Cytopathies involving mitochondrial complex II

Molecular Aspects of Medicine 23 (2002) 369–384 www.elsevier.com/locate/mam

Chapter 2

Cytopathies involving mitochondrial complex II Brian A.C. Ackrell

*

Department of Veterans Affairs Medical Center, Molecular Biology Division, San Francisco, CA 94121, USA

Abstract Complex II (succinate–ubiquinone oxidoreductase) is the smallest complex in the respiratory chain and contains four nuclear-encoded subunits SdhA, SdhB, SdhC, and SdhD. It functions both as a respiratory chain component and an essential enzyme of the TCA cycle. Electrons derived from succinate can thus be directly transferred to the ubiquinone pool. Major insights into the workingks of complex II have been provided by crystal structures of closely related bacterial enzymes, which have also been genetically manipulated to answer questions of structure-function not approachable using the mammalian system. This information, together with that accrued over the years on bovine complex II and by recent advances in understanding in vivo synthesis of the non-heme iron co-factors of the enzyme, is allowing better recognition of improper functioning of human complex II in diseased states. The discussion in this review is thus limited to cytopathies arising because the enzyme itself is defective or depleted by lack of iron–sulfur clusters. There is a clear dichotomy of effects. Enzyme depletion and mutations in SDHA compromise TCA activity and energy production, whereas mutations in SDHB, SDHC, and SDHD induce paraganglioma. SDHC and SDHD are the first tumor suppressor genes of mitochondrial proteins. Ó 2002 Elsevier Science Ltd. All rights reserved.

1. Introduction Mammalian complex II (succinate-ubiquinone oxidoreductase; SQR; EC 1.3.5.1.) is a complex multi-subunit enzyme that straddles the mitochondrial inner membrane in a manner allowing dual functionality as both a component of the electron transport chain and an essential enzyme of the TCA cycle. The enzyme is thus ideally situated to help gear TCA cycle activity to the energy demands of the cell. The peripheral domain (succinate dehydrogenase; SDH) of complex II, which projects into the mitochondrial *

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matrix and is comprised of a flavoprotein (FP; SdhA) and an iron–sulfur (IP; SdhB) subunit, is anchored by two membrane-spanning polypeptides, SdhC and SdhD. The active site of the enzyme containing covalently bound FAD (8a-histidyl-FAD) is located in FP. Electrons derived from oxidation of succinate and destined for the ubiquinone pool in the membrane are carried from the reduced flavin to a quinone binding site(s) on the anchor polypeptides by three linearly aligned iron–sulfur clusters ð½2Fe–2S2þ;1þ ; ½4Fe–4S2þ;1þ , and ½3Fe–4S1þ;0 Þ in IP. Two stabilized and interacting semiubiquinone radicals ðQ Q Þ are detected during ubiquinone reduction by bovine SQR. The mechanism of reduction suggested by the Q Q pair is that an electron is transferred from the reduced [3Fe–4S]0 cluster of IP to one bound quinone and then immediately on to a second bound quinone, which when fully reduced dissociates into the ubiqinone pool. The proximal binding site (QP ) is located next to the [3Fe–4S]1þ;0 cluster and contains a b-type heme, cytochrome b558 , liganded by histidines provided by SdhC and SdhD. The location of the distal binding site (QD ) towards the cytoplasmic side of the membrane is controversial. For detailed descriptions of the properties of the enzyme and assay procedures see Ackrell et al. (1992), H€ agerh€ all (1997). Other considerations include the tissue variance in control of oxidative phosphorylation, which is not executed at a single step of the respiratory chain and phosphorylating apparatus but over multiple steps that change according to tissue and its physiological state. Loss of SQR activity due to a genetic defect or oxidative stress would markedly impact energy production should the depletion exceed the threshold (excess capacity) of the enzyme for that particular tissue. Further, post-mitotic tissues such as brain and heart have the highest metabolic rates and oxygen consumption and, hence, potential for superoxide production and deleterious Fenton chemistry. Whether oxyradicals act by damaging native SQR in the membrane or by inhibiting biosynthesis and assembly or one of the ancillary processes such as iron–sulfur cluster production is presently unclear. It is known, for example, that the [3Fe–4S]1þ;0 cluster in the Ip subunit is particularly vulnerable to oxidative disruption unless protected by the presence of the anchor polypeptides of SQR (Vinogradov et al., 1975). There is by now little question that defective energy output and aberrant glutamate neurotransmission are intimately linked in the development of acute and chronic neurological disease. That human SQR deficiency should be rare is not surprising considering the central role played by the enzyme in metabolism and energy production. The condition, alone or in combination with deficiencies of other Fe–S enzymes, is observed in <20% of patients with respiratory chain defects (Vladutiu and Heffner, 2000). Mutations have now been identified in all subunits. Those in SDHA decrease the activity of SQR in the membrane and those in SDHB, SDHC, and SDHD are projected to destabilize, even disassemble, the structure of the complex as a prelude to tumor formation (Baysal et al., 2001). SQR depletions without mutations in coding sequences must be secondary effects of problems with biogenesis, assembly, and/or maintenance. Clinical symptoms involving skeletal and cardiac muscles and the central nervous system are thus wide ranging. They can also be tissue-specific, but probably not as a result of differential expression of isoforms, since the nuclearencoded subunits are distributed universally and apparently expressed from single

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copy genes: SDHA (chromosome 5p15) (Bourgeron et al., 1995), SDHB (1p35-36.1) (Leckschat et al., 1993), SDHC (1q21) and SDHD (11q23) (Hirawake et al., 1997).

2. Complex II deficiency 2.1. Selective depletion of SQR Selective loss of SQR (70%) has been reported in a case of sporadic Kearns–Sayre syndrome (Table 1) (Rivner et al., 1989), in two brothers presenting with hypertrophic cardiopathy and skeletal myopathy (Reichman and Angelini, 1993), and in a patient with hypertrophic cardiopathy, who showed no loss of SQR activity in any other tissue (Rustin et al., 1993). In the absence of evidence for isoforms, such a tissuespecific pattern of expression is the likely result of a tissue-specific irregularity in transcription and/or translation. The symptom in common in these patients is cardiopathy. Intriguingly, therefore, SQR is a major target in Huntington’s disease (HD), which is an autosomal-dominant neurodegenerative disorder characterized in adulthood by chorea, personality changes and dementia. The mutation in the Huntingtin gene (chromosome 4) is an unstable CAG repeat expansion that introduces glutamines into the N-terminus of the protein (see Albin and Tagle, 1995, for review). While both wild type and mutant proteins are expressed throughout the nervous system and in non-neuronal tissues, patients exhibit an almost exclusive loss of striatal spiny neurons. Examination of postmortem samples from HD patients has confirmed a 30–70% depletion of SQR and complexes III and IV in the caudate (Table 1) and putamen, but not in other tissues (Browne et al., 1997; Gu et al., 1996). The fact that these complexes, but not complex I, are heme-containing proteins has not, however, been explored. Rat and non-human primate models are obtained by the simple expediency of systemic administration of the SQR suicide inhibitor 3-nitropropionate (3NP) or intra-striatal injection of the competitive inhibitors malonate or methylmalonate (Beal et al., 1993; Greene et al., 1993). Semiquantitative histological analysis has confirmed that 60% inhibition of brain SDH (activity was assessed with a dye as electron acceptor) over a 24 h period is sufficient to produce the severe motor symptoms and start of striatal degeneration reminiscent of the human disease (Greene et al., 1993). However, the special vulnerability of the striatum suggests a confluence of factors, since neither the amount of SDH nor its susceptibility to 3NP is particularly different from elsewhere in the brain (Brouillet et al., 1998). One possibility is that spiny neurons are exquisitely sensitive to impaired energy production, which is detected by NMR even before gross striatal atrophy appears (Jenkins et al., 1993). Support for this is that the more obvious signs of striatal SQR inhibition by 3NP include increased N-methyl-D -aspartate (NMDA) toxicity and intracellular Caþþ and lower membrane potentials. This long-term potentiation of NMDA-mediated synaptic transmission, specific to striatal spiny neurons, is dependent on endogenous dopamine (DA) (Calabresi et al., 2001). Compared to other parts of the brain, the striatum is heavily innervated with dopaminergic neurons

372

Table 1 Mitochondrial enzyme activities compared to controls (%) Fe deficiency (rats)

Myopathy with

Kearns–Sayre syndrome

Huntington’s disease

Friedreich’s ataxia

Muscle

Myoglobinuria Muscle

Encephalopathy Muscle

Muscle

Caudate

Heart

Inner-membrane Complex I Complex II Complex III Complex IV

23 16 – 100

Deficienta 22 37 100

24 9 20 73

100 18–34b – 100

100 29b – 34

16 23b – 71d

Matrix Aconitase (i) Before activation (ii) After activatione Malate dehydrogenase Citrate synthase Fumarase

22 43 100 – 100

Deficienta – – 100–150 –

– – – – –

– – – – –

– – – 100 –

14 – – 73 –

References

–c

Hall et al. (1993)

Desnuelle et al. (1989)

Rivner et al. (1989)

Browne et al. (1997)

Bradley et al. (2000)

a b c d e

By immunoblotting. Assayed as succinate-cytochrome c oxidoreductase. Ackrell et al. Unpublished. Not significant. By incubation with Feþþ , sulfide, and reducing agent.

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Enzyme activities

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from the substantia nigra. This raises the specter of superoxide particularly since expression of brain-derived neurotrophic factor, which inhibits apoptosis and DAinduced free radical production, is decreased exclusively in the striatum of HD patients (Petersen et al., 2001). 2.2. Depletion of SQR with other Fe–S enzymes 2.2.1. Iron deficiency Iron deficiency affects 20–50% of the world’s population, being most prevalent among children and pregnant women of developing countries. Studies of iron-deficient rats documented dramatic decreases in activity by SQR, complexes I and III, and matrix aconitase (Table 1). The selective nature of this effect is apparent from the fact that under conditions where SQR and complex I activities decrease by some 80%, the specific contents of individual cytochromes decrease less than 40%, and ATP’ase increases slightly (Maguire et al., 1982). By analyzing for the respective co-factors, FMN and histidyl-flavin, it was demonstrated that the lower activities were due to decreased amounts of complex I and SQR in Fe-deficient membranes (Ackrell et al., 1984). Otherwise each complex functioned normally. Complexes being assembled in iron-deficient membranes were thus fully functional and replete with intact clusters. That modified complexes were absent suggested that cluster-deficient subunits are never assembled, modified complexes are proteolyzed, or there is coordinate repression of subunits linked to low Fe–S cluster supply. Coordinate expression is indicated by binding elements for the nuclear respiration factor NRF-1 in the promoter regions of human SDHA (Parfait et al., 2000), SDHB (Au and Scheffler, 1998), SDHC and SDHD (Hirawake et al., 1997), and the gene for the ubiquitous form of 5-aminolevulinate synthase (Braidotti et al., 1993), the initial enzyme in the synthesis of heme. Lack of heme (Hederstedt and Rutberg, 1980), or the enzyme ferrochelatase responsible for inserting Fe into protoporphyrin IX (Nihei et al., 2001), has been shown in Bacillus subtilis and Escherichia coli, respectively, to prevent the anchor polypeptides from binding FP and IP, which therefore remain in the cytoplasm. That subunit production is tied to ambient Fe concentration gains credence from a recent identification of non-consensus iron-responsive elements (IREs) (Theil and Eisenstein, 2000) in the 50 UTRs of mammalian SDHA and SDHB (Leckschat et al., 1993; Parfait et al., 2000; Au and Scheffler, 1998), and the fact that a non-consensus IRE in the mRNA of the [Fe–S]-containing 75 kD subunit of complex I is functional (Lin et al., 2000). 2.2.2. Limited cluster availability Multiple defects of the respiratory chain have been reported for two patients, one presenting with myopathy and encephalopathy (Desnuelle et al., 1989), the other with severe exercise intolerance and episodic myoglobinuria (Hall et al., 1993). In each case the deficiency of SQR matched those of complexes I and III and aconitase (Table 1). Complex IV and ATPase were unaffected. The fact that blood iron levels in these patients appeared normal and loss of Fe–S enzymes was selective clearly pointed to problems with cluster synthesis or delivery. Although clusters can be synthesized non-enzymically (Holm, 1977) and have been transferred in vitro from SDH to

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Table 2 Depletion of SQR and aconitase caused by defective Fe–S cluster synthesis Deficient protein

Aconitase % Depletion

SQR % Depletion

Mitochondrial Fe Approach Fold increase

References

Nfs1p Isu1p Isu2p Nfu1p Ssq1p Jac1p

83–67 38 9 53 89 75

82–47 56 42 52 92 70

0–15 1.2 1.3 1.3 10.6 6.7

Li et al. (1999) Schilke et al. (1999) Schilke et al. (1999) Schilke et al. (1999) Schilke et al. (1999) Voisine et al. (2001)

Isa1p

>97

>95

8.0

Missense mutation Deletion Deletion Deletion Deletion Temperature mutant Deletion

Isa2p

>97

>95

8.0

Deletion

Yah1p

93

80

30.0

Yfh1p Atm1p

87 <5

40 <15

15 12.0

Regulated expression Deletion Regulated expression

Jensen and Culatta (2000) Jensen and Culatta (2000) Lange et al. (2000) Foury (1999) Kispal et al. (1999)

apoferredoxins by the ‘‘core extrusion’’ technique (Coles et al., 1979), only recently have insights been gained of the enzyme-mediated processes in mitochondria. Synthesis of iron–sulfur clusters in eukaryotes occurs in the mitochondrial matrix by a group of proteins sharing considerable homology with their bacterial counterparts of the Nif (nitrogen fixing) and housekeeping Isc (iron–sulfur cluster assembly) systems (elaborated in a recent review article by M€ uhlenhoff and Lill, 2000). Only brief descriptions are provided here to identify the yeast proteins enumerated in Table 2, which summarizes the discordant iron movement and debilitating effect on complex II assembly should the proteins be defective: (i) Nfs1p, an essential protein, is a desulfurase converting cysteine to alanine and either sulfane (S0 ) or, in the presence of reducing potential, sulfide (S2 ) (Zheng et al., 1993). Depleted Nfs1p activity results in decreased activity of Fe–S enzymes including SQR (Li et al., 1999; Kispal et al., 1999). (ii) In analogy to the role found for the NifU and IscU proteins, yeast Isu1p and Isu2p are projected to accept persulfide from Nfs1p and act as scaffolds for cluster formation (Urbina et al., 2001; Yuvaniyama et al., 2000). While deletion of either of the ISU genes results in significant loss of Fe–S enzyme activities (Schilke et al., 1999), the double deletion is lethal (M€ uhlenhoff and Lill, 2000). (iii) Nfu1p is not essential and possibly has an auxiliary role (Schilke et al., 1999). (iv) Based on studies of Nif IscA of A. vinelandii, yeast Isa1p and Isa2p are alternate scaffolds for [2Fe–2S] and [4Fe-4S] cluster syntheses (Krebs et al., 2001). Deletion of the yeast genes results in dramatic losses of aconitase and SQR activities (Jensen and Culatta, 2000). (v) The major loss of Fe–S enzyme activity incurred by depletion of the yeast matrix ferredoxin Yah1p (Lange et al., 2000) is suggestive of a need in the assembly

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process for electrons donated via Yah1p and the andrenodoxin reductase Arh1p. Accordingly, the E. coli ferredoxin homolog has been shown to complex the metal-bound form of IscA (Ollagnier-de-Choudens et al., 2001). (vi) Binding studies suggest that the primary cellular function of the Hsp66/hsc20 chaperone system in E. coli is to interact with IscU (Isu1p and Isu2p in yeast), presumably by aiding protein folding and unfolding during cluster formation and/or its subsequent insertion into apoenzymes. Depletion of the homologous chaperones in yeast, Ssq1p (Schilke et al., 1999) and Jac1p (Voisine et al., 2001), respectively, considerably lowers aconitase and SQR activities. (vii) Yfh1p, clearly involved in iron homeostasis, is of unknown function (Foury, 1999). Defects in the human homologue frataxin are responsible for the disease Friedreich’s ataxia (see below). The only two genes with the same phylogenetic distribution as YFHI/FRATAXIN are SSQ1 and JAC1 (Huynen et al., 2001). Yfh1p/frataxin would thus be predicted to participate with this chaperone system incluster synthesis or insertion into apo-subunits of respiratory enzymes. (viii) Atm1p is an ATP-binding cassette (ABC) transporter required for cluster export from the mitochondrial matrix into the cytosol. Its deletion has little effect on synthesis of matrix Fe–S proteins (Kispal et al., 1999). It is clear from Table 2 that defects in any of the proteins listed other than Yfh1p/ frataxin, which results in Friedreich’s ataxia, could explain the general loss of Fe–S enzymes in the patients cited above. The iron deposits noted in the second patient’s mitochondria (Hall et al., 1993) are also consistent with the massive accumulation of iron associated with defects of Nfs1p, Yah1p, Ssq1p, Atm1p, and the ISA proteins. The hallmarks of inhibited cluster synthesis, i.e., depletion of SQR and other Fe–S enzymes, lower respiration and growth, DNA damage, and accumulation of iron, are clearly evident in yeast models deleted for Yfh1p. Mitochondrial iron overload occurs at the expense of cytosolic iron, which causes, in turn, the induction of the high affinity iron-uptake system of the plasma membrane that is regulated at the level of transcription by Aft1p, an iron sensor. Although cellular iron levels in Dyfh1 cells are consequently about double that in wild type, the mitochondria show the massive accumulation (10–15 excess) and accretion of iron (Babcock et al., 1997) sometimes seen in patients with Friedreich’s ataxia (Bradley et al., 2000). That matrix Yfh1p is needed for the mobilization of iron to the cytoplasm is evident in the fact that its appearance in cells carrying a YFH1 allele on an inducible promoter correlates with decreased levels of both mitochondrial iron and proteins of the high iron-uptake system (Radisky et al., 1999). While iron overload persists the cells are hypersensitive to oxidative stress. SQR and aconitase appear particularly vulnerable to a YFH1 disruption, suffering decreases in activity (Foury, 1999) approaching 40% and 80%, respectively, similar to those occurring in cardiac tissue of patients with Friedreich’s ataxia (Table 1). Prevention of iron accumulation in Dyfh1 cells (Foury, 1999; Chen and Kaplan, 2000) has been shown to restore in large measure the activities of respiratory complexes except for aconitase. Thus, while Yfh1p deficiency undoubtedly fosters an excess of mitochondrial iron and deleterious Fenton chemistry, it is uncer-tain in yeast whether the protein is totally indispensable for cluster

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synthesis and en-zyme assembly. In contrast, deletion of frataxin in mice is lethal (Cossee et al., 2000). Friedreich’s ataxia is an autosomal recessive disease usually occurring before adulthood and characterized clinically by progressive ataxia, lower limb weakness, and large fiber sensory loss. Frataxin-deficient cells undergoing neurogenesis are particularly susceptible to apoptosis and cell death compared to those directed, for example, into cardiomycetes (Santos et al., 2001). Most patients have hypertrophic cardiomyopathy, while skeletal deformities are seen in two-thirds of the patients, and a third of patients have diabetes. Ninety-six percent of patients with Friedreich’s ataxia are homozygous for a GAA triplet expansion in the first intron of the frataxin gene (chromosome 9q13) that lowers transcriptional efficiency (Campuzano et al., 1996). The larger the number of GAA repeats in the smaller allele, the lesser the frataxin produced and the quicker and more severe the clinical expression of the disease. Depletions of SQR, complexes I and III, and aconitase are most obvious in endomyocardial biopsy samples (Table 1). Compared to deficient yeast, frataxindeficient skin fibroblasts and lymphoblasts accumulate only modest mitochondrial levels of iron, even in the presence of high extra-cellular concentrations. This is attributable in part to differences in regulating iron uptake (Babcock et al., 1997; Tan et al., 2001). That a higher fraction of the mitochondrial iron is in a filterable (free) form is of immediate interest being simultaneously a possible consequence of restricted Fe–S cluster synthesis and the cause of increasing sensitivity to oxidative stress. The cells also have a lower mitochondrial membrane potential (Tan et al., 2001) consistent with their lower respiratory capacity (Table 1) and the lower ATP production seen in patients (Lodi et al., 2001). All of these cellular phenotypes, central to disease development, can be rescued by over-expression of frataxin (Tan et al., 2001). That lack of clusters spirals into these secondary effects would be consistent with findings that cardiomyopathy, sensory nerve defect, and major depletions (80%) of SQR and aconitase in mouse strains heterozygous (+/)) for frataxin all precede intra-mitochondrial iron accumulation and deposition (Puccio et al., 2001). That iron-catalyzed oxygen radical damage subsequently contributes to the etiology of the disease is implicit in the protection afforded by antioxidants (Lodi et al., 2001) and the fact that patients with isolated vitamin E deficiency caused by a mutation in the atocopherol transfer protein exhibit similar clinical symptoms to those with Friedreich’s ataxia (Ouahchi et al., 1995). The problem is magnified in Friedreich’s ataxia by the fact that normal induction of matrix superoxide dismutase in response to excessive superoxide is suppressed by frataxin deficiency (Chantrel-Groussard et al., 2001) and the ability of neurotrophins to protect neurons during development or injury by inhibiting superoxide production (Dugan et al., 1997) may simply be overwhelmed. 3. Effect of mutations in SQR 3.1. Flavoprotein subunit (FP) The first mutation in human SQR was identified in 1995 as a homozygous SdhA Arg554Trp replacement (Table 3) in two siblings presenting with Leigh syndrome, a

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Table 3 Mutations in human SQR and corresponding diseases Subunit

Protein change

Phenotype

FP (SdhA) Initiating codon

M1I

Leigh Syndrome

In mature protein

R451C

In mature protein

A524V

In mature protein

R554W

IP (SdhB) In mature protein

R90X

In mature protein

P197R

In mature protein

Frameshift after P197

Pheochromocytoma

SdhC Initiating codon

M1V

SdhD Initiating codon

Inheritance

References

Autosomal recessive Optic atrophy, ataxia, Autosomal myopathy dominant? Leigh Syndrome Autosomal recessive Leukodystrophy with Leigh Autosomal Syndrome recessive

Parfait et al. (2000) Birch-Machin et al. (2000) Parfait et al. (2000) Bourgeron et al. (1995)

Hereditary paraganglioma+pheochromocytoma Pheochromocytoma

Autosomal dominant Autosomal dominant

Astuti et al. (2001a) Astuti et al. (2001a) Astuti et al. (2001a)

Hereditary paraganglioma

Autosomal dominant

Niemann and Muller (2000)

M1I

Hereditary paraganglioma

Badenhop et al. (2001)

In pre-sequence

Frameshift after A18

Hereditary paraganglioma

In pre-sequence

R22X

Hereditary paraganglioma

Autosomal dominant with imprinting Autosomal dominant with imprinting Autosomal dominant with imprinting

In pre-sequence

Q36X

Hereditary paraganglioma

In pre-sequence

R38X

Hereditary paraganglioma

In pre-sequence In pre-sequence

R38X Frameshift after P41

Pheochromocytoma Hereditary paraganglioma

In mature protein

Frameshift after L64?

In mature protein

Frameshift after W66

Autosomal dominant with imprinting Autosomal dominant with imprinting

Autosomal dominant with imprinting Hereditary paraganglioma Autosomal dominant with imprinting Hereditary Pheochromocy- Autosomal toma dominant

Taschner et al. (2001) Taschner et al. (2001), GimenezRoqueplo et al. (2001) Baysal et al. (2000) Baysal et al. (2000) Gimm et al. (2000) Taschner et al. (2001) Badenhop et al. (2001) Astuti et al. (2001b)

(continued on next page)

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Table 3 (continued) Subunit

Protein change

Phenotype

Inheritance

References

In mature protein

R70G

Hereditary paraganglioma

Taschner et al. (2001)

In mature protein

P81L

Hereditary paraganglioma

Autosomal dominant with imprinting Autosomal dominant with imprinting

In mature protein In mature protein

P81L D92Y

Pheochromocytoma Hereditary paraganglioma

In mature protein

Del Y93

Hereditary paraganglioma

In mature protein

L95P

Hereditary paraganglioma

In mature protein

H102L

Hereditary paraganglioma

In mature protein

Hereditary paraganglioma

In mature protein

Aberrant splicing? L139P

Hereditary paraganglioma

In mature protein

G12S

Polymorphism

In mature protein

S68S

Polymorphism

Autosomal dominant with imprinting Autosomal dominant with imprinting

Autosomal dominant with imprinting

Autosomal dominant with imprinting

Baysal et al. (2000) Gimm et al. (2000) Taschner et al. (2001), Baysal et al. (2000) Badenhop et al. (2001) Taschner et al. (2001) Baysal et al. (2000) Taschner et al. (2001) Taschner et al. (2001) Taschner et al. (2001), Gimm et al. (2000) Taschner et al. (2001), Baysal et al. (2000)

Table adapted from Baysal et al. (2001).

progressive neurodegenerative disease usually with onset in infancy. Skeletal muscle, cultured skin fibroblasts, and circulating lymphocytes all showed a 70% depletion of SQR. Activity was not impaired in the healthy consanguineous parents heterozygous for the mutation (Bourgeron et al., 1995). Based on sequence alignments and information provided by the structures for bacterial L -aspartate oxidase and fumarate reductases (Maltevi et al., 1999; Lancaster et al., 1999; Leys et al., 1999; Iverson et al., 1999; Taylor et al., 1999), human Arg-554 is located in the C-terminal domain of SdhA that folds around the highly conserved core structure and active site made by the other domains. The effects of the Arg554Trp substitution, namely, loss of activity without changes in the binding of succinate or malonate, are the first to indicate that the C-terminal domain has an active role in catalytic cycling during which the active site ‘‘opens’’ and ‘‘closes’’ (Ackrell, 2000). This conclusion has support in the discovery of a second mutation, Ala524Val, in the C-terminal domain, which causes a similar loss of activity to that seen with Arg 554Trp. The patient, also with Leigh syndrome, had a compound heterozygous mutation where the Ala524Val replacement was in-

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herited in the allele from the father and the mother’s allele contained an altered initiating codon (Met1Leu) and produced unstable transcript (Parfait et al., 2000). A fourth mutation (Birch-Machin et al., 2000) causing a somewhat lower decrease (50%) of SQR activity than by mutations in the C-terminal domain (70%) gives rise to late-onset optic atrophy, ataxia, and myopathy as opposed to Leigh syndrome. The affected siblings, but not their sister, carry an Arg408Cys substitution in the active site of the enzyme that prevents catalysis. It is now clear from the structures of the related bacterial fumarate reductases (Lancaster et al., 1999; Leys et al., 1999; Iverson et al., 1999; Taylor et al., 1999) that bonding between Arg-408 and one of the carboxylates is required to position the succinate above and in parallel to the isoalloxine ring of FAD to facilitate proton abstraction. Equally catastrophic is that absence of Arg-408 prevents the FAD from forming a covalent linkage with the protein (Ackrell, 2000; Birch-Machin et al., 2000), which is needed to raise the potential of free flavin (Em0 ¼ 219 mVÞ sufficiently (i.e., to )79 mV for the FAD/ FADH2 couple in beef SDH) (Ohnishi et al., 1981) for succinate to be oxidized (Blaut et al., 1989). Subunit expressed from the mutated allele is thus completely inactive and the patient survives by virtue of good subunit from the second allele. 3.2. Iron–sulfur subunit (IP) Multiple screenings of patients with familial or sporadic pheochromocytoma and/ or paraganglioma have illuminated several heterozygous mutations in the SDHB, SDHC, and SDHD genes (Baysal et al., 2001), which further supports a link between bioenergetic defects and tumorogenesis. Hereditary paragangliomas are highly vascularized, non-chromaffin tumors usually arising in the parasympathetic ganglia in the head and neck, commonly at the carotid body, which senses the systemic oxygen levels. Pheochromocytomas share similar embryological origin to paragangliomas and are catecholamine producing, chromaffin tumors that arise in 90% of cases in the adrenal medulla. Mutations have been found in all three genes (Table 3). Of the SDHB mutations (Astuti et al., 2001a), one was germline in three affected families and introduced the stop codon Arg91X after 62 of 252 residues in the mature protein. A fourth family carried a Pro198Arg replacement and one person with sporadic pheochromocytoma had a frameshift deletion (delC) causing truncation immediately following the same residue. Pro-198 and the adjacent cysteine-197 serving as a ligand to the [3Fe–4S]1þ;0 cluster are both located at the interface formed by SdhB with the anchor polypeptides (Iverson et al., 1999). Any misfolding of SdhB stemming from the Pro198Arg mutation would be expected to disrupt center 3 and/or misalign the doublet of conserved lysine residues Lys241 Lys242 at the C-terminus, either of which prevents the assembly of SQR (Vinogradov et al., 1975; Schmidt et al., 1992). The membrane would then contain a mixture of native SQR and complexes of anchor polypeptides SdhC and SdhD. 3.3. The anchor polypeptides C and D Familial paraganglioma associated with SDHD mutations is inherited through the fathers but not affected mothers. Disease transmission in familial pheochromocytoma

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is also consistent with this imprinting. In contrast, only maternal transmission has been found for SDHB and SDHC mutations. Significantly, patients carrying SDHC and SDHD mutations (Table 3) suffer loss of heterozygosity (LOH) during tumorogenesis that results in loss of the maternal allele specifically. The presence of inactivating mutations and loss of the wild type allele earmark SDHC and SDHD as the first tumor suppressor genes of mitochondrial proteins. The mutations in SDHC and SDHD lead to different degrees of complex dismantling. The Met1Val replacement prevents assembly of SdhC into a complex, since the next initiating codon in the sequence is within the coding region of the mature protein (Met-9) and the truncated protein would lack the pre-sequence necessary for transport into the mitochondrion. Because LOH occurs and both anchor polypeptides are required for binding of the catalytic subunits FP and IP, the membrane will contain only SdhD provided it survives proteolysis. The first seven SDHD mutations listed in Table 3 are also within the presequence and yield nonviable proteins, as would the frameshift mutations occurring after Leu64 and Trp66 (Leu-8 and Trp-10 of the mature protein). By similar reasoning, these mutations allow only SdhC to be incorporated into the membrane, consistent with the absence of catalytic activity (Gimenez-Roqueplo et al., 2001). The other mutations in SDHD would not be expected to completely inactivate the complex. Based on sequence alignments and models of anchor polypeptides (H€agerh€all and Hederstedt, 1996), His-102 is a ligand for the heme at quinone-binding site QP . Substitution of this residue (Maklashina et al., 2001) in E. coli SQR converts the heme from a hexacoordinate low spin to pentacoordinate high spin form, but only slightly affects the activity of the mutant complex. Asp 92 and Leu 95 lie just below His 102 in the same a-helix in a position to influence the properties of the heme and/or bound semiquinone (H€ agerh€ all and Hederstedt, 1996). Substitution of the same histidine in B. subtilis SQR causes a greater instability (H€ agerh€all et al., 1995). Structural perturbations are expected from the Pro81Leu and Leu139Pro substitutions in SDHD, since they both involve proline. Even loss of the complete a-helix containing Leu139 does not completely inactivate the yeast complex (Oyedotun and Lemire, 2001). Since chronic hypoxic stimulation can induce the same tumors as mutations in SDHB, SDHC, and SDHD, it has been hypothesized that SQR has a role in sensing O2 levels in the carotid body and its loss, tantamount to zero oxygen, to signal cell proliferation (Baysal et al., 2000). How this might be accomplished remains enigmatic. One view holds that the signal for stabilization of the hypoxia-inducible factor (HIF-a) is the superoxide produced under hypoxic conditions by the respiratory chain (Chandel et al., 1998). HIF-a up-regulates such target genes as erythropoietin for increased proliferation of erythrocytes, the angiogenic factor (VEGF) for increased blood supply, and glycolytic enzymes to enhance survival under hypoxic conditions. First, the heme in SQR does not bind oxygen (Yang et al., 1997; Yu et al., 1987) and, second, the production of superoxide at SQR is modest compared to that at complex III (Chandel et al., 2000). A mechanism would have to be in place to distinguish sources of superoxide and accommodate the fact that superoxide is a membrane impermeant and would be produced from Q generated by SQR towards the matrix side of the inner membrane.

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381

The question arising is whether the mutations in SDHB, SDHC, and SDHD are designed to flood the system with superoxide as found with the mev1 mutation in SdhC of Caenorhabditis elegans (Senoo-Matsuda et al., 2001). Similarly, an SdhC Glu29Leu mutation in E. coli fumarate reductase, which results in a 70% loss of succinate-quinone oxidoreductase activity, increases the stability of the intermediate semiquinone (H€ agerh€ all et al., 1999) and, hence, chances of univalent reduction of O2 . While this possibility remains to be tested for the mutated complexes retaining some activity, it would seem remote for fragments of the complex lacking FP/IP, although these retain modified heme and bind quinone (Yu et al., 1987; Shenoy et al., 1999). Nonetheless, heme in preparations of bovine anchor polypeptides is reducible by NADH dehydrogenase type II (Yu et al., 1987), suggesting that ‘‘rear end’’ reduction could perhaps occur in the membrane via NADH and a reduced quinone pool if the polypeptides escape proteolysis. Against this, inactivation of SQR will progressively arrest TCA cycle activity and the major flow of reducing equivalents to the respiratory chain, the source of superoxide proposed to signal HIF-a stabilization. The first step in HIF-a degadation has recently been reported to be its hydroxylation by a prolyl hydroxylase-type enzyme (Ivan et al., 2001). The co-factors for such oxygenases are Feþþ , O2 , ascorbate, and 2-oxoglutarate, which is decarboxylated during the reaction. Any modification or disruption of SQR by the SDHB, SDHC, and SDHD mutations would be expected to attenuate TCA cycle activity and thus promote HIF-a stabilization by interrupting the supply of 2-oxoglutarate. Acknowledgements The author thanks Bruce Cochran for his excellent technical assistance, and the Department of Veterans Affairs and the National Institutes of Health (Grant HL16251) for their support. References Ackrell, B.A.C., 2000. FEBS Lett. 466, 1–5. Ackrell, B.A.C., Johnson, M.K., Gunsalus, R.P., Cecchini, G., 1992. In: M€ uller, F. (Ed.), Chemistry and Biochemistry of Flavoenzymes, vol. III. CRC Press, Boca Raton, FL, pp. 229–297. Ackrell, B.A.C., Maguire, J.J., Dallman, P.R., Kearney, E.B., 1984. J. Biol. Chem. 259, 10053–10059. Albin, R.L., Tagle, D.A., 1995. Trends Neurosci. 18, 11–14. Astuti, D., Latif, F., Dallol, A., Dahia, P.L., Douglas, F., George, E., Skoldberg, F., Husebye, E.S., Eng, C., Maher, E.R., 2001a. Am. J. Hum. Genet. 69, 49–54. Astuti, D., Douglas, F., Lennard, T.W.J., Aligianis, I.A., Woodward, E.R., Evans, D.G.R., Eng, C., Latif, F., Maher, E.R., 2001b. The Lancet 357, 1181–1182. Au, H.C., Scheffler, I.E., 1998. Eur. J. Biochem. 251, 164–174. Babcock, M., deSilva, D., Oaks, R., Davis-Kaplan, S., Jiralerspong, S., Montermini, L., Pandolfo, M., Kaplan, J., 1997. Science 276, 1709–1712. Badenhop, R.F., Cherian, S., Lord, R.S.A., Baysal, B.E., Taschner, P.E., Schofield, P.R., 2001. Genes Chromosomes Cancer 31, 255–263. Baysal, B.E., Ferrell, R.E., Willett-Brozick, J.E., Lawrence, E.C., Myssiorek, D., Bosch, A., van der Mey, A.G.L., Taschner, P.E., Rubinstein, W.S., Myers, E.N., Richard, III, C.W., Cornelisse, C.J., Devilee, P., Devlin, B., 2000. Science 287, 848–851.

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