Mitochondrial network complexity and pathological decrease in complex I activity are tightly correlated in isolated human complex I deficiency

Werner J. H. Koopman,1,2 Henk-Jan Visch,2,3 Sjoerd Verkaart,2,3 Lambertus W. P. J. van den Heuvel,3 Jan A. M. Smeitink,3 and Peter H. G. M. Willems2

1Microscopical Imaging Center and 2Department of Biochemistry, Nijmegen Center for Molecular Life Sciences, and 3Department of Pediatrics, Nijmegen Center for Mitochondrial Disorders, Radboud University Nijmegen, Nijmegen, The Netherlands

Submitted 8 March 2005 ; accepted in final form 12 May 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Complex I (NADH:ubiquinone oxidoreductase) is the largest multisubunit assembly of the oxidative phosphorylation system, and its malfunction is associated with a wide variety of clinical syndromes ranging from highly progressive, often early lethal, encephalopathies to neurodegenerative disorders in adult life. The changes in mitochondrial structure and function that are at the basis of the clinical symptoms are poorly understood. Video-rate confocal microscopy of cells pulse-loaded with mitochondria-specific rhodamine 123 followed by automated analysis of form factor (combined measure of length and degree of branching), aspect ratio (measure of length), and number of revealed marked differences between primary cultures of skin fibroblasts from 13 patients with an isolated complex I deficiency. These differences were independent of the affected subunit, but plotting of the activity of complex I, normalized to that of complex IV, against the ratio of either form factor or aspect ratio to number revealed a linear relationship. Relatively small reductions in activity appeared to be associated with an increase in form factor and never with a decrease in number, whereas relatively large reductions occurred in association with a decrease in form factor and/or an increase in number. These results demonstrate that complex I activity and mitochondrial structure are tightly coupled in human isolated complex I deficiency. To further prove the relationship between aberrations in mitochondrial morphology and pathological condition, fibroblasts from two patients with a different mutation but a highly fragmented mitochondrial phenotype were fused. Full restoration of the mitochondrial network demonstrated that this change in mitochondrial morphology was indeed associated with human complex I deficiency.

mitochondria; oxidative phosphorylation; rhodamine 123; fibroblast


MITOCHONDRIA ARE PRESENT in virtually all eukaryotic cells, and in higher animals they are responsible for generation of the vast majority of the principal carrier of chemical energy, ATP (37). Apart from this, mitochondria harbor essential parts of the urea cycle and are crucial for the breakdown of fatty acids, the generation of heat, and the biosynthesis of heme, pyrimidines, amino acids, phospholipids, and nucleotides. Many aspects of mitochondrial function are regulated through the interplay between Ca2+ signals, transduced from the cytosol into the mitochondrial matrix (13), nitric oxide (6), and superoxide (7) generated within the matrix itself. Mitochondria, in turn, play a critical role in the spatiotemporal shaping of cytosolic Ca2+ signals (17, 47).

Alterations in mitochondrial activity can occur within seconds, as is the case for Ca2+-stimulated mitochondrial ATP production (24, 54), where Ca2+ is thought to act by rapidly increasing the activity of Ca2+-sensitive dehydrogenases (e.g., pyruvate dehydrogenase, NAD+-dependent isocitrate dehydrogenase, and 2-oxoglutarate dehydrogenase; Ref. 12) and possibly also of Ca2+-sensitive metabolite carriers of the inner mitochondrial membrane (Ca2+-sensitive aspartate/glutamate carrier; Ref. 29). In addition to this rapid accommodation to higher demands for ATP production in stimulated cells, mitochondria can also perform adaptive responses to meet long-term changes in cellular energy demand. These responses involve changes in structural architecture of the mitochondrial network (4, 22), which are generally accompanied by changes in mitochondrial content, ultrastructure, and enzyme levels (40).

Mitochondria have been demonstrated to possess protective mechanisms to balance, for example, excessive superoxide production through superoxide-induced activation of mitochondrial uncoupler proteins (36). Despite this, dysregulation of mitochondrial signaling processes involving Ca2+, reactive oxygen species, and/or reactive nitrogen species is now recognized to play a key role in controlled cell death or apoptosis (8, 23) as well as a wide variety of human pathologies (46, 57, 59).

Most, if not all, mitochondrial functions require the presence of an intact potential difference across the inner mitochondrial membrane ({Delta}{Psi}) that is maintained by the electron transport chain (ETC). Together with the F0/F1-ATPase (complex V), complexes I, II, III, and IV of the ETC constitute the oxidative phosphorylation (OXPHOS) system. Complex I or NADH:ubiquinone oxidoreductase (EC 1.6.5.3 [EC] ) is the largest OXPHOS complex and forms the entry point for electrons into the OXPHOS system. Structurally, this complex consists of 46 known subunits, 14 of which are evolutionarily conserved and constitute the minimal form of the complex (55). On the basis of detergent studies, complex I subunits have been classified as part of a flavoprotein (FP), an iron-sulfur protein (IP), or a hydrophobic protein (HP) fraction. The complex I core consists of two FP subunits (encoded by the NDUFV1 and NDUFV2 genes in humans), five IP subunits (NDUFS1, NDUFS2, NDUFS3, NDUFS7, and NDUFS8), and seven HP subunits (ND1ND6 and ND4L). The latter are all encoded by the mitochondrial DNA, whereas the remainder are encoded by the nuclear genome.

Complex I deficiency (OMIM 252010 [OMIM] ) is the most common enzymatic deficiency of the OXPHOS system (44, 49, 55) and is associated with a broad spectrum of clinical presentations ranging from early-onset progressive fatal multisystem disorders with lactic acidosis, most often Leigh syndrome, to relatively mild pure myopathic phenotypes (43, 59). In recent years, we have identified (9, 10, 3032, 41, 43, 48, 51) several mutations in nuclear encoded subunits of complex I that are associated with human complex I deficiency. Evidence for a causal relationship between mutation and reduced complex I activity was obtained in Yarrowia lipolytica, a yeast model for complex I deficiency (1).

The effects of complex I mutations on mitochondrial and cellular functioning, which are at the basis of the clinical symptoms, are poorly understood. Earlier studies suggested that the efficiency of the OXPHOS system is somehow linked to mitochondrial shape and/or number (22). Alterations in mitochondrial number and shape are frequently encountered during mitochondrial and cellular dysfunction (2, 15, 18, 20, 25, 35, 38, 52).

Recently, Robinson and coworkers (39) performed a qualitative analysis of mitochondrial shape in fibroblasts from patients with a range of metabolic defects. It was observed that fibroblasts from patients with Leigh syndrome, hypertrophic cardiomyopathy, or fatal infantile lactic acidosis contained higher proportions of "swollen" mitochondrial filaments.

To enhance our understanding of the pathophysiology of mitochondrial complex I deficiency, we studied genetically characterized human complex I-deficient fibroblasts. In doing so, we recently showed (54) that agonist-induced mitochondrial Ca2+ accumulation and ensuing ATP production are significantly decreased in skin fibroblasts from patients with an isolated complex I deficiency caused by mutations in nuclear encoded structural subunits of the complex. Experiments using a newly developed protocol for quantitative analysis of mitochondrial morphology revealed a significant increase in both mitochondrial length and branching in skin fibroblasts from healthy subjects during chronic inhibition of complex I by rotenone (28). This effect of rotenone on the complexity of the mitochondrial network was accompanied by a sustained increase in mitochondrial superoxide production. Most important, the rotenone-induced increase in mitochondrial length and branching, but not the rotenone-induced increase in mitochondrial superoxide production, was completely prevented on cotreatment with the mitochondria-targeted antioxidant mitoquinone (MitoQ). This finding demonstrates that fibroblasts from healthy subjects respond to a reduction in OXPHOS activity through a superoxide-mediated increase in mitochondrial length and branching. In the present study we combined video-rate confocal microscopy and automated image analysis to assess the complexity of the mitochondrial network in living skin fibroblasts and show a tight relationship with the activity of complex I in isolated human complex I deficiency.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chemicals. Culture materials were obtained from Invitrogen (Carlsbad, CA), and rhodamine 123 was purchased from Molecular Probes (Leiden, The Netherlands). All other reagents were from Sigma (St. Louis, MO).

Patient skin fibroblasts. Primary fibroblast cell lines were derived from skin biopsies of 4 healthy subjects (C1–C4) and 13 patients with an isolated complex I deficiency (P1–P13) in the age range of 0–5 yr after informed parental consent was obtained and protocols were approved by the relevant Institutional Review Boards. From each subject, one cell line was derived and stored in liquid nitrogen. The relevant parameters of the cell lines are listed in Table 1. The deficiency was confirmed in both muscle tissue and cultured skin fibroblasts. Complex I activity measurements were performed in a mitochondrion-enriched fraction from cultured skin fibroblasts as described previously (45). The activity of the complex was normalized against that of complex IV (cytochrome-c oxidase), which was measured in the same fraction and expressed as a percentage of the lowest control (0.11 mU/mU cytochrome-c oxidase). Cells were cultured in medium 199 with Earle's salt supplemented with 10% (vol/vol) FCS, 100 IU/ml penicillin, and 100 µg/ml streptomycin in a humidified atmosphere of 95% air-5% CO2 at 37°C. Measurements were performed within five passages after the start of the culture. The passage number at the onset of the culture is given in Table 1. The patient fibroblasts harbored mutations in the NDUFV1 (R59X/T423M; Ref. 41), NDUFS1 (R557X/D618N; manuscript in preparation), NDUFS2 (R228Q, P229Q, S413P; Ref. 32), NDUFS4 (K158fs, VPEEHI67/VEKSIstop, R106X: two unrelated patients; Refs. 9, 32, 51 and manuscript in preparation), NDUFS7 (V122M; Ref. 48), or NDUFS8 (P79L/R102H, R94C; Ref. 30 and manuscript in preparation) subunit of complex I. Also included was one patient with a defective complex I assembly in which no mutation was found in any of the structural subunits (49). All patients were negative with respect to mitochondrial DNA alterations associated with complex I deficiency.


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Table 1. Fibroblast cell lines, mutations, complex I activity, and mitochondrial morphology

 
Cell cycle analysis and cell fusions. For cell cycle analysis, cells were grown to 70% confluence, trypsinized, and stored on ice. After being stained with propidium iodide, cell suspensions were analyzed with flow cytometry (53). For cell fusion, fibroblasts from two patients were mixed in a 1-to-1 ratio and cocultured on coverslips (diameter 22 mm) for 24 h. Subsequently, cells were washed twice with serum-free medium and incubated with 50% (wt/vol) polyethylene glycol (PEG-1000) in medium for 1 min. After two more washes, cells were cultured for another 24 h.

Mitochondrial staining, video-rate confocal imaging, and image analysis. Quantitative analysis of mitochondrial morphology in living cells was performed as described previously (28). Briefly, fibroblasts were grown on coverslips and pulse loaded with rhodamine 123 (200 µM) for 40 s at 20°C. After being loaded, cells were thoroughly washed with HEPES-Tris medium (in mM: 132 NaCl, 4.2 KCl, 1 CaCl2, 1 MgCl2, 5.5 D-glucose, and 10 HEPES, pH 7.4). For confocal imaging, coverslips were mounted in an incubation chamber placed on the stage of an inverted microscope (Nikon Diaphot, Tokyo, Japan) attached to a NORAN OZ laser scanning confocal system (Noran Instruments, Middleton, WI) with a x40 oil immersion planapochromat objective (numerical aperture 1.4; Nikon). Measurements were performed at 20°C. Given the flat morphology of the fibroblasts (size <3 µm in the axial direction), slit settings were chosen in such a way that axially each cell was entirely present within the confocal volume (27). This prevented exclusion of mitochondrial structures from the image and guaranteed an optimal fluorescence signal at minimal laser intensity. Images (512 x 480 pixels) were collected at 30 Hz with a pixel dwell time of 100 ns. To reduce random noise, images were averaged in real time with the running average algorithm of Intervision Acquisition software (Noran) with a window size of 30. This acquisition protocol prevents distortion of the image by mitochondrial movement. Images were recorded from a cross-shaped area transecting the center of the coverslip and converted to TIFF format. Quan-titative analysis of mitochondrial morphology was performed with Image Pro Plus 5.1 software (Media Cybernetics, Silver Spring, MD) as described previously (28).

Statistics. Numerical results were visualized with Origin Pro 7.5 software (OriginLab, Northampton, MA) and are presented as means ± SE. Statistical differences were determined with an independent two-sample Student's t-test (Bonferroni corrected). Pearson's R was calculated to determine the degree of correlation between two sets of data. This parameter approaches 1 or –1 when the linear correlation is directly or inversely proportional, respectively.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Automated analysis of mitochondrial shape and number in human skin fibroblasts. In contrast to earlier studies with human skin fibroblasts (5, 19, 20, 25, 35, 52), the present study was aimed at a more quantitative analysis of mitochondrial shape and number. To this end, we applied our recently developed three-step protocol (28) for automated analysis of mitochondrial morphology. Briefly, cells are first pulse loaded with 200 µM rhodamine 123 for 40 s at room temperature, thoroughly washed, and visualized using video-rate confocal microscopy. Confocal images are then linearly contrast optimized, subjected to a 7 x 7 "top hat" spatial filter, and thresholded. Finally, the resulting binary images are used for automated image analysis yielding the number of mitochondria per cell (Nc), their aspect ratio (AR; ratio between major and minor axes of an ellipse equivalent to the mitochondrion), and their form factor (F; perimeter2/4{pi}·area). Whereas AR is a measure of mitochondrial length, F is a combined measure of both mitochondrial length and degree of branching. Both AR and F are independent of image magnification and have a minimal value of 1 (corresponding to a circular mitochondrion). Therefore, 1 was routinely subtracted from the experimental values obtained for AR and F.

The above procedure is illustrated in Fig. 1, where AR and F are determined in a single fibroblast from a control individual (Fig. 1A) and patient 3 (P3) (Fig. 1B). Figure 1, C (control) and D (P3), depict the mitochondrial structures sorted in descending order from the upper left to the lower right according to their F values. Calculation of the average values of AR and F revealed that both were significantly decreased in the P3 fibroblast (Fig. 1E). Supervised cluster analysis was used to arbitrarily subdivide the mitochondria into three morphological groups, designated groups I, II, and III (Fig. 1, C and D). This allowed determination of which morphological group contributed most to the decrease in AR and F. Figure 1E shows that the average value of F was significantly lower for P3 in all three groups, whereas the average value of AR was significantly lower only in group III.



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Fig. 1. Quantitative analysis of mitochondrial shape and number in human skin fibroblasts. A: control fibroblasts pulse-loaded with rhodamine 123. The asterisk marks the cell used for the illustration of the automated quantitative image analysis procedure. B: similar to A but for fibroblasts from patient 3 (Table 1) harboring a mutation in the NDUFS2 gene. C: mitogram of the control cell marked in A. Individual mitochondria were separated from background and sorted in descending order from top left to bottom right according to their form factor (F). D: similar to C but for the fibroblast of patient 3 marked in B. Note that mitochondria are smaller and appear less branched than in C. E: average values of F and aspect ratio (AR) for all mitochondria (total) and three classes of mitochondria obtained by supervised cluster analysis (classes I, II, and III). Filled bars, control mitochondria; open bars, mitochondria of patient 3. Values significantly lower than control: *P < 0.05, **P < 0.01, ***P < 0.001.

 
Mitochondrial shape and number are altered in complex I-deficient fibroblasts. With the above approach, AR, F, and Nc were compared among 4 control subjects (C1–C4) and 13 complex I-deficient patients (P1–P13; Table 1). The latter cohort consisted of 12 patients with mutations in nuclear encoded subunits of complex I and 1 patient with a complex I assembly defect but without any known mutation in a complex I subunit (P13). Figure 2 shows that F (Fig. 2A), AR (Fig. 2B), and Nc (Fig. 2C) were identical for the four nonrelated control subjects. This finding suggests that genetic variation has no major effect on mitochondrial shape and number in control subjects and justifies pooling of the control data. Seven patients showed either an increased (P1, P6, P10) or a decreased (P2, P3, P4, P5) F, whereas AR was decreased in patients P2, P3, P4, P5, P7, P11, and P13. A significant increase in Nc was observed in patients P4, P5, P8, P9, and P12.



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Fig. 2. Mitochondrial shape and number, but not cell cycle phase, differ greatly between fibroblasts from complex I-deficient patients. A: average values of F. B: average values of AR. C: average values of no. of mitochondria/cell (Nc). C1–C4, individual control cell lines (shaded bars); C, pooled control (filled bar); 1–13, patients 1–13 (open bars). On each day, the average value obtained for control fibroblasts was set at 100%, to which the other values were related. Numerical values are given in Table 1. D: distribution of cell cycle phases in control subjects and 6 representative patients. *Significantly different from control (P < 0.05).

 
To validate our rhodamine 123-based approach for automated analysis of mitochondrial morphology in patient fibroblasts, we used a different staining protocol utilizing mitochondria-targeted enhanced yellow fluorescent protein (mitoEYFP). The analysis involved one control cell line (C4) and two patient cell lines (P1 and P3). To express mitoEYFP, fibroblasts were infected with a baculovirus containing the cDNA of this protein fused with the leader sequence of COX8. This virus was produced essentially as described by Visch et al. (54). After image processing, automated analysis revealed that P1 fibroblasts displayed a significantly (P < 0.01) increased F (138 ± 8%, n = 30) and normal AR (98 ± 2) and Nc (103 ± 2), whereas P3 fibroblasts showed a significantly (P < 0.001) decreased F (64 ± 2, n = 27) and AR (72 ± 3) and a normal Nc (103 ± 2). Comparison with the values presented in Table 1 shows that these values were not different from those obtained with rhodamine 123.

To rule out the possibility that the differences in mitochondrial shape and number were due to alterations in cell cycle phase, patient fibroblasts were stained with propidium iodide and analyzed using flow cytometry. Figure 2D shows that there were no apparent differences in cell cycle phase distribution between control and patient fibroblasts. Moreover, AR, F, and Nc were not correlated with cell cycle phase. Similarly, no correlation between passage number and mitochondrial morphology was observed (Table 1).

Plotting AR as a function of F for all patient cell lines used in this study revealed a linear correlation (Fig. 3A; R = 0.90, P < 0.001). This shows that the relative changes in mitochondrial length and degree of branching occurred in parallel in these cell lines. In contrast, no correlation was observed between Nc and F (Fig. 3B; R = –0.08; P = 0.79) or between Nc and AR (not shown; R = 0.012; P = 0.70), demonstrating that alterations in mitochondrial morphology and number are independent of each other.



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Fig. 3. Mitochondrial length correlates with the degree of mitochondrial branching but not with Nc. A: linear correlation between F and AR. B: lack of correlation between F and Nc. Numerals indicate patient numbers (Table 1). See RESULTS for details.

 
Differences in mitochondrial shape and number are independent of the complex I subunit that is affected. The data presented in Table 1 show that there is no direct connection between the affected subunit and the mitochondrial phenotype. For instance, patients with a defect in the NDUFS4 gene displayed increased branching (P6), shortening (P7), or increased number (P8, P9). Similarly, patients with a mutation in the NDUFS2 gene showed either reduced branching and shortening (P3) or fragmentation (P4, P5), whereas patients with a mutation in the NDUFS8 gene displayed either shortening (P11) or an increased number of mitochondria (P12).

Altered mitochondrial shape is restored by genetic complementation. To provide evidence for a causal connection between altered mitochondrial phenotype and isolated complex I deficiency, we performed cell fusion experiments between fibroblasts from patient 2 (P2), carrying a mutation in the NDUFS1 gene (chromosome 2q33–q34), and patient 3 (P3), with a mutation in the NDUFS2 gene (chromosome 1q23). Both patients displayed a significant reduction in F, whereas Nc was normal (Table 1). After equal amounts of fibroblasts from both patients were mixed, cells were seeded on coverslips and grown for 24 h. Next, cell fusion was induced by 1-min treatment with PEG-1000. After another 24 h of culturing, cells were stained with rhodamine 123 for analysis of mitochondrial morphology. The number of nuclei per cell was used to discriminate between fused and nonfused cells. Values of F were normalized to those of non-PEG-1000-treated control cells recorded on the same day. In addition to the heterologous fusions of patients 2 and 3 (P2 x P3), we also performed homologous fusions between individual control cells (C x C), cells of patient 2 (P2 x P2), and cells of patient 3 (P3 x P3). On average, fused cells contained two to three nuclei [C x C: 2.8 ± 0.4 nuclei, n = 13 fused cells (bar g); P2 x P2: 2.4 ± 0.3, n = 12 fused cells (bar h); P3 x P3: 2.1 ± 0.1, n = 7 fused cells (bar i); and P2 x P3: 2.4 ± 0.2, n = 28 fused cells (bars k and l); Fig. 4A]. During homologous fusions (Fig. 4A; C x C, P2 x P2, and P3 x P3), PEG-1000 treatment induced a small reduction in F in both nonfused (bars d, e, f) and fused (bars g, h, i) cells. Only during heterologous fusions (P2 x P3) was F found to be increased to control values in 8 of 28 fused cells (29%; Fig. 4A, bar l). Figure 4B shows representative cells with a restored (left) and a nonrestored (right) mitochondrial network. PEG-1000 treatment did not alter Nc (data not shown).



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Fig. 4. Restoration of mitochondrial shape by genetic complementation in human complex I deficiency. A: average values of F 24 h after polyethylene glycol (PEG)-induced fusion of control cells (C), cells from patient 2 (P2), cells from patient 3 (P3), and a mixture of equal amounts of cells from patients 2 and 3 (P2 x P3). Only in the P2 x P3 fusion experiments were fused cells with a mitochondrial morphology identical to controls observed (bar l), whereas the remainder displayed a patient phenotype (bar k). B: binuclear fibroblasts observed in a cell fusion experiment between fibroblasts of patient 2 (P2) and patient 3 (P3) displaying a restored (left) and a nonrestored (right) mitochondrial morphology. Number of cells analyzed: n = 29 (a), n = 27 (b), n = 32 (c), n = 16 (d), n = 9 (e), n = 20 (f), n = 13 (g), n = 12 (h), n = 7 (i), n = 11 (j), n = 20 (k), and n = 8 (l). Statistical difference with respect to the indicated condition (a, d, or g): *P < 0.05, **P < 0.01, ***P < 0.001; n.s., not significant.

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Alterations in mitochondrial shape and number have been associated with cell death, protection against metabolic stress, adaptation to changes in mitochondrial substrates, nuclear transport, and mitochondrial pathology (2, 14, 16, 25, 35, 38, 52, 46). In this study we applied a recently developed protocol for automated analysis of mitochondrial shape and number in living cells (28) and found that either one or both were significantly altered in skin fibroblasts from 13 patients with an isolated complex I deficiency.

Validation of rhodamine 123-based protocol for automated analysis of mitochondrial morphology in complex I-deficient human skin fibroblasts. Specific staining of mitochondria with rhodamine 123 is based on the highly negative mitochondrial membrane potential ({Delta}{psi}). Importantly, the first step during image processing (linear contrast optimization) enhances all signals that are above background and thereby minimizes the effect of regional differences in {Delta}{psi} on the outcome of our automated analysis. Thus reductions in {Delta}{psi}, as might be expected in complex I-deficient fibroblasts, will only interfere with our analysis when they lead to a dramatic loss of {Delta}{psi}. Using the mitochondrial potential sensor JC-1, we previously showed (54) that {Delta}{psi} is only slightly decreased in complex I-deficient patient fibroblasts. Moreover, we recently showed (28) that rotenone, added at a concentration at which it reduced the activity of complex I by 80%, did not alter rhodamine 123 staining, and therefore {Delta}{psi}, but significantly increased F. Finally, automated analysis of the mitochondrial morphology of two patient cell lines displaying distinct morphological phenotypes gave identical results after expression of mitoEYFP or staining with rhodamine 123. Although our approach has not been verified by serial sectioning electron microscopy, preliminary data from fluorescence loss in photobleaching (FLIP) experiments with mitoEYFP-expressing human fibroblasts indicate that the mitochondrial structures identified by the automated analysis protocol indeed form a continuum.

Aberrant mitochondrial phenotypes in complex I-deficient human fibroblasts. To compare mitochondrial morphology between control and patient cells, we determined three parameters: the aspect ratio AR, which is a measure of mitochondrial length, the form factor F, which is a combined measure of both mitochondrial length and degree of branching, and Nc, which is a measure of the number of mitochondria per cell (28). All patient cell lines (P1–P13) displayed alterations in at least one of these parameters compared with four healthy control subjects (Fig. 2 and Table 1). Importantly, these changes did not correlate with passage number (Table 1). This is in agreement with previous findings showing that mitochondrial morphology is independent of passage number in these cells (20). Recent work revealed a connection between cell cycle phase and mitochondrial structure in human osteosarcoma (143B) cells (34). However, no such connection was found for human skin fibroblasts in the present study.

Our large-scale analysis of mitochondrial shape and number in ~2,500 individual cells containing ~250,000 mitochondria allowed identification of 5 aberrant phenotypes (Table 1). Three patients with mutations in the NDUFV1 (P1), NDUFS4 (P6), or NDUFS7 (P10) gene displayed a single increase in F (phenotype I: increased branching). Three patients (P7, P11, P13) displayed a single decrease in AR (phenotype IV: shortening). The latter patients carried a mutation in either the NDUFS8 (P11) or the NDUFS4 (P7) gene, whereas P13 had an assembly defect of which the genetic cause is not yet known. Two patients, one with a mutation in the NDUFS1 gene (P2) and one with a mutation in the NDUFS2 gene (P3), displayed a significant decrease in F and AR, whereas Nc was unaltered (phenotype II: decreased branching and shortening). Two patients with a different mutation in the NDUFS2 gene (P4, P5) showed a decrease in F and AR and an increase in Nc, suggesting that this phenotype might be associated with mitochondrial fragmentation (phenotype III: fragmentation). Finally, in three patients with mutations in either the NDUFS4 (P8, P9) or the NDUFS8 (P12) gene, a single increase in Nc was observed (phenotype V: increase in number). In conclusion, our analysis reveals two types of mitochondrial morphology change within our cohort of complex I-deficient patient fibroblasts: 1) a parallel change in length and degree of branching (Fig. 3A) and 2) an increase in number.

Concerning the other OXPHOS complexes, we have thus far analyzed the mitochondrial morphology in primary skin fibroblasts from five patients with an isolated complex II deficiency and two patients with an assembly defect in complex IV. It was found that two of the five complex II-deficient fibroblasts displayed only a small increase in F, whereas this parameter was unaltered in both complex IV-deficient fibroblasts. The latter observation is in agreement with a previous report in the literature (35). These preliminary findings suggest that as far as respiratory chain deficiencies are concerned, the aberrations in mitochondrial morphology may be restricted to cells with a deficiency in complex I. Clearly, more patient cell lines must be analyzed to assess the extent to which mitochondrial morphology is changed in human complex II deficiency and to prove the absence of such alterations in human complex IV deficiency.

Linear relationship between mitochondrial network complexity and residual complex I activity in human complex I deficiency. Complex I activity measurements were performed in a mitochondria-enriched fraction from cultured skin fibroblasts. The activity of the complex was normalized against that of complex IV (cytochrome-c oxidase), which was measured in the same fraction, and expressed as a percentage of the lowest control (0.11 mU/mU cytochrome-c oxidase; Ref. 45). Importantly, all patients who participated in this study had normal complex IV activity. Close examination of the normalized complex I activities (Table 1) revealed relatively low values for patients in whom F was markedly decreased (P2, P3) and/or Nc was markedly increased (P4, P5, P8, P12).

To investigate the relationship between complex I activity and F and Nc for the whole cohort of patient fibroblasts used in this study, we plotted the ratio of complex I to complex IV activity against that of F to Nc (Fig. 5). Figure 5 demonstrates a linear relationship between the normalized complex I activity and the ratio of F to Nc (R = 0.90, P < 0.0001). A similar correlation was found between normalized complex I activity and the ratio of AR to Nc (R = 0.72, P = 0.005). Importantly, Fig. 5 shows that the higher activity ratios are associated with a single increase in F (none of the patients displayed a significant reduction in Nc), whereas the lower activity ratios are associated with a decrease in F and/or an increase in Nc. Because a decrease in F and an increase in Nc reflect a condition of fragmentation of the mitochondrial network, whereas a single increase in F represents an increase in complexity of this network, our data provide first experimental evidence for a tight correlation between complex I activity and mitochondrial network complexity in complex I-deficient patient fibroblasts.



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Fig. 5. Linear relationship between residual complex I activity and F-to-Nc ratio in human complex I-deficient fibroblasts. Numerical values of F, Nc, and complex I activity are given in Table 1. Roman numerals indicate different mitochondrial morphological phenotypes. C, pooled control value.

 
Differences in mitochondrial morphology are independent of affected complex I subunit. Analysis of the aberrant mitochondrial phenotypes revealed that there was no correlation between shape and/or number and the affected complex I subunit. The fact that mitochondrial shape was also aberrant in a complex I-deficient patient with a defective complex I assembly but without any known defect in a complex I gene (P13) supports this idea. Patients with mutations in the NDUFS4 gene display a marked clinical heterogeneity, even in (unrelated) patients with the same mutation (10). It is tempting to speculate that the distinct mitochondrial morphologies revealed in this study are somehow related to the diverse clinical presentations of human complex I deficiency (43, 44).

Restoration of altered mitochondrial shape by somatic fusion. Our imaging approach gave us the unique opportunity to assess whether mitochondrial morphology was restored on complementation of the genetic defect by somatic fusion. Recently, genetic complementation was shown to restore complex I activity, leading to the identification of mutations in a nuclear complex I gene, the NDUFS6 gene, not previously associated with complex I deficiency (26). In the present study, two primary patient cell lines, one with a mutation in the NDUFS1 gene (P2) and one with a mutation in the NDUFS2 gene (P3), both displaying a reduction in F but normal Nc, were chosen for the cell fusion experiment. The low F values allowed optimal demonstration of morphological restoration. Mitochondrial morphology was fully restored in 29% of the fused cells. This finding is in agreement with a situation in which each of the two cell lines harbors a different mutation, leading to the aberrant mitochondrial morphology.

Full restoration of the mitochondrial network was observed at 24 h after cell fusion. This kinetics is in agreement with our recent work (50) showing that steady-state levels of fully assembled complex I are achieved within 24 h after induction in 143B osteosarcoma cells. In contrast, another study required 48 h for the assembly of complex I (58), whereas in agreement with the present findings, all small and spherical {rho}0 mitochondria were found to take a normal elongated shape within a few hours after introduction of normal mitochondria into {rho}0 HeLa cells (21).

Possible mechanisms underlying aberrations in mitochondrial morphology in complex I-deficient patient fibroblasts. Our findings support the existence of a link between complex I deficiency and mitochondrial network complexity. Using primary skin fibroblasts from a healthy subject, we recently showed (28) that inhibition of complex I by rotenone is paralleled by a significant increase in F but not AR and Nc. This suggests that an increase in mitochondrial branching may constitute a "default" adaptive response to counterbalance the effects of a reduction in complex I activity. In agreement with this idea, preliminary data suggest that the rotenone-induced increase in mitochondrial complexity is associated with an increased expression of complex I. Increases in mitochondrial network complexity may lead to improved ATP delivery (42) and/or enhanced intermitochondrial signaling through reactive oxygen species (3).

Similarly to results with rotenone, one group of patient fibroblasts showed an increase in F but not AR and Nc. This suggests that these fibroblasts were able to execute the default adaptive response. Intriguingly, these fibroblasts showed a relatively small decrease in complex I activity compared with the healthy fibroblasts. In contrast, a second group of patient fibroblasts showed a decrease in F and/or increase in Nc, suggesting that these fibroblasts were unable to perform the adaptive response. Compared with healthy fibroblasts, these patient fibroblasts showed a large decrease in complex I activity. The rotenone concentration that maximally increased F decreased the activity of complex I by 80% (28). The same decrease was observed in fibroblasts of P12, which displayed the lowest activity of all fibroblasts used in this study. However, these patient fibroblasts did not show an increase in F but an increase in Nc. This indicates that factors other than the mere reduction in complex I activity are involved in determining mitochondrial morphology in patient fibroblasts.

One possible explanation is that the rate of mitochondrial superoxide production is increased physiologically in the adapting patient fibroblasts but pathologically in the nonadapting fibroblasts. Indeed, preliminary data show that superoxide production is significantly but to a variable degree increased in our cohort of complex I-deficient patient fibroblasts. In contrast, fibroblasts from patients with an isolated complex II (5 patients) or complex IV (2 patients) deficiency did not show such an increase and, as discussed above, also did not show a major change in F. Evidence that mitochondrial superoxide production may play a role in mitochondrial morphology changes comes from our recent observation (28) that the mitochondria-targeted antioxidant mitoQ abolishes the rotenone-induced increase in F in healthy fibroblasts. In addition, mitoQ abolished the rotenone-induced increase in lipid peroxidation, suggesting that increased mitochondrial superoxide production acts through an increase in lipid peroxidation to alter mitochondrial morphology. In agreement with the above findings, lipid peroxidation products were found to be increased in skin fibroblasts from complex I-deficient patients (33). Alternatively, the increase in oxygen free radicals observed in complex I-deficient patient fibroblasts may alter the activity of redox-sensing nuclear transcription factors (11).

In summary, the present study shows that mitochondrial network complexity and complex I activity are tightly coupled in complex I-deficient patient fibroblasts and that the aberrations in mitochondrial network complexity in these cells are indeed due to the presence of the disease-causing mutation.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by equipment grants of Netherlands Organization for Health Research and Development (ZON, No. 903-46-176) and Netherlands Organization for Scientific Research (NWO, No. 911-02-008) and the European Community's Sixth Framework Program for Research, Priority 1 "Life sciences, genomics and biotechnology for health," contract number LSHM-CT-2004-503116.


    ACKNOWLEDGMENTS
 
We are grateful to Prof. O. Elpeleg (Metabolic Disease Unit, Shaare-Zedek Medical Center, Jerusalem, Israel), Prof. H. Mandel (Dept. of Pediatrics, Rambam Medical Center, Technion-Faculty of Medicine, Haifa, Israel), Prof. S. Stöckler-Ipsiroglu (Dept. of Pediatrics, General Hospital and University Hospital, Vienna, Austria), Dr. B. Plecko (Universitätsklinik fur Kinder- und Jugendheilkunde, Universität Graz, Austria), and Dr. F. A. Wijburg (Dept. of Pediatrics, Academic Medical Center, Amsterdam) for providing the patient cell lines. We also thank the technicians of the Nijmegen Center for Mitochondrial Disorders for complex I activity measurements and cell culture and A. Pennings (Dept. of Hematology, Radboud University Nijmegen Medical Centre) for cell cycle analysis.


    FOOTNOTES
 

Address for reprint requests and other correspondence: J. A. M. Smeitink, Nijmegen Center for Mitochondrial Disorders, Dept. of Pediatrics, Radboud Univ. Nijmegen Medical Center, PO Box 9101, 6500 HB Nijmegen, The Netherlands (e-mail: j.smeitink{at}cukz.umcn.nl)

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.


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