Mitochondrial Medicine, Biochemical Evaluation of Mitochondrial Function - CHEMICAL BIOLOGY


Mitochondrial Medicine, Biochemical Evaluation of Mitochondrial Function

Richard J.T. Rodenburg and Jan A.M. Smeitink, Nijmegen Center for Mitochondrial Disorders, Radboud University Nijmegen Medical Centre, The Netherlands

doi: 10.1002/9780470048672.wecb343

Mitochondrial medicine is an emerging field in medicine that focuses on diseases in which the mitochondrial energy generating system plays a central role. These diseases include the ''classical'' mitochondrial disorders, in which a (genetic) defect in the mitochondrial energy generating system is the primary cause of the disease, but also more common disorders such as Parkinson and cancer, in which mitochondrial energy metabolism plays an important role in the pathogenesis. In this review, we present an overview of the mitochondrial energy generating system, starting at the conversion of pyruvate into acetyl-CoA by pyruvate dehydrogenase, via the TCA cycle in which reduction equivalents are formed, to the oxidative phosphorylation system, where the reduction equivalents are used to convert ADP into ATP. The mitochondrial energy generating system can be examined by global assays that measure the rate of pyruvate conversion, the rate of oxygen consumption, or the rate of ATP production. In addition, an overview is given of the spectrophotometric assays that are available to measure PDHc, several TCA cycle enzymes, and the enzymes of the oxidative phosphorylation. In the diagnostic analysis of patients suspected to suffer from a mitochondrial disorder, these assays are applied to evaluate the functioning of the mitochondrial energy generating system in muscle biopsies and other types of patient samples. In addition, the results provide clues for further investigations at the molecular genetic level. Thus, the biochemical analysis of patient material is an important step in establishing the diagnosis of a mitochondrial disorder.

The importance of mitochondria in health and disease has given rise to a new area in medicine, called mitochondrial medicine. The foundation for this medical discipline lies in the recognition of diseases in which disturbances in one of the many steps of mitochondrial energy production are present. The role of mitochondria in energy metabolism disorders is well recognized and has been the subject of study for many decades. The increasing awareness of the relationship between mitochondria and several more common disorders, like Parkinson disease and cancer, makes a thorough understanding of the chemistry of the mitochondrial energy generating system, and the analytical methods to examine the functioning of this system, necessary for an increasing variety of medical and biochemical specialists. Here, we review a) how the cell’s energy currency, ATP, is produced, and b) assays to determine the overall capacity of the system as well as single enzymes in relation to genetic disorders of energy production.

Biological Background

Mitochondrial disorders

Mitochondrial disorders can be defined as disorders that are caused by a defect in the mitochondrial energy generating system (MEGS). The clinical spectrum is very broad, but in almost all cases involve one or more tissues that have a high energy demand, in particular skeletal muscle and neuronal tissue. In addition, heart, liver, kidney, and other tissues can be involved as well. The severity and the course of the disease are extremely diverse, ranging from severe neonatal systemic disorders to mild exercise intolerance presenting at a high age. In addition to these “classical” mitochondrial disorders, it has become apparent that the mitochondrion and its energy generating system plays a role in other, more frequently occuring diseases as well, such as Parkinson (1), diabetes (2), and cancer (3). This review will be restricted to “classical” mitochondrial disorders, although the assays and principles described here could be of value for the investigation of other disorders as well.

The mitochondrial energy generating system

The production of ATP by the MEGS is a complex process involving many different transporters and enzymes (4). Mitochondrial ATP is the end product of the oxidation of pyruvate (alpha-keto propionic acid). Pyruvate is the final product of the glycolysis, the anaerobic catabolism of glucose. Other substrates for the MEGS are fatty acids and several amino acids, in particular glutamine. Pyruvate is transported into the mitochondria where it is metabolized into acetyl CoA. The acetyl CoA is oxidized in the tricarboxylic acid (TCA) cycle, during which both NADH and FADH2 are produced. These reducing equivalents are oxidized by the respiratory chain, which leads to the translocation of protons out of the mitochondrial matrix. The mitochondria use the resulting proton-motive force to generate ATP from ADP and phosphate. The ATP is released in the mitochondrial matrix, and can be exported to the cytosol. The MEGS is described in more detail in the next chapter.

The Chemistry of the OXPHOS System

The conversion of pyruvate to acetyl-CoA

The conversion of pyruvate into acetyl-CoA involves two key players. Pyruvate is imported into the mitochondrion by a specific transporter. To date, only one case has been described in which a pyruvate carrier deficiency could be shown by functional assays (5). However, no genetic defect responsible for pyruvate carrier deficiency has been identified yet. After pyruvate has entered the mitochondrial matrix, it is converted into acetyl-CoA by the pyruvate dehydrogenase complex, PDHc. This large enzyme complex has a molecular mass of approximately 9 MDa and contains multiple copies of three enzymatic entities: 20-30 copies of alpha-ketoacid dehydrogenase (E1; EC, 60 copies of dihydrolipoamide acyltransferase (E2; EC, and 6 copies of dihydrolipoamide dehydrogenase (E3; EC, as well as 12 copies of the structural building block E3 binding protein (6). The E1 subcomplex consists of a tetramer of two E1a and two E1β subunits. PDHc decarboxylates pyruvate and esterifies the resulting acetyl-group to CoA. During this reaction, NAD+ is reduced to NADH. PDHc requires 5 different cofactors, namely NAD+, CoA, thiamine pyrophosphate, FAD and lipoic acid. The activity of PDHc is tightly regulated. First of all, the activity is controlled in an allosteric manner by the reaction products acetyl-CoA and NADH. In addition, two specific enzymes regulate PDHc activity via a phosphorylation site in the E1 subunits. PDH kinase (EC is ATP dependent and inactivates PDHc by phosphorylating the E1 component when the ATP/ADP ratio is high (7). This enzyme is also activated by high NAD+ and acetyl-CoA levels, and inactivated by high pyruvate levels. Four isoforms of PDH kinase have been identified, each having a different tissue distribution. By contrast, when the ATP/ADP ratio is low and the pyruvate levels are high, PDH kinase is not active and PDHc E1 is dephosphorylated by PDH phosphatase (EC, of which two isoforms are known (8). Due to this tight regulation, the oxidation rate of pyruvate by PDHc is directly coupled to the mitochondrial ATP production rate.

The generation of reduction equivalents in the TCA cycle

The acetyl-CoA generated by PDHc fuels the TCA cycle. Three dehydrogenases of the TCA cycle are responsible for the reduction of NAD+ into NADH. These are isocitrate dehydrogenase (EC, α-ketoglutarate dehydrogenase (consisting of 3 enzymatic subunits E1 (EC, E2 (, and E3 (EC, and malate dehydrogenase (EC In addition, the TCA cycle enzyme succinate dehydrogenase (EC converts FAD into FADH2. Furthermore, succinate-CoA ligase generates GTP or ATP, depending on the isotype of this enzyme complex (EC and EC, respectively). The formation of these high-energy molecules is accompanied by the release of CO2 in two of the reactions of the TCA cycle. In addition to acetyl-CoA, other metabolites can fuel the TCA cycle as well, This includes glutamine, which enters the TCA cycle at the site of 2-oxoglutarate. In tissues that catabolize fatty acids, the end product of the beta-oxidation is acetyl-CoA, which is converted into the ketone bodies 3-hydroxybutyrate and acetoacetate (in the liver) or enters the TCA cycle (in most other tissues) and in this way contributes to the synthesis of ATP.

The oxidative phosphorylation system

NADH and FADH2 are oxidized by the OXPHOS system to generate ATP. This is a coordinated multistep process that involves 5 large enzyme complexes: the respiratory chain complexes I, II, III, and IV, and ATP synthase (complex V). Complex I (NADH:ubiquinone oxidoreductase; EC is by far the largest respiratory chain enzyme complex. It consists of 45 different subunits and has a molecular weight of approximately 1 MDa. Complex I oxidizes NADH and the electrons that are released from NADH are transferred to a flavin mononucleotide present in complex I and subsequently via a channel of 8 iron sulfur clusturs within the peripheral arm of the complex towards CoQ10 that is present in the innner mitochondrial membrane (9). The redox reaction of complex I is directly coupled to the pumping of protons by complex I from the mitochondrial matrix across the mitochondrial inner membrane to the mitochondrial intermembrane space, which is in direct connection with the cytosol for small ions. Complex II (succinate dehydrogenase; EC oxidizes FADH2 and translocates electrons towards CoQio-Complex II is the only respiratory chain complex that does not contribute to the mitochondrial proton gradient. Complex III (ubiquinol:cytochrome c oxidoreductase; EC translocates the electrons from CoQ10 to cytochrome c, a small, haeme-containing protein that acts as an electron carrier between complexes III and IV of the respiratory chain. The last step of the respiratory chain is complex IV (cytochrome c oxidase; EC, that oxidizes cytochrome c and transfers the electrons to molecular oxygen, which leads to the generation of water. The pumping of protons by the respiratory chain complexes I, III, and IV across the inner mitochondrial membrane increases the pH of the mitochondrial matrix and generates a potential difference between matrix and inter membrane space. Complex V (adenosine triphosphatase; EC utilizes the potential energy of the proton gradient to convert ADP and phosphate into ATP.

Defects in the mitochondrial energy generating system

In theory, a defect in any of the transport proteins and enzymes mentioned above could result in a reduced mitochondrial energy generating capacity. To date, primary defects at the protein and DNA level have been found in PDHc, fumarase (EC (10), α-ketoglutarate dehydrogenase (2-oxoglutarate dehydrogenase) (11), succinate-CoA ligase (12, 13), complexes I (9), II (14), III (15), IV (16), and V (17), the phosphate carrier (18), and ANT. A functional defect in the pyruvate carrier has been identified as well (5). In addition to these structural components of the mitochondrial energy generating system, there are many additional proteins involved in the production of these structural components. These include the chaperones required to assemble the enzyme complexes of the respiratory chain. The structural building blocks of the OXPHOS system are encoded by multiple genes. Except for complex II, these genes are located in both the nuclear and the mitochondrial genomes. Most nuclear genetic defects in these structural genes result in an isolated enzyme deficiency, although several examples have been described of patients with a mutation in a complex I gene that also result in reduced enzyme activities of other enzyme complexes, e.g. complex III and PDHc, indicating that these complexes have a higher order of organisation that can be disturbed by defects in one of the complexes. The existence of the so-called supercomplexes has become more apparent by functional and structural studies in the last few years. Many different pathogenic point mutations and rearrangements in the mtDNA have been found in the last two decades. Depending on the type of mutation, these can cause either isolated or combined deficiencies of complexes I, III, IV, and V (19). The mitochondrial genome (or mtDNA) is replicated by a dedicated polymerase, POLG. Defects in the POLG gene leads to depletion of, and/or deletions in, the mtDNA. This causes isolated or combined deficiencies of the mtDNA encoded OXPHOS complexes I, III, IV, and/or V. Furthermore, defects in proteins involved in nucleotide metabolism can lead to mtDNA depletion and OXPHOS deficiencies as well. Depletion has also been observed in patients suffering from a defect in the TCA cycle enzyme succinate-CoA ligase, although the underlying mechanism is not yet fully understood (12, 20). The transcription and translation of mtDNA encoded proteins involves several mitochondria-specific proteins, in which defects also lead to combined OXPHOS enzyme deficiencies (21, 22). A special class of defects are those leading to a CoQ10 deficiency. The biosynthesis of CoQi0 is a multistep process that has been completely elucidated in yeast. In humans, several steps of this process have been shown to exist as well, and three different genetic defects in this pathway have been identified to date (23).

Tools and Techniques to Study the OXPHOS System

There are several approaches to perform biochemical analyses of the OXPHOS system. This chapter will focus on assays to perform structural analyses, enzyme activity assays, and ATP production, oxygen consumption and substrate oxidation assays.

Structural analysis of the OXPHOS system

Blue-Native PAGE is a technique that is very suitable to study the relative amounts and the assembly status of OXPHOS complexes (24, 25). It can be performed as either a 1D or a 2D assay. In the 1-dimensional approach, the complexes are separated on a non-denaturing acrylamide gel containing the Commassie dye Serva Blue G. All 5 complexes can be visualised in this way. In the 2-dimensional approach, the second dimension is a denaturing SDS-PAGE, resulting in the separation of the OXPHOS complexes into their individual protein building blocks. After blotting, more specific staining methods can be performed, e.g. using anti-OXPHOS complex antibodies. This can provide detailed information on the assembly status of the OXPHOS complexes. Another powerful tool to perform structural analysis of OXPHOS complexes is by immunopurification followed by mass spectrometric analysis of the isolated complexes. Using this approach, it has been shown that bovine complex I (and presumably also human complex I) consists of 45 different subunits (26). Recently, this technique has lead to the identification of Ecsit as a protein involved in complex I assembly, as it was found to be associated with complex I (27). This is quite an unexpected finding, as Ecsit was previously known as a cytosolic protein involved in a pro-inflammatory signal-transduction pathway from a Toll-like receptor and in embryonic development (28). By immunopurification and mass spectrometry, it could be shown that Ecsit associates with complex I, and that the N-terminal part of Ecsit appears to be required for mitochondrial import (27).

Enzyme activity assays

The traditional way to determine OXPHOS enzyme activities is by spectrophotometry. Several assays have been described for all 5 OXPHOS complexes. The assays are performed in homogenates of tissue samples or cultured cells, in crude mitochondria-enriched 600 g supernatants of tissue/cell homogenates, or in mitochondrial preparations from 14000 g pellets derived from 600 g supernatants. Obviously, the higher the purity of the mitochondrial preparation, the higher the specific activity of the enzymes measured. Therefore, very high specific activities can be achieved by using immunopurified complexes

(29), although this approach is not widely applied yet. In addition to enzyme activity assays in solution, BN-PAGE can be used to estimate OXPHOS enzyme activities by in-gel activity assays (30). In-gel activities are particularly suitable to monitor relative activities, e.g. to evaluate changes in enzyme activities under different experimental conditions. For quantitative enzyme activity measurements, spectrophotometric analysis is the method of choice. Table 1 contains a summary of the most commonly used spectrophotometric enzyme assays for measurement of the OXPHOS enzymes. Below, a brief description of these assays is given.

Complex I

Spectrophotometric assays for measuring the activity of complex I (or NADH:ubiquinone oxidoreductase) are usually based on measurements of NADH, which is oxidized by complex I to NAD+. In addition, the assay requires CoQ as a cosubstrate for complex I. Usually CoQ1 or decylubiquinone are used because these have better solubility than CoQ10. Assays usually contain bovine serum albumine, which probably is required to stabilize the protein sample and to aid the solubilization of CoQ analogues. The specific activity of complex I can be determined by measuring the rate of NADH conversion at 340 nm in the absence and presence of the specific complex I inhibitor rotenone (31). Recently, our lab described a new assay that measures complex I by including 2,6-dichlorophenolindophenol (DCIP) as a terminal acceptor of electrons that are derived from the oxidation of NADH and the subsequent reduction of the CoQ-analogue decylubiquinone (32). DCIP reduction can be followed spectrophotometrically at 600 nm. As the molar absorption coefficient of DCIP at 600 nm is more than 3 times higher than that of NADH at 340 nm, this new assay has a much higher sensitivity than the assay that measures NADH, with similar specificity. Complex I can also be measured as NADH:cytochrome c oxidoreductase, in which the combined activity of complex I + CoQ10 + III is measured by addition of NADH and oxidized cytochrome c as substrates. The assay measures the rotenone-sensitive reduction of cytochrome c, which can be followed spectrophotometrically at 550 nm.

Complex II

Complex II is usually measured in two ways: either as succinate:ubiquinone oxidoreductase or as succinate:cytochrome c oxidoreductase. The most commonly used assay for succinate:ubiquinone oxidoreductase (or isolated complex II) uses DCIP in the same way as described above for the new complex I assay, only in this case succinate is added as a substrate (instead of NADH). The specificity of DCIP reduction can be determined by measuring in the presence or absence of malonate, a specific inhibitor of complex II. The assay for succinate:cytochrome c oxidoreductase (or complex II + III) uses succinate and oxidized cytochrome c as substrates and measures the reduction of cytochrome c, which can be followed spectrophotometrically at 550 nm. The assay is also suitable to screen for coenzyme Q deficiencies, as it is dependent on the endogenously present CoQ10. In case of a CoQ deficiency, a reduced succinate:cytochrome c oxidoreductase activity will be observed that can be normalized by addition of exogenous CoQ to the reaction mixture (33).

Complex III

Complex III (ubiquinol:cytochrome c oxidoreductase) reduces cytochrome c and oxidizes reduced CoQ. In addition to these substrates, the assay contains a strong complex IV inhibitor (e.g. potassium cyanide) to prevent re-oxidation of reduced cytochrome c. The complex III activity can be derived from the rate of cytochrome c reduction, which can be followed at 550 nm. The specificity of the assay is determined by measuring in the absence or presence of antimycin A, a specific inhibitor of complex III activity.

Complex IV

Complex IV (cytochrome c oxidase) is measured by addition of reduced cytochrome c to the reaction mixture. The oxidation of cytochrome c can be followed at 550 nm. This assay has a low background activity, and measurement in the absence and presence of a selective inhibitor is not necessary.

Complex V

The activity of the ATP-forming enzyme complex V is usually assessed by determining the reverse reaction ATP → ADP + Pi. The reaction is coupled to reactions catalyzed by pyruvate kinase (ADP + phosphoenolpyruvate → pyruvate + ATP) and lactate dehydrogenase (pyruvate + NADH → lactate + NAD+). This final reaction can be followed spectrophotometrically by measuring NADH at 340 nm. The activity of complex V (ATPase) can be derived from the rate of NADH conversion in the presence and absence of the specific complex V inhibitor oligomycin.

Other enzymes

In addition to assays for the OXPHOS enzymes, assays have been described for many other enzymes involved in the MEGS. For PDHc, various types of diagnostic assays have been described and are widely used (33). Also for the TCA cycle enzymes various assays have been described, although to date, pathogenic defects have only been found in α-ketoglutarate dehydrogenase (38), succinate-CoA ligase (12), and fumarase (39).

Table 1. Summary of respiratory chain, complex V and PDHc activity assays. All assays are spectrophotometric assays except for the PDHc assay with CO2 detection, which is a radiochemical assay. The specific inhibitor indicated is used for blank measurements. Non-standard abbreviations: UQ1: ubiquinone-Q1; DQ: decylubiquinone; DCIP: 2,6-dichlorophenolindophenol, PK: pyruvate kinase; LDH: lactate dehydrogenase; AABS: p-[p-(aminophenyl)azo]benzene sulfonic acid; ArAt: arylamine acetyltransferase


Activity measured


Specific inhibitor



complex I





NADH (340 nm)

[31, 33, 35]



DCIP (600 nm)


complex I + III

NADH:cytochrome c



cytochrome c



cytochrome c

(550 nm)

complex II


succinate, DQ


DCIP (600 nm)

[33, 35]


complex II + III




cytochrome c

[33, 35]

c oxidoreductase

cytochrome c

(550 nm)

complex III


DQ (red),

antimycin A

cytochrome c

[33, 35]

c oxidoreductase

cytochrome c

(550 nm)

complex IV

cytochrome c oxidase

cytochrome c (red) none

cytochrome c

[33, 35]

(550 nm)

complex V




NADH (via

[33, 35]

PK/LDH at 340 nm)



pyruvate, NADH,


CO2 (radiochemically)

NADH (340 nm) Acetyl-CoA (via

AABS/ArAt at 460 nm)

[35, 37]



The mitochondrial energy generating system

As outlined above, the generation of ATP from pyruvate by the mitochondrion involves many different transporters and enzymes. A subset of the individual components can be assayed individually (e.g. the respiratory chain enzymes). In addition, the process of ATP generation can be studied by using assays that provide information of the mitochondrial energy generating system (MEGS) in toto, and moreover, on the functioning of individual enzymes in the context of the intact mitochondrion. Several types of assays have been developed and are described below. For all these assays, a crucial factor is the integrity of the mitochondria, since the generation of ATP is dependent on an inner mitochondrial membrane potential. Therefore, it is important to include control experiments that test the coupling of the ATP synthesis to the respiratory chain, as this will provide information on the integrity of the inner mitochondrial membrane.

Two types of ATP production assays can be discriminated. The first type is performed in cultured cells permeabilized by digitonin (40), or in mitochondria-enriched fractions from tissue homogenates (41). Different substrates can be added to monitor different metabolic routes that lead to mitochondrial ATP synthesis. For example, by addition of pyruvate and malate, pyruvate enters the mitochondria via a pyruvate carrier and is subsequently metabolized by pyruvate dehydrogenase to acetyl-CoA that is further metabolized in the citric acid cycle where it is coupled to oxalate that is the product of malate dehydrogenase. As the ATP production rate is under the control of the ADP/ATP ratio, an excess of ADP should be present in the assay. In this way, the pathway from the pyruvate carrier to ATP can be monitored. Any primary defect in this pathway will result in a decreased ATP production rate. Therefore, this type of assay is very suitable as a diagnostic tool to screen for a defect in the MEGS. The second type of ATP production assay is restricted to cultured cells. In this assay, luciferase is used as an ATP sensor in vivo. Luciferase, an enzyme derived from the firefly, converts its substrate luciferin under the emission of light. This reaction requires ATP, and thus, the amount of light produced is a measure for the amount of ATP. Cells can be stably transfected with an expression vector encoding luciferase (42), infected with a virus that encodes luciferase (43), or microinjected with plasmid DNA encoding luciferase (44). By addition of an appropriate targeting sequence, luciferase can be intramitochondrial ATP. The ATP levels can be monitored by luminometry. A more sophisticated aproach is to determine subcellular (e.g. intramitochondrial) ATP levels by a microscope coupled to a CCD camera. When cells are placed in a flow cell under the microscope/CCD camera system, they can be exposed to different stimuli that regulate ATP production, in particular hormones that lead to intracellular Ca2+ fluxes, which stimulate mitochondrial ATP production (43). This can be monitored real-time by means of the camera. In contrast to the first type of ATP assay, this real-time ATP assay is not only dependent on the integrity of the mitochondrial ATP generating machinery, but also on the intracellular mechanisms that regulate ATP production. This set-up is very suitable to perform functional studies of the mitochondrial energy generating system in vivo and the factors that affect the activity of this system, but is less suitable as a diagnostic tool.

Substrate oxidation rate measurements

Substrate oxidation rate measurements provide detailed information on the functioning of the MEGS (41). The MEGS performs three decarboxylation reactions, at the level of PDHc, isocitrate dehydrogenase, and 2-oxoglutarate dehydrogenase. By using radiolabeled substrates in which the radiolabel is present at a carboxyl residue, the activity of the MEGS can be followed by measuring the amount of released radiolabeled CO2. Usually, combinations of radiolabeled substrates and unlabeled co-substrates are used, either in the presence or absence of specific inhibitors. An example is the use of radiolabeled pyruvate, which is decarboxylated by PDHc. To force the reaction to proceed at maximum rate, the reaction contains an excess of ADP, in order to maintain a high ADP/ATP ratio. In addition, the acetyl-CoA formed by PDHc has to be removed by addition of an appropriate co-substrate. When carnitine is used as cosubstrate, it will be coupled to the acetyl group of acetyl-CoA by carnitine-acetyl transferase. When malate is used as a co-substrate, this will be converted to oxalate in the TCA cycle which is subsequently coupled to the acetylgroup of acetyl-CoA by citrate synthase. When these assays are performed in a control sample, e.g. a muscle sample from a healthy individual, the ratio of the pyruvate oxidation in the presence of carnitine or malate will be approximately 1. Also in a sample from a PDHc deficient patient, this ratio will be near 1, however, in that case both reaction rates will be equally reduced due to the PDHc defect. Interestingly, in case of a respiratory chain defect, the ratio between these two reactions will be around 2 in favour of the reaction with carnitine (41). As this latter reaction does not directly involve a TCA cycle enzyme, an unfavourable NADH/NAD+ ratio will have much less effect on the PDHc activity when carnitine is used as a co-substrate compared to the reaction with malate as a co-substrate. As a final example, in case of a complex V defect, the reaction of pyruvate + malate will have a lower rate than in a control sample. Addition of the uncoupler CCCP will result in a normalization of the reaction rate, a phenomenon that is also observed in case of a defect in the phosphate carrier or the ATP:ADP antiporter ANT.

Oxygen consumption assays

In principle, oxygen consumption rate assays can be used for the same purposes as substrate oxidation rate assays. Also in this case, the pathway from the substrate of choice down to molecular oxygen conversion by complex IV can be evaluated, and even the steps beyond complex IV that are of influence on the activity of complex IV, such as complex V (45). The assay can be performed either in cellular or tissue extracts or in whole cells in vivo or ex vivo. The classical way to detect molecular oxygen is electrochemically by using a Clark-type oxygen electrode (45). More recently, molecular probes have been developed that made it possible to perform oxygen consumption rate measurements by fluorimetry (46, 47). This latter type of assay has the advantage that small volumes can be tested in 96-well plates using relatively simple laboratory equipment. By using appropriate combinations of substrates, the oxygen consumption rate measurements are suitable to locate primary defects in the MEGS, in a similar manner as with radiochemical substrate oxidation rate assays.

Diagnostic Biochemical Analysis of the Mitochondrial Energy Generating System

The mitochondrial energy generating system requires efficient interplay between a large number of different proteins and protein complexes, which by themselves can be made up of large numbers of individual subunits that have to be assembled in an ordered manner. This complexity of the mitochondrial energy generating system is probably one of the main reasons for the heterogeneity of mitochondrial disorders. Due to this clinical diversity, establishing the diagnosis “mitochondrial disorder” usually requires a combination of clinical, chemical, biochemical, and genetic examination of the patient suspected for a mitochondrial disorder (48-50). The biochemical analysis of a muscle biopsy is the corner stone of the diagnostic examination for mitochondrial disorders. The results will show whether or not the MEGS functions properly in this tissue. Unfortunately, the number of (potentially) mitochondrial disease causing candidate genes is very large, and molecular genetic techniques to rapidly sequence hundreds of candidate genes in a diagnostic setting are not yet available. The biochemical results are not only diagnostic in their own right, but, in combination with the clinical features of the patient, also provide important clues that are used to select candidate genes for molecular genetic analysis. Nevertheless, the diagnosis mitochondrial disorder can not always be made at the molecular genetic level, in particular in those cases in which a comprehensive biochemical analysis has not been performed.

The biochemical diagnostic analysis of a patient suspected for a mitochondrial disorder is usually performed on a muscle biopsy, as this tissue has a very high energy demand and is more likely to exhibit signs of mitochondrial dysfunctioning than tissues with a relatively low energy conversion rate. Depending on the clinical features, it could be considered to examine other types of tissue, such as liver or heart. It has been shown that in mitochondrial depletion syndromes with liver involvement, e.g. due to mutations in DGUOK or MPV17, muscle tissue may not always show biochemical abberations while liver shows clear signs of enzyme deficiencies (51, 52). Obvious drawbacks of organ or muscle biopsies are that an invasive procedure is required to obtain the tissue sample. Technically, it is possible to measure in fibroblasts or even blood samples, but these do not always express the mitochondrial defect. Nevertheless, these types of samples do have a very important added value to the biochemical diagnosis. First of all, a positive or a negative fibroblast result in combination with a positive muscle result has consequences for the selection of candidate genes for subsequent molecular genetic analysis. For example, in case of a mitochondrial depletion syndrome due to mutations in the POLG gene, very severe enzyme deficiencies can be observed in muscle and/or liver, whereas fibroblasts often show normal enzyme activities (53). By contrast, in case of a mitochondrial translation defect, respiratory chain enzyme deficiencies are observed both in muscle and in fibroblasts. A second important aspect of fibroblast (or lymphocyte) analysis is that a systemic expression of a biochemical defect may allow for prenatal diagnosis on the basis of biochemical analysis of chorionic tissue or amniocytes in families of mitochondrial patients in which the underlying molecular genetic defect has not (yet) been identified (54).


1. Bogaerts V, Theuns J, and van Broeckhoven C. Genetic findings in Parkinson’s disease and translation into treatment: a leading role for mitochondria? Genes Brain Behav. 2007; (Epub).

2. Rabol R, Bushel R, Dela, F. Mitochondrial oxidative function and type 2 diabetes. Appl. Physiol. Nutr. Metab. 2006; 31:675-683.

3. Modica-Napolitano JS, Kulawiec M, Singh KK. Mitochondria and human cancer. Curr. Mol. Med. 2007; 7:121-131.

4. Smeitink J, van den Heuvel L, Dimauro S. The genetics and pathology of oxidative phosphorylation. Nat. Rev. Genet. 2001; 2:342-352.

5. Brivet M, Garcia-Cazorla A, Lyonnet S, Dumez Y, Nassogne MC, Slama A, Boutron A, Touati G, Legrand A, Saudubray JM. Impaired mitochondrial pyruvate importation in a patient and a fetus at risk. Mol. Genet. Metab. 2003; 78:186-192.

6. Smolle M, Prior AE, Brown AE, Cooper A, Byron O, Lindsay JG. A new level of architectural complexity in the human pyruvate dehydrogenase complex. J. Biol. Chem. 2006; 281:19772-19780.

7. Ling M, McEachern G, Seyda A, MacKay N, Scherer SW, Bratinova S, Beatty B, Giovannucci-Uzielli ML, Robinson BH. Detection of a homozygous four base pair deletion in the protein X gene in a case of pyruvate dehydrogenase complex deficiency. Hum. Mol. Genet. 1998; 7:501-505.

8. Huang B, Gudi R, Wu P, Harris RA, Hamilton J, Popov KM. Isoenzymes of pyruvate dehydrogenase phosphatase. DNA-derived amino acid sequences, expression, and regulation. J. Biol. Chem. 1998; 273:17680-17688.

9. Janssen RJ, Nijtmans LG, van den Heuvel LP, Smeitink JA. Mitochondrial complex I: structure, function and pathology. J. Inherit. Metab Dis. 2006; 29:499-515.

10. Gellera C, Uziel G, Rimoldi M, Zeviani M, Laverda A, Carrara F, DiDonato S. Fumarase deficiency is an autosomal recessive encephalopathy affecting both the mitochondrial and the cytosolic enzymes. Neurology 1990; 40:495-499.

11. Bonnefont JP, Chretien D, Rustin P, Robinson B, Vassault A, Aupetit J, Charpentier C, Rabier D, Saudubray JM, Munnich A. Alpha-ketoglutarate dehydrogenase deficiency presenting as congenital lactic acidosis. J. Pediatr. 1992; 121:255-258.

12. Elpeleg O, Miller C, Hershkovitz E, Bitner-Glindzicz M, Bondi-Rubinstein G, Rahman S, Pagnamenta A, Eshhar S, Saada A. Deficiency of the ADP-forming succinyl-CoA synthase activity is associated with encephalomyopathy and mitochondrial DNA depletion. Am. J. Hum. Genet. 2005; 76:1081-1086.

13. Ostergaard E, Christensen E, Kristensen E, Mogensen B, Duno M, Shoubridge EA, Wibrand F. Deficiency of the alpha Subunit of Succinate-Coenzyme A Ligase Causes Fatal Infantile Lactic Acidosis with Mitochondrial DNA Depletion. Am. J. Hum. Genet. 2007; 81:383-387.

14. Briere JJ, Favier J, El Ghouzzi V, Djouadi F, Benit P, Gimenez AP, Rustin P. Succinate dehydrogenase deficiency in human. Cell. Mol. Life Sci. 2005; 62:2317-2324.

15. www.mitomap. org.

16. Pecina P, Houstkova H, Hansikova H, Zeman J, Houstek J. Genetic defects of cytochrome c oxidase assembly. Physiol. Res. 2004; 53:213-223.

17. Houstek J, Pickova A, Vojtiskova A, Mracek T, Pecina P, Jesina P. Mitochondrial diseases and genetic defects of ATP synthase. Biochim. Biophys. Acta 2006; 1757:1400-1405.

18. Mayr JA, Merkel O, Kohlwein SD, Gebhardt BR, Bohles H, Fotschl U, Koch J, Jaksch M, Lochmuller H, Horvath R, Freisinger P, Sperl W. Mitochondrial phosphate-carrier deficiency: a novel disorder of oxidative phosphorylation. Am. J. Hum. Genet. 2007; 80:478-484.

19. Taylor RW, Turnbull DM. Mitochondrial DNA mutations in human disease. Nat. Rev. Genet. 2005; 6:389-402.

20. Carrozzo R, Dionisi-Vici C, Steuerwald U, Lucioli S, Deodato F, Di Giandomenico S, Bertini E, Franke B, Kluijtmans LA, Meschini MC, Rizzo C, Piemonte F, Rodenburg R, Santer R, Santorelli FM, van Rooij A, Vermunt-de Koning D, Morava E, Wevers RA. SUCLA2 mutations are associated with mild methylmalonic aciduria, Leigh-like encephalomyopathy, dystonia and deafness. Brain 2007; 130:862-874.

21. Jacobs HT, Turnbull DM. Nuclear genes and mitochondrial translation: a new class of genetic disease. Trends Genet. 2005; 21:312-314.

22. Spinazzola A, Zeviani M. Disorders of nuclear-mitochondrial intergenomic communication. Biosci. Rep. 2007; 27:39-51.

23. Dimauro S, Quinzii CM, Hirano M. Mutations in coenzyme Q10 biosynthetic genes. J. Clin. Invest. 2007; 117:587-589.

24. Schagger H, von Jagow G. Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal. Biochem. 1991; 199:223-231.

25. Nijtmans LG, Henderson NS, Holt IJ. Blue Native electrophoresis to study mitochondrial and other protein complexes. Methods 2002; 26:327-334.

26. Carroll J, Fearnley IM, Skehel JM, Shannon RJ, Hirst J, Walker JE. Bovine complex I is a complex of 45 different subunits. J. Biol. Chem. 2006; 281:32724-32727.

27. Vogel RO, Janssen RJ, van den Brand MA, Dieteren CE, Verkaart S, Koopman WJ, Willems PH, Pluk W, van den Heuvel LP, Smeitink JA, Nijtmans LG. Cytosolic signaling protein Ecsit also localizes to mitochondria where it interacts with chaperone NDUFAF1 and functions in complex I assembly. Genes Dev. 2007; 21:615-624.

28. Kopp E, Medzhitov R, Carothers J, Xiao C, Douglas I, Janeway CA, Ghosh S. ECSIT is an evolutionarily conserved intermediate in the Toll/IL-1 signal transduction pathway. Genes Dev. 1999; 13:2059-2071.

29. Murray J, Schilling B, Row RH, Yoo CB, Gibson BW, Marusich MF, Capaldi RA. Small scale immunopurification of cytochrome c oxidase for a high throughput, multiplexing analysis of enzyme activity, and amount. Biotechnol. Appl. Biochem. 2007; 48:167-178.

30. Zerbetto E, Vergani L, Dabbeni-Sala F. Quantification of muscle mitochondrial oxidative phosphorylation enzymes via histochem- ical staining of blue native polyacrylamide gels. Electrophoresis 1997; 18:2059-2064.

31. Fischer JC, Ruitenbeek W, Trijbels JM, Veerkamp JH, Stadhouders AM, Sengers RC, Janssen AJ. Estimation of NADH oxidation in human skeletal muscle mitochondria. Clin. Chim. Acta 1986; 155:263-273.

32. Janssen AJ, Trijbels FJ, Sengers RC, Smeitink JA, van den Heuvel LP, Wintjes LT, Stoltenborg-Hogenkamp BJ, Rodenburg RJ. Spectrophotometric assay for complex I of the respiratory chain in tissue samples and cultured fibroblasts. Clin. Chem. 2007; 53:729-734.

33. Kirby DM, Thorburn DR, Turnbull DM, Taylor RW. Biochemical assays of respiratory chain complex activity. Methods Cell Biol. 2007; 80:93-119.

34. Reisch AS, Elpeleg O. Biochemical assays for mitochondrial activity: assays of TCA cycle enzymes and PDHc. Methods Cell Biol. 2007; 80:199-222.

35. Rustin P, Chretien D, Bourgeron T, Gerard B, Rotig A, Saudubray JM, Munnich A. Biochemical and molecular investigations in respiratory chain deficiencies. Clin. Chim. Acta 1994; 228:35-51.

36. Lopez LC, Schuelke M, Quinzii CM, Kanki T, Rodenburg RJ, Naini A, Dimauro S, Hirano M. Leigh syndrome with nephropathy and CoQ10 deficiency due to decaprenyl diphosphate synthase subunit 2 (PDSS2) mutations. Am. J. Hum. Genet. 2006; 79:1125-1129.

37. Sterk JP, Stanley WC, Hoppel CL, Kerner J. A radiochemical pyruvate dehydrogenase assay: activity in heart. Anal. Biochem. 2003; 313:179-182.

38. Bonnefont JP, Chretien D, Rustin P, Robinson B, Vassault A, Aupetit J, Charpentier C, Rabier D, Saudubray JM, Munnich A. Alpha-ketoglutarate dehydrogenase deficiency presenting as congenital lactic acidosis. J. Pediatr. 1992; 121:255-258.

39. Zinn AB, Kerr DS, Hoppel CL. Fumarase deficiency: a new cause of mitochondrial encephalomyopathy. N. Engl. J. Med. 1986; 315:469-475.

40. Wanders RJ, Ruiter JP, Wijburg FA, Zeman J, Klement P, Houstek J. Prenatal diagnosis of systemic disorders of the respiratory chain in cultured chorionic villus fibroblasts by study of ATP-synthesis in digitonin-permeabilized cells. J. Inherit. Metab Dis. 1996; 19:133-136.

41. Janssen AJ, Trijbels FJ, Sengers RC, Wintjes LT, Ruitenbeek W, Smeitink JA, Morava E, van Engelen BG, van den Heuvel LP, Rodenburg RJ. Measurement of the energy-generating capacity of human muscle mitochondria: diagnostic procedure and application to human pathology. Clin. Chem. 2006; 52:860-871.

42. Gajewski CD, Yang L, Schon EA, Manfredi G. New insights into the bioenergetics of mitochondrial disorders using intracellular ATP reporters. Mol. Biol. Cell. 2003; 14:3628-3635.

43. Visch HJ, Rutter GA, Koopman WJ, Koenderink JB, Verkaart S, de Groot T, Varadi A, Mitchell KJ, van den Heuvel LP, Smeitink JA, Willems PH. Inhibition of mitochondrial Na+ -Ca2+ exchange restores agonist-induced ATP production and Ca2+ handling in human complex I deficiency. J. Biol. Chem. 2004; 279:40328-40336.

44. Kennedy HJ, Pouli AE, Ainscow EK, Jouaville LS, Rizzuto R, Rutter GA. Glucose generates sub-plasma membrane ATP microdomains in single islet beta-cells. Potential role for strategically located mitochondria. J. Biol. Chem. 1999; 274:13281-13291.

45. Hofhaus G, Shakeley RM, Attardi G. Use of polarography to detect respiration defects in cell cultures. Methods Enzymol. 1996; 264:476-483.

46. Hynes J, Floyd S, Soini AE, O’Connor R, Papkovsky DB. Fluorescence-based cell viability screening assays using water-soluble oxygen probes. J. Biomol. Screen. 2003; 8:264-272.

47. Will Y, Hynes J, Ogurtsov VI, Papkovsky DB. Analysis of mitochondrial function using phosphorescent oxygen-sensitive probes. Nat. Protoc. 2006; 1:2563-2572.

48. Bernier FP, Boneh A, Dennett X, Chow CW, Cleary MA, Thorburn DR. Diagnostic criteria for respiratory chain disorders in adults and children. Neurology 2002; 59:1406-1411.

49. Morava E, van den Heuvel L, Hol F, de Vries MC, Hogeveen M, Rodenburg RJ, Smeitink JA. Mitochondrial disease criteria: diagnostic applications in children. Neurology 2006; 67:1823-1826.

50. Wolf NI, Smeitink JA. Mitochondrial disorders: a proposal for consensus diagnostic criteria in infants and children. Neurology 2002; 59:1402-1405.

51. Mandel H, Szargel R, Labay V, Elpeleg O, Saada A, Shalata A, Anbinder Y, Berkowitz D, Hartman C, Barak M, Eriksson S, Cohen N. The deoxyguanosine kinase gene is mutated in individuals with depleted hepatocerebral mitochondrial DNA. Nat. Genet. 2001; 29:337-341.

52. Spinazzola A, Viscomi C, Fernandez-Vizarra E, Carrara F, D’Adamo P, Calvo S, Marsano RM, Donnini C, Weiher H, Strisciuglio P, Parini R, Sarzi E, Chan A, Dimauro S, Rotig A, Gasparini P, Ferrero I, Mootha VK, Tiranti V, Zeviani M. MPV17 encodes an inner mitochondrial membrane protein and is mutated in infantile hepatic mitochondrial DNA depletion. Nat. Genet. 2006; 38:570-575.

53. de Vries MC, Rodenburg RJ, Morava E, van Kaauwen EP, Ter Laak H, Mullaart RA, Snoeck IN, van Hasselt PM, Harding P, van den Heuvel LP, Smeitink JA. Multiple oxidative phosphorylation deficiencies in severe childhood multi-system disorders due to polymerase gamma (POLG1) mutations. Eur. J. Pediatr. 2007; 166:229-234.

54. Niers L, van den Heuvel L, Trijbels F, Sengers R, Smeitink J. Prerequisites and strategies for prenatal diagnosis of respiratory chain deficiency in chorionic villi. J. Inherit. Metab Dis. 2003; 26:647-658.

Further Reading

Oxidative Phosphorylation in Health and Disease. Edited by: Jan Smeitink, Rob C.A. Sengers and J.M. Frans Trijbels. ISBN: 0-30648232-0.

Mitochondria, 2nd edition. Methods in cell biology, vol 80. Edited by: Liza A. Pon and Eric A. Schon. ISBN: 0-12-544173-8.

Neuromuscular Disease Center website:

GeneReview of mitochondrial disorders website:

Uppsala University Human Mitochondrial Genome Database:

See Also

Electron Transfer Chain, Chemistry of

Metabolic Diseases, Chemical Biology of

Mitochondria, Structural Dynamics of

Mitochondrial Proteomics

Oxidative Metabolism, Chemistry of