2.4.1  Starch and sucrose degradation

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Figure 2.19 Pathways of starch metabolism. Numbers refer to the following enzymes: 1, ß-amylase; 2, α-amylase; 3, starch phosphorylase; 4, glucosidase; 5, hexose kinase; 6, phosphoglucomutase; 7, glucose 6-phosphate isomerase. (Original drawing courtesy David Day)

Starch is the major carbon reserve in most plants. It is a mixture of amylose and amylopectin and is deposited as granules inside plastids (chloroplasts in leaves, amyloplasts in non-photosynthetic tissues). The initial attack on starch granules in leaves and non-photosynthetic tissues is by a-amylase and a debranching enzyme. Oligosaccharides released during starch degradation, such as maltose, maltotriose and maltotetraose, are hydrolysed to glucose by a-glucosidase. These processes are summarised in Figure 2.19.

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Figure 2.20 A scheme for breakdown of sucrose in plants. Broken lines indicate possible interconnecting reactions. Numbers refer to the following enzymes: 1, invertase; 2, sucrose synthase 3, UDP-glucose pyrophosphorylase; 4, hexokinase; 5, fructokinase; 6, phosphogluconiutase; 7 phosphohexose isomerase; 8, phosphofructophosphotransferase; 9, phosphofructokinase. (Original drawing courtesy David Day)

Reactions in plant tissues leading to degradation of sucrose to hexose monophosphates are outlined in Figure 2.20. The first step is cleavage of the glycosidic bond by either invertase (Equation 2.1) or sucrose synthase (Equation 2.2).

Sucrose + H₂O → D-glucose + D-fructose (2.1)

Sucrose + UDP → UDP-glucose + D-fructose (2.2)

Plant tissues contain two types of invertases which hydrolyse sucrose to glucose and fructose in an essentially irreversible reaction: acid invertase, which has optimum activity near pH 5, and is present

In vacuoles, the free space outside cells, and may be associated with the cell wall; and alkaline or neutral invertase, which is maximally active at about pH 7 to 7.5 and is located in the cytosol. Sucrose synthase is a cytosolic enzyme that catalyses a readily reversible reaction, but probably acts only in the breakdown of sucrose in vivo. Sucrose appears to be partitioned between alkaline invertase and sucrose synthase in the cytosol on the basis of differences in affinity of the two enzymes for the substrate. (K_m values for sucrose of alkaline invertase and sucrose synthase generally fall within the ranges 10–15 mM and 20–30 mM, respectively.) Glucose and fructose are metabolised further following phosphorylation to the corresponding hexose-6-P, probably by separate enzymes for the two hexoses. Plant tissues contain several hexose kinases that have specificity towards either glucose or fructose. A substantial portion of the glucose kinase in plant tissues is associated with the outer surface of the outer mitochondrial membrane, while fructokinases appear to be soluble in the cytosol.

 Two molecules of ATP are required to metabolise the hexoses formed upon cleavage of sucrose by invertase. How-ever, when sucrose is cleaved by sucrose synthase, part of the energy in the glycosidic bond is conserved in the UDP-glucose formed and only one molecule of ATP is required for the further metabolism of fructose. UDP-glucose may be converted to glucose-1-P by UDP-glucose pyrophosphorylase. Glucose-1-P is converted to glucose-6-P, and glucose-6-P to fructose-6-P, by phosphoglucomutase and phosphohexose isomerase, respectively.

2.4.2  The glycolytic pathway

Glycolysis originally described the sequence of reactions that convert glycogen to lactic acid in muscle and is usually considered to include the metabolism of hexose phosphates to pyruvate. In plant tissues, starch takes the place of glycogen in this scheme, and there is probably a second end-product, either oxaloacetate or malate (Figure 2.21).

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Figure 2.21 Interrelationships between glycolysis and the pentose phosphate pathway. Enzymes abbreviated are: HK, hexokinase; HPI, hexose phosphate isomerase; PFK, phosphofructokinase; ALD, aldolase; TPI, triose phosphate isomerase; GAPDI-I, glyceraladehyde 3-phosphate dehydrogenase; PGA, phosphoglycerate; ENO, enolase; PK, pyruvate kinase; PDC, pyruvate decarboxylase; ADH, alcohol dehydrogenase; LDH, lactate dehydrogenase; G6PDH, glucose-6-phosphate dehydrogenase; 6-PGDH, 6-phosphogluconate dehydrogenase; PRI,
phosphoriboisomerase; PPE, pentose phosphate epimerase; TK, transketolase; TA, transaldolase; PEPC, phosphoenolpyruvate carboxylase; MDH, malate dehydrogenase. Note that lactate and ethanol are formed only when mitochondrial function is inhibited, as under anaerobiosis. (Original drawing courtesy David Day)

Moreover, in plants glycolysis occurs in both cytosol and plastids, with reactions in the different compartments catalysed by separate isoenzymes. The first step in the pathway is phosphorylation of fructose-6-P to fructose-1,6-P₂. Plant tissues contain two enzymes capable of catalysing this step: an ATP-dependent phosphofructokinase (PFK), which catalyses an essentially irreversible reaction and occurs in the cytosol and plastids, and phosphofructophosphotransferase (PFP) (now called PP_i-dependent phosphofructokinase, PP_i-PFK), which occurs only in the cytosol and utilises PP_i as the phosphoryl donor in a reaction that is readily reversible.

Regulation of PFK is achieved by a combination of mechanisms, including pH, the concentration of substrates and effector metabolites, changes in the state of aggregation, and covalent modification by phosphorylation/dephos-phorylation of the protein. The relative importance of these mechanisms varies depending on the organism. In plants phosphoenolpyruvate (PEP) is probably the most potent regulator, inhibiting at µM concentrations, but 3-PGA and 2-PGA also strongly inhibit. P_i activates the cytoplasmic PFK, and to a lesser extent that from plastids, and overcomes the inhibition by PEP. The enzyme is also activated by Cl^– and other anions. The regulatory metabolite fructose-2,6-P₂, a powerful activator of PFK from animals, has no effect on the enzyme from plants.

PFP, discovered subsequently, is ubiquitous in plants and has a catalytic potential higher than that of PFK. Fructose-2,6-P₂ strongly activates PFP, but the physiological significance of this activation and, indeed, the role of PFP, in plants have not yet been clearly established. Fructose-2,6-P₂ is a potent inhibitor of cytosolic fructose-1,6-bisphosphatase which is an important control point of sucrose biosynthesis regulating the partitioning of photosynthate between sucrose and starch in leaves. Whether fructose-2,6,-P₂ has a role in the control of glycolysis through its activation of PFP is not clear.

Since the reaction catalysed by PFP is reversible and the concentration of fructose-2,6-P₂ in the cytosol is usually high enough to maintain PFP in an activated form, the direction of this reaction in vivo is likely to depend on availability of substrates. In tissues where sucrose breakdown is occurring, PFP may function to generate PP_i to facilitate the conversion of UDP-glucose to glucose-1-P (Figure 2.20). Under these conditions, the simultaneous and opposing action of PFK and PFP in the cytosol could set up a potentially wasteful substrate cycle between fructose-6-P and fructose-1,6-P₂. The operation of such a cycle may be a cost of having a mechanism to generate PP_i and, ultimately, UDP for the breakdown of sucrose by sucrose synthase. PFP may also act as an inducible enzyme in some plant tissues, providing increased glycolytic capacity when required during certain stages of plant development or during adjustment to adverse environmental conditions.

Fructose-1,6-P₂ is cleaved by aldolase to form dihydroxyacetone-P and glyceraldehyde 3-P, and these triose phosphates are interconverted in a reaction catalysed by triose phosphate isomerase. Glyceraldehyde 3-P is oxidised to glycerate-1,3-P₂ by an NAD-dependent glyceraldehyde 3-P dehydrogenase in the cytosol and an NADP-linked enzyme in plastids. Glyceraldehyde 3-P dehydrogenase is sensitive to inhibition by the reduced pyridine nucleotide cofactor, which must be reoxidised to maintain the flux through the glycolytic pathway. In chloroplasts, the reactions catalysed by fructose-1,6-P₂ aldolase, triose phosphate isomerase and NADP-dependent glyceraldehyde 3-P dehydrogenase also form part of the PCR cycle. The remaining steps for PEP formation are shown in Figure 2.21; all steps from fructose-1,6-P₂ to PEP are reversible.

The end-product of glycolytic reactions in the cytosol of plants is determined by the relative activities of the two enzymes that can utilise PEP as a substrate: pyruvate kinase, which forms pyruvate and a molecule of ATP, and PEP carboxylase, which forms oxaloacetate (Figure 2.21). Both of these reactions are essentially irreversible and there are fine controls that regulate the partitioning of PEP between these reactions. Pyruvate kinase requires monovalent cations and is inhibited by ATP (and therefore is sensitive to the energy status of the cell), whereas PEP carboxylase is inhibited by malate and is independent of cell energy status. The sensitivity of PEP carboxylase to malate is regulated by phosphoryl-ation of the enzyme by a protein kinase: the phosphorylated form is less sensitive to malate inhibition. This phosphorylation may form part of an important diurnal regulatory cycle in the leaves of crassulacean acid metabolism plants (see Section 2.1).

Oxaloacetate is reduced by malate dehydrogenase to malate which, along with pyruvate, can be taken up into mitochondria and metabolised further (see below). The reduction of oxaloacetate in the cytosol could provide a cytosolic mechanism for oxidising NADH formed by glyceraldehyde 3-P dehydrogenase (Figure 2.21).

In chloroplasts glycolysis is most active in conjunction with the breakdown of starch to form sucrose for export to non-photosynthetic tissues. There is some doubt about the occurrence of phosphoglycerate mutase in chloroplasts, and therefore the main products of the glycolytic reactions may be triose phosphates and 3-PGA. These could be exported through the P_i translocator in the chloroplast envelope to the cytosol, where sucrose synthesis takes place. In photosynthetic cells, the triose P exported to the cytosol for sucrose synthesis (Section 2.1.8) could also enter the glycolytic pathway directly to provide mitochondrial substrates.

Studies of changes in the content of glycolytic intermediates in plant tissues that undergo an altered rate of
respiration (e.g. during the climacteric in ripening fruits or in plant tissues placed under low-O₂ stress) indicate that the conversion of fructose-6-P to fructose-1,6-P₂, and PEP to pyruvate, are major regulatory steps. For example, a decrease in activity of PEP carboxylase and pyruvate kinase (the latter in response to a lower energy demand as indicated by an increase in the cytosolic ATP/ADP ratio) can lead to an increase in the concentration of inhibitory metabolites of PFK and, consequently, a decrease in the rate of glycolysis. The rate of oxidation of NAD(P)H is also likely to have a bearing on the glycolytic flux at the glyceraldehyde 3-P dehydrogenase step.

2.4.3  Pentose phosphate pathway

An alternative route for the breakdown of glucose-6-P is provided by the pentose phosphate pathway (Figure 2.21, right side). This pathway functions mainly to generate NADPH and precursors for various biosynthetic processes. These include ribose-5-P, which provides the ribosyl moiety of nucleotides and is a precursor for the biosynthesis of the purine skeleton, and erythrose-4-P, for the biosynthesis of aromatic compounds in the shikimic acid pathway. Glyceraldehyde 3-P and fructose-6-P formed in the pentose phosphate pathway may be metabolised further in the glycolytic pathway. Alternatively, fructose-6-P may be converted back to glucose-6-P by phosphohexose isomerase and recycled through the pentose phosphate pathway. The pentose phosphate pathway may account for between 15 and 30% of the hexose phosphate oxidised to glyceraldehyde 3-P and CO₂.

As with glycolysis, reactions of the pentose phosphate pathway are catalysed by different sets of isoenzymes that occur either in the cytosol or in plastids. The reactions in the non-oxidative phase of the pentose phosphate pathway are readily reversible and also form part of the PCR cycle of chloroplasts.

2.4.4  Mitochondria and organic acid oxidation

βOrganic acids produced in the cytosol by processes described above are further oxidised in mitochondria via the tricarboxylic acid (TCA) or Kreb’s cycle and subsequent respiratory chain. Energy released by this oxidation is used to synthesise ATP which is then exported to the cytosol for use in biosynthetic events.

(a)  Mitochondrial structure

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Figure 2.22 Transmission electron micrograph of a parenchyma cell in a floral nectary of broad bean (Vicia faba) showing an abundance of mitochondria, generally circular in proiile and varying between about 0.75 and 1.5 µm in diameter. Each mitochondrion is encapsulated by an outer and inner membrane which is in turn infolded to form cristae. Under higher magnification, ‘knobs’ are seen to protrude from this inner membrane and are now recognised as ATP synthase complexes (refer to Figure 1 in Case study 2.1). Cytoplasmic ribosomes are also apparent, many of which have been organised into polyribosomal helices. Scale bar = 0.5 µm. (Original electron micrograph courtesy Brian Gunning)

Plant mitochondria (Figure 2.22) are typically double-membrane organelles where the inner membrane is invaginated to form folds (‘cristae’). The outer membrane contains relatively few proteins and is permeable to most compounds of less than 5 kDa molecular weight, by virtue of a pore-forming protein (‘porin’). This outer membrane also contains an NADH dehydrogenase and a b-type cytochrome whose function is not understood. The inner membrane is the main permeablity barrier of the organelle and controls the movement of molecules by means of a series of carrier proteins. The inner membrane also houses the redox carriers of the respiratory chain and delineates the soluble matrix which contains the enzymes of the TCA cycle, other soluble proteins and the protein synthesising machinery.

Mitochondria are semi-autonomous organelles with their own DNA and protein synthesising machinery. However, the mitochondrial genome codes for only a small portion of the proteins which make up the mitochondrion; the rest are encoded on nuclear genes and synthesised in the cytosol. These proteins are transported into the mitochondrion and assembled with the mitochondrially synthesised proteins to form respiratory complexes. The number of mitochondria per cell varies with tissue type (from a few hundred in mature differentiated tissue to some thousands in specialised cells such as those in the infected zone of N₂-fixing nodules; Millar
et al. 1995b). Understandably, more active cells such as those in meristems are generally equipped with larger numbers and consequently show faster respiration rates.

(b)  Mitochondrial substrates

Two substrates are produced from glycolytic PEP for oxidation in mitochondria: malate and pyruvate (Figure 2.21). These compounds are thought to be the most abundant mitochondrial substrates in vivo. However, amino acids may also serve as substrates for mitochondrial respiration in some tissues, particularly in seeds rich in stored protein. This oxidation may be preceded by transamination within the mitochondrion to produce a TCA cycle intermediate, or in some cases may occur directly. For example, most mitochondria contain glutamate dehydrogenase which oxidises glutamate to α-ketoglutarate and produces NADH (as well as ammonia). Some mitochondria also contain proline and glycine dehydrogenases, enzymes that feature in photorespiration and are largely confined to leaf mitochondria (Section 2.3). β-oxidation of fatty acids can occur in plant mitochondria, although this oxidation is slow compared to that in animal mitochondria (most fatty acid oxidation in plants occurs in microbodies).

(c) Carbon metabolism in mitochondria

Malate and pyruvate enter the mitochondrial matrix across the inner membrane via separate carriers. Malate is then oxidised by two enzymes: malate dehydrogenase (a separate isoenzyme from that in the cytosol), which yields OAA and NADH, and NAD-linked malic enzyme, which yields pyruvate and NADH and releases CO₂ (Figure 2.23).

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Figure 2.23 Tricarboxylic acid cycle with appendages. Glycolysis in the cytoplasm yields pyruvate of malate as substrates. Malic enzyme catalyses the decarboxylation of malate to pyruvate within the mitochondria. (Original drawing courtesy David Day)

Pyruvate formed either from malate or transported directly from the cytosol is oxidised by the key enzyme pyruvate dehydrogenase to form CO₂, acetyl-CoA and NADH. This enzyme, which requires coenzyme A, thiamine pyrophosphate and lipoic acid as cofactors, effectively links the TCA cycle to glycolysis. It consists of a complex of three enzymes: pyruvate dehydrogenase itself, dihydrolipoyl transacetylase and the flavoprotein dihydrolipoyl dehydrogenase. The pyruvate dehydrogenase complex has a molecular weight in the millions and is subject to sophisticated regulatory mechanisms, including phosphorylation/dephosphorylation by a kinase/phosphatase couple. Reversible phosphorylation by the kinase inactivates the enzyme and various factors regulate the kinase (e.g. pyruvate inhibits it whereas ammonia stimulates it). Pyruvate dehydrogenase is also subject to feedback inhibition from acetyl-CoA and NADH.

(d) Tricarboxylic acid (TCA) cycle

The TCA cycle proper begins with a condensation of acetyl-CoA and OAA, to form the six-carbon molecule citrate and release CoA (Figure 2.23) (a reaction catalysed by citrate synthase). Aconitase catalyses the next step, converting citrate to isocitrate. Both of these enzymes occur as isoenzymes in other cellular compartments, citrate synthase in glyoxisomes of oil seeds and aconitase in the cytosol.

NAD-linked isocitrate dehydrogenase then oxidatively decarboxylates isocitrate to form CO₂ and a-ketoglutarate, and reduce NAD⁺. The a-ketoglutarate thus formed is oxidised further to succinyl-CoA in a reaction catalysed by the enzyme α-ketoglutarate dehydrogenase. This enzyme is a complex that has similarities to pyruvate dehydrogenase and the reaction is analogous to the formation of acetyl-CoA from pyruvate. The reaction mechanisms are also very similar but α-ketoglutarate dehydrogenase is not subject to the complicated control of pyruvate dehydrogenase. Succinyl-CoA synthase then catalyses the conversion of succinyl-CoA to succinate, with the concomitant phosphorylation of ADP to ATP, the only substrate-level phosphorylation step in the mitochondrion. This enzyme in plants differs from its mammalian counterpart in that it is specific for ADP rather than GDP.

Succinate dehydrogenase (SDH), which catalyses the oxidation of succinate to fumarate, is the only membrane-bound enzyme of the TCA cycle and is part of the respiratory electron transport chain (complex II, Figure 2.24). SDH is a complex consisting of a flavoprotein and several other subunits; the former has FAD as a covalently bound cofactor and the enzyme also contains two bi-nuclear iron–sulphur clusters. Electrons from FADH₂ are passed on to ubiquinone.

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Figure 2.24 Electron transport chain of plant mitochondria. Roman numerals indicate respiratory complexes equivalent to mammalian counterparts. Complex I, NADH-UQ oxido-reductase; complex II, succinate dehydrogenase; complex III, Cyt b/c₁ complex; complex IV, Cyt c oxidase; complexV, ATP synthetase. Arabic numerals indicate special plant features: 1, external NADPH dehydrogenase; 2, internal, rotenone-insensitive NADH dehydrogenase; 3, alternative (cyanide-insensitive) oxidase; UQ, ubiquinone. Unbroken arrows indicate pathways of electron flow; broken arrows indicate proton translocation sites. (Original drawing courtesy David Day)

Fumarase catalyses the conversion of fumarate to malate and is unique to the mitochondrion, making it a convenient marker for the mitochondrial matrix. Malate dehydrogenase catalyses the final step of the TCA cycle, oxidising malate to OAA and producing NADH. The reaction is freely reversible, although the equilibrium constant strongly favours the reduction of OAA, necessitating rapid turnover of OAA and NADH to maintain this reaction in a forward direction.

Overall, during one turn of the cycle, three carbons of pyruvate are released as CO₂, one molecule of ATP is formed directly, and four NADH and one FADH₂ are produced. The latter strong reductants are oxidised in the respiratory chain to reduce O₂ and produce ATP. Although most of the TCA cycle enzymes in plant mitochondria are NAD linked, NADP-dependent isoforms of isocitrate and malate dehydrogenases also exist, and these may play a role in a protective reductive cycle in the matrix.

Regulation of carbon flux through the TCA cycle probably occurs via phosphorylation/dephosphorylation of pyruvate dehydrogenase, which will depend in turn on mitochondrial energy status and feedback inhibition of various enzymes by NADH and acetyl-CoA. The rate of cycle turnover thus depends on the rate of electron flow through the respiratory chain (to reoxidise NADH) and utilisation of ATP. TCA cycle turnover will also depend on the rate of substrate provision by reactions in chloroplasts and cytosol, and in pea leaves this may be a major limitation on the rate of respiration in vivo. For example, in an experiment to estimate respiratory chain capacity, Wiskich and Dry (1985) isolated mitochondria from pea leaves and resuspended them in a known volume of reaction medium. A large proportion of organelles is usually ruptured during isolation, and it is important to estimate yield of intact mitochondria. Therefore the volume of the leaf homogenate obtained upon disruption of the leaves was also measured, and a small aliquot set aside. A mitochondrial marker for enzyme activity (e.g. fumarase) was measured for both isolated mitochondria preparation and crude homogenate. The percentage yield of intact mitochondria was inferred from their comparative activities. Respiration rate by isolated mitochondria was then measured in an oxygen electrode with a mixture of substrates. That value was extrapolated to a mitochondrial capacity of a whole leaf by correcting for the yield of intact mitochondria. In a parallel experiment, the in vivo rate of respiration by intact leaf tissue was measured in a second oxygen electrode. Comparative values were as follows:

Rate of respiration by mitochondrial suspension: 220 µmol O₂ h^–1
Per cent yield of mitochondria: 50
Fresh mass of leaf tissue: 10 g
Respiratory capacity of tissue: 44 µmol O₂ h^–1 g^–1
Respiratory rate of intact leaf tissue: 23 µmol O₂ h^–1 g^–1

Respiratory capacity as inferred from the activity of isolated mitochondria exceeded actual measured rates of intact leaf tissue. Respiration in vivo is therefore constrained, and such restriction might be due to either substrate supply and/or the ATP/ADP ratio.

2.4.5  Respiratory chain

The respiratory chain of mitochondria consists of a series of membrane-bound redox centres which catalyse the transfer of electrons from NADH and FADH₂ to O₂, forming H₂O and translocating protons across the inner membrane (Figure 2.24). Translocation of protons is made possible by release of redox energy that accompanies electron transfer from the strong reductant NADH to the strong oxidant O₂, and is functionally linked to electron transfer. (This electron transfer involves a release in redox energy of 1.14 V which is equivalent to 52.7 kcal of chemical energy, enough to drive the synthesis of three ATP molecules.) In this way, a protonmotive force (Dµ_H⁺) is created across the inner membrane and is used to drive phosphorylation of ADP via the ATP synthase complex (Figure 2.24, right side).

Plant mitochondria have a respiratory chain which is more complicated than that of animals and contains additional NADH dehydrogenases and an alternative oxidase which catalyses cyanide-insensitive O₂ consumption. These additional enzymes (which are also found in most fungi) do not translocate protons and therefore are not linked to ATP synthesis; they are often referred to as the non-phosphorylating bypasses of the plant respiratory chain. The other complexes of the chain are common to all mitochondria and have been extensively studied in animals and fungi and to some extent in plants. They have been assigned Roman numerals by researchers of mammalian respiration (Figure 2.24).

According to structural arrangements that underlie electron transport in plant mitochondria, large protein-containing complexes of the respiratory chain are immersed in the inner membrane by virtue of their hydrophobic subunits, and interact with one another via two smaller molecules: ubiquinone and cytochrome c. The lipid-soluble ubiquinone is small enough to move rapidly along and across the membrane, and participates in H⁺ transport across the membrane as well as shuttling electrons from complexes I and II to complex III. Location and oxidation–reduction status are shown in Figure 2.25. Cytochrome c is a small haem-containing protein located on the outer surface of the inner membrane, which shuttles electrons between complexes III and IV. In this respect, the respiratory chain is similar in layout to the photosynthetic electron transport chain: three large complexes which communicate by a quinone and a small mobile protein (Cyt c or plastocyanin). However, orientation of components in the membrane is inverted and the net reaction catalysed is opposite to that in chloroplasts (Figure 1.11).

(a) NADH oxidation

Complex I, NADH-ubiquinone oxido-reductase (Figure 2.24), is a large multi-subunit complex of 30–40 polypeptides, seven of which are synthesised in the mitochondrion. One of the subunits, a 50 kDa protein, contains flavinmononucleotide as a cofactor and is the dehydrogenase which oxidises NADH and passes electrons to FeS-containing proteins of the complex, and eventually to ubiquinone. The passage of electrons through the complex is accompanied by H⁺ translocation across the membrane (mechanism poorly understood). Complex I is inhibited specifically by the flavonoid rotenone and analogues. The NADH-binding site is exposed to the matrix and the complex oxidises NADH produced by the TCA cycle and other NAD-linked enzymes.

Plant mitochondria contain another matrix-located NADH dehydrogenase which is insensitive to rotenone and does not pump protons across the membrane (called the ‘rotenone-insensitive bypass’). Plant mitochondria also oxidise NADH and NADPH by two dehydrogenases on the outside of the inner membrane. This oxidation is not inhibited by rotenone and is not linked to H⁺ translocation. These external dehydrogenases are presumed to oxidise NAD(P)H produced in the cytosol.

(b) Succinate oxidation

Complex II (Figure 2.24) is succinate dehydrogenase, which also spans the membrane and has its active site exposed to the matrix. It consists of five subunits, one of which is encoded by the mitochondrial genome in plants while the rest are synthesised in the cytosol. SDH also contains FeS and haem centres which participate in electron transfer from succinate to ubiquinone. Unlike complex I, complex II does not pump H⁺ and succinate oxidation is therefore linked to the synthesis of less ATP (see below). Malonate, an analogue of succinate, is a strong competitive inhibitor of succinate dehydrogenase.

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Figure 2.25 The protonmotive Q cycle mechanism of proton translocation at complex III of the respiratory chain. Oxidised quinone (UQ) accepts an electron from Cyt b₅₆₂ of complex III and a proton from the matrix to form the semiquinone (UQH).The semiquinone accepts another proton from the matrix and an electron from complex I (or II) to form the quinol which then diffuses across the membrane (broken arrows) to interact with Cyt b₅₆₆ and the FeS protein of complex III near the outside of the inner membrane. UQH₂ is oxidised to the semiquinone by Cyt b₅₆₆, losing a proton to the external medium in the process, which then reduces b₅₆₂. The semiquinone is oxidised by the FeS protein (not shown), losing another proton which then reduces Cyt c₁ and thence Cyt c. The quinone formed then diffuses back across the membrane to interact with Cyt b₅₆₂. In this way, one electron is transferred from complex I (or II) on one side of the membrane to Cyt c on the other, with another electron and UQ shuttling back and forth across the membrane. Concurrently, two H⁺ are translocated from the matrix to the external medium for each electron flowing to Cyt c. Broken arrows indicate diffusion of both fully oxidised and fully reduced ubiquinone; unbroken arrows indicate electron flow. (Original drawing courtesy David Day)

(c) Cytochromes

Complex III (Figure 2.24) is the cytochrome b/c₁ complex, consisting of two b-type cytochromes, b₅₆₆ and b₅₆₂, cytochrome c₁, an FeS protein named the Rieske iron–sulphur protein and several other polypeptides. The complex contains eight subunits, one of which is synthesised in the mitochondrion. Electron flow from ubiquinol to cytochrome c is accompanied by the translocation of four H⁺ per electron pair, across the membrane, via the so-called Q cycle. According to this mechanism, ubiquinone is reduced on the matrix side of the membrane by one electron from complex I or II and one from Cyt b₅₆₂. The quinol then diffuses across to the outside of the membrane to reduce Cyt c₁ and the Rieske FeS centre; the electron from c₁ is passed on to c and then in turn to cytochrome oxidase, while the FeS electron is handed on to Cyt b₅₆₆, then to b₅₆₂, which reduces ubiquinone. Thus the b cytochromes participate in the movement of electrons across the complex but do not participate in the reduction of Cyt c (Figure 2.25). Various inhibitors of complex III have been discovered, with antimycin A and myxothiazol most widely used in research.

The final complex of the main respiratory chain (Figure 2.24) is complex IV, cytochrome c oxidase. As the name implies, cytochrome c oxidase accepts electrons from cytochrome c on the outside of the inner membrane and transfers them to the inside of the membrane where O₂ is reduced to form H₂O. The complex contains 7–9 polypeptides (three of which are synthesised in the mitochondrion). Two cytochrome haem centres, a and a₃, and two copper atoms make up its redox active components. Like complex I, cytochrome oxidase is a proton pump, but the mechanism is still poorly defined.

2.4.6  Oxidative phosphorylation

When electrons are transferred from NADH to O₂, a large release of redox energy enables ATP formation in complex V of the respiratory chain (ATP synthase in Figure 2.24). Energy release associated with electron transport is conserved by H⁺ translocation across the membrane to form a protonmotive force (Δµ_H⁺) which has both an electrical (ψ) and a pH component (Δµ_H⁺ = Δψ + ΔpH). This forms the basis of the chemiosmotic theory proposed by Mitchell in 1960 and now widely accepted. See Equations 6.16a and 6.16b in Nobel 1983.

In plant mitochondria, Δµ_H⁺ exists mainly as a Δψ of 150–200 mV, with a pH gradient (ΔpH) of 0.2–0.5 units. ATP synthesis occurs as H⁺ move from a compartment of high potential (outside the membrane) to one of low potential (the mitochondrial matrix) through the ATP synthase complex. Oxidation of NADH via the cytochrome pathway has three H⁺ translocation sites associated with this process, and is linked to synthesis of up to three ATP molecules for each molecule of NADH oxidised (i.e. three ATP formed per two electrons). By contrast, both succinate and external NADH oxidation, or NADH oxidation via the rotenone-insensitive bypass, are linked to the synthesis of only two ATP molecules per two electrons, as these events are associated with only two H⁺ pumping sites.

The number of H⁺ translocated for each pair of electrons transferred from NADH to O₂ remains controversial; it will depend, in the final analysis, on the mechanism(s) by which H⁺ translocation is coupled to electron flow and this has still to be elucidated. Given the magnitude of free energy change during ATP synthesis and the magnitude of measured Δµ_H⁺, at least three H⁺ are needed per ATP synthesised. Consequently, the H⁺/2e^– ratio for oxidation of NADH must be at least 9.

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Figure 2.26 Stylised O₂ electrode recording of respiring plant mitochondria illustrating respiratory control. Oxygen consumption is measured as a function of time. The isolated mitochondria are depleted of substrates and are therefore dependent on added substrate. Addition of ADP (inorganic phosphate is in the reaction medium) allows oxidative phosphorylation to proceed, dissipating some of the protonmotive force and thereby stimulating electron transport; the enhanced rate of O₂ uptake is called State 3. When all added ADP is phosphorylated, electron transport slows to what is known as State 4 ('resting' state). Addition of more ADP stimulates O₂ uptake further, but addition of oligomycin, which blocks the ATP synthetase, lowers O₂ uptake to the State 4 rate. Addition of an uncoupler (protonophore) fully dissipates the protonmotive force and stimulates O₂ uptake; no ATP synthesis occurs in the presence of the uncoupler. When the O₂ concentration falls to zero, respiration ceases (Original drawing courtesy David Day)

(a) ATP synthase

ATP synthase is another multi-subunit complex (Figure 1 in Case study 2.1) with at least nine polypeptides, some present in multiple copies, four of which are synthesised in the mitochondrion. This massive complex spans the inner mitochondrial mem-brane and comprises two major parts: F₀, which consists of hydrophobic subunits embedded in the inner membrane and which acts as an H⁺ pore or channel; and F₁, which is hydrophilic and extends into the matrix on a ‘stalk’. F₁ contains the active site of the ATP synthase and is a reversible ATPase. When connected to F₀, F₁ can either hydrolyse ATP and drive H⁺ translocation into the intermembrane space, or, when Δµ_H⁺ drives H⁺ back into the matrix through F₀, it can synthesise ATP from bound ADP and P_i. The stalk contains a protein known as the oligomycin-sensitivity-conferring protein (OSCP) because it binds the antibiotic oligomycin which then prevents H⁺ translocation through F₀ and inhibits ATP synthesis. Therefore, adding oligomycin to mitochondria oxidising a substrate in the presence of ADP restricts O₂ uptake (Figure 2.26).

The reaction mechanism of the ATP synthase remains controversial but the most favoured hypothesis is a
‘con-formational’ model. According to this model, F₁ has three nucleotide-binding sites which can exist in three con-figurations: one with loosely bound nucleotides, one with tightly bound nucleotides and the third in a nucleotide-free state. H⁺ movement through F₀ results in rotation of F₁, causing a conformational change during which the site with loosely bound ADP and P_i is converted to one which binds them tightly in a hydrophobic pocket in which ATP synthesis occurs. Further H⁺ movement then causes another rotation of F₁ and the ATP binding site is exposed and releases the nucleotide. In the meantime, the other nucleotide-binding sites are undergoing similar changes, with ADP and P_i being bound and converted to ATP. Thus H⁺ translocation drives the three sites through three different configurations and the main expenditure of energy is in the induction of a conformational change that releases tightly bound ATP, rather than in ATP synthesis itself.

(b) Respiratory control

Electron transport through the respiratory chain, and therefore rate of O₂ uptake, is controlled by availability of ADP and P_i, a phenomenon described as ‘respiratory control’. In the absence of ADP or P_i, the proton pore of ATP synthase is blocked and a Δµ_H⁺ builds up to a point where it restricts further H⁺ translocation across the inner membrane. Since electron transport is functionally linked to H⁺ translocation, this elevated Δµ_H⁺ will also restrict O₂ uptake. That outcome is easily seen with isolated mitochondria (Figure 2.26) where O₂ uptake is stimulated by adding ADP (‘State 3’ respiration). When all of the added ADP has been consumed, O₂ uptake decreases again (‘State 4’). In steady state, the rate of electron flow is determined by the rate of flow of H⁺ back across the membrane: when ADP and P_i are available the backflow is rapid and occurs via ATP synthase; in the absence of these compounds, backflow is no more than a slow leak. The ratio of State 3 to State 4 (the respiratory control ratio) is thus an indication of coupling between ADP phosphorylation and electron transport. Larger values represent tighter coupling. The proton leak can be dramatically stimulated by some compounds which act as protonophores or proton channels; these compounds collapse the Δµ_H⁺ and increase O₂ uptake up to the State 3 rate (Figure 2.26). However, no ATP is formed and these compounds are called uncouplers because they uncouple the linked processes of electron transport and phosphorylation.

CASE STUDY 2.1  Knobs and ATP synthase

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Figure 1 Nature’s tiniest rotary motor shown here as a highly diagrammatic version of an ATP synthase complex — a device for generating ATP from free energy stored as a transmembrane protonmotive force (a gradient in electrochemical potential of protons, consisting of a pH gradient and a gradient in electric potential). The ATP synthase complex of chloroplasts was referred to earlier as a ‘coupling factor’ (Figure 1.11). (a) In mitochondria, an asymmetric portion F₀ spans the inner membrane and provides a conduit for proton movement across this inner membrane (from intermembrane space to matrix), movement that imparts a torque between two adjacent subunits ‘a’ and ‘c’.Torque so generated is transmitted to F1 via the shaft ‘γ’ plus the ‘ε’ subunit. (b) ‘a’, ‘b’ and ‘δ’ subunits are linked to the F1 hexainer (α₃β₃) and constitute the ‘stator’ of this rotary motor. The ‘rotor’ itself is represented here as a shaded portion of the overall complex, and comprises ‘c’, ‘γ’ and ‘ε’ subunits. The ‘γ’ subunit behaves as a rotating shaft that mediates an exchange of energy derived from proton flow through F₀ for ATP synthesis via the cooperative activity of three catalytic sites within F₁ (three ATP are generated for every 12 protons that pass through this rotary motor).

In a widely acclaimed technical achievement, Hiroyuki Noji and colleagues at Yokohama (Noji et. al. 1997) attached a fluorescent actin filament to the tip of a 'γ' subunit and recorded continuous rotation during synthesis of ATP, thus confirming a rotary motion that had been predicted on biophysical grounds. Unrestrained by a long actin filament, rotation rate in vivo would peak around 150 revs per second. Significantly, when provided with a source of ATR this self same device (an ATP synthase complex) can draw energy from ATP hydrolysis to pump protons against a gradient. Now working as an ATPase, these rotary motors sustain energy-dependent processes including nutrient ion uptake and salt exclusion by plant roots. (Based on Elston et al. 1998; reproduced with permission of Nature)

During the mid-1950s Marjorie Wilkins and I, both of CSIRO Food Preservation and Transport, were collaborating with John Farrant, CSIRO Industrial Chemistry. Using his electron microscope, we had seen what we then called ‘knobs’ protruding from the inner surfaces of plant mitochondria (Farrant et al. 1956).

Happily, the molecular structure of such bodies is now so well understood, thanks to the experiments of many workers around the world, that we have a good understanding of their function at a molecular level. As a retired plant physiologist, I have become specially interested in the membrane-bound F₀F₁ATPase of E. coli, also an ATP synthase. Here my thanks are due to the Frank Gibson (1991) and Graham Cox Group in John Curtain School at ANU, whose work led to my understanding the hypothesis of its structure and function.

The ‘knobs’ attached to a membrane-bound portion (Figure 1) are thought to be analogous to motors, with internal subunits that rotate at about 150 times per second relative to adjacent subunits, and with the stalk complex acting as a kind of ‘stator’. According to this rotational model, protons are pumped across the membrane. Proton pumping in one direction results in formation of ATP from ADP and phosphate; in the other direction, ATP is hydrolysed to ADP, liberating H⁺ ions, to which the movement of other ions is linked. Thanks to the techniques of molecular biology and to many workers, much of the structure is now understood. The ‘knobs’ in chloroplast membranes are similar in structure to those in mitochondria, but point the opposite way.

2.4.7  An alternative oxidase

In both plants and animals, cytochrome oxidase is sensitive to a number of inhibitors, the best known of which are carbon monoxide and cyanide. Plants, however, show resistance to both carbon monoxide and cyanide because they are equipped with an alternative oxidase. This enzyme does not translocate H⁺ and therefore is not linked to ATP formation. The enzyme is a quinol oxidase and appears to consist of one to three polypeptides of about 25–35 kDa, depending on the plant species. The proteins are encoded in nuclear genes which show tissue-specific expression. Cyanide-insensitive O₂ uptake is inhibited by hydroxamic acids (SHAM — salicyl hydroxamic acid — is the most commonly used) and n-propylgallate.

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Figure 2.27 Regulation of alternative oxidase. (a) The alternative oxidase exists as a dimer linked together by disulphide bonds. Reduction of these bonds is required to allow activation of the oxidase; this activation occurs by a direct interaction between pyruvate and the oxidase. Oxidation inactivates the oxidase even when pyruvate is present. (b) Putative feed-forward regulation of the alternative oxidase to prevent fermentation from accumulated pyruvate and formation of reactive oxygen intermediates (ROI) frona over-reduced ubiquinone (Q). In vivo, reduction of the oxidase is apparently achieved via NAD(P)H which will accumulate when carbon flux through the TCA cycle is high and/ or the cytochrome chain is inhibited. Small solid arrows indicate activation pathways; dashed arrows indicate potentially deleterious side reactions. (Original drawing courtesy David Day)

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Figure 2.28 Experimental evidence for schemes in Figure 2.27. (a) and (b) represent oxygen electrode recordings obtained with a suspension of mitochondria isolated from tobacco leaves. NADH is added as the electron donor, and ADP ensures that electron transport is not restricted by the rate of oxidative phosphorylation. Addition of KCN inhibits cytochrome oxidase and thus allows alternative oxidase activity to be measured. At times indicated by (1)-(4), samples of mitochondria were taken from the reaction vessel, their proteins separated by SDS-PAGE in the absence of reductant, and alternative oxidase protein (AOX) bands visualised by immunoblotting (shown schematically as inserts). At (1), pyruvate has been added to the mitochondria but the rate of cyanide-sensitive respiration remains low because AOX protein is largely oxidised (the covalently linked dimer is evident at 70 kD in the blot); however, when isocitrate is added at (2) to generate NADPH in the mitochondrial matrix, AOX becomes reduced (major band at 35 kD is seen in the blot) and cyanide—insensitive O₂ uptake becomes rapid as AOX is activated. At (3), isocitrate has been added and AOX is reduced, but respiration remains slow because pyruvate is not present to activate the enzyme fully. At (4), the redox status,of AOX does not change, but AOX activity increases dramatically upon pyruvate addition. Thus both a reductant such as NADPH and an activator such as pyruvate must be present before the alternative oxidase is fully engaged. In this case NADPH was generated from isocitrate via NADP-linked isocitrate dehydrogenase in the mitochondrial matrix. (Based on Vanlerberghe et al. 1995)

Activity of the alternative oxidase is regulated by a complicated feed-forward mechanism (Figure 2.27, with experimental evidence in Figure 2.28). The oxidase exists in mitochondria as a dimer which can be inactivated by covalent linkage via disulphide bonds. Activation of the enzyme involves reduction of that bond, probably via matrix NADPH in a thioredoxin-mediated reaction. The reduced (but not the oxidised) enzyme is stimulated allosterically by pyruvate and some other 2-oxo acids (such as glyoxylate), which interact directly with the oxidase.

This activation by pyruvate was discovered almost by accident during collaborative research involving Harvey Millar and David Day (Australian National University) and Joe Wiskich (University of Adelaide). It began when Jim Siedow (Duke University, USA) emailed the ANU group to say that he could not repeat their published results showing that soybean root mitochondria have alternative oxidase activity. As became apparent, this difference was due to substrate. Harvey Millar (a fourth year honours student at the time) had been using a mixture of malate plus pyruvate as TCA cycle substrates for his isolated mitochondria, whereas Jim Siedow was using succinate. Harvey and David had gone to Adelaide to use the so-called ‘Q’ electrode of Joe Wiskich to investigate apparent differences between substrates in more detail. However, they could not repeat their results from Canberra. Much to their consternation, soybean root mitochondria in Adelaide were completely sensitive to cyanide. However, the Wiskich group routinely used a mixture of malate plus glutamate to drive the TCA cycle, and when pyruvate was added to the reaction vessel, mitochondrial preparations showed a dramatic stimulation of respiration despite the presence of cyanide. Significantly, the level of ubiquinol remained unchanged, implying that this effect was not simply due to oxidation of added pyruvate. A few frantic weeks of experimentation followed, resulting in a publication by Millar et al. (1993) on organic acid activation of the alternative oxidase of plant mitochondria, and illustrating the value of global communication between research groups.

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Figure 2.29 Electron flux via either cytochrome oxidase (Cyt ox) or the alternative oxidase (Alt ox) varies according to the oxidation/reduction status of the quinone pool (Q). At low levels of reduced Q (i.e. a QH₂/Q ratio of 0.25 or thereabouts) electron flow via Cyt ox is near capacity, whereas Alt ox contributes little to the overall flux of electrons to molecular oxygen, and thus production of H₂0. The Alt ox pathway becomes progressively engaged at higher values of the QH₂/ Q ratio, and the extent of that engagement is greatly enhanced by pyruvate due to direct activation ofthe Alt ox pathway
(Original drawing courtesy Harvey Millar; derived from data in Hoefhagel et al. 1995 and fitted to the Q pool model of Van den Bergen et al. 1994).

Ubiquinol (Figure 2.23) is substrate for the alternative oxidase whose activity is also governed partly by the degree of reduction of the quinone pool. Where pyruvate is absent, the ratio of Q_reduced/Q_total (QH₂/Q ratio) must be very high to activate the oxidase; when pyruvate is present, the alternative oxidase becomes active at a much lower QH₂/Q ratio (Figure 2.29) and can compete with the cytochrome chain for electrons, and efficiency of ATP generation diminishes. In Figure 2.29, ubiquinone reduction is presented as the ratio of fully reduced ubiquinol to total ubiquinone (oxidised or reduced). Kinetics of the cytochrome pathway are effectively linear with respect to ubiquinone reduction. Kinetics of the alternative pathway are sigmoidal with significant activity only apparent once the ubiquinone pool is half reduced. When pyruvate is present, alternative pathway kinetics shift to the left, allowing greater electron flux at lower ubiquinone pool reduction levels. Under those circumstances, the alternative pathway will compete with the cytochrome pathway for reduced ubiquinone. Saturation of electron flux via either pathway is not observed in plant mitochondria as the mitochondrial dehydrogenases that provide the driving force for ubiquinone reduction usually become limiting long before either oxidase capacity is saturated. Consequently, the rate of electron flux at the steady-state level of ubiquinone reduction (ordinate in Figure 2.29) will be at the point of intersection between dehydrogenase and oxidase pathway kinetics (open circles in Figure 2.29).

These different controlling factors seem to form part of a regulatory mechanism that ensures that the alternative oxidase is ‘turned on’ when carbon flux through the cell is high or when the cytochrome chain is inhibited (e.g. by high cytosolic ATP/ADP). Under these conditions, pyruvate and reduced pyridine nucleotide levels will be relatively high, ensuring reduction and activation of the oxidase. This mechanism may point to a protective role for the alternative oxidase, preventing accumulation of pyruvate (which may lead to fermentation) and over-reduction of respiratory chain components (which may cause generation of damaging reactive oxygen species such as superoxide ions). Exposure of plants to low temperatures may cause disruption of the cytochrome path (probably via lipid phase changes) and the alternative oxidase may play a role here, since cold treatment stimulates its synthesis. Alternative oxidase synthesis is also induced by other conditions of stress including pathogen attack and ethylene-triggered processes such as fruit ripening, as well as by cytochrome chain inhibitors, all suggesting a protective role.

FEATURE ESSAY 2.2 Thermogenesis

Joe Wiskich

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Figure 1 Joe Wiskich (Botany Dept, University of Adelaide) examines spadices of Arum italicum (left) and Dracunculus vulgare (right) collected from the Adelaide Botanic Garden. Both species heat their spadices via 'futile cycles' of respiratory metabolism to volatilise aromatic compounds and attract pollinating insects.

Respiration is a combustive process and about 75% of its free energy is used to phosphorylate ADP to ATP. Cells tend to maintain very high ATP/ADP ratios and this places a limitation on the rate of respiration via allosteric inhibition of glycolysis and via respiratory control applied to the mitochondrial electron transfer chain.

Heat generation is a useful phenomenon in plants, and is used to melt snow or volatilise aromatic compounds to attract insects. To generate heat it is necessary to have inefficient ATP-utilising processes and a rapid rate of respiration. This can be achieved by ‘futile cycles’ whereby ATP and a kinase phosphorylate a metabolite and a phosphatase regenerates it. The net result of this process is the hydrolysis of ATP to ADP and P_i, producing both heat and ADP which allows for rapid respiration. Bumblebees use such a cycle in cold weather to warm up their muscles before ‘take off’. If we are cold, we tend to shiver which produces the same effect.

Another way of achieving heat production is not to make ATP in the first place. Hibernating animals have brown adipose tissue whose mitochondria contain ‘thermogenin’ (a proton-translocating protein) in their inner membranes. When this protein is functional it prevents the establishment of a protonmotive force across the inner mitochondrial membrane and the free energy of respiration is liberated as heat. We are born with some of this brown adipose tissue but lose it early in life.

Plants also produce heat, as was observed by Lamarck in 1788. The family Araceae has more than 100 genera and about 3000 species. In many aroids the fertile flowers are clustered around a central spadix which is full of respiratory fuel. This can be mainly carbohydrate (as in Arum) or a mixture of carbohydrate and fat (as in Philodendron). In these plants the spadix can increase its respiratory rate 100-fold and raise its temperature above 40°C. The actual increase can be 20–30°C depending on the ambient temperature. For example, the inflorescence of the skunk cabbage (Symplocarpus foetidus) melts the snow as it grows up through it. Generally, the aroids generate heat to volatise aromatic compounds which attract insects to facilitate cross-pollination. The process plants use for this purpose differs from the ‘thermogenin’ protein of brown adipose tissue but achieves the same result. The mitochondria of thermogenic plants contain an ‘alternative oxidase’ which branches from the mitochondrial electron transfer chain at ubiquinone. This is the alternative path and differs from the cytochrome path in that it does not conserve energy (to produce ATP) and is insensitive to cyanide and other inhibitors of cytochrome oxidase.

The aroid spadices with their heat production, rapid respiration rates and complete insensitivity to cyanide were considered to be the ‘type-specimens’ for cyanide-insensitive respiration. However, other plants showed only partial sensitivity to cyanide and these were regarded as ‘cyanide resistant’. Even those tissues completely sensitive to cyanide could be ‘induced’ to become cyanide resistant, for example fresh potato tuber slices are sensitive but become resistant on ageing. Thus, the alternative path is a feature potentially available to all higher plant mitochondria and at some stage during the development of any tissue is likely to become important. Inducing the alternative path really means inducing or activating the terminal ‘alternative oxidase’.

In 1987 Pierre Rustin questioned the nature of the alternative oxidase; was it a ubiquinol oxidase? Were we measuring quinol auto-oxidation? Could it be a lipoxygenase or was it a ‘free radical’ mechanism? Some of the problems we had at that stage were: first, the reactivity of the alternative oxidase to some substrates was different in mitochondria from thermogenic compared to non-thermogenic tissues; second, the activity could only be solubilised from thermogenic mitochondria; third, attempts to purify it were only partially successful; and, finally, some results were being interpreted in terms of different ubiquinone pools existing within the membranes so that the NADH from some matrix enzymes (NAD-malic enzyme) had better access to the alternative oxidase than others (NAD-malic dehydrogenase). Since then a number of significant events have occurred.

My laboratory was investigating photorespiratory metabolism in pea leaf mitochondria and we had difficulty interpreting some of our results. We could explain our data if we assumed either a mixed population of mitochondria or a differential access of NADH generated by the oxidation of glycine to the electron transport chain compared to the NADH generated by malate oxidation. We eliminated the first possibility with some immunogold-labelling studies, and it was to test the second that Tony Moore (University of Sussex, UK) come to my laboratory with a so-called Q electrode. This measured the redox state of the inner-membrane pool of ubiquinone-10, using a more water soluble quinone analogue as a mediator.

David Day (Australian National University) also visited at this time and brought with him some soybean mitochondria; these have a reasonable degree of alternative path activity. So for the first time we were able to measure simultaneously the rate of alternative oxidase activity and the redox state of its substrate. The initial results were quite clear — the alternative oxidase showed little activity until the ubiquinone pool was about 50% reduced and increased quite markedly above that. The cytochrome path became active as soon as some ubiquinone was reduced and reached apparent saturation at 20–30% reduction. These results appeared to validate, in general — but not precise — terms, the Bahr and Bonner hypothesis that the cytochrome path had to be saturated before any flux through the alternative path could be observed.

However, we still had problems. In soybean mitochondria succinate and NADH reduced the ubiquinone pool to the same extent, yet the alternative path oxidised succinate much more rapidly. Reduced quinone analogues were also poor substrates. It was Harvey Millar in David Day’s laboratory who noticed that malate oxidation via the alternative oxidase was much faster if pyruvate, rather than glutamate, was used to remove the oxaloacetate. It was soon established that pyruvate and other 2 oxo-acids such as glyoxylate, oxaloacetate and 2-oxoglutarate activated the alternative oxidase. In the presence of pyruvate, soybean mitochondria oxidised NADH and quinol analogues via the alternative oxidase. So the real problem was the availability of pyruvate — substrates which could produce pyruvate and activate the alternative oxidase were more readily oxidised than those that didn’t. This solution to one problem highlighted another. When we repeated our analyses on the relationship between the redox state of the ubiquinone pool and rates of O₂ uptake it became obvious that the alternative oxidase was very active at a relatively low level of ubiquinone reduction. This meant that the alternative path was now competing with the cytochrome path and electron flow could ‘switch’ from one pathway to the other. A wealth of data on estimating the contribution of the alternative oxidase to tissue respiration (so-called ‘engagement’ or ‘rho’ determinations) now had to be treated with great caution. This is because the technique used inhibitors and assumed electron flow could not switch from the alternative to the cytochrome path. Further, any inhibition of the electron transfer chain in tissues could lead to an increase in the concentration of pyruvate.

We believe that pyruvate activation eliminates the problems associated with preferential oxidation of substrates via the alternative oxidase. There are still differences in kinetics of the alternative oxidase with respect to the redox state of ubiquinone among mitochondria from different tissues. However, I feel that these should be considered in terms of the balance between input and output from the ubiquinone pool. When Peter Rich (Glynn Research Institute, Bodmin, UK) visited my laboratory he brought some very potent alternative oxidase inhibitors with him. Using these we estimated that Arum and soybean mitochondria contain 720 and 58 pmol alternative oxidase mg^–1 mitochondrial protein. Clearly a mitochondrion with a high amount of the enzyme will maintain faster rates of O₂ uptake at lower levels of reduced ubiquinone.

Meanwhile, advances were being made in North America. Tom Elthon, in Lee McIntosh’s laboratory (Michigan State University,) had produced an antibody to alternative oxidase which cross-reacted with proteins from a wide range of plants. The alternative oxidase, when reduced, was detectable on electrophoresis gels in the range of 32 to 37 kDa. There appears to be three separate protein forms which exist in varying combinations in different tissues. Subsequently, David Rhoads and Lee McIntosh isolated the genes from both thermogenic and non-thermogenic tissues. Thus, the alternative oxidase finally reached the status of being a real protein, a nuclear-encoded enzyme. From Jim Siedow’s laboratory (Duke University, North Carolina), Ann Umbach reported that the enzyme could exist as an inactive oxidised dimer bringing attention to regulation by its redox state, involving sulphydryl–disulphide interactions. This has physiological implications. David Day and I received an Australian government DITAC Collaborative Research Award to visit Lee McIntosh, who had produced transgenic tobacco plants over- and under-expressed in alternative oxidase protein. Although mitochondria isolated from the leaves of over expressed plants contained more alternative oxidase protein than did wild type, their activity was the same. However, full activity was elicited by reducing the enzyme, which could be achieved by adding citrate. We suggest that NADP-isocitrate dehydrogenase produces NADPH which reduces the enzyme, most probably via a thioredoxin-type process.

We now have a feed-forward system to activate the alternative oxidase. It depends on an increase in the supply of mitochondrial substrate which can both reduce the alternative oxidase and activate it. In normal metabolism pyruvate appears to be important and this is usually considered to arise from pyruvate kinase but it must be remembered that plant mitochondria contain NAD-malic enzyme and can produce their own pyruvate from any Kreb’s cycle intermediate. During photorespiration in C₃ leaves the supply of glycine to mitochondria is very rapid. Assuming the photorespiratory rate to be about 25% of net CO₂ fixation the generation of reducing power within the mitochondria would be two to three times that of dark respiration. The fate of this reducing power is problematical — some finding its way to the peroxisomes for hydroxypyruvate reduction and some being oxidised by the electron transport chain. If there are problems in eliminating the reducing power, glyoxylate concentrations would rise and activate the alternative oxidase.

The role of the alternative path in thermogenic aroids has been mentioned. In other plants, its gene expression is induced by cold, so it may have a general role in warming plant tissues. However, it is present in all plants and can be induced by heat, drought, nutrient deficiency, insect and fungal attack, treatment with poisons — in fact any stressful situation — so it must have a more general role as well.

I recall some scientists considering the alternative path to be ‘wasteful’ respiration and who grew plants in the presence of alternative oxidase inhibitors. Presumably, they expected to get bigger and better plants; another great idea ruined by an ugly fact — the plants died. It seems to me that the alternative oxidase is present perhaps to generate some heat (certainly among some plant scientists if not plants), but also to maintain the redox state of the cell at some maximum level of reduction. What causes the tissues to become over-reduced is secondary and of little consequence. Once the tissue reaches a critical value of reduction the alternative oxidase swings into action and if there is insufficient enzyme the gene is signalled to produce more. Over-reduction can lead to the production of deleterious superoxides. The alternative path allows the respiratory pathways to produce intermediates without being subjected to severe adenylate control.

Further reading

Day, D.A., Whelan, J., Millar, A.H., Soedow, J.N. and Wiskich, J.T. (1995). ‘Regulation of the alternative oxidase in plants and fungi’, Australian Journal of Plant Physiology, 22, 497–509.

McIntosh, L. (1994). ‘Molecular biology of the alternative oxidase’, Plant Physiology, 105, 781–786.

Meeuse, B.J.D. (1975). ‘Thermogenic respiration in aroids’, Annual Review of Plant Physiology, 26, 117–126.

2.4.8  Interactions between mitochondria and chloroplasts

One dominant interaction between mitochondria and chloroplasts involves metabolite exchange as part of the photorespiratory carbon/nitrogen cycle, but more subtle interactions also occur. For example, respiratory ATP production in light is required to maintain maximum photosynthetic activity (Krömer 1995). Experimentally, if oligomycin is used to disrupt oxidative phosphorylation of mitochondria in leaf cells, photosynthesis is inhibited even though oligomycin is specific to mitochondrial ATP synthase at the concentrations applied (Krömer 1995). Clearly, a link must exist between mitochondrial ATP synthesis and photosynthesis via the energy demands of sucrose synthesis (because of sucrose phosphate synthase). A decline in supply of ATP decreases the rate of sucrose synthesis and this affects the rate of photosynthetic metabolism in chloroplasts. Using a different approach to this same issue, Shyam et al. (1993) inhibited production of mitochondrial ATP with azide and/or the uncoupler FCCP and exacerbated photoinhibition in pea leaves. Their results confirm an interplay between chloroplasts and mitochondria, and suggest a protective role for mitochondria, perhaps by provision of energy for chloroplast repair.

Leaf respiration rates are commonly measured in darkness, as either CO₂ release or O₂ consumption. Such measurements are complicated in light by reverse gas exchange from photosynthesis. However, a family of A : p_i curves can be constructed to reveal a convergence point where leaf gas exchange is independent of photon irradiance, leading to a conclusion that CO₂ release is inhibited in light relative to that in dark (Brooks and Farquhar 1985). Such inhibition of CO₂ release in light involves a decrease in carbon flow through the TCA cycle and does not seem to involve photorespiration. In contrast, respiratory O₂ consumption (excluding that associated with glycine decarboxylation) is stimulated in light compared to that in darkness. Put another way, while TCA cycle activity is decreased in light, electron transport may increase.

This apparent paradox can be explained. During active photosynthesis (non-photoinhibitory conditions), mitochondria in a leaf cell are able to oxidise surplus redox equivalents arising from photosynthetic electron transport. Those redox equivalents are then exported from chloroplasts via the malate–OAA shuttle or the DHAP–PGA shuttle (Krömer and Heldt 1991). Under these conditions, mitochondria would oxidise cytosolic NAD(P)H rather than that generated by the TCA cycle.

Coincidentally, the importance of exporting photosynthetic reducing equivalents to the mitochondria appears to increase during cold hardening (Hurry et al. 1995) and may assist in preventing photoinhibition. It is also possible that the export of α-ketoglutarate from the mitochondrion for nitrate reduction (see above) may lead to a decrease in CO₂ release from the TCA cycle; nitrate reduction is greater in light because of a need for photosynthetic reduction of nitrite, the next step in this reaction sequence leading to amino nitrogen.

2.4.9  Energetics of respiration

(a)  Efficiency

Respiration represents a substantial loss of carbon from a plant, and under adverse conditions can be as high as two-thirds of the carbon fixed daily in photosynthesis. Both the rate and the efficiency of respiration will therefore affect plant growth significantly. The overall process of respiration results in the release of a substantial amount of energy which may be harnessed for metabolic work. In theory, the energy released from the complete oxidation of one molecule of glucose to CO₂ and H₂O in respiratory reactions leads to the synthesis of a net equivalent of 36 molecules of ATP. However, in plants, because there are alternative routes for respiration, this yield can be greatly reduced.

Mechanisms for regulating respiration rates in whole plants remain unclear. Convention has it that the rate of respiration is matched to the energy demands of the cell through feed-back regulation of glycolysis and electron transport by cytosolic ATP/ADP. However, since plants have non-phosphorylating bypasses in their respiratory chain that are insensitive to ATP levels, and since PEP carboxylase and PFP might be involved in sucrose degradation, the situation in vivo is not so simple. For example, the rotenone-insensitive bypass of complex I requires high concentrations of NADH in the matrix before it can operate and seems to be active only when substrate is plentiful and electron flow through complex I is restricted by lack of ADP. Alternative oxidase activity also depends on carbon and ADP availability. In other words, non-phosphorylating pathways act as carbon or reductant ‘overflows’ of the main respiratory pathway and will only be active in vivo when sugar levels are high and the glycolytic flux rapid, or when the cytochrome chain is inhibited during stress. In glycolysis, the interaction between environmental signals and key regulatory enzymes, as well as the role of PFP and its activator fructose-2,6-P₂, will be important.

(b)  Allocation of respiratory energy to process physiology

One way of viewing respiratory cost for plant growth and survival is by subdividing measured respiration into three components associated with (1) nutrient acquisition, (2) growth and (3) maintenance. Such conceptual distinctions are somewhat arbitrary, and these categories of process physiology must not be regarded as three discrete sets of biochemical events. Such energy-dependent processes are all interconnected to some extent because ATP represents a universal energy currency for all three, while a common pool of substrates is drawn upon in sustaining production of that ATP (Amthor 1989). Nevertheless, cells do vary in their respiratory efficiency, while genotype × environment interactions are also evident in both generation and utilisation of products from oxidative metabolism. Such variation has implications for growth and resource use efficiency (Section 6.5).