Gas regulation of complex II reversal via electron shunting to fumarate in the mammalian ETC

Abstract

The electron transport chain (ETC) is a major currency converter that exchanges the chemical energy of fuel oxidation to proton motive force and subsequently, ATP, generation, using O₂ as a terminal electron acceptor. Discussed herein, two new studies reveal that the mammalian ETC is forked. Hypoxia or H₂S exposure promotes the use of fumarate as an alternate terminal electron acceptor. The fumarate/succinate and CoQH₂/CoQ redox couples are nearly iso-potential, revealing that complex II is poised for facile reverse electron transfer, which is sensitive to CoQH₂ and fumarate concentrations. The gas regulators, H₂S and ^•NO, modulate O₂ affinity and/or inhibit the electron transfer rate at complex IV. Their induction under hypoxia suggests a mechanism for how traffic at the ETC fork can be regulated.

The electron transport chain is a major currency converter

Oxidation-reduction (or redox) reactions (see Glossary) constitute life’s currency, harnessing energy released from the thermodynamically favorable movement of low-potential donor electrons to high-potential acceptors. The electron transport chain (ETC) is a major currency converter in the cellular marketplace of metabolically generated low-potential reductants, namely NADH, FADH₂ and CoQH₂. At the redox donor end of the ETC, proton-coupled electron transfer generates a proton motive force across the inner mitochondrial membrane that is dispensed as ATP via oxidative phosphorylation and stored as potential energy to power cellular functions such as membrane transport. At the redox acceptor end, O₂, a high-potential oxidant, is churned to water in four one-electron steps. The cellular profit from glucose via ETC-fueled ATP generation can be ~18-fold greater than from aerobic glycolysis [1].

The high ATP exchange rate of oxidative phosphorylation is not, however, the sole determinant of cellular metabolic choice. It is well known that various cell types such as pluripotent stem cells [2], proliferating vascular endothelial cells [3], and malignant cells [4] are highly glycolytic. In these instances, the lucrative ATP yield from oxidative phosphorylation is sacrificed for repurposing mitochondria as anabolic hubs, supporting macromolecular precursor synthesis [4]. The complex regulatory mechanisms governing the extent to which the aerobic glycolysis versus the oxidative phosphorylation currency exchange counter is utilized, continue to be elucidated.

O₂ availability represents a plausible mechanism for substrate-level regulation of the ETC, governing the relative usage of glycolysis versus oxidative phosphorylation. Tissues and cell types are exposed to variable O₂ tensions, ranging from a low of 0.4% in the colonic lumen to 13% in regions of lung [5]. Even within an organ, O₂ tensions can vary widely, e.g., from 1.9% in the renal medulla to 9% in the cortex [5]. However, the potential for O₂ concentration per se to substantially influence ETC flux is questionable since the K_M(O2) for cytochrome c oxidase is very low. Depending on the rate of electron throughput, K_M(O2) is estimated to range from <0.2 to 1 μM, corresponding to <0.02–0.1% O₂ tension [6]. Since the O₂ tension in most tissues exceeds the K_M(O2) for cytochrome c oxidase, alternative cellular strategies must exist to regulate O₂ affinity and/or proton-coupled electron transfer rate through the ETC. The concentration of electron donors, the proton motive force and ADP levels are other factors that influence the rate of electron transfer to cytochrome c oxidase. In this review, we discuss how the gas regulators, i.e., hydrogen sulfide (H₂S), nitric oxide (^•NO), and O₂ interact to impact the ETC exchange rate by complex II reversal, using fumarate as an alternate terminal electron acceptor [7, 8]. We also discuss how the redox properties of complex II make it uniquely poised for bidirectional currency transactions in the ETC that is sensitive to fluctuations in the metabolic marketplace.

ETC architecture

The ETC is a marvel of cellular engineering and one that is perhaps unrivaled in complexity. Some 69 subunits encoded by the nuclear and mitochondrial genomes come together for the assembly of the ETC core, comprising complexes I, III, and IV (Fig. 1A). Complex I is an NADH:ubiquinone oxidoreductase, moving electrons from NADH to ubiquinone or coenzyme Q₁₀ (CoQ) as protons are translocated from the matrix to the inner mitochondrial membrane space. Complex II, comprising 4 subunits, is a component of the Krebs cycle (succinate dehydrogenase) and transfers electrons from FADH₂ to CoQ as succinate is oxidized to fumarate. Other mitochondrial dehydrogenases (dihydroorotate dehdydrogenase, glycerol 3-phosphate dehydrogenase, and proline dehdydrogenase) and oxidoreductases (sulfide quinone oxidoreductase (SQOR) and the electron transfer flavoprotein quinone oxidoreductase (ETF-QO)) also exchange redox currency at the ETC, using CoQ as an acceptor, and move electrons from organic (e.g., proline, dihydroorotate) or inorganic (H₂S) donors. Electrons from CoQH₂ (or ubiquinol) are funneled to complex III (cytochrome bc₁ complex) and, via cytochrome c, to complex IV or (cytochrome c oxidase) where O₂ serves as the terminal electron acceptor. Complexes III and IV also pump protons as they transfer electrons. An ~1160 mV potential difference between the NADH/NAD⁺ (E°′= −340 mV) and 0.5 H₂O/O₂ (+820 mV) redox couples translates to a maximal transmembrane protonmotive force of 180–220 mV [9]. Proton reentry to the mitochondrial matrix through complex V (ATP synthase) drives ATP synthesis. The ETC leaks both protons and electrons, with the latter generating reactive oxygen species, which can have beneficial signaling roles at low, but are damaging at high concentrations [10].

Figure 1. The mammalian ETC is forked.

A. Various electron donors support electron flux through the ETC via proton-coupled electron transfer. CoQ is an intermediate electron carrier in the membrane and accepts electrons from complexes I and II and donates electrons to complex III. CoQ also accepts electrons from various housekeeping enzymes like dihydroorotate dehydrogenase (DHOD), proline dehydrogenase (PRODH), glycerol 3-phosphate dehydrogenase (G3PDH), electron transfer flavoprotein ubiquinol oxidoreductase (ETF-QO), and sulfide quinone reductase (SQOR). Proton translocation occurs at complexes I, III and IV, building up a proton motive force. B. Modulation of O₂ affinity and/or electron transfer activity at complex IV by ^•NO, H₂S or O₂ restriction due to hypoxia can inhibit O₂-dependent forward electron transfer through the ETC, leading to a reductive shift in the NADH/NAD⁺ and CoQH₂/CoQ pools (red up arrows). Under these conditions, complex II can reverse direction, using fumarate as a terminal electron acceptor. For simplicity, the establishment of redox cycles between complex II operating in reverse and DHOD and SQOR are shown here, sustaining de novo pyrimidine synthesis and H₂S oxidation, respectively. In principle, these redox cycles can be established between other redox couples that use CoQ as an electron acceptor. C. The bifurcation in the mammalian ETC allows electron flow to either complex II or complex IV where fumarate and O₂, respectively are utilized as terminal electron acceptors. Only complex I is shown as the electron donor for simplicity although any CoQ-dependent enzyme can use the forked path.

The ETC is forked and can use fumarate as a terminal electron acceptor

Metabolic reprogramming is an adaptive strategy that increases network diversity in response to extracellular cues (e.g., O₂ or nutrient availability) or cellular needs (e.g., proliferation). Classic examples include diverting pyruvate to lactate thereby disconnecting glycolysis from oxidative phosphorylation (i.e., the Warburg effect [11]), and reversing direction within the Krebs cycle (e.g., reductive carboxylation [12]). Two recent studies have illuminated the capacity for reprogramming within the mammalian ETC, leading to electron shunting to fumarate either constitutively or under stress, or when complex III or IV is inhibited [7, 8] (Fig. 1B). Fumarate is the two-electron oxidation product of succinate in a reaction catalyzed by succinate dehydrogenase that concomitantly reduces FAD to FADH₂. Inhibition of complex III or IV leads to a reductive shift upstream in the ETC as recycling of CoQH₂ to CoQ is slowed, creating an electron acceptor insufficiency with potentially widespread metabolic ramifications. Succinate accumulation is a signature of cells exposed to H₂S [7] or hypoxia, as well as of cells lacking complex III or complex IV activity [8]. Succinate also accumulates in other physiological states such as in the ischemic heart [13] and in post exercise muscle [14].

Electron shunting to fumarate from complex I and DHOD (Fig. 1B) is observed when O₂-dependent electron flux is impaired [8]. By using fumarate in lieu of O₂, cells sustain de novo pyrimidine biosynthesis, supporting proliferation. In cell culture, fumarate reduction increases progressively as O₂ concentration decreases from 20% to 1%, plateauing at ~3%. Elegant in vivo isotope tracing studies reveal that tissues exhibit considerable heterogeneity in their capacity to reduce fumarate [8]. Thus, while some tissues (liver, brain, and kidney) display fumarate reduction constitutively and at levels that exceed succinate oxidation, others use this option scarcely, if at all (e.g., lung, heart, white adipose tissue, gastrocnemius muscle, pancreas, thymus). Exercise on the other hand, increases fumarate reduction in heart and white adipose tissue. Hypoxia (1%) increases fumarate reduction in most tissues, ex vivo [8].

The variable extent to which tissues use fumarate as a terminal electron acceptor is not simply correlated with either fumarate levels or the CoQH₂/CoQ ratio [8] and suggests that other strategies are at play for regulating electron flow at the fork in the ETC (Fig. 1B,C). One such regulator is H₂S, an endogenously produced gas, which at low concentrations, stimulates respiration but at high concentrations, inhibits complex IV [15]. The rapidity of the cellular response to H₂S [16] is consistent with the hydrophobicity of H₂S and its tendency to concentrate in membrane versus aqueous solvent [17]. While the pH dependence for inhibition of cytochrome c oxidase by H₂S has not been reported, its reactivity with other heme proteins is enhanced at lower pH, implying that the protonated rather than the anionic form of sulfide interacts with their metal centers [18–20].

H₂S induces a reductive shift in the NADH/NAD⁺ pool that is accompanied by increased glucose consumption and lactate generation [16, 21]. H₂S also increases lipid biogenesis, which is fueled by glutamine-dependent reductive carboxylation [22], hallmarks of reductive stress. A shift in the mitochondrial but not cytoplasmic NADH/NAD⁺ pool triggers these H₂S-driven metabolic changes and is attenuated by dissipation of the mitochondrial NADH pool using the water-forming NADH oxidase LbNOX [23]. The paradoxical observation that H₂S clearance continues even at very low O₂, first hinted at the existence of a complex IV-independent pathway for its oxidation [24] and led to the discovery that complex II reversal is a mechanism for prioritizing H₂S clearance when respiration is inhibited [7]. In H₂S treated cells, the malate-aspartate shuttle and the purine nucleotide cycle furnish fumarate to support reverse complex II activity [7], which are also the sources of fumarate leading to ischemic succinate accumulation [13].

Fumarate reduction has been previously reported under conditions where complex IV activity is greatly diminished or absent. For instance, fumarate reduction is used as an adaptive strategy in retina that is exposed to a chronically hypoxic environment, which leads to downregulation of a complex IV subunit [25]. In an ex vivo study, fumarate reduction was described as a component of a malate-succinate shuttle for the transfer of electron equivalents (as succinate) across an O₂ gradient, i.e., from O₂-poor retina to O₂-rich retinal pigment epithelium-choroid [25]. Reverse complex II activity has also been described in ischemia when complex IV activity is precluded by the lack of O₂ [13]. The relative contributions of the Krebs cycle versus reverse complex II activity to ischemic succinate accumulation is controversial [13, 26].

H₂S and NO•: Gas regulators of the ETC

H₂S and ^•NO are metabolically generated gases derived from amino acids and function as signaling molecules that elicit varied physiological effects [27, 28]. H₂S is synthesized from cysteine and/or homocysteine by the action of cystathionine β-synthase, γ-cystathionase, or mercaptopyruvate sulfurtransferase [29] while arginine is the source of ^•NO synthesized by nitric oxide synthases [30] (Fig. 2A,B). A third gas signaling molecule, CO, is a synthesized from heme by heme oxygenases, but unlike ^•NO and H₂S, its effects on the ETC are relatively poorly characterized [31]. ^•NO and H₂S are reversible inhibitors of complex IV. Adding complexity to their interaction however, ^•NO and H₂S also serve as substrates and are oxidized by complex IV to nitrate [32] and a putative hydrodisulfide species [33], respectively. As a competitive inhibitor, ^•NO increases the K_Mapp for O₂ for cytochrome c oxidase. As O₂ concentrations vary from 10 to 145 μM, the IC₅₀ value for ^•NO increases from ~20 to 270 nM, which is within the range of estimated tissue ^•NO concentrations (10–450 nM) [34]. The IC₅₀ value is comparable to the ED₅₀ (10–20 nM) for soluble guanylate cyclase, a major ^•NO receptor that signals via the second messenger cGMP [35, 36].

Figure 2. Scheme showing biogenesis of ^•NO and H₂S.

A. H₂S is synthesized by three enzymes of which cystathionine β-synthase (CBS) and γ-cystathionase (CSE) function canonically in the transsulfuration pathway. At cellular concentrations of reactants, the transsulfuation reactions (blue arrows) are preferentially catalyzed. Depending on cysteine availability, the enzymes can switch to H₂S biogenesis as shown. CBS and CSE catalyze multiple H₂S-generating reactions but for simplicity, only the major reactions at physiologically relevant concentrations of substrate are shown. Mercaptopyruvate sulfur transferase (MPST) converts 3-mercaptopyruvate to pyruvate and H₂S in the presence of a reductant (not shown). B. Arginine is the source of ^•NO catalyzed by nitric oxide synthases (NOS) in the presence of reducing equivalents, which are not shown here for simplicity. Hcy and Cys denote homocysteine and cysteine, respectively.

An H₂S sensor analogous to soluble guanylate cyclase has not been identified. However, SQOR could potentially function in this capacity, generating CoQH₂ and reactive sulfur species as potential second messengers [37]. SQOR is a membrane anchored flavoprotein with an unusual cysteine trisulfide redox cofactor that oxidizes H₂S to glutathione persulfide (GSSH), concomitantly reducing CoQ to CoQH₂ (Fig. 3) [38]. Electrons from H₂S oxidation can enter the ETC at the level of complex III, making sulfide an inorganic substrate for ATP generation [39]. GSSH is further oxidized to sulfite via the action of ETHE1, a persulfide dioxygenase [40], or can donate its sulfane sulfur for thiosulfate synthesis catalyzed by thiosulfate sulfurtransferase (or rhodanese) [41]. The reported K_i for the inhibition of cytochrome c oxidase by H₂S is 200 nM [42]. SQOR shields complex IV from respiratory poisoning by H₂S and its expression levels are correlated with the cellular sensitivity to this gas [16]. The steady-state concentration of H₂S ranges between 10–50 nM in tissues where reliable estimates exist and is considerably higher (~1 μM) in aorta [24, 43]. Some cell types, like the epithelial cells lining the large intestine, are exposed to high concentrations of exogenous H₂S (0.2–2.4 mM) derived from microbial metabolism [44, 45] and express high levels of the sulfide oxidation pathway enzymes for its detoxification [16].

Figure 3. Sulfide oxidation connects H₂S to the ETC.

H₂S is oxidized via the mitochondrial sulfide oxidation pathway comprising SQOR in the mitochondrial inner membrane, which converts H₂S to glutathione persulfide (GSSH) while reducing CoQ to CoQH₂. GSSH is further oxidized to sulfite by ETHE1, a persulfide dioxygenase, or serves as a sulfane sulfur donor, forming thiosulfate in a reaction catalyzed by thiosulfate sulfur transferase (TST, also known as rhodanese). In some cells, thiosulfate is further oxidized to sulfate in a reaction catalyzed by sulfite oxidase (not shown).

By increasing the K_M for O₂ by ^•NO or by decreasing the rate of O₂ reduction by H₂S [42, 46], both gases can dial down ETC flux, increase local O₂ levels, and potentially modulate the O₂ threshold for hypoxia sensing. ^•NO induces metabolic hypoxia, a paradoxical state in which O₂ increases due to cytochrome c oxidase inhibition and leads to destabilization of HIF1α under hypoxic conditions [47]. Under these conditions, O₂ can be diverted to other enzymes whose activities might otherwise be limited under hypoxia. For example, ^•NO-enhanced O₂ availability (that is independent of cGMP), stimulates luciferase activity and is important for bioluminescent communication during firefly courtship [48].

HIF and ATF4 regulation of H₂S and ^•NO

^•NO and H₂S biogenesis are upregulated by hypoxia [49], nutrient stress, and the unfolded protein response [50, 51] pathway (Fig. 4). A molecular intersection between ^•NO (and CO) and H₂S signaling occurs at cystathionine β-synthase, the first enzyme in the transsulfuration pathway, which in turn switches the substrate preference of γ-cystathionase, the second enzyme in the pathway, from cystathionine to cysteine [52]. While cystathionine β-synthase and γ-cystathionase are both promiscuous [53], kinetic control renders catalysis of cystathionine and cysteine generation, respectively as their dominant reactions at physiologically relevant substrate concentrations (Fig. 2A). The activity of cystathionine β-synthase is inhibited when ^•NO or CO binds to the regulatory ferrous heme cofactor [54, 55]. With reduced competition from cystathionine under these conditions, the metabolic track preference of γ-cystathionase flips from cystathionine to H₂S biogenesis [52] (Fig. 4). Reprogramming of the transsulfuration pathway upregulates H₂S synthesis under endoplasmic reticulum stress conditions [52] and could also be induced in response to other triggers such as hypoxia.

Figure 4. Regulation of gas modulator synthesis.

CSE is regulated by the transcription factor, ATF4 which is activated by amino acid starvation or during the unfolded protein response, leading to increased H₂S synthesis (red arrow). Under hypoxia, NOS and heme oxygenase are induced leading to ^•NO and CO biosynthesis, respectively, which inhibit CBS, a hemeprotein, leading to reduced cystathionine. Under these conditions, the transsulfuration pathway enzymes switch metabolite tracks from the canonical cystathionine and cysteine producing reactions (white arrows) to H₂S biogenesis primarily by CSE (red arrow). Thus, the three cellular triggers shown here have the potential to upregulate synthesis of the ETC gas regulators, H₂S and ^•NO. Cyst denotes cystathionine.

Complex II is tuned for reversible electron transfer

A sizeable potential difference (ΔE°′) exists between CoQH₂/CoQ and the CoQ-dependent mitochondrial oxidoreductases except for complex II and ETF-QO (Table 1), and provides the thermodynamic driving force for CoQ reduction. The large ΔE°′ also precludes ready reversal of electron transfer from CoQH₂ at these redox junctions. In contrast, the CoQH₂/CoQ and fumarate/succinate couples are almost iso-potential, rendering the direction of electron transfer catalyzed by complex II sensitive to the mitochondrial CoQH₂/CoQ and fumarate/succinate ratios. Consistent with the predicted facile reversibility of succinate dehydrogenase, is its ability to support E. coli growth on fumarate as a terminal electron acceptor under anaerobic conditions, in a strain that lacks fumarate reductase [56].

Table 1.

Redox potentials for reactions coupling to the mitochondrial CoQH₂/CoQ pool

Enzyme	Redox Couple	E°′	Reference
DHOD	Dihydroorotate/Orotate	−252 V	[62]
PRODH	Proline/Δ1-pyrroline-5-carboxylate	−123 mV	[63]
ETF-QO	1FADsq/FADox	+28 mV	[64]
SQOR	2H2S/HSSH	−230 mV	[65]
G3PDH	Glycerol 3-phosphate/dihydroxyacetone phosphate	−190 mV	[66]
Complex II	Fumarate/succinate	+30 mV
Complex I	3NADH/NAD+	−340 mV	[67]
	CoQH2/CoQ	+40 to +60 mV	[68]

¹The one-electron redox potential for FAD in ETF-QO is reported as CoQ reduction by ETF-QO reportedly occurs in two one-electron transfer steps.

²The two-electron redox potential reported for the H₂S/HSSH couple is listed. The reaction catalyzed by SQOR leads to the synthesis of the alkyl persulfide, GSSH.

³Value for pH 7.8

Conditions that increase CoQH₂ levels (e.g. reductive stress triggered by H₂S, hypoxia, complex III or complex IV dysfunction) and fumarate (via metabolic reprogramming), favor complex II reversal as seen in human cells exposed to H₂S [7] or hypoxia [8] or inhibited at complex III or IV [8]. Consistent with this prediction, dissipation of the mitochondrial CoQH₂ pool by ectopic expression of the alternate oxidase suppresses fumarate reduction in cells with impaired complex III or IV activity [8]. In principle, allosteric regulation of complex II, for example via posttranslational modification, could also influence its propensity to reverse the direction of electron flow but such a mechanism, if it exists, remains to be identified. In ETF-QO, the one-electron redox potential for the proximal electron donor to CoQ, i.e., the flavin semiquinone/flavin couple, is high (Table 1). ETF-QO is a component of a multiprotein electron wire comprising ETF and a family of acyl-CoA dehydrogenases. Unlike complex II, reverse electron transfer through this system is unlikely to be facile.

We posit that the concentration of gas regulators, H₂S and ^•NO relative to O₂, modulates complex IV activity, underlying the variable capacity of tissues to siphon off electrons to fumarate under steady-state conditions or during an acute response. The physiological relevance of H₂S-induced electron rerouting from CoQH₂ to fumarate is supported by the substantial decrease in sulfide oxidation in mice with a targeted deletion of complex II in intestinal epithelial cells, which are exposed to the gas from gut microbes [7].

Prokaryotes adapted to growth in O₂ limiting or anaerobic environments, and eukaryotes like parasitic helminths and freshwater snails that are facultative anaerobes, have long been known to use fumarate as an alternate electron. In these organisms, fumarate reduction is correlated with the use of lower potential quinone derivatives such menaquinone and rhodoquinone [57]. In mammals, the establishment of an overall redox cycle between CoQ and fumarate versus O₂ as a terminal electron acceptor, shaves off a sizeable thermodynamic advantage (ΔΔE°′ ~770 mV). Nevertheless, the redox couples remain highly favorable for substrate oxidation and fumarate reduction e.g., ΔE°′ is ~ +302 mV for DHOD and ~+280 mV for SQOR, corresponding to ΔG°′ values of −13.9 kcal/mol and −12.9 kcal/mol, respectively. Similarly, the ΔE°′ for coupling NADH oxidation by complex I to fumarate reduction by complex II is ~ +390 mV, corresponding to a ΔG°′ of −18 kcal/mol, rendering electron flow to either O₂ or fumarate highly favorable (Fig. 1C). Importantly, the CoQ-fumarate-dependent redox cycle provides an avenue for the continued operation of CoQ-dependent reactions when forward electron transfer to complex IV is impeded and aerobic glycolysis assumes a more significant role in ATP generation.

Concluding Remarks

Two recent studies have revealed a homeostatic role for complex II reversal in tissue physiology (e.g., liver, kidney, brain, and colon) under conditions that lead to a reductive shift in the CoQH₂/CoQ redox pool [7, 8]. Based on the similar potentials of the CoQH₂/CoQ and fumarate/succinate redox couples, complex II appears to be uniquely poised to bifurcate the ETC, increasing its functional versatility. The gas regulators, ^•NO and H₂S, can serve as valves for tuning O₂ affinity and/or inhibiting the electron transfer rate in complex IV, which in turn, influences partitioning upstream at the ETC fork (Figs. 1B,C).

The contribution of fumarate reduction to metabolic physiology awaits fuller elucidation and might be particularly relevant in the context of pathologies that are marked by reductive stress such as diabetes, cancer, ischemia and obesity. H₂S is cytoprotective when co-administered during reperfusion [58], consistent with its ability to inhibit complex IV and drive complex II reversal, thereby reducing reactive oxygen species generation that is initiated by oxidation of succinate, which accumulates during ischemia. Dissipation of the NADH pool by LbNOX improves glucose tolerance in mice on a high fat diet, suggesting the therapeutic potential of a general strategy for targeting reductive stress [59]. Hereditary mutations in complex II subunits are associated with familial paraganglioma and Leigh’s syndrome [60]. Considering the bidirectional capacity of complex II in the cellular milieu, the metabolic vulnerabilities resulting from the loss of fumarate reduction in addition to succinate oxidation will have to be considered to understand the underlying disease mechanisms. Interestingly, among the known risk factors for nonfamilial paraganglioma are chronic hypoxia, including high altitude living and cardiorespiratory diseases [60, 61]. Future studies will unravel the molecular details of how electron flow is regulated at the ETC fork and the bioenergetic and signaling contexts for and the consequences of fumarate versus O₂ utilization (see Outstanding Questions).

OUTSTANDING QUESTIONS.

Why is fumarate a significant electron acceptor in some but not other tissues?

Is the varying use of fumarate by the ETC correlated with varying levels of H₂S-, ^•NO-induced or other type of reductive stress?

What is the primary source of fumarate in tissues that utilize it as a significant electron acceptor?

What is the metabolic fate of succinate formed via fumarate reduction?

What are the regulatory mechanisms that coordinate electron flux at the ETC fork?

Does hypoxia promote utilization of fumarate as an electron acceptor by enhancing H₂S and/or ^•NO biogenesis?

How is H₂S synthesis regulated in normal and in disease states?

Does the dependence on fumarate as an electron acceptor change in disease states such as diabetes, obesity and cancer?

How much of the CoQ pool is free in the mitochondrial membrane?

How does the composition and proportion of respiratory supercomplexes influence electron flux at the ETC fork?

Does reverse electron transfer through complex II support intercellular redox cycles in tissues other than retina?

What is the therapeutic potential for modulating fumarate reduction? For example, could enhancing fumarate reduction during the reperfusion phase decrease ischemic injury?

HIGHLIGHTS.

• Two recent studies reported the discovery that fumarate can serve as a terminal electron acceptor in mammalian tissues, revealing that the electron transport pathway is forked.

• The similar redox potentials of the fumarate/succinate and reduced/oxidized coenzyme Q couples, indicate that complex II is poised to operate either in the direction of succinate oxidation or fumarate reduction.

• Interaction between the gas regulators, i.e., O₂, hydrogen sulfide and nitric oxide, could modulate flux at the electron transport chain fork, siphoning electrons to complex II or complex III.

• It is critically important to understand the underlying mechanisms for tissue and cell-specific variations in the use of fumarate as a terminal electron acceptor in normal and disease states.

Glossary

Cytochrome c

A hemeprotein that transfers electrons between complex III and complex IV in the electron transport chain.

Coenzyme Q₁₀ (CoQ)

A ubiquinone derivative with ten isoprene units attached to the benzoquinone head group. CoQ is a redox active membrane cofactor and can exist in three oxidation states: fully reduced (CoQH₂), radical semiquinone (CoQH^•), and fully oxidized (CoQ).

Dihydroorotate dehydrogenase (DHOD)

A mitochondrial enzyme that catalyzes CoQ-dependent oxidation of dihydroorotate to orotate, in the de novo pyrimidine biosynthetic pathway.

Electron transport chain

Comprises a series of protein complexes I–IV that transfer electrons while concomitantly generating a proton gradient across the mitochondrial inner membrane. In mammals, O₂ and fumarate serve as terminal electron acceptors.

Electron transfer flavoprotein quinone oxidoreductase (ETF-QO)

Transfers electrons to CoQ from ETF, which in turn is reduced by a family of acyl-CoA dehydrogenases that are involved in the β-oxidation of fatty acids.

Fumarate

A C4-dicarboxylate intermediate in the Krebs cycle and a terminal electron acceptor for the ETC.

Glycolysis

A cytoplasmic pathway for glucose oxidation to pyruvate that generates ATP and NADH.

Hypoxia

A state in which the tissue O₂ tension is restricted and leads to stabilization of hypoxia inducible factors (HIFs), which initiate a cascade of signaling events.

K _M

The substrate concentration at which half maximal activity of an enzyme is seen (also referred to as the Michaelis constant).

Malate-aspartate shuttle

An intercompartmental metabolic shuttle that allows the transfer of cytosolic reducing equivalents (from NADH) as malate, which crosses the mitochondrial inner membrane via a reversible α–ketoglutarate/malate carrier. Mitochondrial aspartate is exchanged for cytosolic glutamate through the aspartate-glutamate carrier.

Mitochondrial glycerol 3-phosphate dehydrogenase (G3PDH)

Catalyzes the CoQ-dependent oxidation of glycerol 3-phosphate to dihydroxyacetone phosphate. Together with cytoplasmic G3PDH, which catalyzes the NADH-dependent reverse reaction, constitutes the glycerophosphate shuttle, linking carbohydrate and lipid metabolism. The shuttle also serves to transfer reducing equivalents from cytosolic NADH to the mitochondrial ETC.

NAD⁺

Nicotinamide adenine dinucleotide is a redox active coenzyme used by many enzymes. NADH is the reduced form.

Proton motive force (PMF)

Refers to the transmembrane proton gradient. In the mitochondrion, proton coupled electron transfers by complexes I, III and IV generate the proton motive force. The movement of protons back to the mitochondrial matrix via complex V powers ATP synthesis in a process referred to as oxidative phosphorylation. The PMF is also used to power other cellular functions such as membrane transport.

Purine nucleotide cycle

A sequence of reactions in which aspartate and inosine monophosphate (IMP) are converted to adenylosuccinate, which is cleaved to fumarate and adenosine monophosphate (AMP). Deamination of AMP releases ammonia and IMP and completes the cycle.

Proline dehydrogenase (PRODH)

A mitochondrial enzyme that catalyzes the CoQ-dependent oxidation of L-proline to Δ1-pyrroline-5-carboxylate.

Redox reaction

Oxidation-reduction (redox) reaction involving an electron donor (reductant) and an electron acceptor (oxidant).

Reductive carboxylation

Conversion of α-ketoglutarate to isocitrate catalyzed by isocitrate dehydrogenase. This reaction is the reverse of oxidative decarboxylation catalyzed by the same enzyme in the Krebs cycle.

Sulfide quinone oxidoreductase (SQOR)

Is a mitochondrial inner membrane enzyme that catalyzes the oxidation of H₂S to glutathione persulfide (GSSH) as it reduces CoQ to CoQH₂.