A spectrophotometric coupled enzyme assay to measure the activity of succinate dehydrogenase

Accepted Manuscript A spectrophotometric, coupled enzyme assay to measure the activity of succi‐ nate dehydrogenase Andrew J.Y. Jones, Judy Hirst PII: DOI: Reference:

S0003-2697(13)00336-9 http://dx.doi.org/10.1016/j.ab.2013.07.018 YABIO 11421

To appear in:

Analytical Biochemistry

Received Date: Revised Date: Accepted Date:

4 June 2013 9 July 2013 12 July 2013

Please cite this article as: A.J.Y. Jones, J. Hirst, A spectrophotometric, coupled enzyme assay to measure the activity of succinate dehydrogenase, Analytical Biochemistry (2013), doi: http://dx.doi.org/10.1016/j.ab.2013.07.018

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A spectrophotometric, coupled enzyme assay to measure the activity of succinate dehydrogenase† Andrew J. Y. Jones and Judy Hirst* The Medical Research Council Mitochondrial Biology Unit, Wellcome Trust / MRC Building, Hills Road, Cambridge, CB2 0XY, U. K.

†This research was funded by The Medical Research Council.

*Author to whom correspondence should be addressed: Medical Research Council Mitochondrial Biology Unit, Wellcome Trust / MRC Building, Hills Road, Cambridge, CB2 0XY, U. K. Tel : +44 1223 252810, Fax : +44 1223 252815, E-mail : [email protected]

Short title: Coupled enzyme assay for succinate dehydrogenase Subject category: Enzymatic assays and analyses

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Abstract Respiratory complex II (succinate:ubiquinone oxidoreductase) connects the tricarboxylic acid cycle to the electron transport chain in mitochondria and many prokaryotes. Complex II mutations have been linked to neurodegenerative diseases and metabolic defects in cancer. However, there is no convenient stoichiometric assay for the catalytic activity of complex II. Here, we present a simple, quantitative, real-time method to detect the production of fumarate from succinate by complex II that is easy to implement and applicable to the isolated enzyme, membrane preparations, and tissue homogenates. Our assay uses fumarate hydratase to convert fumarate to malate, and oxaloacetate decarboxylating malic dehydrogenase to convert malate to pyruvate and NADP+ to NADPH; the NADPH is detected spectrometrically. Simple protocols for the high-yield production of the two enzymes required are described; oxaloacetate decarboxylating malic dehydrogenase is also suitable for accurate determination of the activity of fumarate hydratase.

Unlike existing spectrometric assay methods for

complex II that rely on artificial electron acceptors (such as 2,6-dichlorophenolindophenol) our coupled assay is specific and stoichiometric (1:1 for succinate oxidation to NADPH formation), so it is suitable for comprehensive analyses of the catalysis and inhibition of succinate dehydrogenase activities in samples with both simple and complex compositions.

Keywords: Coupled assay Fumarate hydratase Oxaloacetate decarboxylating malic enzyme Succinate:ubiquinone oxidoreductase

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Introductory statement In mammalian mitochondria, succinate:ubiquinone oxidoreductase (succinate dehydrogenase, complex II, EC 1.3.5.1) catalyzes the oxidation of succinate to fumarate in the matrix, and the reduction of ubiquinone-10 in the inner membrane (1, 2). Thus, it links the tricarboxylic acid cycle to the respiratory electron transport chain. Complex II comprises four subunits (1-3). In the hydrophilic domain, the largest subunit houses a covalently-bound flavin adenine dinucleotide (FAD) cofactor and the succinate-binding site, and a smaller subunit contains three iron-sulfur clusters that transfer electrons from the flavin to ubiquinone. The binding site for the ubiquinone substrate is located close to a heme cofactor, in the membraneassociated domain that is composed of two hydrophobic subunits. Homologues of mammalian complex II are widely conserved in aerobic organisms, in both eukaryotes and prokaryotes. Some prokaryotes use fumarate as a terminal electron acceptor in anaerobic respiration, by using enzymes that are closely-related to complex II to catalyze the reverse reaction, reduction of fumarate to succinate (4). Some parasites, such as Ascaris suum, switch between succinate oxidation and fumarate reduction during different life stages, and the same switch has been proposed to occur in mammalian cells during hypoxia (5).

Mutations in complex II have been identified as a cause of Leigh syndrome, a

neurodegenerative disease (6), and complex II has recently been identified as a significant source of mitochondrial reactive oxygen species production (7). Complex II mutations have been linked also to hereditary paragangliomas, tumors in the head and neck (8), and there is currently significant interest in how mitochondrial succinate and fumarate levels are linked to the stabilization of HIF-1 and thus to cell proliferation (9). Due to its central role in metabolism and medicine, measurements of complex II activity have long been used to assess the activity of both the enzyme itself and of the mitochondrial electron transport chain as a whole. However, there is no convenient and

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reliable method for testing complex II activity. Clark oxygen electrodes are often used (10) (see for example (7, 11)) but they are expensive in their sample requirements, and as they measure succinate:O2 oxidoreduction they can only assess the combined activity of respiratory complexes II, III and IV. Alternative real-time assays of complex II rely on artificial electron acceptors which change color upon reduction and can therefore be monitored spectrophotometrically. 2,6-dichlorophenolindophenol (DCPIP)1 and 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyl tetrazolium chloride (INT) are reduced directly by complex II, and also by the quinol product of complex II catalysis (see, for example (12-14)). DCPIP and other tetrazolium salts (which do not react efficiently with complex II directly) have also been used as secondary electron acceptors, with phenazine methosulfate (PMS) to mediate electron transfer from complex II (see, for example, (12, 14, 15)). However, all of these assays lack specificity. When either the electron acceptor or PMS react with the enzyme directly they fail to focus on the complete catalytic cycle of succinate:ubiquinone oxidoreduction, the reactivity is not specific to complex II, and O2 interferes with the results (16). On the other hand, the reaction with ubiquinol is inefficient, electrons are not scavenged stoichiometrically when the respiratory chain is turning over, and the measured rate is always a combination from all possible pathways. Here, we present a convenient, stoichiometric and precise coupled enzyme assay for succinate oxidation (see Scheme 1).

Our assay uses two enzymes: fumarate hydratase

(FumC, EC 4.2.1.2) converts the fumarate product of succinate oxidation to malate (17), and then oxaloacetate decarboxylating malic dehydrogenase (MaeB, EC 1.1.1.40) couples the conversion of malate to pyruvate to the reduction of NADP+ to NADPH (18); the NADPH is detected spectrophotometrically. Because our assay quantifies succinate oxidation directly it is suitable for measuring succinate:ubiquinone oxidoreduction by isolated complex II, as well as succinate consumption in membrane and tissue preparations.

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Materials and Methods Preparation of FumC and MaeB Fumarate hydratase (FumC) and oxaloacetate decarboxylating malic dehydrogenase from Escherichia coli (MaeB) were over-expressed in E. coli. Plasmid constructs encoding the fumC and maeB genes were supplied by Professor Todd Weaver (University of Wisconsin, La Crosse) (17) and Professor María Drincovich (National University of Rosario, Argentina) (18), respectively. The same protocol, adapted from refs. (17) and (18), was used for expression and purification of both proteins unless otherwise stated. BL21(DE3) E. coli cells (New England Biolabs) were transformed with the maeB or fumC plasmid. Cells were grown at 32 °C in LB media supplemented with 50 µg mL-1 ampicillin until A600 reached ~0.6, then protein expression was induced using isopropyl -D-1-thiogalactopyranoside (IPTG, 1 mM for FumC and 0.1 mM for MaeB) for 18 hours at 20 °C.

Cells were harvested by

centrifugation (10 mins., 3500 × g), resuspended in buffer A (50 mM potassium phosphate, 300 mM NaCl, pH 7.8 at 4 °C) for FumC or buffer B (20 mM Tris-Cl, 100 mM NaCl, 25 mM imidazole and 10 % w/v glycerol, pH 7.4 at 4 °C) for MaeB, then lyzed in one passage through a Constant Systems Ltd. model B 2.2 kW Z cell disruptor at 30 PSI. The lysates were centrifuged (45 mins., 150,000 × g), the pellets discarded, and the supernatants filtered (0.45 µm) before being loaded onto a pre-equilibrated 25 mL Ni-sepharose 6 fast flow column (GE Healthcare). The column was washed using buffer A + 60 mM imidazole for FumC or buffer B for MaeB, then the required proteins eluted in buffer A + 400 mM imidazole for FumC or buffer B + 300 mM imidazole for MaeB. Fractions were identified by activity assays, pooled and concentrated (Amicon Ultra-15 50 kDa centrifugal filter units), then dialyzed overnight in buffer C (10 mM Tris-SO4, 5 mM EDTA, 1 mM dithiothreitol, pH 7 at 4 °C) for FumC or buffer D (60 mM Tris-SO4, 20 mM β-mercaptoethanol, 25 mM imidazole and 10 % w/v

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glycerol, pH 7.4 at 4 °C) for MaeB. Typical yields were 36 and 60 mg per liter of culture for FumC and MaeB, respectively.

Preparation of complex II-containing samples Submitochondrial particles (SMPs) were prepared from bovine heart mitochondria as described previously (11). Complex II was solubilized from bovine heart mitochondrial membranes (~10 mg protein mL-1 in 10 mM Tris-SO4 pH 7.4, 250 mM sucrose, 1 mM EDTA, 0.005% PMSF) by the addition of 0.5% n-dodecyl-β-D-maltopyranoside (DDM); the sample was stirred on ice for 30 mins., then centrifuged (30 mins., 48000 x g) and the supernatant collected. Complex I was prepared from bovine heart mitochondria as described previously (19). Tissue samples were prepared from rat skeletal muscle using a procedure modified from ref. (20). Briefly, the leg muscle was stripped of fat and connective tissue and chopped finely (<1 mm pieces), then homogenized in 15 volumes of buffer (320 mM sucrose, 10 mM TrisHCl, pH 7 at 4 °C) using a tight fitting glass Teflon homogenizer.

The sample was

centrifuged at 1000 x g for 5 mins. at 4 °C and the supernatant collected.

Kinetic activity measurements All assays were carried out in 10 mM Tris-SO4 (pH 7.4 at 32 °C), 250 mM sucrose, 2 mM MgSO4 and 1 mM K2SO4 at 32 ˚C, unless otherwise stated. 20 µg mL-1 gramicidin was included to dissipate the proton motive force in SMPs. Standard concentrations were 5 mM succinate, 2 mM NADP+, 60 µg mL-1 FumC and 300 µg mL-1 MaeB. NADPH concentration was followed at 340-380 nm (ε = 4.81 mM-1 cm-1) using a Molecular Devices Spectramax Plus 384 platereader. The reduction of DCPIP (100 µM, 600 nm (ε = 21 mM-1 cm-1, blue to colorless) (14) and INT (100 µM, 500 nm (ε = 19 mM-1 cm-1, colorless to red) (13) were measured in the presence and absence of 100 µM decylubiquinone, 5 mM succinate, or 100

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µM NADH. When required, ubiquinol oxidation by the respiratory chain was inhibited by 400 µM NaCN, and atpenin A5 (Santa Cruz Biotechnology Inc.) was added from a concentrated stock solution in DMSO. O2 consumption by SMPs was measured using a Clark electrode in a stirred 1 mL perspex cell held at 32 °C (Rank Brothers Ltd, Cambridge).

Results and Discussion Kinetic characterization of FumC and MaeB Figure 1 shows how the rates of catalysis by the FumC and MaeB enzymes used here depend on the concentrations of fumarate, and malate and NADP+, respectively. First, the assay buffer used for all experiments contained 2 mM Mg2+ because MaeB requires Mg2+ for catalysis (18); as expected, no catalysis was observed in the absence of Mg2+, and concentrations above 4 mM have been shown previously to inhibit (18). 2 mM K+ was also included in the assay buffer, as it was shown previously to activate MaeB by ~30% (18). The rate of MaeB catalysis is easily measured by the accumulation of NADPH at 340-380 nm; Figure 1B shows that the apparent KM for malate is relatively high (7.3 ± 0.9 mM) and the Vmax is poor (8.37 ± 0.08 mol min-1 mg-1 or 92 ± 0.93 s-1) and Figure 1C shows that for NADP+ the KM is lower (52 ± 2.6 µM). Both KM values were measured with the second substrate at high enough concentration for Vmax. Our values are comparable to those reported previously (KM(malate) = 3.4 mM, KM(NADP+) = 41.5 µM, Vmax = 67 s-1) (18). Figure 1A shows how the rate of conversion of fumarate to pyruvate, in the presence of FumC and MaeB, depends on the concentration of fumarate present; the KM for fumarate is 64 ± 7.8 µM and Vmax is 293 ± 7 mol min-1 mg-1 or 975 ± 21.7 s-1. The values are similar to values reported previously (207 µM and 1149 s-1) (21). For these measurements the concentration of MaeB was set to 300-600 µg mL-1 (0.45-0.90 µM) to overcome its poor Vmax value; increasing the MaeB concentration further did not increase the observed rate of catalysis (see below also). 7

The concentration of NADP+ was set to 2 mM, ~20 times higher than KM. Along with the release of CO2, the high NADP+/NADPH ratio precludes the reverse reaction, conversion of pyruvate to malate, which has been demonstrated to occur slowly under some conditions (18). The lower KM and higher Vmax values observed for FumC allowed it to be used at lower concentrations, 60-120 µg mL-1 (0.3-0.6 µM), than MaeB in subsequent experiments (see below also). Previously, the decarboxylating malic enzyme from pigeon liver (equivalent to MaeB) was used in an assay for fumarate hydratase activity (22). Similar assays have been used recently in studies of fumarate hydratase deficiencies (see, for example, (23)). However, commercially-available decarboxylating malic enzymes (EC 1.1.1.38, .39 or .40) are prohibitively expensive, precluding them being applied extensively in kinetic studies. Alternatively, malate dehydrogenases (EC 1.1.1.37 or .82) are much cheaper, but they catalyze reversibly to reach an equilibrium position and thus can only provide a comparative evaluation of enzyme activities. The MaeB preparation described here compares favorably on both criteria: it catalyses irreversibly, and can be prepared easily and cheaply in large amounts.

Measurement of the rate of succinate oxidation by submitochondrial particles Figure 2A shows a typical assay trace recorded to follow succinate:O2 oxidoreduction by respiratory complexes II, III and IV in SMPs. The rate of NADPH formation is not constant throughout the assay: it begins slowly then speeds up to reach a constant value after approximately 100 s. During the assay ‘lag phase’ the intermediate fumarate and malate concentrations accumulate to their steady-state values. Thus, the lag phase can be eliminated by increasing the concentrations of FumC and MaeB. However, the final rate measurement is unchanged, and because of the increased enzyme requirements we consider this unnecessary.

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To confirm the specificity of the coupled assay for the conversion of succinate to fumarate, rates of NADPH formation were measured in the absence of various assay components. In the absence of SMPs, succinate, MaeB, or NADP+ the observed rates were negligible, but in the absence of FumC a significant rate, ~10% of the control, was observed. Based on the other control experiments we conclude that this results from a low level of endogenous fumarate hydratase in the SMP preparation ⎯ which does not affect the assay results because it merely reduces the load on the exogenous FumC. Figure 2B shows that our standard working concentrations of MaeB (300 µg mL-1) and FumC (60 µg mL-1) are adequate for the quantitative detection of succinate oxidation by SMPs (provided the lag phases are discarded), and Figure 2C shows that the rate of NADPH reduction by the detection system depends directly on the SMP concentration over the whole range tested. Therefore, the detection system is not rate limiting in any of these experiments, and the observed rates accurately define the rates of succinate oxidation.

Comparison of the coupled assay system with alternative methods Succinate oxidation by complex II in SMPs is coupled to the reduction of O2 by complex IV, via the electron transport chain, in a 2:1 succinate:O2 stoichiometry.

Therefore,

measurements of O2 consumption, using a Clark electrode, are the ‘gold standard’ for confirming the accuracy of any new assay. Figure 3A shows that measurements of the rate of succinate oxidation by SMPs using our coupled assay system agree very well with Clark electrode measurements. Note that our assay system can be applied to measurements on isolated complex II, whereas the Clark electrode can only be used when complex II catalysis is linked to O2 reduction by the respiratory chain. Our assay has the additional advantages of being a spectrophotometric method, with significantly lower sample requirements, improved convenience, and quicker and easier data accumulation.

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To investigate alternative methods which are applicable to isolated complex II, as well as to membrane and tissue preparations, we compared our coupled assay for succinate oxidation to an assay using DCPIP (12, 14) (see Table 1). The rate of succinate oxidation by detergent-solubilized bovine heart mitochondrial membranes was measured in the presence of a short-chain ubiquinone, decylubiquinone (the solubilization isolates complex II from the other enzymes and precludes significant turnover from endogenous ubiquinone-10). Using 30 µg mL-1 of solubilized membranes, 5 mM succinate and 100 µM decylubiquinone, the observed coupled-assay rate was 0.41 ± 0.023 µmol min-1 mg-1 (the rate was negligible in the absence of any of the components). With 100 µM DCPIP, a substantial rate of DCPIP reduction (0.062 ± 0.003 µmol min-1 mg-1) was observed even in the absence of decylubiquinone. A similar result (0.088 ± 0.005 µmol min-1 mg-1) was obtained when 100 µM NADH was added instead of succinate. Therefore, DCPIP is reduced directly by the respiratory chain enzymes without the participation of ubiquinone, so the assay does not address the complete complex II turnover cycle. When 100 µM decylubiquinone and 100 µM DCPIP were present (the DCPIP should be reduced by the decylubiquinol product), the rate of DCPIP reduction increased to 0.20 ± 0.007 µmol min-1 mg-1. This value is significantly less than the value from our coupled assay, even if the ubiquinone-independent rate is not subtracted. Furthermore, when the full respiratory chain is present DCPIP is unable to compete with complex III for ubiquinol, so complex IV must be inhibited to allow ubiquinone reduction to be measured. For example, when complex I NADH:ubiquinone oxidoreduction was measured with 30 µg mL -1 SMPs and 100 µM NADH, in the absence of a complex IV inhibitor, the rate of NADH oxidation measured directly was 0.38 ± 0.005 µmol min-1 mg-1, but the rate of DCPIP reduction was much lower, 0.18 ± 0.009 µmol min-1 mg-1. Rates of succinate oxidation by solubilized complex II determined using 100 µM INT (13) instead of DCPIP were similarly lower than the control value (see Table 1).

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Measurement of complex II activity in tissue samples To detect biochemical defects in complex II in patients with mitochondrial diseases, rates of complex II catalysis are typically measured in tissue homogenates from biopsy samples. Specifically, the rate of DCPIP reduction is monitored in the presence of succinate, and in the presence and absence of ubiquinone-1 (24). Therefore, measurements from our coupled assay were compared with DCPIP measurements on rat skeletal muscle homogenates (see Table 1). Our coupled assay, with 5 mM succinate, 50 µM ubiquinone-1, and 1.5 mM NaCN, gave a rate of 0.12 ± 0.006 µmol min-1 mg-1, that was fully sensitive to 2 µM atpenin A5 (a complex II inhibitor (14)). Using the same substrate concentrations the rate measured using 100 µM DCPIP was only 0.06 ± 0.001 µmol min-1 mg-1. In the absence of both ubiquinone-1 and NaCN the rates were 0.054 ± 0.002 µmol min-1 mg-1 (coupled assay) and 0.019 ± 0.001 µmol min-1 mg-1 (DCPIP). As expected, in the absence of ubiquinone-1 NaCN inhibited succinate oxidation in the coupled assay fully, but the DCPIP assay gave a rate of 0.018 ± 0.001 µmol min-1 mg-1. Typical rates reported using succinate, ubiquinone-1, NaCN and DCPIP in human tissue samples are 0.061 0.079 µmol min-1 mg-1 (24), comparable to the rate detected here in rat tissue by the same method. Thus, the same picture emerges for tissues samples as for solubilized membranes ⎯ that rates of succinate oxidation measured using DCPIP are a significant underestimate of the true rates. However, we are aware of the importance of standardized measurements for the comparison of values from tissue biopsy samples (24), and that continuing with established assay protocols may thus be preferable in this case.

Determination of kinetic parameters for succinate dehydrogenase catalysis The coupled assay was used to determine the KM and Vmax values for complex II in SMPs (see Figure 3A), and the IC50 values for two complex II inhibitors (Figures 3B and 3C). The KM value obtained, 410 ± 55 µM, is within the range of values reported previously (using DCPIP)

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of 130 µM (25), 303 µM (26), 1.3 mM (27) and 1.5 mM (28). By using the published value of 0.19 nmol of complex II per mg of protein in a mitochondrial inner membrane preparation (29) our Vmax of 1.2 ± 0.03 µmol min-1 mg-1 (although depending strongly on the assay conditions and composition), equates to a turnover number of 100 ± 2.7 s-1. Note that our value is similar to that reported previously from a Clark electrode measurement in 5 mM succinate (0.94 µmol min-1 mg-1 (30)); both values are higher than a previous value from this laboratory (0.264 µmol min-1 mg-1 (11)) due to improvements in mitochondria preparation. Figure 3B shows that atpenin A5, an inhibitor that bindings in the complex II ubiquinonebinding site, has an IC50 value of 2.4 ± 1.2 nM, in good agreement with the published value of 3.6 nM (14) measured using DCPIP in mitochondria. Figure 3C shows that malonate, an inhibitor that binds in the complex II flavin site, has an IC50 value of 96 ± 1.3 µM, in line with the values of 42 µM in rat mitochondria (31) and 180 210 µM in human cell lines (32) measured using DCPIP. Inhibition of complex II by the flavin site inhibitor oxaloacetate could not be quantified as it is an intermediate of the MaeB reaction and so interferes with our detection system.

Acknowledgements We thank Professor Todd Weaver (University of Wisconsin, La Crosse) and Professor María Drincovich (National University of Rosario, Argentina) for the fumC and maeB plasmids.

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Abbreviations footnote:

The abbreviations used are: DCPIP, 2,6-dichlorophenolindophenol; FumC, fumarate hydratase; INT, 2-(4-iodo-phenyl)-3(4-nitrophenyl)-5-phenyl tetrazolium chloride; MaeB, oxaloacetate decarboxylating malic dehydrogenase; PMS, phenazine methosulfate; SMP, submitochondrial particle.

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References 1.

B. A. C. Ackrell, Progress in understanding structure-function relationships in respiratory chain complex II, FEBS Lett. 466 (2000) 1-5.

2.

G. Cecchini, Function and structure of complex II of the respiratory chain, Annu. Rev. Biochem. 72 (2003) 77-109.

3.

F. Sun, X. Huo, Y. Zhai, A. Wang, J. Xu, D. Su, M. Bartlam & Z. Rao, Crystal structure of mitochondrial respiratory membrane protein complex II, Cell 121 (2005) 1043-57.

4.

J. J. van Hellemond & A. G. M. Tielens, Expression and functional properties of fumarate reductase, Biochem. J. 304 (1994) 321-31.

5.

C. Sakai, E. Tomitsuka, H. Esumi, S. Harada & K. Kita, Mitochondrial fumarate reductase as a target of chemotherapy: from parasites to cancer cells, Biochim. Biophys. Acta 1820 (2012) 643-51.

6.

B. A. C. Ackrell, Cytopathies involving mitochondrial complex II, Mol. Aspects Med. 23 (2002) 369-84.

7.

C. L. Quinlan, A. L. Orr, I. V. Perevoshchikova, J. R. Treberg, B. A. Ackrell & M. D. Brand, Mitochondrial complex II can generate reactive oxygen species at high rates in both the forward and reverse reactions, J. Biol. Chem. 287 (2012) 27255-64.

8.

C. Bardella, P. J. Pollard & I. Tomlinson, SDH mutations in cancer, Biochim. Biophys. Acta 1807 (2011) 1432-43.

9.

C. Frezza, P. J. Pollard & E. Gottlieb, Inborn and acquired metabolic defects in cancer, J. Mol. Med. 89 (2011) 213-20.

10.

E. A. Jones & H. Gutfreund, Oxidation of succinate and the control of the citric acid cycle in the mitochondria of guinea-pig liver, mammary gland and kidney, Biochem. J. 87 (1963) 639-48.

13

11.

K. R. Pryde & J. Hirst, Superoxide is produced by the reduced flavin in mitochondrial complex I: a single, unified mechanism that applies during both forward and reverse electron transfer, J. Biol. Chem. 286 (2011) 18056-65.

12.

Y. Hatefi & D. L. Stiggall, Preparation and properties of succinate:ubiquinone oxidoreductase (complex II), Methods Enzymol. 53 (1978) 21-7.

13.

P. Munujos, J. Coll-Cantí, F. González-Sastre & F. J. Gella, Assay of succinate dehydrogenase

activity

by

a

colorimetric-continuous

method

using

iodonitrotetrazolium chloride as electron acceptor, Anal. Biochem. 212 (1993) 506-9. 14.

H. Miyadera, K. Shiomi, H. Ui, Y. Yamaguchi, R. Masuma, H. Tomoda, H. Miyoshi, A. Osanai, K. Kita & S. Omura, Atpenins, potent and specific inhibitors of mitochondrial complex II (succinate-ubiquinone oxidoreductase), Proc. Natl. Acad. Sci. USA 100 (2003) 473-7.

15.

M. M. Nachlas, S. I. Margulies & A. M. Seligman, A colorimetric method for the estimation of succinic dehydrogenase activity, J. Biol. Chem. 235 (1960) 499-503.

16.

M. Worsfold, M. J. Marshall & E. B. Ellis, Enzyme detection using phenazine methosulphate and tetrazolium salts: interference by oxygen, Anal. Biochem. 79 (1977) 152–6.

17.

T. M. Weaver, D. G. Levitt, M. I. Donnelly, P. P. Wilkens Stevens & L. J. Banaszak, The multisubunit active site of fumarase C from Escherichia coli, Nature Struct. Biol. 2 (1995) 654–62.

18.

F. P. Bologna, C. S. Andreo & M. F. Drincovich, Escherichia coli malic enzymes: two isoforms with substantial differences in kinetic properties, metabolic regulation, and structure, J. Bacteriol. 189 (2007) 5937–46.

14

19.

M. S. Sharpley, R. J. Shannon, F. Draghi & J. Hirst, Interactions between phospholipids and NADH:ubiquinone oxidoreductase (complex I) from bovine mitochondria, Biochemistry 45 (2006) 241-8.

20.

M. Spinazzi, A. Casarin, V. Pertegato, L. Salviati & C. Angelini, Assessment of mitochondrial respiratory chain enzymatic activities on tissues and cultured cells, Nature Protocols 7 (2012) 1235-46.

21.

M. Estévez, J. Skarda, J. Spencer, L. Banaszak & T. M. Weaver, X-ray crystallographic and kinetic correlation of a clinically observed human fumarase mutation, Protein Sci. 11 (2002) 1552–7.

22.

M. D. Hatch, A simple spectrophotometric assay for fumarate hydratase in crude tissue extracts, Anal. Biochem. 85 (1978) 271-5.

23.

A. A. Alam, A. J. Rowan, N. C. Wortham, P. J. Pollard, M. Mitchell, J. P. Tyrer, E. Barclay, E. Calonje, S. Manek, S. J. Adams, P. W. Bowers, N. P. Burrows, R. Charles-Holmes, L. J. Cook, B. M. Daly, G. P. Ford, L. C. Fuller, S. E. HadfieldJones, N. Hardwick, A. S. Highet, M. Keefe, S. P. MacDonald-Hull, E. D. A. Potts, M. Crone, S. Wilkinson, F. Camacho-Martinez, S. Jablonska, R. Ratnavel, A. MacDonald, R. J. Mann, K. Grice, G. Guillet, M. S. Lewis-Jones, H. McGrath, D. C. Seukeran, P. J. Morrison, S. Fleming, S. Rahman, D. Kelsell, I. Leigh, S. Olpin & I. P. M. Tomlinson, Genetic and functional analyses of FH mutations in multiple cutaneous and uterine leiomyomatosis, hereditary leiomyomatosis and renal cancer, and fumarate hydratase deficiency, Hum. Mol. Genet. 12 (2003) 1241-52.

24.

F. Medja, S. Allouche, P. Frachon, C. Jardel, M. Malgat, B. Mousson de Camaret, A. Slama, J. Lunardi, J. P. Mazat & A. Lombès, Development and implementation of standardized respiratory chain spectrophotometric assays for clinical diagnosis, Mitochondrion 9 (2009) 331-9.

15

25.

V. G. Grivennikova, E. V. Gavrikova, A. A. Timoshin & A. D. Vinogradov, Fumarate reductase activity of bovine heart succinate-ubiquinone reductase. New assay system and overall properties of the reaction, Biochim. Biophys. Acta 1140 (1993) 282-92.

26.

S. McLean, S. M. Richards, S.-L. Cover, S. Brandon, N. W. Davies, J. P. Bryant & T. P. Clausen, Papyriferic acid, an antifeedant triterpene from birch trees, inhibits succinate dehydrogenase from liver mitochondria, J. Chem. Ecol. 35 (2009) 1252–61.

27.

T. E. King, Preparation of succinate dehydrogenase and reconstitution of succinate oxidase, Methods Enzymol. 10 (1967) 322–31.

28.

Y. Yang, H.-R. Wang, J.-X. Xu & H.-M. Zhou, Kinetics of modification of the mitochondrial succinate-ubiquinone reductase by 5,5'-dithiobis-(2-nitro-benzoic acid), J. Protein Chem. 15 (1996) 169-76.

29.

H. Schägger & K. Pfeiffer, The ratio of oxidative phosphorylation complexes I-V in bovine heart mitochondria and the composition of respiratory chain supercomplexes, J. Biol. Chem. 276 (2001) 37861-7.

30.

D. S. Kalashnikov, V. G. Grivennikova & A. D. Vinogradov, Submitochondrial fragments of brain mitochondria: general characteristics and catalytic properties of NADH:ubiquinone oxidoreductase (complex I). Biochemistry (Mosc.) 76 (2011) 20916.

31.

A. P. Wojtovich & P. S. Brookes, The endogenous mitochondrial complex II inhibitor malonate regulates mitochondrial ATP-sensitive potassium channels: implications for ischemic preconditioning, Biochim. Biophys. Acta 1777 (2008) 882-9.

32.

E. Tomitsuka, Y.-I. Goto, M. Taniwaki & K. Kita, Direct evidence for expression of type II flavoprotein subunit in human complex II (succinate-ubiquinone reductase), Biochem. Biophys. Res. Commun. 311 (2003) 774-9.

 

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Table Table 1. Comparison of values measured using the different assay methods1. Preparation

Substrates

Coupled assay

DCPIP

INT

Complex II2

Succinate + DQ

0.41 ± 0.023

0.20 ± 0.007

0.25 ± 0.024

Complex II2

Succinate only

0.002 ± 0.001

0.062 ± 0.003

0.036 ±0.003

Tissue3 + NaCN4

Succinate + Q1

0.12 ± 0.006

0.06 ± 0.001

Tissue3 + NaCN4

Succinate

0 ± 0.001

0.018 ± 0.001

Tissue3

Succinate

0.054 ± 0.002

0.019 ± 0.001

Table footnotes: 1: all values are reported in mol min-1 mg-1, and measured with 5 mM succinate and 100 M decylubiquinone (DQ) or 50 M ubiquinone-1 (Q1) as stated. 2: detergent-solubilized bovine heart mitochondrial membranes in which complex II is functionally isolated. 3: rat skeletal muscle homogenate. 4: NaCN inhibits cytochrome c oxidase; it prevents reoxidation of endogeneous ubiquinol-10 so that no ubiquinone-10 is present.

17

Figure and Scheme Legends

Scheme 1. Reaction scheme for the coupled assay, illustrating the use of FumC and MaeB to couple the oxidation of succinate to the reduction of NADP+ in a 1:1 stoichiometry. Figure 1. Kinetic characterization of the FumC and MaeB enzymes. A). Rate of conversion of fumarate to malate by FumC, detected by the production of NADPH that is driven by the conversion of malate to pyruvate by MaeB. The concentrations were 1.7 µg mL-1 FumC, 300 µg mL-1 MaeB and 2 mM NADP+. B) and C). Rate of conversion of malate to pyruvate by MaeB, detected by the production of NADPH. The concentrations were 44.2 µg mL-1 MaeB and 4 mM NADP+ (B) or 40 mM malate (C). In all cases assays were carried out in triplicate with standard error bars reported, at 32 °C, in buffers containing 10 mM TrisSO4 at pH 7.4, 250 mM sucrose, 2 mM MgSO4, and 1 mM K2SO4. Figure 2.

Quantifying the rate of succinate oxidation by the respiratory chain in

submitochondrial particles. A). Assay trace measuring succinate oxidation by SMPs (40 µg mL-1), detected by the coupled FumC-MaeB assay system as the formation of NADPH (300 µg mL-1 MaeB, 60 µg mL-1 FumC). The trace becomes linear after approximately 100 s. B). Observed rates of succinate oxidation by SMPs (40 µg mL-1), detected by the coupled FumC-MaeB assay system as the formation of NADPH, over a range of FumC and MaeB concentrations. C). Rates of succinate oxidation by SMPs, detected by the coupled FumCMaeB assay system as the formation of NADPH, over a range of SMP concentrations (300 µg mL-1 MaeB, 60 µg mL-1 FumC). In B and C assays were carried out in triplicate with standard error bars reported. In all cases assays were at 32 °C, in buffers containing 10 mM Tris-SO4 at pH 7.4, 250 mM sucrose, 5 mM succinate, 2 mM NADP+, 2 mM MgSO4, and 1 mM K2SO4.

18

Figure 3. Kinetic parameters describing succinate oxidation and its inhibition by the respiratory chain in SMPs. A). Rate of succinate oxidation by SMPs detected by the coupled assay system (solid symbols) or by the Clark O2 electrode (open symbols). B) and C). Rate of succinate oxidation by SMPs detected by the coupled assay in the presence of varying concentrations of atpenin A5 (B) or malonate (C). Inhibition curves were fit using the standard dose-effect relationship with Hill slope = 1. The assays contained 120 µg mL-1 FumC, 600 µg mL-1 MaeB, 5 mM succinate (B and C), 2 mM NADP+, 2 mM MgSO4 and 1 mM K2SO4 in 10 mM Tris-SO4 at pH 7.4 and 250 mM sucrose. The measurements were carried out in triplicate at 32 °C and are reported with standard error bars.

19

(μmol min-1 mg-1)

Rate

Figure

300 200 100 0 0

A 5

10

15

20

(μmol min-1 mg-1)

Rate

[fumarate] (mM) 8

8

6

6

4

4

2

2

0

B 0 10 20 30 40 50 60

[malate] (mM)

0

C 0

1

2

3

[NADP+] (mM)

4

Abs. 340-380 nm

Figure

0.5 0.4 0.3 0.2 0.1

A

0

100

Rate

0

200

300

Time (s)

μmol min-1 mg-1

Abs min-1

0.8

0.3

0.6

0.2 MaeB = 300 μg mL-1 FumC = 60 μg mL-1

0.4 0.2 0

0.1

B 0

1

2

3

Relative enzyme concentration

4

0

C 0

100

[SMP] (μg

200 mL-1)

(μmol min-1 mg-1)

Rate

Figure

1.2 0.8 0.4 0

A 0

5

10

15

20

Rate (%)

[succinate] (mM) 100

100

80

80

60

60

40

40

20 0

B 0

20

40

60

80 100

[atpenin] (nM)

20 0

C 0

1

2

3

4

[malonate] (mM)

5

Scheme

oxaloacetate succinate fumarate decarboxylating dehydrogenase hydratase malic enzyme + NADPH NADP Q H2O succinate fumarate malate pyruvate QH2

-O

-O

+ CO2

-O

O

O

O

OH O

O O-

O O-

O O

O-

O-