondria.
Blocking mitochondrial movement with Noc did not affect the rate of intramatrix protein diffusion in both healthy and patient fibroblasts. This shows that mitochondrial velocity and intramatrix protein diffusion are mutually independent. Although the two patient fibroblast lines analyzed in the present study represent opposite extremes in terms of residual CI activity and F (28), no differences in fraction and velocity of moving mitochondria and/or intramatrix protein diffusion were observed. This finding suggests the absence of a quantitative relationship between these latter parameters and either residual CI activity or mitochondrial shape for the patient fibroblasts.
Mitochondrial velocity is increased in patient fibroblasts.
On average, 80% of the mitochondria present in healthy and patient fibroblasts displayed two EYFP translational diffusion coefficients (Dfast2 and Dslow2), whereas the remainder exhibited only a single EYFP translational diffusion coefficient (Dfast1). Treatment with Noc abolished mitochondrial movement and caused complete disappearance of the slow diffusion component (Dslow2). The same result was obtained before with healthy fibroblasts and led to the conclusion that Dslow2 is a quantitative measure of microtubule-mediated mitochondrial movement (30). At first sight, our finding that the fraction of moving mitochondria is not altered in patient fibroblasts seems to contradict our observation that chronic CI inhibition by rotenone (100 nM, 72 h) decreased this fraction in healthy fibroblasts (30). However, the residual CI activity was lower in rotenone-treated healthy fibroblasts (20% of control) in comparison with the patient fibroblasts used in the present study (Table 1). This may suggest that, in living fibroblasts, mitochondria stop to move when their CI activity decreases to below a critical level. In accordance with this idea, the velocity of moving mitochondria was lower in rotenone-treated healthy fibroblasts (30). Alternatively, in view of recent work showing that mitochondrial movement is inhibited at high cellular ADP levels (36), our FCS data may be interpreted as an indication that CI-deficient patient fibroblasts have metabolically adapted to the CI-deficient state, whereas, in contrast, healthy fibroblasts chronically treated with rotenone have not. Such an adaptation may also explain the unexpected finding that mitochondrial velocity was two times higher in patient compared with healthy fibroblasts. Our data furthermore show the lack of any correlation between fraction and velocity of moving mitochondria and mitochondrial length and degree of branching (F). The latter parameter was previously found to be increased in rotenone-treated healthy fibroblasts (27) and to be quantitatively related to residual CI activity for a large cohort of patient fibroblasts (28).
Intramatrix protein diffusion is faster in moving mitochondria of patient fibroblasts. EYFP diffusion within the mitochondrial matrix was quantified by Dfast1 in stationary mitochondria, Dfast2 in moving mitochondria and DNoc in mitochondria of cells treated with Noc. In healthy fibroblasts, Dfast1, Dfast2, and DNoc were equal, demonstrating the absence of any relation between intramatrix protein diffusion and mitochondrial movement. Similarly, Dfast1, Dfast2, and DNoc did not differ between the two patient fibroblast lines, suggesting that intramatrix protein diffusion is not quantitatively related to residual CI activity and/or mitochondrial shape in CI-deficient patient fibroblasts. In both patient fibroblast lines, however, Dfast2 was three times larger than Dfast1, showing that intramatrix protein diffusion is faster in moving compared with stationary mitochondria. Together, these results are compatible with the idea that, in patient fibroblasts, primarily mitochondrial velocity and intramatrix protein diffusion are increased but that these increases are mutually independent.
The protein concentration in the mitochondrial matrix is estimated to be 300 mg/ml (23), which makes it the most crowded compartment of the cell. It has been demonstrated that the mitochondrial matrix can assume two configurations, the orthodox state, which is characterized by a large volume, a low protein concentration, and fast metabolite diffusion, and the condensed state, with a small volume, a high protein concentration, and slower metabolite diffusion (18). EM analysis of mitochondrial ultrastructure revealed that the density of the mitochondrial matrix was not detectably different between control and patient fibroblasts. Similarly, cristae morphology, previously predicted to affect matrix protein diffusion (39, 56), was unaltered in patient cells. Although 80% of all mitochondria were moving in our life cell FCS experiments, we do not know how quickly matrix protein diffusion increases when stationary mitochondria start moving or decreases when moving mitochondria become stationary. Obviously, mitochondrial movement will immediately seize during fixation prior to EM, and this may explain why no difference in matrix density and/or cristae morphology between control and patient cells was observed. Alternatively, the change in matrix density toward a less dense state may only be relatively small and therefore not detectable by EM analysis.
In light of the above findings, the marked increase in matrix protein diffusion in moving patient fibroblast mitochondria is compatible with the idea that, in these mitochondria, the matrix has adopted a less dense configuration. It has been demonstrated that mitochondria with a more orthodox matrix have a lower metabolic activity than mitochondria with a condensed matrix (44). Accordingly, the mitochondrial metabolic activity may be decreased in moving mitochondria of patient fibroblasts. This conclusion is in agreement with the CI-deficient state of these mitochondria and is supported by our previous finding that intramatrix EYFP diffusion is markedly increased in mitochondria of healthy fibroblasts chronically treated with rotenone (30). Recent work revealed that the matrix configuration of cancer cell mitochondria changed from condensed to orthodox upon replacement of galactose with glucose in the culture medium, thereby forcing the cells to change their mode of energy production from oxidative into glycolytic (41). Extrapolation of this result to the present study supports our idea that patient fibroblasts have also become more glycolytic. This is supported by previous findings revealing an increased lactate-to-pyruvate ratio in cultured skin fibroblasts harboring respiratory chain defects (61). In agreement with this idea, cells with impaired OXPHOS function were found to generate ATP via the glycolytic pathway when grown in a medium containing both D-glucose and pyruvate (16). Similarly, it was demonstrated that glycolysis was increased in 143B cybrid cells with pathogenic mtDNA point mutations (60). Finally, mouse hearts depleted of mtDNA by ablation of the tfam gene displayed a global switch from oxidative to glycolytic metabolism (20).
Conclusions. Taken together, our findings are compatible with a model in which inherited CI deficiency causes two mutually independent changes, first, a change toward a less dense configuration of the mitochondrial matrix, and second, a change toward a more glycolytic mode of energy production associated with an increase in mitochondrial velocity. According to this model, the latter change is not (yet) achieved in healthy fibroblasts treated with rotenone for 3 days. Most likely, the observed change in matrix protein diffusion reflects an adaptation to the CI-deficient state of the mitochondria in that it may serve to reduce diffusion bottlenecks, thus promoting exchange of metabolites across the inner mitochondrial membrane and diffusion of matrix constituents.
|
GRANTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Present addresses: M. A. Hink, Max Planck Institute for Molecular Physiology, Dortmund, Germany; S. Verkaart, Department of Cell Physiology, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands.
|
FOOTNOTES |
|---|
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 |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
2. Benard G, Bellance N, James D, Parrone P, Fernandez H, Letellier T, Rossignol R. Mitochondrial bioenergetics and structural network organization. J Cell Sci 120: 838–848, 2007.
3. Benit P, Chretien D, Kadhom N, de Lonlay-Debeney P, Cormier-Daire V, Cabral A, Peudenier S, Rustin P, Munnich A, Rotig A. Large-scale deletion and point mutations of the nuclear NDUFV1 and NDUFS1 genes in mitochondrial complex I deficiency. Am J Hum Genet 68: 1344–1352, 2001.[CrossRef][Web of Science][Medline]
4. Benit P, Beugnot R, Chretien D, Giurgea I, de Lonlay-Debeney P, Issartel JP, Corral-Debrinski M, Kerscher S, Rustin P, Rotig A, Munnich A. Mutant NDUFV2 subunit of mitochondrial complex I causes early onset hypertrophic cardiomyopathy and encephalopathy. Hum Mutat 21: 582–586, 2003.[CrossRef][Web of Science][Medline]
5. Benit P, Slama A, Cartault F, Giurgea I, Chretien D, Lebon S, Marsac C, Munnich A, Rotig A, Rustin P. Mutant NDUFS3 subunit of mitochondrial complex I causes Leigh syndrome. J Med Genet 41: 14–17, 2004.
6. Bernardi P. Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev 79: 1127–1155, 1999.
7. Boxma B, de Graaf RM, van der Staay GW, van Alen TA, Ricard G, Gabaldon T, van Hoek AH, Moon-van der Staay SY, Koopman WJH, van Hellemond JJ, Tielens AG, Friedrich T, Veenhuis M, Huynen MA, Hackstein JH. An anaerobic mitochondrion that produces hydrogen. Nature 434: 74–79, 2005.[CrossRef][Medline]
8. Brandt U. Energy converting NADH:quinone oxidoreductase (complex I). Annu Rev Biochem 75: 69–92, 2006.[CrossRef][Web of Science][Medline]
9. Brock R, Hink MA, Jovin TM. Fluorescence correlation spectroscopy of cells in the presence of autofluorescence. Biophys J 75: 2547–2557, 1998.[Web of Science][Medline]
10. Carrol J, Fearnley IM, Skehel JM, Shannon RJ, Hirst J, Walker JE. Bovine complex I is a component of 45 different subunits. J Biol Chem 281: 32724–32727, 2006.
11. Coenen MJ, Antonicka H, Ugalde C, Sasarman F, Rossi R, Heister JG, Newbold RF, Trijbels FJ, van den Heuvel LP, Shoubridge EA, Smeitink JAM. Mutant mitochondrial elongation factor G1 and combined oxidative phosphorylation deficiency. N Engl J Med 351: 2080–2086, 2004.
12. Duchen MR. Mitochondria in health and disease: perspectives on a new mitochondrial biology. Mol Aspects Med 25: 365–451, 2004.[Medline]
13. Dunning CJ, McKenzie M, Sugiana C, Lazarou M, Silke J, Connelly A, Fletcher JM, Kirby DM, Thorburn DR, Ryan MT. Human CIA30 is involved in the early assembly of mitochondrial complex I and mutations in its gene cause disease. EMBO J 26: 3227–3237, 2007.[CrossRef][Web of Science][Medline]
14. Enderlein J, Gregor I, Patra D, Fitter J. Art and artifacts of fluorescence correlation spectroscopy. Curr Pharm Biotechnol 5: 155–161, 2004.[CrossRef][Web of Science][Medline]
15. Fernandez-Moreira D, Ugalde C, Smeets R, Rodenburg RJ, Lopez-Laso E, Ruiz-Falco ML, Briones P, Martin MA, Smeitink JAM, Arenas J. X-linked NDUFA1 gene mutations associated with mitochondrial encephalomyopathy. Ann Neurol 61: 73–83, 2007.[CrossRef][Web of Science][Medline]
16. Gajewski CD, Yang L, Schon EA, Manfredi G. New insights into the bioenergetics of mitochondrial disorders using intracellular ATP reporters. Mol Biol Cell 14: 3628–3635, 2003.
17. Gennerich A, Schild D. Fluorescence correlation spectroscopy in small cytosolic compartments depends critically on the diffusion model used. Biophys J 79: 3294–3306, 2000.[Web of Science][Medline]
18. 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.
19. Halestrap AP. The regulation of the matrix volume of mammalian mitochondria in vivo and in vitro and its role in the control of mitochondrial metabolism. Biochim Biophys Acta 973: 355–382, 1989.[Medline]
20. Hansson A, Hance N, Dufour E, Rantanen A, Hultenby K, Clayton DA, Wibom R, Larsson NG. A switch in metabolism precedes increased mitochondrial biogenesis in respiratory chain-deficient mouse hearts. Proc Natl Acad Sci USA 101: 3136–3141, 2004.
21. Hink MA, Visser AJ. Characterization of membrane mimetic systems with fluorescence. In: Applied Fluorescence in Chemistry, Biology, and Medicine, edited by Strehmel W, Schrader B, and Seifert H. New York: Springer Verlag, 1998.
22. Hink MA, Griep RA, Borst JW, van Hoek A, Eppink MH, Schots A, Visser AJ. Structural dynamics of green fluorescent protein alone and fused with a single chain Fv protein. J Biol Chem 275: 17556–17560, 2000.
23. Jakobs S. High resolution imaging of live mitochondria. Biochim Biophys Acta 1763: 561–575, 2006.[Medline]
24. Kim Schwille P SA. Intracellular application of fluorescence correlation spectroscopy: prospects for neuroscience. Curr Opin Neurobiol 13: 583–590, 2003.[CrossRef][Web of Science][Medline]
25. Kirby DM, Salemi R, Sugiana C, Ohtake A, Parry L, Bell KM, Kirk EP, Boneh A, Taylor RW, Dahl HH, Ryan MT, Thorburn DR. NDUFS6 mutations are a novel cause of lethal neonatal mitochondrial complex I deficiency. J Clin Invest 114: 837–845, 2004.[CrossRef][Web of Science][Medline]
26. Koopman WJH, Hink MA, Visser AJ, Roubos EW, Jenks BG. Evidence that Ca2+-waves in Xenopus melanotropes depend on calcium-induced calcium release: a fluorescence correlation microscopy and line scanning study. Cell Calcium 26: 59–67, 1999.[CrossRef][Web of Science][Medline]
27. Koopman WJH, Verkaart S, Visch HJ, van der Westhuizen FH, Murphy MP, van den Heuvel LW, Smeitink JAM, Willems PHGM. Inhibition of complex I of the electron transport chain causes oxygen radical-mediated mitochondrial outgrowth. Am J Physiol Cell Physiol 288: C1440–C1450, 2005.
28. Koopman WJH, Visch HJ, Verkaart S, van den Heuvel LW, Smeitink JAM, Willems PHGM. Mitochondrial network complexity and pathological decrease in complex I activity are tightly correlated in isolated human complex I deficiency. Am J Physiol Cell Physiol 289: C881–C890, 2005.
29. Koopman WJH, Visch HJ, Smeitink JAM, Willems PHGM. Simultaneous quantitative measurement and automated analysis of mitochondrial morphology, mass, potential, and motility in living human fibroblasts. Cytometry A 69: 1–12, 2006.[Medline]
30. Koopman WJH, Hink MA, Verkaart S, Visch HJ, Smeitink JAM, Willems PHGM. Partial complex I inhibition decreases mitochondrial motility and increases matrix protein diffusion as revealed by fluorescence correlation spectroscopy. Biochim Biophys Acta 1767: 940–947, 2007.[Medline]
31. Koopman WJH, Verkaart S, Visch HJ, Nijtmans LG, Smeitink JAM, Willems PHGM. Human NADH:ubiquinone oxidoreductase deficiency: radical changes in mitochondrial morphology? Am J Physiol Cell Physiol 293: C22–C29, 2007.
32. Lloyd D, Salgado EJ, Turner MP, Suller MTE, Murray D. Cycles of mitochondrial energization driven by the ultradian clock in a continuous culture of Saccharomyces cerevisiae. Microbiology 148: 3715–3724, 2002.
33. Loeffen JL, Smeitink JAM, Triepels RH, Smeets RJ, Schuelke M, Sengers R, Trijbels F, Hamel B, Mullaart R, van den Heuvel LW. The first nuclear-encoded complex I mutation in a patient with Leigh syndrome. Am J Hum Genet 63: 1598–1608, 1998.[CrossRef][Web of Science][Medline]
34. Loeffen JL, Elpeleg O, Smeitink JAM, Smeets RJ, Stöckler-Ipsiroglu S, Mandel H, Sengers R, Trijbels F, van den Heuvel LP. Mutations in the complex I NDUFS2 gene of patient with cardiomyopathy and encephalomyopathy. Ann Neurol 49: 195–201, 2001.[CrossRef][Web of Science][Medline]
35. Mannella CA. The relevance of mitochondrial membrane topology to mitochondrial function. Biochim Biophys Acta 1762: 140–147, 2006.[Medline]
36. Mironov SL. ADP regulates movements of mitochondria in neurons. Biophys J 92: 2944–2952, 2007.[CrossRef][Web of Science][Medline]
37. Müller M, Mironov S, Ivannikov MV, Schmidt J, Richter DW. Mitochondrial organization and motility probed by two-photon microscopy in cultured mouse brainstem neurons. Exp Cell Res 303: 114–127, 2005.[Web of Science][Medline]
38. Ogilvie I, Kennaway NG, Shoubridge EA. A molecular chaperone for mitochondrial complex I assembly is mutated in a progressive encephalopathy. J Clin Invest 115: 2784–2792, 2005.[CrossRef][Web of Science][Medline]
39. Ölveczky BP, Verkman AS. Monte Carlo analysis of obstructed diffusion in three dimensions: application to molecular diffusion in organelles. Biophys J 74: 2722–2730, 1999.[Web of Science]
40. Robinson BH. Lactic acidemia and mitochondrial disease. Mol Genet Metab 89: 3–13, 2006.[CrossRef][Web of Science][Medline]
41. Rossignol R, Gilkerson R, Aggeler R, Yamagata K, Remington SJ, Capaldi A. Energy substrate modulates mitochondrial structure and oxidative capacity in cancer cells. Cancer Res 64: 985–993, 2004.
42. Rube DA, van der Bliek AM. Mitochondrial morphology is dynamic and varied. Mol Cell Biochem 256/257: 331–339, 2004.
43. Saada A, Edvardson S, Rapoport M, Shaag A, Amry K, Miller C, Lorberboum-Galski H, Elpeleg O. C6ORF66 is an assembly factor of mitochondrial complex I. Am J Hum Genet 82: 32–38, 2008.[CrossRef][Web of Science][Medline]
44. Scalettar BA, Abney JR, Hackenbrock CR. Dynamics, structure, and function are coupled in the mitochondrial matrix. Proc Natl Acad Sci USA 88: 8057–8061, 1991.
45. Schuelke M, Smeitink JAM, Mariman E, Loeffen JL, Plecko B, Trijbels F, Stockler-Ipsiroglu S, van den Heuvel LW. Mutant NDUFV1 subunit of mitochondrial complex I causes leukodystrophy and myoclonic epilepsy. Nat Genet 21: 260–261, 1999.[CrossRef][Web of Science][Medline]
46. Schwille P, Kummer S, Heikal AA, Moerner WE, Webb WW. Fluorescence correlation spectroscopy reveals fast optical excitation-driven intramolecular dynamics of yellow fluorescent proteins. Proc Natl Acad Sci USA 97: 151–156, 2000.
47. Sengupta P, Garai K, Balaji J, Periasamy N, Maiti S. Measuring size distribution in highly heterogeneous systems with fluorescence correlation spectroscopy. Biophys J 84: 1977–1984, 2003.[Web of Science][Medline]
48. Smeitink JAM, van den Heuvel LW, DiMauro S. The genetics and pathology of oxidative phosphorylation. Nat Rev Genet 2: 342–352, 2001.[CrossRef][Web of Science][Medline]
49. Smeitink JAM, Sengers R, Trijbels F, van den Heuvel LW. Human NADH:ubiquinone oxidoreductase. J Bioenerg Biomembr 33: 259–266, 2001.[CrossRef][Web of Science][Medline]
50. Smeitink JAM, van den Heuvel LW, Koopman WJH, Nijtmans LG, Ugalde C, Willems PHGM. Cell biological consequences of mitochondrial NADH: ubiquinone oxidoreductase deficiency. Curr Neurovasc Res 1: 29–40, 2004.[CrossRef][Web of Science][Medline]
51. Triepels RH, van den Heuvel LP, Loeffen JL, Buskens CA, Smeets RJ, Rubio Gozalbo ME, Mariman EC, Wijburg FA, Barth PG, Trijbels JM, Smeitink JAM. Leigh syndrome associated with a mutation in the NDUFS7 (PSST) nuclear encoded subunit of complex I. Ann Neurol 45: 787–790, 1999.[CrossRef][Web of Science][Medline]
52. Ugalde C, Janssen RJ, van den Heuvel LW, Smeitink JAM, Nijtmans LG. Differences in assembly or stability of complex I and other mitochondrial OXPHOS complexes in inherited complex I deficiency. Hum Mol Genet 13: 659–667, 2004.
53. Van den Heuvel LP, Ruitenbeek W, Smeets R, Gelman-Kohan Z, Elpeleg O, Loeffen JL, Trijbels F, Mariman EC, de Bruijn D, Smeitink JAM. Demonstration of a new pathogenic mutation in human complex I deficiency: a 5-bp duplication in the nuclear gene encoding the 18-kD (AQDQ) subunit. Am J Hum Genet 62: 262–268, 1998.[CrossRef][Web of Science][Medline]
54. Verkaart S, Koopman WJH, van Emst-de Vries SE, Nijtmans LG, van den Heuvel LW, Smeitink JAM, Willems PHGM. Superoxide production is inversely related to complex I activity in inherited complex I deficiency. Biochim Biophys Acta 1772: 373–381, 2007.[Medline]
55. Verkaart S, Koopman WJH, Cheek J, van Emst-de Vries SE, van den Heuvel LW, Smeitink JAM, Willems PHGM. Mitochondrial and cytosolic thiol redox state are not detectably altered in isolated human NADH:ubiquinone oxidoreductase deficiency. Biochim Biophys Acta 1772: 1041–1051, 2007.[Medline]
56. Verkman AS. Solute and macromolecular diffusion in cellular aqueous compartments. Trends Biochem Sci 27: 27–33, 2002.[CrossRef][Web of Science][Medline]
57. Visch HJ, Rutter GA, Koopman WJH, Koenderink JB, Verkaart S, de Groot T, Varadi A, Mitchell KJ, van den Heuvel LW, Smeitink JAM, Willems PHGM. Inhibition of mitochondrial Na+-Ca2+ exchange restores agonist-induced ATP production and Ca2+ handling in human complex I deficiency. J Biol Chem 279: 40328–40336, 2004.
58. Visch HJ, Koopman WJH, Leusink A, van Emst-de Vries SE, van den Heuvel LW, Willems PHGM, Smeitink JAM. Decreased agonist-stimulated mitochondrial ATP production caused by a pathological reduction in endoplasmic reticulum calcium content in human complex I deficiency. Biochim Biophys Acta 1762: 115–123, 2005.
59. Visser AJ, Hink MA. New perspectives of fluorescence correlation spectroscopy. J Fluorescence 9: 81–87, 1999.[CrossRef]
60. Von Kleist-Retzow JC, Hornig-Do HT, Schauen M, Eckertz S, Dinh TA, Stassen F, Lottmann N, Bust M, Galunska B, Wielckens K, Hein W, Beuth J, Braun JM, Fischer JH, Ganitkevich VY, Maniura-Weber K, Wiesner RJ. Impaired mitochondrial Ca2+ homeostasis in respiratory chain-deficient cells but efficient compensation of energetic disadvantage by enhanced anaerobic glycolysis due to low ATP steady state levels. Exp Cell Res 313: 3076–3089, 2007.[CrossRef][Web of Science][Medline]
61. Wijburg FA, Feller N, Scholte HR, Przyrembel H, Wanders RJ. Studies on the formation of lactate and pyruvate from glucose in cultured skin fibroblasts: implications for detection of respiratory chain defects. Biochem Int 19: 563–570, 1989.[Web of Science][Medline]
62. Willems PHGM, Valsecchi F, Distelmaier F, Verkaart S, Visch HJ, Smeitink JAM, Koopman WJH. Mitochondrial Ca2+ homeostasis in human NADH:ubiquinone oxidoreductase deficiency. Cell Calcium. In press.
63. Yi M, Weaver D, Hajnóczky G. Control of mitochondrial motility and distribution by the calcium signal: a homeostatic circuit. J Cell Biol 167: 661–672, 2004.
This article has been cited by other articles:
|
|
 |
|
  G. Benard and R. Rossignol Mitochondrial fluidity matters. Focus on "Inherited complex I deficiency is associated with faster protein diffusion in the matrix of moving mitochondria" Am J Physiol Cell Physiol, May 1, 2008; 294(5): C1123 - C1123. [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
