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Critical Reviews in Plant Sciences, 25:159–198, 2006 c Taylor & Francis Group, LLC Copyright ISSN: 0735-2689 print / 1549-7836 online DOI: 10.1080/07352680600563876

The Functional Organization and Control of Plant Respiration William C. Plaxton Departments of Biology and Biochemistry, Queen’s University, Kingston, Ontario, K7L 3N6, Canada

Florencio E. Podestá CEFOBI, University of Rosario, Suipacha 531, Rosario 2000, Argentina Referee:

Dr. Greg C. Vanlerberghe, University of Toronto at Scarborough, Department of Life Sciences and Department of Botany, 1265 Military Trail, Scarborough, ON Canada M1C 1A4

Table of Contents I.

INTRODUCTION ............................................................................................................................................. 160 A. The Multi-Faceted Functions of Plant Respiration .......................................................................................... 161 1. ATP Production ................................................................................................................................. 161 2. Production of Biosynthetic Precursors ................................................................................................. 161 3. Carbon-Nitrogen Interactions .............................................................................................................. 162 4. Optimizing Photosynthesis ................................................................................................................. 162 5. Stress Acclimation ............................................................................................................................. 163 6. Reactive Oxygen Species and Programmed Cell Death .......................................................................... 164 7. Fruit Ripening ................................................................................................................................... 164 8. Thermogenesis .................................................................................................................................. 164 B. Mitochondria Are Dynamic Structures in Living Plant Cells ........................................................................... 165

II.

THE FUNCTIONAL ORGANIZATION OF PLANT RESPIRATION ............................................................... 165 A. Pyrophosphate: An Alternate Energy Donor of Plant Cells .............................................................................. 166 1. Cytosolic PPi Metabolism .................................................................................................................. 166 2. Mitochondrial PPi Metabolism ............................................................................................................ 169 B. Plant Mitochondria Can Respire Alternative Substrates to Pyruvate .................................................................. 170 1. Malate .............................................................................................................................................. 170 2. Formate ............................................................................................................................................ 170 3. Amino Acids and Fatty Acids ............................................................................................................. 170 C. Non-Energy Conserving Bypasses of Mitochondrial Electron Transport Contribute to the Unique Flexibility of Plant Metabolism ..................................................................................................................................... 171 1. Non-Proton Pumping Alternate NAD(P)H Dehydrogenases ................................................................... 171 2. Non-Proton Pumping Alternative Oxidase ............................................................................................ 172 3. Uncoupling Protein ............................................................................................................................ 172 D. The Adaptive Value of Adenylate- and Pi-Independent Bypasses of Cytosolic Glycolysis and Mitochondrial Respiration Is Illustrated by Pi-Starved Plants ...................................................................................................... 173 1. Glycolytic Bypasses of Pi-Starved Plants ............................................................................................. 173 2. Mitochondrial Electron Transport Chain Bypasses of Pi-Starved Plants .................................................. 174 3. Transcript Profiling Indicates that Pi Starvation Inducibility of Glycolytic and Mitochondrial Electron Transport Bypass Proteins Is Widespread ............................................................................................. 174 E. Efficient Plant Respiration May Be Promoted by Glycolytic and TCA Cycle Multi-Enzyme Metabolons, and Mitochondrial Electron Transport Protein Supercomplexes .............................................................................. 174 1. Glycolytic and TCA Cycle Metabolons ................................................................................................ 175

Address correspondence to William Plaxton, Department of Biology, Queen’s University, Kingston, Ontario, K7L 3N6, Canada. E-mail: [email protected]

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´ W. C. PLAXTON AND F. E. PODESTA

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F.

2. Respirasome and ATP Synthasome Supercomplexes of the Mitochondrial Inner Membrane ..................... 176 Tissue and/or Developmental-Specific Isozymes of Plant Respiratory Enzymes ................................................. 176

III.

METABOLIC CONTROL OF PLANT RESPIRATION .................................................................................... 177 A. Coarse versus Fine Control of Plant Respiration ............................................................................................. 177 1. Coarse Control .................................................................................................................................. 177 2. Fine Control ...................................................................................................................................... 178 B. Metabolic Control Analysis .......................................................................................................................... 178 C. Key Control Enzymes of Plant Respiratory Carbon Metabolism ....................................................................... 179 D. Specific Mechanisms of Fine Metabolic Control as Applied to Plant Respiration ............................................... 180 1. Fine Control #1: Variation in Substrate Concentration ........................................................................... 181 2. Fine Control #2: Variation in pH ......................................................................................................... 181 3. Fine Control #3: Allosteric Effectors ................................................................................................... 181 4. Fine Control #4: Reversible Covalent Modification ............................................................................... 183 4.1. Phosphorylation-Dephosphorylation ..................................................................................... 184 4.2. Disulfide-Dithiol Interconversion ......................................................................................... 186 4.3. S-Nitrosylation ................................................................................................................... 187 E. Overview of the Fine Control of Plant Respiratory Carbon Metabolism ............................................................ 187 1. Glycolysis ......................................................................................................................................... 187 2. PDC and TCA Cycle .......................................................................................................................... 188

IV.

CONCLUDING REMARKS ............................................................................................................................. 188

ACKNOWLEDGMENTS ........................................................................................................................................... 189 REFERENCES .......................................................................................................................................................... 189

The respiratory pathways of glycolysis, the tricarboxylic acid (TCA) cycle, and mitochondrial electron transport chain (miETC) are central features of carbon metabolism and bioenergetics in aerobic organisms. Respiration is essential for growth, maintenance, and carbon balance of all plant cells. Although the majority of respiratory enzymes are common to all organisms, plant respiration has evolved as a complex metabolic network endowed with a wide variety of unique characteristics. Plants have the option of employing alternative enzymes that bypass several of the conventional steps in cytosolic glycolysis, the TCA cycle, and miETC. The extent and conditions under which these bypasses operate is the subject of intensive research. The highly flexible nature of respiratory metabolism in plants has likely evolved in response to the crucial biosynthetic role played by respiration beyond its role in ATP generation; both functions must proceed if plants are to survive under varying and often stressful environmental and nutritional conditions. Additional complexity arises due to the existence of tissue- and/or developmental-specific isozymes of many plant respiratory enzymes, as well as the extensive interactions between photosynthesis and respiration, and plastidic, cytosolic, and mitochondrial metabolism in general. Recent progress in biochemistry, physiology, cell biology, genomics, transcriptomics, proteomics, metabolomics, and in vivo flux analyses have resulted in exciting new insights into many aspects of plant respiratory metabolism. Experiments on transgenic or mutant plants possessing significantly elevated or reduced levels of respiratory enzymes are augmenting

our understanding of the functions, organization, and control of plant respiration. Metabolic engineering of plant respiration is of significant practical interest as it provides both an important approach to enhancing crop yields, as well as a potential mechanism for mitigating global climate change due to elevated atmospheric CO2 levels. Keywords

I.

glycolysis, mitochondrial electron transport chain, phosphoenolpyruvate, pyruvate dehydrogenase complex, regulatory protein phosphorylation, thioredoxin, tricarboxylic acid cycle

INTRODUCTION Plants employ photosynthesis to harness light energy for the oxidation of H2 O into O2 and the simultaneous reduction of CO2 into carbohydrates, typically stored as the insoluble polysaccharide starch or soluble disaccharide sucrose. Conversely, respiration usually involves the controlled oxidation of these highly reduced carbohydrates via the sequential pathways of glycolysis and the mitochondrial tricarboxylic acid (TCA) cycle, producing CO2 and reducing equivalents (NAD(P)H and FADH2 ). In aerobic respiration, the NAD(P)H and FADH2 generated during carbohydrate oxidation transfer their electrons to O2 via the mitochondrial electron transport chain (miETC), resulting

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in respiratory O2 consumption and ATP production (Siedow and Day, 2000). Metabolite movement across the mitochondrial inner membrane (i.m.) is catalyzed by a series of integral i.m. transporter proteins that operate as specific exchangers or cotransporters for phosphate (Pi), adenine nucleotides, mono-, di-, and tricarboxylates, amino acids, and cofactors such as NAD+ and coenzyme-A (CoA) (Picault et al., 2004; Tegeder and Weber, 2005). Respiration forms the core of plant intermediary metabolism and thus plays a pivotal role in the growth and metabolism of all photosynthetic organisms (Krömer, 1995; Raghavendra and Padmasree, 2003). There are many distinguishing attributes in the organization and control of plant respiratory metabolism. These include the existence of parallel glycolytic pathways in the cytosol and plastid, alternative “bypass” enzymes in cytosolic glycolysis and the TCA cycle, and the presence of both phosphorylating and non-energy conserving (i.e., non-ATP producing) pathways of miETC (Plaxton, 1996, 2004a, 2005; Givan, 1999; Siedow and Day, 2000; Podestá, 2004; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). The remarkable flexibility of plant respiration has been suggested to have evolved as an essential adaptation that helps plants to acclimate to unavoidable stresses that they are exposed to in their natural environment (Black et al., 1987; Theodorou and Plaxton, 1993; Møller, 2001; Podestá, 2004; McDonald and Vanlerberghe, 2005; Plaxton, 2005). Plant biomass production is ultimately determined by the ratio between photosynthetic CO2 assimilation and respiratory CO2 release. Depending upon the plant species and growth conditions, as much as 70 percent of photosynthetically generated carbohydrate may be respired in the same day (Millenaar and Lambers, 2003). Respiration plays a major role in the global carbon cycle as 40 to 60 percent of total CO2 assimilated annually by photosynthetic terrestrial plants is released back into the atmosphere by respiring plants, representing about 50 percent of the annual input of CO2 from terrestrial ecosystems (Amthor, 1997, 2000; Atkin and Tjoelker, 2003; Gifford, 2003; Gonzalez-Meler et al., 2004; Atkin et al., 2005). Terrestrial plant respiration releases about 10 times more CO2 per year than anthropogenic fossil fuel combustion (Amthor, 1997; Gifford, 2003). Therefore, the metabolic engineering of plant respiration provides both an important practical approach to enhancing crop yields as well as a potential mechanism for mitigating global climate change due to elevated atmospheric CO2 levels. Despite the opposing nature of photosynthesis and respiration, tight metabolic controls ensure that futile cycling between these pathways is minimized (Krömer, 1995; Raghavendra and Padmasree, 2003). Numerous advances in our understanding of plant respiration have been and will continue to be achieved by following the classical reductionist approach of isolating and characterizing its individual components. However, analysis of the organization and control of plant respiration in the emerging post-genomic era has been complicated by the unexpectedly huge number of annotated genes that encode the constituent proteins (Fernie et al., 2004). Recent progress in biochemistry, physiology, cell biol-

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ogy, genomics, proteomics, metabolomics, in vivo flux analyses, and metabolic modeling have resulted in exciting new insights into plant respiratory metabolism (Amthor, 2000; Giersch, 2000; Roessner et al., 2002; Fernie et al., 2004; Schwender et al., 2004; Sweetlove and Fernie, 2005). A number of reviews and book chapters concerning various aspects of plant respiration have appeared over the past decade or so (Huppe and Turpin, 1994; Krömer, 1995; Plaxton, 1996, 2004a, 2005; Hill, 1997; Givan, 1999; Plaxton and Carswell, 1999; Cannell and Thornley, 2000; Siedow and Day, 2000; Siedow and Umbach, 2000; Møller, 2001; Vanlerberghe and Ordog, 2002; Atkin and Tjoelker, 2003; Gifford, 2003; Millenaar and Lambers, 2003; Raghavendra and Padmasree, 2003; Fernie et al., 2004; Gonzalez-Meler et al., 2004; Hourton-Cabassa et al., 2004; Podestá, 2004; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). The purpose of this review is to provide a critical and up-to-date overview of the roles, organization, and control of plant respiration. We emphasize how the unique features of plant respiratory metabolism may facilitate plant survival in an ever-changing and frequently stressful natural environment. The models presented are intended to provide a framework for future studies in this important field. A. The Multi-Faceted Functions of Plant Respiration 1. ATP Production As in photosynthesis, respiratory electron transfer through the miETC releases a large amount of free energy, which may be used to generate a proton electrochemical gradient across the inner mitochondrial i.m. This “proton motive force” (PMF) can be harnessed by the ATP synthase complex of the i.m. to drive the otherwise endergonic conversion of ADP and inorganic phosphate (Pi) into ATP. Plant respiratory O2 consumption is partly controlled by the availability of ADP and Pi, and by the presence of additional protein complexes that can allow respiration to proceed without generating PMF across the i.m. or that uncouple PMF dissipation from ATP synthesis (Siedow and Day, 2000; Møller, 2001; Hourton-Cabassa et al., 2004; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). 2.

Production of Biosynthetic Precursors A significant proportion of the carbon that enters the primary pathways of plant respiration is not oxidized to CO2 and H2 O, but may be withdrawn for the biosynthesis of numerous compounds such as secondary metabolites, isoprenoids, amino acids, and fatty acids. The anabolic role of glycolysis and the TCA cycle is particularly important in actively growing tissues (Hill, 1997). Export of TCA cycle intermediates requires import of substrates that can generate both acetyl-CoA and oxaloacetate (OAA) (Tegeder and Weber, 2005) (Figure 1). If pyruvate were supplied as the only substrate, export of TCA cycle intermediates would negate OAA regeneration and bring the TCA cycle to a halt. Phosphoenolpyruvate (PEP) carboxylase (PEPC) is a ubiquitous and tightly regulated plant cytosolic enzyme that catalyzes a crucial anaplerotic reaction to replenish

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FIG. 1. Interactions between carbon and nitrogen metabolism involves three compartments in plants. This scheme highlights the pivotal role of the two terminal enzymes of glycolysis, PKc and PEPC, in controlling the provision of the mitochondria with respiratory substrates as well as for generating TCA cycle C-skeletons for NH4 + assimilation via GS/GOGAT in plastids and aspartate aminotransferase (AAT) in the cytosol. The coordinate control of PKc and PEPC by allosteric effectors, particularly L-glutamate and L-aspartate, provides a mechanism for the control of cytosolic glycolytic flux and PEP partitioning during and following NH+ 4 -assimilation as discussed elsewhere in the text. Abbreviations are as defined in text and as follows: Pyr, pyruvate; Cit, citrate; Mal, malate; Fdred and Fdox , reduced and oxidized ferredoxin, respectively.

TCA cycle intermediates that are withdrawn for biosynthesis or N-assimilation (Figure 1) (Vanlerberghe et al., 1990; Huppe and Turpin, 1994; Chollet et al., 1996; Plaxton, 1996; Moraes and Plaxton, 2000; Blonde and Plaxton, 2003; Lepiniec et al., 2003; Izui et al., 2004; Britto and Kronzucker, 2005; Foyer et al., 2005; Nimmo, 2005). Carbon-Nitrogen Interactions The assimilation of inorganic N, in the form of ammonia (NH+ 4 ), into organic nitrogenous molecules is a pivotal metabolic process in plants. Because NH+ 4 is toxic it must be rapidly assimilated into the biocompatible amino acids glutamine (Gln) and glutamate (Glu) via Gln synthetase (GS) and Glu 2-oxoglutarate (2-OG) aminotransferase (GOGAT) (Lancien et al., 2000; Foyer et al., 2005). N-assimilation via GS/GOGAT is essential to life, since the Glu and Gln produced are primary amino donors for the biosynthesis of all nitrogenous compounds needed by growing plants, including amino acids/proteins, nucleic acids, chlorophylls, polyamines, and many important secondary metabolites such as alkaloids. N-assimilation must closely interact with respiratory carbon metabolism, since GS requires ATP and GOGAT requires C-skeletons and reductant in the form of 2OG and reduced ferredoxin or NADH, respectively (Huppe and Turpin, 1994; Lancien et al., 2000; Stitt et al., 2002; Britto and Kronzucker, 2005; Foyer et al., 2005) (Figure 1). Over 50 percent of net plant carbon may be committed to N-assimilation in some tissues in order to generate C-skeletons and energy (ATP and reductant) required for the GS/GOGAT system (Huppe and

Turpin, 1994). Respiration of stored and/or translocated photosynthate provides energy in non-green tissues, whereas photosynthesis usually supplies energy in photosynthetic cells. In all cells, however, an increased rate of N-assimilation necessitates enhanced carbohydrate flux through respiratory pathways since the TCA cycle is the only net source of C-skeletons (2-OG) for the GOGAT enzyme (Figure 1).

3.

4.

Optimizing Photosynthesis Illuminated leaves simultaneously assimilate CO2 through the photosynthetic carbon reduction (Calvin-Benson) cycle and lose CO2 through photorespiration and day respiration. In darkness, leaves can no longer assimilate CO2 via the Calvin-Benson cycle but produce CO2 and ATP through dark respiration. However, mitochondrial metabolism, particularly the miETC and oxidative phosphorylation, are essential for optimizing photosynthetic CO2 fixation in the light (Krömer, 1995; Raghavendra and Padmasree, 2003; van Lis and Atteia, 2004). Many studies examining the interactions of photosynthesis with respiration have focused on how photosynthesis might modulate the rate of respiration. For example, light directly influences the miETC via photoreceptor-mediated transcriptional control, likely for supporting photosynthetic metabolism (Escobar et al., 2004). Respiration optimizes chloroplast photosynthesis in numerous ways including: (i) facilitating export of excess reducing equivalents from the chloroplast, (ii) accelerating photosynthetic induction, and (iii) supplying ATP for cytosolic anabolic processes (Krömer, 1995; Raghavendra and Padmasree, 2003). Moreover,

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there is a large discrepancy between the generation and utilization of reductant through photosynthetic electron transport chain and the Calvin-Benson cycle, respectively. The miETC plays an essential role to dissipate the excess reducing equivalents produced by photochemical reactions, thereby preventing over-reduction of photosynthetic electron transport components and consequent oxidative damage to thylakoid membranes (Raghavendra and Padmasree, 2003; van Lis and Atteia, 2004). Examination of transgenic or mutant plants defective in key components of the miETC has provided definitive evidence of the crucial in vivo role that the miETC plays in photosynthetic metabolism. For example, a tobacco CMSII mutant, in which Complex I (the major NADH dehydrogenase of the miETC) is nonfunctional, was demonstrated to exhibit impaired photosynthetic performance, particularly under photorespiratory conditions (Dutilleul et al., 2003). It was concluded that Complex I is required to avoid redox disruptions of photosynthesis under conditions where leaf mitochondria must simultaneously oxidize both respiratory and photorespiratory substrates (i.e., pyruvate and glycine, respectively). A key function of mitochondrial respiration during photosynthesis, particularly in C3 leaves, is the oxidation of glycine produced in the photorespiratory pathway. During photorespiration, a massive flux of carbon passes through the glycine decarboxylase complex within the mitochondrial matrix, producing large amounts of NADH (Douce et al., 2001; Raghavendra and Padmasree, 2003; van Lis and Atteia, 2004; McDonald and Vanlerberghe, 2005). Although a significant proportion of this NADH may be exported to the cytosol via redox shuttles, much of it must be reoxidized to NAD+ by the miETC. Transgenic potato plants with an antisense reduction in glycine decarboxylase were recently employed to assess the interaction between photorespiration and respiration (Bykova et al., 2005). As expected, mitochondria isolated from these plants exhibited a decreased capacity for glycine oxidation and glycine accumulated in the leaves. The lower capacity for leaf photorespiration was compensated for by increased respiratory decarboxylations (i.e., TCA cycle flux) in the light. This was interpreted as a decreased light suppression of the TCA cycle in the glycine decarboxylase-deficient plants (Bykova et al., 2005). Tcherkez and co-workers (2005) employed 12 C/13 C stable isotope techniques to assess the in vivo respiratory metabolism of illuminated Phaseolus vulgaris leaves fed with 13 C-enriched glucose (Glc) or pyruvate. It was shown that the TCA cycle was reduced by 95 percent in the light compared to the dark decarboxylation rate. These data are in accordance with those of Hanning and Heldt (1993) who reported that mitochondria extracted from illuminated leaves exhibited a low metabolic flux through the TCA cycle. Analysis of transgenic plants exhibiting reduced expression of TCA cycle enzymes has underscored the importance of TCA cycle function in the control of photosynthesis (Carrari et al., 2003; Fernie et al., 2004; Nunes-Nesi et al., 2005). Leaves containing reduced levels of mitochondrial aconitase or malate de-

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hydrogenase have both been characterized by reduced levels of TCA cycle intermediates and an elevated rate of photosynthesis and total plant dry matter accumulation (Carrari et al., 2003; Nunes-Nesi et al., 2005). The results indicate that: (i) the TCA cycle normally competes with the sucrose biosynthetic pathway for carbon, (ii) TCA cycle flux coupled with oxidative phosphorylation is not an important ATP source for cytosolic sucrose synthesis, and (iii) the influence of respiration on photosynthesis is greater than once imagined. Mitochondrial functions in leaves in the light are important components of a highly flexible metabolic system in plants. This flexibility is a necessary adaptation to the variable conditions that plants typically encounter during growth in their natural environment. For example, owing to the action of various fine metabolic controls (i.e., allosteric effectors and reversible covalent modification of key control enzymes) glycolytic and TCA cycle flux and overall respiration rate of illuminated leaves or green algae can be rapidly and dramatically enhanced in response to various environmental perturbations (such as following resup+ ply of NO− 3 or NH4 to N-limited leaves/algae) (Paul et al., 1978; Hammel et al., 1979; Vanlerberghe et al. 1992; Huppe and Turpin, 1994)). 5.

Stress Acclimation Owing to their sessile lifestyles, plants have evolved numerous unique and sophisticated adaptations to help them cope with the inevitable abiotic and biotic stresses that are imposed upon them. Respiratory metabolism plays a crucial role in the ability of plants to acclimate at the biochemical level to a wide variety of stresses including wounding, low temperature, hypoxia, UV irradiation, heavy metal exposure, salinity and water stress, nutrient deprivation, oxidative stress, and pathogen infection (Norman et al., 1994; Plaxton, 1996, 2004a, 2005; Plaxton and Carswell, 1999; Givan, 1999; Siedow and Umbach, 2000; Møller, 2001; Seki et al. 2001; Tiwari et al., 2002; Millenaar and Lambers, 2003; Vance et al., 2003; McDonald and Vanlerberghe, 2005; Ribas-Carbo et al., 2005). Various intermediates of glycolysis and the TCA cycle may be withdrawn to facilitate stress acclimation. For example, the roots of many plants excrete large amounts of citric and malic acids into the rhizosphere to alleviate nutritional Pi or Fe2+ deficiency, or Al3+ toxicity (Dinkelaker et al., 1989; Hoffland et al., 1992; Johnson et al., 1994; Neumann and Rümheld, 1999; López-Millàn et al., 2000; Ma et al., 2001; Massonneau et al., 2001; Ryan et al., 2001; Tesfaye et al., 2001; Schulze et al., 2002; Vance et al., 2003; Plaxton, 2004a; Shane et al., 2004; Cramer et al., 2005). The amount of carbon exuded in these two organic acids can be enormous, ranging from 10% to greater than 25% of the total plant dry weight (Vance et al., 2003). Root excretion of organic acids allows for the chelation of Al3+ , Fe3+ , and Ca2+ , and subsequent displacement and root uptake of Pi from bound or precipitated/insoluble forms (Ryan et al., 2001), and may also cause organic P to become more susceptible to hydrolysis by secreted acid phosphatases (Vance et al., 2003). In Al-toxic soils, root exudation of organic acids

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protects plants by chelating Al3+ ions in the rhizosphere, thus preventing their entry into the root cytoplasm (Ryan et al., 2001; Tesfaye et al., 2001). A striking feature of primary plant metabolism is that the same step in a metabolic pathway can frequently be accomplished in a variety of different ways. This metabolic flexibility is perhaps best exemplified by genetic engineering experiments in which post-transcriptional gene silencing technologies have been utilized to partially or fully eliminate an enzyme traditionally considered to be “essential,” and yet the resulting transgenic plants were able to grow and develop more or less normally (Plaxton, 1996, 2005; Fernie et al., 2004). A surprising example of the flexibility of plant primary metabolism was the enhanced dark respiration rate of transgenic tobacco leaves lacking cytosolic pyruvate kinase (PKc ) (Grodzinski et al., 1999). By contrast, animals or microbial mutants lacking this key glycolytic enzyme cannot respire carbohydrates. Later we consider examples of how the unusual flexibility of plant respiratory metabolism may help plants to cope with unavoidable stresses that are imposed upon them in their natural environment. 6.

Reactive Oxygen Species and Programmed Cell Death While the photosynthetic electron transport chain represents a major source of reactive oxygen species (ROS) in plants, the miETC also generates a significant amount of ROS that may be the dominant ROS source in non-green tissues, as well as photosynthetic tissues in the dark (Møller, 2001; McDonald and Vanlerberghe, 2005). Wheat leaf mitochondria contain significantly more ROS-modified proteins than do the corresponding chloroplasts or peroxisomes (Bartoli et al., 2004). This is because the production of harmful ROS is an unavoidable consequence of aerobic respiration. During respiration, O2 may undergo a univalent reduction at sites of ROS generation in Complexes I and III of the miETC, forming superoxide (O2 − ) which subsequently dismutates to H2 O2 . ROS production by purified mitochondria is increased under ADP-limiting conditions that increase the membrane potential and decreased by uncouplers that dissipate the PMF (McDonald and Vanlerberghe, 2005). Enhanced in vivo ROS generation by the miETC has been implicated in many plant biotic and abiotic stresses (Møller, 2001; McDonald and Vanlerberghe, 2005). For example, exposure of Arabidopsis cells to chronic oxidative stress increased miETC flux and O2 uptake, leading to elevated ROS production, ATP depletion, and programmed cell death (PCD) (Tiwari et al., 2002). PCD is an active, genetically controlled process that is accompanied by a suite of distinctive morphological and biochemical changes. It ultimately leads to the removal of cells no longer needed by the organism, and occurs during normal plant growth and development, and in response to biotic stresses (such as pathogen attack), as well as abiotic stresses including temperature extremes, UV irradiation, sugar or Pi starvation, hypoxia, and oxidative, water, or salinity stress (Krishnamurthy et al., 2000; Singh et al., 2003). PCD also occurs as the end point of

senescence, facilitating active nutrient recycling for use by other organs. In animals, PCD is triggered by mitochondrial events; namely the translocation of cytochrome c, a mobile electron carrier of the miETC, into the cytosol (Krishnamurthy et al., 2000). In the cytosol, cytochrome c binds to effector proteins that subsequently bring about a cascade of reactions that result in the controlled destruction of the cell. It also appears that plant PCD is associated with cytochrome c release from the mitochondrial i.m. (Balk et al., 1999; Tiwari et al., 2002). However, the role that mitochondrial cytochrome c release may play in initiating PCD in plant cells remains to be established. 7.

Fruit Ripening Many climacteric fruit store imported photosynthate in the form of starch in amyloplasts and as ripening proceeds this carbon is exported to the cytosol and converted into sugars and CO2 . The initiation of fruits ripening is frequently associated with a marked and rapid increase in respiratory CO2 evolution, termed the respiratory climacteric (Seymour, 1993). This respiratory increase is closely followed by the massive conversion of starch, which in the case of bananas comprises approximately 20 percent of the fresh weight of unripe fruit, into sucrose with up to 5 percent lost as CO2 in respiration (Seymour, 1993). Enhanced glycolytic flux and the associated rise in mitochondrial respiration at the climacteric are believed to generate ATP for the conversion of starch to sucrose and associated substrate (futile) cycles (Beaudry et al., 1989; Hill and ap Rees, 1994), as well as carbon skeletons needed for transamination reactions and other anabolic processes within this tissue (Law and Plaxton 1995, 1997; Turner and Plaxton, 2000, 2003). Due to the highly predictable pattern of carbohydrate metabolism during ripening, and the ease with which abundant amounts of fruit at various stages of ripening can be obtained, the banana represents an ideal model system in which to investigate the control of glycolytic and gluconeogenic carbon flux in vascular plants (Ball and ap Rees, 1988; Ball et al., 1991; Hill and ap Rees, 1994, 1995; Law and Plaxton, 1995, 1997; Turner and Plaxton, 2000, 2003). Control of initiation of the respiratory climacteric is desirable for increasing the storage life of climacteric fruits. 8.

Thermogenesis Certain plant species, particularly members of the family Araceae such as the skunk cabbage and voodoo lily, produce a thermogenic club-like structure known as a spadix on the developing floral apex. These groups are often associated with insect pollinators. During anthesis a profusion of mitochondria in the spadix cells contribute to massive respiration rates largely via the non-energy conserving alternative oxidase (AOX; discussed below), which diverts electrons from the cytochrome pathway and uncouples the miETC from ATP production (Seymour, 1999; Siedow and Day, 2000; Millenaar and Lambers, 2003; McDonald and Vanlerberghe, 2005). The free energy is consequently released as heat, raising tissue temperatures 10 to 25◦ C above ambient. Heat production is thought to enhance

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the production and volatilization of floral scents that help to attract insect pollinators (Seymour, 1999). The AOX has also been suggested to participate in limited thermoregulation in nonspecialized plant tissues in order to help plants survive cold or freezing stress (Moynihan et al., 1995). However, it has been argued that the resultant rate of heat production is insufficient to cause significant temperature increases of physiological importance (Breidenbach et al., 1997). Enhanced AOX activity during low temperature stress may have more to do with limiting chillinginduced ROS production by the miETC than with thermoregulation. Additional research is warranted to fully assess the relationships between respiratory electron flow to the non-energy conserving AOX and low temperature tolerance in plants.

B.

Mitochondria Are Dynamic Structures in Living Plant Cells Although identified over 50 years ago as the eukaryotic organelle responsible for oxidative energy metabolism, it is only very recently that yeast, animal, and plant cell biologists have begun to fully appreciate the complex mechanisms that mediate mitochondrial shape, size, and numbers in eukaryotic cells (Catlett and Weismann, 2000; Logan, 2003; Moyes and Hood, 2003; Wada and Suetsugu, 2004). When observed under the electron microscope (EM) mitochondria often appear as oval structures measuring several µm in length and 0.5 to 1 µm wide. This has fostered the “textbook” view that mitochondria exist as distinct organelles whose number per eukaryotic cell reflects the cell’s respiratory capacity. Calculations based upon crosssectional views of mitochondria in EMs suggest that as in animals, the number of mitochondria per plant cell varies according to the metabolic activity of the particular tissue (Logan, 2003). However, the notion that mitochondrial profiles seen in EMs of eukaryotic cells represent discrete and static organelles of known size and abundance has been disproved. Correct positioning and active movement of organelles within eukaryotic cells are essential for cellular homeostasis and adaptation to external stress (Wada and Suetsugu, 2004). Recent evidence indicates that the mitochondrial profiles observed in thin-section EMs of yeast and many animal tissues actually represent portions of a larger interconnected mitochondrial network, known as the mitochondrial reticulum (Catlett and Wisman, 2000; Moyes and Hood, 2003). Such results imply that the number of mitochondria in some eukaryotic cells may be considerably smaller and that their size is much larger than originally thought. Moreover, living animal cells contain large branched mitochondria in a dynamic state of flux, with segments of one mitochondrion frequently pinching off and fusing with another mitochondrion (Moyes and Hood, 2003). These dynamic interactions appear to reflect differing metabolic states of the cell (i.e., O2 or Glc availability, etc.) and indicate that the concept of the number of mitochondria present in a given cell may be irrelevant. This fascinating aspect of mitochondrial structure-function has only been recently addressed in plant systems, in which com-

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paratively little is known about the genes, proteins, and mechanisms controlling mitochondrial dynamics. Homology searches of the Arabidopsis genome failed to identify homologues of many yeast mitochondrial development genes suggesting that fundamental differences exist in the control of vascular plant mitochondrial morphology and dynamics (Logan, 2003). Whereas mitochondrial transport in plant cells is dependent upon actin filaments (Van Gestel et al., 2002), animal and yeast mitochondrial movement is mediated by microtubules (Wada and Suetsugu, 2004). Examination of hypocotyls and roots of transgenic Arabidopsis seedlings expressing mitochondrial-targeted green fluorescent protein (GFP) via epifluorescent microscopy revealed that plant mitochondria are highly dynamic structures, changing in size and shape within seconds (Logan and Leaver, 2000). Spherical, sausage- and worm-shaped Arabidopsis mitochondria were observed, with the sausage- and worm-shaped mitochondria being particularly dynamic (Logan and Leaver, 2000). Hypoxia stress was correlated with the formation of giant mitochondria in Arabidopsis leaves and cultured tobacco cells (Ramonell et al., 2001; Van Gestel and Verbelen, 2002). Forward and reverse molecular genetic approaches will be particularly instrumental to further establish the physiological and metabolic basis for mitochondrial structural dynamics in living plant cells. For example, Logan and co-workers (2003) screened mutagenized Arabidopsis seedlings whose mitochondria were labeled with GFP to isolate mutants defective in mitochondrial morphology and distribution. Four of the five mutants that were described resulted from the mutation of novel genes (Logan et al., 2003). These types of mutants constitute a powerful resource that will help to further establish the mechanisms that mediate this vital feature of plant cell biology and bioenergetics.

II.

THE FUNCTIONAL ORGANIZATION OF PLANT RESPIRATION Plants utilize sucrose and starch as the principle substrates for respiration, and can fully oxidize these fuels to CO2 and H2 O via the standard pathways of glycolysis, the TCA cycle, and the miETC. However, there are several remarkable attributes in the organization and associated bioenergetic features of plant respiratory metabolism that are not commonly seen in non-plant systems. Firstly, plant glycolysis exists in the plastid and cytosol, with the parallel reactions catalyzed by distinct nuclear-encoded isozymes (Plaxton, 1996; Givan, 1999; Podestá, 2004). Plastidic and cytosolic glycolysis “communicate” through the action of highly selective metabolite transporters that are present in the inner plastid envelope (Tegeder and Weber, 2005). The prime functions of glycolysis in darkened chloroplasts and nonphotosynthetic plastids are to participate in the breakdown of starch as well as to generate carbon skeletons, reductant and ATP for anabolic pathways such as fatty acid synthesis (Plaxton, 1996). Although plastids from several nonphotosynthetic tissues, including developing wheat and castor seeds have been found to possess all the enzymes of glycolysis from Glc to pyruvate,

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some chloroplasts may lack one or several of the enzymes of the lower half of glycolysis (i.e., enolase and phosphoglyceromutase) (Emes and Tobin, 1993; Plaxton, 1996). Secondly, the plant cytosolic glycolytic pathway is a complex network containing parallel enzymatic reactions at the level of sucrose, fructose6-phosphate (Fru-6-P), glyceraldehyde-3-phosphate, and PEP metabolism (Figure 2) (Black et al., 1987; Plaxton, 1996, 2004a, 2005; Givan, 1999; Podestá, 2004). As discussed below, these nonconventional glycolytic bypass enzymes may be especially important under conditions of metabolic stress. Third, a number of bypass reactions in the plant TCA cycle exist and include: (i) a NADP+ -specific isocitrate dehydrogenase (ICDH) within the mitochondrial matrix that can circumvent the NAD+ specific ICDH of the conventional TCA cycle (Igamberdiev and Gardeström, 2003), (ii) the glyoxylate cycle of germinating oil seeds which necessitates two key enzymes (isocitrate lyase and malate synthase) that collectively bypass both decarboxylating reactions of the TCA cycle, thereby allowing gluconeogenesis from stored fats to occur (Falk et al., 1998; Siedow and Day, 2000; McDonald and Vanlerberghe, 2005), and (iii) the γ -amino butyric acid (GABA) shunt which may function as an alternative, NAD+ -independent, mechanism for Glu entry into the TCA cycle (Shelp et al., 1999) (Figure 4). Fourthly, respiratory O2 consumption by the plant miETC can be mediated by the phosphorylating cytochrome pathway or the non-energy conserving pathway involving the rotenone insensitive NAD(P)H dehydrogenase bypasses to Complex I, and the cyanide resistant AOX bypass to Complex III and IV (Figures 2 and 3) (Siedow and Day, 2000; Siedow and Umbach, 2000; Møller, 2001; Millenaar and Lambers, 2003; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). Parallel reactions of glycolysis, the TCA cycle, and miETC are believed to endow plants with crucial metabolic flexibility that facilitates their development and acclimation to unavoidable abiotic stresses.

anoxia or Pi starvation, or following the addition of respiratory poisons, that elicit significant reductions in cellular ATP pools (Duff et al., 1989b; Dancer et al., 1990; Rychter and Randall, 1994; Plaxton, 1996, 2004a, 2005; Rea and Poole, 1997; Stitt, 1998). The significance of PPi in plant metabolism was demonstrated by the introduction of the Escherichia coli inorganic PPiase gene into tobacco and potato plants under control of a constitutive promoter (Jellito et al., 1992). Expression of the E. coli PPiase in the plant cytosol caused PPi levels to be reduced by up to 3-fold and led to a dramatic inhibition of plant growth. To assess the relative importance of PPi versus ATP as an energy donor in the plant cytosol, Davies et al. (1993) computed the standard free energy changes for PPi and ATP hydrolysis under a variety of cytosolic conditions. The results indicate that PPi would be particularly favored as a phosphoryl donor, relative to ATP, under cytosolic conditions known to accompany stresses such as anoxia or nutritional Pi deprivation. This underscores the importance of PPi as an autonomous energy donor of the plant cytosol. Consistent with this view was the finding that tolerance of acclimated maize root tips to anoxia is not critically dependent upon high energy charge (Xia et al., 1995). How stressed versus non-stressed plant cells maintain a relatively constant level of cytosolic PPi remains enigmatic. Anabolism, and hence the rate of PPi production, would generally be more prevalent under non-stressed conditions. However, even during stresses such as anoxia or Pi starvation, PPi would continue to be generated (albeit at a lower rate) during the synthesis of essential macromolecules (i.e., proteins, nucleic acids, membranes, etc.) needed to support diminished growth, and/or replace those that may have become damaged or worn out. This is indicated by the fact that cellular PPi levels remain fairly stable in plants that have been exposed to abiotic stresses that elicit significant reductions in adenylate pools (Duff et al., 1989b; Dancer et al., 1990; Plaxton, 1996, 2004a, 2005; Rea and Poole, 1997; Stitt, 1998).

A.

1.

Pyrophosphate: An Alternate Energy Donor of Plant Cells Inorganic pyrophosphate (PPi) is a byproduct of a host of anabolic reactions, including the polymerization reactions involved in the final steps of macromolecule synthesis. One doctrine of cellular bioenergetics (stemming from animal studies) is that the anhydride bond of PPi is never utilized to perform cellular work since it is immediately hydrolyzed by an inorganic pyrophosphatase (PPiase) as soon as it is produced (Plaxton, 1996, 2004a, 2005; Rea and Poole, 1997). However, as was recognized by Baltscheffsky (1967) almost four decades ago, the large amounts of PPi generated during biosynthesis are not always wasted, but may enhance the energetic efficiency of several cellular processes in plants and various anaerobic microorganisms. The significant PPi concentration of the plant cytosol (up to 0.5 mM) is due to the absence of soluble inorganic PPiase in this compartment (Stitt, 1998; Farre et al., 2001). PPi levels of plant cells are remarkably insensitive to abiotic stresses such as

Cytosolic PPi Metabolism The discovery some 25 years ago of the strictly cytosolic PPi-dependent phosphofructokinase (PPi-PFK; reaction 10, Figure 2) in plants (Carnal and Black, 1979), and its potent allosteric activation by low concentrations of the regulatory metabolite fructose-2,6-bisphosphate (Fru-2,6-P2 ) (Sabularse and Anderson, 1981) led to a surge of research on the role of PPi in plant metabolism (Stitt, 1990, 1998). It is now evident that PPi-PFK is an adaptive plant enzyme whose activity and subunit composition are dependent upon a variety of environmental, developmental, species and tissue-specific cues (Duff et al., 1989b; Mertens et al., 1990; Wong et al., 1990; Botha and Botha, 1991; Mertens, 1991; Blakeley et al., 1992; Theodorou et al., 1992; Theodorou and Plaxton, 1994; Plaxton, 1996, 2004a; Murley et al., 1998; Stitt, 1998). Although transgenic potato plants with significantly reduced PPi-PFK levels showed no detectable growth or phenotypic difference from control plants, there was a slight shift in carbon partitioning between starch and sucrose

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FIG. 2. Alternative pathways of cytosolic glycolysis, miETC, and tonoplast H+ -pumping processes (indicated by bold arrows) that may facilitate respiration and vacuolar pH maintenance by Pi-deprived plant cells. These bypasses negate any dependence of respiration and tonoplast H+ -pumping on adenylates and Pi, the levels of which become markedly depressed during severe Pi starvation. Organic acids produced by PEPC may also be excreted by roots to increase the availability of mineral bound Pi. A key component of this model is the critical secondary role played by “metabolic Pi recycling systems” during Pi deprivation. Enzymes that catalyze the numbered reactions are as follows: 1, sucrose synthase; 2, invertase; 3, NDP kinase; 4, fructokinase; 5, hexokinase; 6, UDP-glucose pyrophosphorylase; 7, P-glucomutase; 8, P-glucose isomerase; 9, ATP-PFK; 10, PPi-PFK; 11, aldolase; 12, triose-phosphate isomerase; 13 and 14, phosphorylating and non-phosphorylating glyceraldehyde 3-phosphate dehydrogenase, respectively; 15, phosphoglycerate kinase; 16, phosphoglyceromutase; 17, enolase; 18, PEP phosphatase; 19, H+ -ATPase; 20, H+ -PPiase; 21, PPDK; 22. MDH; 23, NAD-ME. Abbreviations are as defined in text or legend for Figure 1 and as follows: DHAP, dihydroxyacetone-phosphate; Ga3P, glyceraldehyde-3-phosphate; 1,3-DPGA, 1,3-diphosphoglycerate; 2-PGA, 2-phosphoglycerate; UQ, ubiquinone.

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FIG. 3. Organization of the electron transport processes occurring within the i.m. of plant mitochondria. The illustration provides detail of the cytochrome pathway (electrons flow to complex IV), which generates the electromotive force for ATP synthesis, and the non-energy conserving pathway (electrons pass directly to O2 through AOX). Abbreviations are as defined in the text and as follows: CI, complex I or NADH dehydrogenase; CII, complex II or succinate dehydrogenase; CIII, complex III or cytochrome bc1 complex; CIV, complex IV or cytochrome oxidase. Crosses inside circles indicate sites of inhibition for different inhibitors of the miETC.

FIG. 4. Glycine, proline, glutamate, and GABA may function as important respiratory substrates in plant mitochondria. Enzymes that catalyze the numbered reactions are as follows: 1 and 2, glycine decarboxylase and serine hydroxymethyl transferase, respectively; 3, proline dehydrogenase; 4, 1 -pyrroline-5-carboxylate dehydrogenase; 5, AAT; 6, glutamate dehydrogenase; 7, glutamate decarboxylase; 8, GABA-transaminase; 9, succinic semialdehyde dehydrogenase. Abbreviations are as defined in the text and as follows: Succ, succinate; Fum, fumarate; P5C, 1 - pyrroline-5-carboxylate; GSA, glutamic gamma-semialdehyde.

PLANT RESPIRATION

and a decrease in the triose-phosphate to hexosephosphate ratio (Hajirezaei et al., 1994). From these data, the authors concluded that PPi-PFK catalyzes a net glycolytic flux in potato tubers. Similar conclusions have been drawn from analysis of transgenic tobacco plants containing elevated levels of the PPi-PFK activator, Fru-2,6-P2 (achieved by over-expressing mammalian 6-phosphofructo-2-kinase) (Scott et al., 1995). Apart from PPi-PFK, PPi could be employed as an alternative energy donor for at least two other processes of the plant cytosol that are normally dependent upon ATP: (i) sucrose conversion to hexose-phosphates can proceed via the ATPdependent invertase pathway or via the PPi-dependent sucrose synthase (SuSy) pathway, and (ii) active transport of protons into the vacuole from the cytosol can be carried out by ATP- or PPi-dependent H+ -pumps of the tonoplast (Figure 2) (Plaxton, 1996; Rea and Poole, 1997; Stitt, 1998; Plaxton and Carswell, 1999). That PPi-powered processes may be a crucial facet of the metabolic adaptations of plants to environmental extremes that lead to depressed ATP (but not PPi) pools is indicated by the significant induction or upregulation of SuSy, UDP-Glc pyrophosphorylase, PPi-PFK, and the tonoplast H+ -PPiase by anoxia or hypoxia, extended Pi-starvation, and/or cold stress (Duff et al., 1989b; Theodorou et al., 1992, 1994; Carystinos et al., 1995; Guglielminetti et al., 1995; Theodorou and Plaxton, 1994, 1996; Plaxton, 1996, 2004a, 2005; Carswell et al., 1997; Rea and Poole, 1997; Ciereszko et al., 1998, 2001; Murley et al., 1998; Plaxton and Carswell, 1999; Zeng et al., 1999; Palma et al., 2000; Massonneau et al., 2001; Geigenberger, 2003; Uhde-Stone et al., 2003; Vance et al., 2003; Wang et al., 2003; Wasaki et al., 2003; Hammond et al., 2004; Misson et al., 2005). The induction of PPi-dependent cytosolic bypasses may help plants to survive certain stresses by circumventing ATPlimited reactions while conserving diminished cellular pools of ATP. Sucrose metabolism of anoxic rice seedlings was shown to proceed mainly through SuSy with nucleoside diphosphate kinase facilitating the cycling of uridilates needed for operation of this pathway (reaction 3, Figure 2) (Guglielminetti et al., 1995). When oxygen levels are low, SuSy activity increases while invertase activity decreases (Guglielminetti et al., 1995; Zeng et al., 1999; Geigenberger, 2003). Assuming that PPi is a byproduct of anabolism, no ATP is needed for the conversion of sucrose to hexose-phosphates via the SuSy pathway, whereas 2 ATPs are needed for the invertase pathway. Mertens (1991) argued that PPi-PFK also functions in glycolysis in hypoxic rice seedlings, since both PPi-PFK activity and the level of its allosteric activator Fru-2,6-P2 are increased, while the activity of ATP-PFK declines (Mertens et al., 1990). Thus, the net yield of ATP obtained during glycolytic fermentation of sucrose to pyruvate is doubled from 4 to 8 if sucrose is metabolized via the SuSy and PPi-PFK bypasses, relative to the invertase and ATP-PFK pathways (Figure 2). Pyruvate, Pi dikinase (PPDK) represents a third enzyme that can theoretically employ PPi to catalyze a glycolytic bypass reaction in the cytosol of plant cells (reaction

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21, Figure 2). This enzyme is highly expressed in the stroma of mesophyll chloroplasts of C4 leaves where it plays a key role in the C4 photosynthetic pathway in which it functions in the PPi producing direction to regenerate PEP from pyruvate. However, PPDKs having a non-photosynthetic function are expressed as both cytosolic and plastidic isoforms in C3 plant cells (Chastain and Chollet, 2003). Although the metabolic roles of PPDK in C3 plants remain obscure, the cytosolic PPDK has been hypothesized to function as a glycolytic bypass to PKc in non-green hypoxic rice tissues (i.e., developing seed endosperm and flooded roots) so as to enhance the ATP yield of glycolysis (Moons et al., 1998; Chastain and Chollet, 2003). PPDK could confer a considerable bioenergetic advantage for ATP-depleted plant cells, since this enzyme catalyzes the net creation of two “ATP equivalents” (i.e., two “high energy” phosphoanhydride bonds) in the glycolytic direction, as opposed to the single ATP generated by the PKc reaction (Plaxton, 2005). Our understanding of the importance of PPi in cellular bioenergetics has been augmented by research on “energy-poor” anaerobic microorganisms such as the bacteria Priopionibacterium shermanii and the parasitic protist Entamoeba histolytica (the latter causing amoebic dysentery in humans). These species have no inorganic PPiase, ATP-PFK, or PK, but instead employ PPi to convert Fru-6-P into fructose-1,6-bisphosphate (Fru-1,6-P2 ) via PPi-PFK, and PEP and AMP into pyruvate, ATP and Pi via PPDK (Plaxton, 2005). Owing to their use of PPi-PFK and PPDK, these organisms are able to yield 5 ATP per hexose oxidized to two molecules of pyruvate, with a net expenditure of 3 PPi recycled from biosynthetic reactions. This clearly represents a considerable bioenergetic advantage for obligate anaerobes such as P. shermanii and E. histolytica, since the net ATP yield for classical glycolysis (as it occurs in animals or yeast) is only two ATP per hexose converted to two molecules of pyruvate. Likewise, the alternative PPi-dependent reactions of the plant cytosol outlined in Figure 2 (i.e., the SuSy/UDP-Glc pyrophosphorylase pathway of sucrose to hexosephosphate conversion, PPi-PFK, the tonoplast H+ -PPiase, and PPDK) would confer a significant bioenergetic benefit that may extend the survival time of plant cells that have become ATP-depleted during stresses such as chilling, hypoxia, or Pi starvation (Carystinos et al., 1995; Plaxton, 1996, 2004b, 2005; Rea and Poole, 1997; Ciereszko et al., 1998, 2001; Massonneau et al., 2001; Palma et al., 2000; Chastain and Chollet, 2003; Geigenberger, 2003; Podestá and Plaxton, 2003). 2.

Mitochondrial PPi Metabolism The miETC-dependent synthesis of PPi was first demonstrated more than 20 years ago (Kowalczyk and Maslowski, 1984). This appears to be catalyzed by a unique H+ -PPiase located in the mitochondrial i.m., which is also capable of coupling PPi hydrolysis to the generation of PMF across the i.m. (Vianello and Macri, 1999). The H+ -PPiase of the plant mitochondrial i.m. has been suggested to contribute to the known capacity of plant mitochondria to survive periods of hypoxia stress (Vianello and

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Macri, 1999). PPi import and synthesis by plant mitochondria was recently evaluated (Casolo et al., 2002). The matrix PPi level in mitochondria isolated from etiolated pea seedlings was about 0.2 mM, and an H+ -PPiase was localized to the matrix side of the mitochondrial i.m., with an H+ /PPi stoichiometry of 2 (Casolo et al., 2002). Whether this mitochondrial H+ -PPiase operates in the forward (PPi synthesizing, PMF dissipating) or reverse (PPi hydrolysis, PMF generating) direction, remains unclear. However, as depicted in Figure 3 it could operate under some conditions as a PPi synthase that utilizes PMF generated by the miETC. The PPi may then accumulate in the mitochondrial matrix, or be exported to the cytosol via the adenine nucleotide translocator, exchanging PPi in antiport with ADP (Casolo et al., 2002). Further research is required to fully assess the metabolism and role(s) of PPi in plant mitochondria. Plant Mitochondria Can Respire Alternative Substrates to Pyruvate Carbohydrates are generally believed to be the major respiratory substrate for plant mitochondria. The end-product of the glycolytic sequence is pyruvate, which can be directly transported into the mitochondrion by a specific transporter embedded in the mitochondrial i.m. (Picault et al., 2004; Tegeder and Weber, 2005). The pyruvate dehydrogenase complex (PDC) of the mitochondrial matrix links glycolysis to the TCA cycle by catalyzing the oxidative decarboxylation of pyruvate, yielding CO2 , acetyl-CoA and NADH (Hill, 1997; Siedow and Day, 2000; Tovar-Mendez et al., 2003). However, apart from pyruvate, plant mitochondria also have the capacity to respire a number of alternative substrates, some of which may become particularly important under certain developmental or environmental situations.

the most significant glycolytically derived substrate for respiration (Edwards et al., 1998; Jenner et al., 2001). Netting (2002) has also summarized evidence arguing against the entry of malate into the mitochondria for oxidation. Nevertheless, there may be specific situations, such as during oilseed embryogenesis (Schwender et al., 2004) or prolonged Pi starvation (Nagano et al., 1994), when the flux of PEP to pyruvate via the PEPC, MDH and NAD-ME bypass to PKc (Figure 2) could be significant. Future studies are required to determine the suite of physiological conditions that may stimulate intramitochondrial pyruvate production via NAD-ME. 2.

Formate One of the most abundant proteins in the mitochondrial matrix of non-green plant tissues is formate dehydrogenase (FDH) which catalyzes: formate + NAD+ → CO2 + NADH.

B.

1.

Malate It has been widely assumed that malate is an important respiratory substrate for plant mitochondria, such that a significant fraction of glycolytic products enter the TCA cycle via the combined reactions of PEPC, malate dehydrogenase (MDH), and NAD-dependent malic enzyme (NAD-ME), rather than PKc (Figure 2). However, most data in support of malate as a respiratory substrate were obtained with isolated mitochondria, complicating interpretations of physiological relevance. In fact, recent NMR and GC-MS studies of plant tissues or suspension cells incubated with 13 C-labelled substrates demonstrated that the flux of malate to pyruvate via mitochondrial NAD-ME is usually very low, relative to the flux of PEP to pyruvate via PKc (Dieuaide-Noubhani et al., 1995; Edwards et al., 1998; Ratcliffe and Sachar-Hill, 2001; Rontein et al., 2002; Schwender et al., 2004). Thus, a marked reduction in mitochondrial NADME in transgenic potatoes had no detectable effect on respiratory metabolism (Jenner et al., 2001). It was concluded that the import of PEPC-derived malate into the mitochondria usually serves an anaplerotic role to support biosynthesis and Nassimilation, whereas pyruvate derived from the PKc reaction is

Our understanding of the function of FDH in plant mitochondrial metabolism is far from complete and awaits determination of formate origins and metabolism in different plant tissues (Igamberdiev et al., 1999; McDonald and Vanlerberghe, 2005). However, potato leaf FDH was markedly upregulated in response to cold or drought stress, and such leaves readily utilize formate as a respiratory substrate in support of oxidative phosphorylation (Hourton-Cabassa et al., 1998). Thus, formate oxidation by FDH may provide an important source of reducing power for the miETC under stress conditions that lead to formate accumulation while simultaneously compromising other respiratory pathways such as the PDC and TCA cycle. 3.

Amino Acids and Fatty Acids Pulse-chase radiolabeling studies have indicated that the TCA cycle is generally involved in the accumulation and interconversion of amino acids, rather than their oxidation (Hill, 1997). In most situations where there is net protein catabolism, such as senescence or seed germination, the amino acids produced are translocated elsewhere rather than being respired. Similarly, although fatty acids are frequently stored as triglycerides (particularly in oilseeds), they are rarely respired, but are instead oxidized to acetyl-CoA in peroxisomes or glyoxysomes and converted via the glyoxylate cycle and gluconeogenesis into sugars for transport (Falk et al., 1998; McDonald and Vanlerberghe, 2005). An important exception is during carbon starvation such as that observed during prolonged darkness (Brouquisse et al., 1998), or sucrose deprivation of heterotrophic suspension cell cultures or excised root tips (Dieuaide-Noubhani et al., 1995, 1997). Natural carbon starvation may arise during abiotic or biotic stresses that lead to a significant decrease in photosynthesis, thus limiting the supply of carbohydrates to sink tissues. Some types of senescence or postharvest situations may also be associated with carbohydrate starvation. Plant metabolism must acclimate to the lack of carbohydrates by substituting protein and lipid catabolism for sugar catabolism (Dieuaide-Noubhani

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et al., 1995, 1997; Brouquisse et al., 1998; Falk et al., 1998). Thus, enzymatic activities related to sugar metabolism and respiration, nitrogen reduction and assimilation, or protein synthesis decrease during sugar starvation. In contrast, enzyme activities related to the catabolism of proteins, amino acids, and mitochondrial β-oxidation of fatty acids increase (Dieuaide-Noubhani et al., 1997). Acyl-CoA dehydrogenases involved in mitochondrial β-oxidation have been characterized from several plants (Bode et al., 1999). Recent studies of Arabidopsis mutants lacking the key glyoxylate cycle enzymes isocitrate lyase and malate synthase have provided novel insights into fatty acid respiration by plant mitochondria (Falk et al., 1998; Eastmond et al., 2000). Amino acid oxidation may be preceded by a transamination reaction that produces a glycolytic or TCA cycle intermediate, or the oxidation may occur directly. A well known example of the latter is the reaction catalyzed by Glu dehydrogenase (reaction 6, Figure 4). Although the subject of some debate, it has been clearly demonstrated by means of 15 N- or 13 C-labelling experiments that the main function of Glu dehydrogenase is to deaminate Glu into 2-OG (Dubois et al., 2003). Plant mitochondria also contain a variety of branched-chain dehydrogenase complexes that are capable of oxidizing the branched chain amino acids (i.e., valine, leucine, and isoleucine) into their corresponding α-keto acids (Mooney et al., 2002; Taylor et al., 2004; McDonald and Vanlerberghe, 2005). These branched-chain dehydrogenase complexes appear to be induced by their substrates, particularly under conditions of carbohydrate starvation (Fujiki et al., 2001, 2002). The potential role of branched-chain amino acids catabolism as an oxidative phosphorylation energy source or as a detoxification pathway during plant stress was recently emphasized (Taylor et al., 2004). Apart from carbohydrate starvation, there are at least three additional situations where amino acid oxidation within the mitochondrial matrix may make a significant contribution to respiration. The first is glycine produced during leaf photorespiration in which two molecules of glycine are converted by the concerted action of glycine decarboxylase and serine hydroxymethyl transferase into one serine, CO2 and NH4 + with the production of NADH, thereby providing an important source of reducing equivalents for the miETC (Douce et al., 2001; Raghavendra and Padmasaree, 2003; McDonald and Vanlerberghe, 2005). The biochemistry and molecular biology of the glycine decarboxylase complex has been thoroughly investigated (Douce et al., 2001). Two additional amino acids that can be oxidized at significant rates in plant mitochondria are proline and GABA (Figure 4). Both are biocompatible solutes that may accumulate to significant levels during stresses such as drought, high salinity, or cold, and then rapidly disappear when the stress is removed (Hill, 1997; Shelp et al., 1999). Proline is oxidized by a threestep process that ultimately feeds 2-OG into the TCA cycle (Figure 4). In barley leaves recovering from drought, the oxidation of proline accounted for up to 20 percent of the rate of respiration (Stewart and Voetberg, 1985). GABA, a four-carbon

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non-protein amino acid, may represent a significant proportion of the free amino acid pool in plant cells subjected to abiotic or biotic stresses (Shelp et al., 1999). Various stresses initiate a signal transduction pathway in which increased cytosolic Ca2+ activates a calmodulin-dependent Glu decarboxylase leading to a marked elevation in intracellular GABA levels (Figure 4). After the stress is relieved, GABA can be transported into the mitochondrion and enter the TCA cycle via its conversion to succinate through the sequential action of GABA transaminase and succinic semialdehyde dehydrogenase (Figure 4). C.

Non-Energy Conserving Bypasses of Mitochondrial Electron Transport Contribute to the Unique Flexibility of Plant Metabolism The plant miETC exhibits similarities to the energyconserving miETC of animal mitochondria insofar as both systems contain a mobile electron carrier (cytochrome c), along with four respiratory chain complexes, namely: the rotenonesensitive Complex I (NADH dehydrogenase), Complex II (succinate dehydrogenase), Complex III (cytochrome c reductase), the cyanide-sensitive Complex IV (cytochrome c oxidase), along with Complex V (ATP synthase). However, as outlined in Figure 3, plant mitochondria are endowed with a highly branched respiratory chain that is characterized by several additional pathways for electron transport (Siedow and Day, 2000; Siedow and Umbach, 2000; Møller, 2001; Millenaar and Lambers, 2002; Vanlerberghe and Ordog, 2002; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). 1.

Non-Proton Pumping Alternate NAD(P)H Dehydrogenases In addition to oxidizing internally generated NADH via the proton-pumping Complex I, plant mitochondria possess a complex series of non-proton motive dehydrogenases located on the outer and inner surface of the i.m., which can oxidize NADH or NADPH in a rotenone insensitive manner (Figure 3) (Møller, 2001; Millenaar and Lambers, 2002; Michalecka et al., 2003, 2004; Moore et al., 2003; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). All of these dehydrogenases will reduce the ATP-yield of respiration since they are non-proton pumping bypasses to the proton-pumping Complex I. The various nonproton motive NAD(P)H dehydrogenases have been well documented in isolated plant mitochondria and sub-mitochondrial particles, and characterization of the corresponding genes has begun (Moore et al., 2003; Michalecka et al., 2003, 2004; Rasmusson et al., 2004). Several of the genes encoding known or putative alternate NAD(P)H dehydrogenases include amino acid sequences similar to EF-hand Ca2+ -binding motifs (Rasmusson et al., 1999). This agrees with biochemical data demonstrating that the activities of the internal and external NADPH dehydrogenases of the plant miETC are Ca2+ -dependent (Rasmusson et al., 2004). NADPH is generated within the mitochondrial matrix by enzymes such as the aforementioned NADP-specific ICDH

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(Møller, 2001; Igamberdiev and Gardeström, 2003; Gray et al., 2004). This NADPH can be used as a source of reductant for various mitochondrial processes, including the rotenone-insensitive NADPH dehydrogenase located on the matrix side of the i.m. (Figure 3) (Moore et al., 2003; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005). Although the precise functions and control of the external facing non-energy conserving NAD(P)H dehydrogenases of the i.m are not well established, they presumably influence the redox balance of the cytosolic pyridine nucleotide pools (i.e., the ratio of [NAD(P)+ ]/[NAD(P)H]) and may thus affect many cytosolic reactions. 2.

Non-Proton Pumping Alternative Oxidase One of the most distinctive features of the plant miETC is its capacity for cyanide-resistant respiration due to the presence of the AOX. Plant AOX exists as a 32 kDa homodimer that depending upon its activation state (see below) can divert electrons from the main respiratory chain at the level of the ubiquinone pool to catalyze the four-electron reduction of O2 to H2 O (Siedow and Umbach, 2000; Vanlerberghe and Ordog, 2002; Millenaar and Lambers, 2003; McDonald and Vanlerberghe, 2005). AOX dramatically reduces the ATP-yield of respiration as it circumvents the proton-pumping sites constituted by Complexes III and IV of the standard miETC (Figure 3), with the resultant free energy released as heat. AOX is encoded by a multigene family, but the significance of the multiple gene products remains enigmatic (Vanlerberghe and Ordog, 2002). Although long believed to be a plant-specific phenomenon, AOX genes and cyanide-resistant respiration have recently been discovered in numerous marine bacteria, some fungi and protists, as well as in several animal invertebrate phyla (McDonald and Vanlerberghe, 2005). Due to its non-energy conserving nature, substantial research has attempted to assess the physiological functions of AOX respiration in plants. Understanding plant AOX functions has been facilitated by technical advances that have made it possible for researchers to discriminate between AOX capacity (i.e., maximum electron flux to AOX) versus AOX engagement (i.e., actual in vivo electron flux through AOX). In isolated mitochondria and some tissues, AOX capacity can be estimated using chemical inhibitors (Juszczuk et al., 2001; McDonald et al., 2002). Although more difficult to quantify, the extent of AOX engagement during whole plant respiration can be directly determined by assessing O2 isotope discrimination (Gonzàlez-Meler et al., 1999; Siedow and Umbach, 2000; McDonald et al., 2002). Such noninvasive measurements rely on the use of mass spectrometry to discriminate between the differential fractionation of 16 O and 18 O isotopes by the alternative versus cytochrome pathways. This technique has helped to establish the in vivo contributions of the respective miETC pathways to overall plant respiration rates under a range of physiologically interesting situations (Gonzàlez-Meler et al., 1999, 2001; McDonald et al. 2002; Millnaar and Lambers, 2003; Guy and Vanlerberghe, 2005). Transgenic plants defective in or overexpressing AOX activity have also been invalu-

able tools for elucidating in vivo AOX roles (Maxwell et al., 1999; Parsons et al., 1999; Robson and Vanlerberghe, 2002; Vanlerberghe et al., 2002; McDonald and Vanlerberghe, 2005; Guy and Vanlerberghe, 2005). Apart from its well defined role in floral thermogenesis in certain species (Seymour, 1999), AOX respiration is induced under many different environmental conditions to help plants survive stress. The non-energy conserving nature of the AOX pathway, along with complex biochemical controls that control electron flux to AOX endow plant mitochondria with considerable metabolic flexibility. This flexibility allows the mitochondria to maintain electron flux to O2 when electron flux through the phosphorylating cytochrome pathway is restricted by high adenylate energy charge, or under a number of stresses such as low temperature or Pi deprivation. AOX transcript profiling and immunoquantification studies have demonstrated its marked induction in response to diverse stress situations including: wounding or pathogen attack, drought and osmotic stress, low temperature, low light, and Pi starvation, in addition to treatment with chemicals such as salicylic acid, H2 O2 , or inhibitors of the cytochrome pathway (Rychter and Mikulska, 1990; Rychter et al., 1992; Hoefnagel et al., 1993, 1994; Van Der Straeten et al., 1995; Breidenbach et al., 1997; Plaxton and Carswell, 1999; Juszczuk et al., 2001; Seki et al., 2002; Millenaar and Lambers, 2003; Shane et al., 2004; McDonald and Vanlerberghe, 2005; Misson et al., 2005; Noguchi et al., 2005). In general, any condition that inhibits or decreases the activity of the standard phosphorylating pathway of miETC induces AOX. It has also been demonstrated that transgenic plants lacking functional AOX have increased amounts of ROS originating from the mitochondrion and that such cells are more susceptible to PCD-inducing treatments (Maxwell et al., 1999; Robson and Vanlerberghe, 2002). By contrast, cells overexpressing AOX exhibited reduced amounts of ROS (Vanlerberghe et al., 2002; McDonald and Vanlerberghe, 2005). Recent studies with purified mitochondria also support a crucial AOX function in limiting ROS generation by the miETC (Camacho et al., 2004). By “draining” excess electrons, AOX appears to help minimize the production of damaging ROS that otherwise arises when ubiquinone becomes over-reduced (Møller, 2001; Vanlerberghe et al., 2002; Millenaar and Lambers, 2003; McDonald and Vanlerberghe, 2005). This could be important if the NADH/NAD+ redox ratio is relatively high (reductant overflow) or if the cytochrome pathway becomes inhibited. At the same time NAD+ is continually turned over so that PDC, and NAD+ -dependent dehydrogenases of glycolysis and the TCA cycle can continue to operate. 3.

Uncoupling Protein A more recent addition to the complexity of the plant miETC is the presence of an uncoupling protein (UCP) (HourtonCabassa et al., 2004). UCP is a homologue of thermogenin, an i.m. protein responsible for non-shivering thermogenesis in mammalian brown fat cells. UCP and thermogenin allow protons to diffuse down their concentration gradient from the

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intermembrane space into the matrix, circumventing the ATP synthase complex and thus uncoupling electron transport from ATP production (Figure 3). Hence, although by different mechanisms, AOX and UCP are both non-energy conserving bypasses of the plant miETC. As is the case with AOX, UCP expression is quite variable, responsive to stress, and dependent upon developmental processes and growth conditions (Hourton-Cabassa et al., 2004; McDonald and Vanlerberghe, 2005). Plant UCP is activated by superoxide suggesting that it may work with AOX to help reduce mitochondrial ROS production (Daley et al., 2003; Smith et al., 2004). The functional significance of the three putative UCPs encoded by the Arabidopsis genome remains to be determined (Hourton-Cabassa et al., 2004). Mitochondria from transgenic plants overexpressing UCP have been examined to provide important insights into UCP function (Brandalise et al., 2003; Smith et al., 2004). An increase in UCP protein content was correlated with a significant reduction in the rate of superoxide production and enhanced tolerance to H2 O2 , and an increase in pyruvate to citrate flux (Brandalise et al., 2003; Smith et al., 2004). The latter result led to the conclusion that UCP may not only influence mitochondrial ROS production, but also TCA cycle flux (Smith et al., 2004). D.

The Adaptive Value of Adenylate- and Pi-Independent Bypasses of Cytosolic Glycolysis and Mitochondrial Respiration Is Illustrated by Pi-Starved Plants Pi plays a central role in virtually all major metabolic processes in plants, including photosynthesis and respiration. Despite its importance, Pi is one of the least available nutrients in many terrestrial and aquatic environments. In soil, Pi is often complexed to Al3+ , Ca3+ , or Fe3+ cations and thus occurs in an insoluble mineral form that renders it unavailable to plants (Plaxton and Carswell, 1999; Vance et al., 2003; Plaxton, 2004a). The massive use of Pi fertilizers in agriculture demonstrates how the free Pi levels of most soils are suboptimal for plant growth. Worldwide reserves of rock phosphate—our major source of Pi fertilizers in agriculture—have been projected to be exhausted within the next 50 to 75 years (Vance et al., 2003). Thus, research on plant biochemical adaptations to Pi deficiency is of significant practical importance. It should serve to facilitate development of molecular tools and rational strategies for the application of biotechnology to reduce or eliminate our current over-reliance on expensive, polluting, and nonrenewable Pi fertilizers. 1.

Glycolytic Bypasses of Pi-Starved Plants As a consequence of the marked decline (up to 50-fold) in cytoplasmic Pi levels that follows severe Pi stress, large (up to 80%) reductions in intracellular levels of ATP, ADP and related nucleoside-Ps also occur, whereas PPi levels remain largely unaffected (Ashihara et al., 1988; Duff et al., 1989b; Lauer et al., 1989; Dancer et al., 1990; Rychter and Mikulska, 1990; Rychter et al., 1992; Rychter and Randall,

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1994; Lee and Ratcliffe, 1993; Rao and Terry, 1995; Plaxton, 1996, 2004a; Plaxton and Carswell, 1999; Fernie et al., 2002; Le Roux et al., 2005). This is expected to inhibit carbon flux through the enzymes of classical glycolysis that are dependent upon adenylates (i.e., hexokinase, ATP-dependent phosphofructokinase (ATP-PFK), 3-phosphoglycerate (3-PGA) kinase, and PKc ) or Pi (i.e., phosphorylating NAD+ -dependent glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as cosubstrates (Figure 2). Despite depleted intracellular Pi and adenylate pools, Pi-deprived plants must continue to generate energy and C-skeletons for key metabolic processes. At least six Piand adenylate-independent glycolytic “bypass” enzymes (i.e., SuSy, UDP-Glc pyrophosphorylase, PPi-PFK, nonphosphorylating NADP-GAPDH, PEPC, PEP phosphatase, and NAD-ME) have been reported to be significantly upregulated following Pi deprivation of plant cells (Duff et al., 1989b; Hoffland et al., 1992; Theodorou et al., 1992; Pilbeam et al., 1993; Johnson et al., 1994; Nagano et al., 1994; Theodorou and Plaxton, 1994; Ciereszko et al., 1998, 2001; Murley et al., 1998; Moraes and Plaxton, 2000; Juszczuk et al., 2001; Massonneau et al., 2001; Fernie et al., 2002; Juszczuk and Rychter, 2002; Hammond et al., 2003, 2004; Toyota et al., 2003; Vance et al., 2003; Plaxton, 2004a; Shane et al., 2004; Ticoni and Abel, 2004; Peñaloza et al., 2005). It has been hypothesized that these enzymes represent Pi starvation-inducible bypasses to the adenylate or Pi-dependent glycolytic enzymes thereby facilitating glycolytic flux during severe Pi stress when the intracellular levels of Pi and adenylates are very low (Duff et al., 1989a; Theodorou and Plaxton, 1993; Plaxton and Carswell, 1999; Vance et al., 2003; Plaxton, 2004a, 2005). The proposal that PPi-PFK substitutes for ATP-PFK under conditions of Pi starvation has been challenged by the observation that the growth characteristic of Pi-limited transgenic tobacco plants with markedly reduced PPi-PFK are indistinguishable from those of wild type plants (Paul et al., 1995). However, although these transgenic plants had greatly reduced amounts of PPi-PFK, the enzyme was not completely abolished. The residual PPi-PFK activity may have still been sufficient to accommodate a reduced glycolytic flux under Pi-limiting conditions. The levels of the PPi-PFK activator Fru-2,6-P2 in the transgenic lines were significantly higher relative to those of the control plants (Paul et al., 1995). This may have allowed the residual PPi-PFK to become more fully activated in vivo. A subsequent study of transformed tobacco lines over-expressing mammalian 6-phosphofructo-2-kinase concluded that the contribution of ATP-PFK and PPi-PFK to glycolysis respectively decreases and increases under conditions of Pi-deficiency (Fernie et al., 2002). As discussed above, PEPC can theoretically function as a glycolytic enzyme by indirectly bypassing the reaction catalyzed by PKc (Figure 2). The PEPC-MDH-ME bypass of PKc may be important during nutritional Pi deprivation when PKc activity may become ADP limited (Duff et al., 1989b; Nagano et al., 1994; Moraes and Plaxton, 2000; Juszczuk et al., 2001; Juszczuk

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and Rychter, 2002; Plaxton, 2004b, 2005; Le Roux et al., 2005). Compared to Pi-sufficient controls, PEPC activity was found to be 5- and 3-fold greater in extracts of Pi-deficient Brassica nigra (black mustard), Brassica napus (rapeseed), and Catharanthus roseus (Madagascar periwinkle) suspension cells, respectively (Duff et al., 1989b, Nagano et al., 1994; Moraes and Plaxton, 2000). Metabolite determinations and kinetic studies of PKc and PEPC in C. roseus suggested that the contribution of PEPC to the metabolism of PEP increased in Pi-starved cells in vivo (Nagano et al., 1994). Further evidence for the operation of this PKc bypass in Pi-deprived C. roseus was provided by the rapid release of 14 CO2 from organic compounds derived from fixed H14 CO3 and from [14 C]-malate (Nagano et al., 1994). Pi is a by-product of the reactions catalyzed by the glycolytic bypass enzymes PPi-PFK, PEPC, PEP phosphatase, PPDK, as well as the H+ -PPiase of the tonoplast (Figure 2). Thus, these enzymes could play a dual role during Pi stress. They may facilitate respiration or active transport of H+ ions across the tonoplast by circumventing an adenylate-dependent reaction, while simultaneously recycling Pi for its reassimilation into the metabolism of the Pi-starved cells (Duff et al., 1989a, 1989b; Theodorou et al., 1992; Theodorou and Plaxton, 1993, 1994, 1996; Kondracka and Rychter, 1997; Palma et al., 2000; Plaxton, 2004a). It should be noted, however, that in contrast to PEPC, PPi-PFK, PEP phosphatase, and the tonoplast H+ -PPiase, we are unaware of any published evidence indicating a glycolytic bypass role for PPDK in Pi-deficient plants. Moreover, we have been unable to detect PPDK activity in extracts of Pi-sufficient or Pi-starved B. napus suspension cells (Podestá and Plaxton, unpublished data). Nevertheless, future research should consider possible situations during which cytosolic PPDK might function with PPi-PFK to utilize PPi while providing dual roles as a glycolytic bypass and Pi-recycling enzymes in Pi-starved plants. Scavenging of Pi from extracellular sources may also be aided by the enhanced excretion of organic acids (i.e., citric and malic acids) brought about by the upregulation of PEPC, citrate synthase, and MDH (Ryan et al., 2001; Vance et al., 2003). Roots and suspension cell cultures of Pi-starved plants have been demonstrated to markedly upregulate PEPC, and this has been correlated with the excretion of significant levels of malic and/or citric acids (Duff et al., 1989b; Hoffland et al., 1992; Johnson et al., 1994; Nagano et al., 1994; Moraes and Plaxton, 2000; Massaonneau et al., 2001; Ryan et al., 2001; Vance et al., 2003; Shane et al., 2004; Plaxton, 2004a; Cramer et al., 2005; Peñaloza et al., 2005). This results in acidification of the rhizosphere, which thereby contributes to the solubilization and assimilation of mineral Pi from the environment.

conserving pathways of miETC provides a mechanism whereby respiratory flux may be maintained under conditions when the availability of ADP and/or Pi are restrictive (i.e., during severe Pi deprivation). Groundbreaking research from Anna Rychter’s laboratory (Rychter and Mikulska, 1990; Rychter et al., 1992) has been augmented by a variety of subsequent studies demonstrating that plants acclimate to Pi stress by the upregulation and/or increased engagement of the non-energy conserving (i.e., rotenone- and/or cyanide-insensitive) pathways of the miETC (Figures 2 and 3) (Hoefnagel et al., 1993, 1994; Gonzàlez-Meler et al., 2001; Juszczuk et al., 2001; Vanlerberghe and Ordog, 2002; Millenaar and Lambers, 2003; Shane et al., 2004; Plaxton, 2004a; McDonald and Vanlerberghe, 2005; Sieger et al., 2005). This allows continued functioning of the TCA cycle and miETC with limited ATP production and may thereby contribute to the survival of Pi-deficient plants. This has been corroborated by the impaired growth and metabolism of Pi-deprived transgenic tobacco plants unable to synthesize a functional AOX protein (Parsons et al., 1999). Lack of AOX under Pi limitation was recently correlated with increased levels of proteins commonly associated with oxidative stress and/or stress injury (Sieger et al., 2005). It was concluded that AOX respiration provides a crucial adaptive mechanism by which plant cells can modulate their growth response to Pi availability and that AOX also has nutrient-specific roles in maintaining cellular redox and carbon balance (Sieger et al., 2005). 3.

Transcript Profiling Indicates that Pi Starvation Inducibility of Glycolytic and Mitochondrial Electron Transport Bypass Proteins Is Widespread The recent application of microarray and related “high throughput” transcript profiling technologies has allowed researchers to simultaneously catalogue the effects of Pi deficiency on the expression of thousands of genes in plants such as Arabidopsis, rice, and white lupin (Hammond et al., 2003, 2004; Vance et al., 2003; Wasaki et al., 2003; Wu et al., 2003; Ticconi and Abel, 2004; Misson et al., 2005). Interestingly, the enhanced expression of genes encoding alternative glycolytic enzymes (such as PPi-PFK) and miETC proteins (such as AOX) that do not require Pi or adenylates as co-substrates has been observed. The differential regulation of genes involved in primary metabolism demonstrates the activation of genes involved in bypassing the ATP- and Pi-dependent enzymes, and changing respiratory metabolism required to generate energy and carbon skeletons during Pi deficiency (Hammond et al., 2004). E.

2.

Mitochondrial Electron Transport Chain Bypasses of Pi-Starved Plants The significant reductions in intracellular Pi and ADP levels that follow extended Pi deprivation will impede respiratory electron flow through the cytochrome pathway at the sites of coupled ATP synthesis. However, the presence of non-energy

Efficient Plant Respiration May Be Promoted by Glycolytic and TCA Cycle Multi-Enzyme Metabolons, and Mitochondrial Electron Transport Protein Supercomplexes Many current biochemistry textbooks continue to depict metabolism as occurring by a series of independent enzymes that are randomly dispersed in a uniform aqueous environment.

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However, accumulating evidence indicates that metabolic pathways thought to be entirely soluble may exist in vivo as organized multi-enzyme complexes (or metabolons) that are loosely associated with membrane fractions or cytoskeletal proteins in the cell (Srere, 1987; Hrazdina and Jensen, 1992; Ovadi and Srere, 2000; Plaxton, 1996, 2004b; Winkel, 2004). Due to high intracellular protein concentrations, such interactions are far more likely in vivo than in dilute in vitro enzymological studies. For example, a massive protein concentration exists within the mitochondrial matrix and ranges from about 250 to 550 mg/ml depending upon the functional state of the organelle (Ovadi and Srere, 2000). The micro-compartmentation of enzymes and metabolic pathways that results from metabolon formation increases metabolic fluxes owing to the direct transfer or channeling of intermediates between active sites of consecutive enzymes. In addition to the kinetic advantages, metabolic channeling may: (i) reduce the concentration of the channeled intermediate in the bulk solution, thus sparing the limited solvent capacity of the cell, (ii) alter enzyme kinetic properties due to conformational changes that occur during binding, and (iii) regulate competition between branch pathways for common metabolites, and (iv) sequester toxic or reactive intermediates (Srere, 1987; Hrazdina and Jensen, 1992; Ovadi and Srere, 2000; Plaxton, 2004b; Winkel, 2004). The existence of known stable multi-enzyme complexes such as mitochondrial PDC and glycine decarboxylase complexes provides favorable evidence for the interaction of soluble enzymes that are metabolically sequential. 1.

Glycolytic and TCA Cycle Metabolons There is a large body of evidence—mostly obtained with non-plant systems—that supports the existence of glycolytic and TCA cycle metabolons (Srere, 1987; Hrazdina and Jensen, 1992; Fothergill-Gilmore and Michels, 1993; Plaxton, 1996, 2004b; Ovadi and Srere, 2000; Haggie and Verkman, 2002). Specific associations have been demonstrated in vitro between purified sequential enzymes of non-plant glycolysis or the TCA cycle, and also between these enzymes and components of the cytoskeleton (glycolysis) or miETC (TCA cycle). The development of a metabolic control theory for the muscle system concluded that flux control coefficients for enzymes of channelled pathways are usually larger than those in the corresponding nonchannelled pathway (Kholodenko et al., 1994). Complexes capable of catalyzing several consecutive steps of glycolysis or the TCA cycle have been isolated following gentle extraction of animal tissues as well as several microorganisms (Srere, 1987; Ovadi and Srere, 2000). Although the existence of a complete glycolytic metabolon is doubtful, it seems probable that physical interactions between groups of sequential glycolytic enzymes such as ATP-PFK, aldolase and NAD+ -dependent GAPDH exist in vivo in animal cells (Fothergill-Gilmore and Michels, 1993; Ovadi and Srere, 2000; Plaxton, 2004b). Analysis of 13 C labeling patterns in metabolites derived from TCA cycle intermediates conclusively demonstrated in vivo channeling within the TCA cycle of the brewer’s yeast, Saccharomyces cerevisiae (Sumegi

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et al., 1990). All of the enzymes of the TCA cycle, including several from plant systems, have been shown to specifically bind to the matrix surface of the mitochondrial i.m. (Laties, 1983; Gross, 1990; Ovadi and Srere, 2000). More recently, four TCA cycle enzymes from mammalian mitochondria were tagged with green fluorescent protein so that their intramitochondrial location could be directly assessed via fluorescence microscopy (Haggie and Verkman, 2002). The results provided convincing additional evidence that the four enzymes were indeed physically associated within the mitochondrial matrix. The molecular mechanisms that control the extent of enzyme: enzyme interactions in vivo are not fully understood. Binding of enzymes in vitro is influenced by numerous factors including pH, concentration of substrates, products and allosteric effectors, and enzyme phosphorylation. For example, during muscle contraction ATP-PFK is phosphorylated in vivo which promotes its physical association with contractile proteins (actin/myosin), and hence other metabolically sequential glycolytic enzymes (Srere, 1987; Ovadi and Srere, 2000; Plaxton, 2004b). This is thought to contribute to the large increase in glycolytic flux (Pasteur effect) characteristic of the initiation of anaerobic muscle work in mammals. Although there is compelling evidence for the direct transfer of substrates between respiratory metabolons of non-plant cells, these possibilities have received relatively scant attention in plants. However, several studies indicate the probable existence of analogous plant metabolons in the Calvin-Benson cycle as well as plant respiratory pathways (Fernie et al., 2004; Plaxton, 2004b; Sweetlove and Fernie, 2005). For example the well characterized stimulation of respiration that accompanies aging of storage root slices may result, in part, from an association of glycolytic enzymes with a particulate fraction of the cell promoting an elevated glycolytic rate (Moorhead and Plaxton, 1988). A co-immunoprecipitation study demonstrated that cytosolic aldolase specifically interacts with the metabolically sequential ATP-PFK and PPi-PFK in carrot storage roots (Moorhead and Plaxton, 1992). Similarly, PEPC and MDH of CAM and C4 leaves appear to form a functional complex that favors PEPC activity (Queiroz-Claret and Queiroz, 1992; Baret et al., 1995). Furthermore, a study of pea leaf mitochondria concluded that metabolic microcompartments containing different complements of photorespiratory versus TCA cycle enzymes exist within the same mitochondrial matrix (Wiskich et al., 1990). This was proposed to help maintain daytime TCA cycle flux in the presence of a large photorespiratory (glycine oxidation) flux. Some glycolytic enzymes such as hexokinase appear to reversibly associate with the external surface of animal mitochondria (Ovadi and Srere, 2000). A recent proteomic study of highly purified mitochondria from Arabidopsis suspension cells demonstrated an unexpected association between this organelle with seven of the ten enzymes that constitute the standard glycolytic pathway (Giegé et al., 2003). Glycolytic enzyme activity and protease protection assays confirmed the presence of the glycolytic pathway on the external surface of the isolated

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Arabidopsis mitochondria. The in vivo mitochondrial association of aldolase and enolase was confirmed by the expression of the corresponding enolase/aldolase-yellow fluorescent protein fusions in Arabidopsis protoplasts. This was corroborated by the ability of highly purified Arabidopsis mitochondria to metabolize 13 C-labeled Glc into 13 C-labeled TCA cycle intermediates when supplied with the appropriate cofactors (Giegé et al., 2003). This microcompartmentation of the glycolytic pathway was suggested to provide pyruvate at high concentrations directly to the mitochondrion while minimizing competition for glycolytic intermediates from other pathways such as the oxidative pentose-phosphate pathway. Further research is required to establish whether the mitochondrial association of the glycolytic pathway observed in vitro (Giegé et al., 2003) actually operates in vivo to channel significant amounts of glycolyticallyderived pyruvate into the mitochondria, or whether this association might have additional and/or unrelated functions. 2.

Respirasome and ATP Synthasome Supercomplexes of the Mitochondrial Inner Membrane The mitochondrial i.m. is one of the most proteinaceous biological membranes (Srere, 1987). It has been estimated that 50 percent of its surface is occupied by protein, and of this about 35 percent of the total protein of the i.m. is composed of the miETC and ATP synthase. According to the standard fluid mosaic model of membrane structure-function, the protein complexes of the miETC are randomly arranged and freely diffuse in a lateral direction within the i.m. However, there is a probable association of miETC components into highly ordered supercomplexes. The use of mild i.m. solubilization techniques and blue-native polyacrylamide gel electrophoresis to separate native protein complexes according to size led to the initial discovery of respiratory supercomplexes in bacteria, and subsequently in yeast and mammalian mitochondria (Schägger and Pfeiffer, 2000). The term respirasome was suggested for these miETC supercomplexes, which can autonomously carry out respiration in the presence of cytochrome c and ubiquinone. miETC supercomplexes are believed to increase the rates of respiratory electron transfer, as well as the capacity for miETC protein insertion into the i.m., while reducing the frequency of unwanted side reactions (Schägger and Pfeiffer, 2000). Supercomplexes of Complex I and III (with a variety of different stoichiometries) from Arabidopsis, bean, barley, rice, and potato mitochondria were recently isolated and analyzed (Eubel et al., 2003, 2004a, b; Krause et al., 2004; Millar et al., 2004a). Although the use of gentler techniques demonstrated that Complex IV also associates in a supercomplex with Complex I and III in a potato mitochondria, a supercomplex of Complex I and III was still the most abundant form (Eubel et al., 2004a). These studies indicate that respirasomes of Complex I, III, and IV is a conserved feature of eukaryotic mitochondria. By contrast, Complex II and AOX were not incorporated into the plant miETC supercomplexes under the conditions applied. It was hypothesized that supercomplex formation between Complex I and III limits ac-

cess of AOX to its substrate ubiquinol and possibly contributes to the control of cyanide resistant respiration in plants (Eubel et al., 2003). To date, there are conflicting reports as to whether plant AOX is present in such supercomplexes (Eubel et al., 2003; Navet et al., 2004), and no evidence for the presence of alternate NAD(P)H dehydrogenases of the plant miETC. The fact that the component respiratory complexes thus far identified can associate together with different stoichiometries suggests that the formation of supercomplexes could be an important mechanism that controls the flow of electrons through miETC (Eubel et al., 2004b; Sweetlove, 2005). Further research is required to establish if and how such respirasome supercomplexes mediate the rate or path of electron transport in plants. Additional studies in animal systems have demonstrated that ATP synthase exists in a supercomplex termed the ATP synthasome in association with both the Pi transporter and adenine nucleotide translocator that exchanges ATP and ADP across the i.m. (Ko et al., 2003). This provides an obvious mechanism whereby the substrates for ATP synthesis (ADP and Pi) are channeled directly to ATP synthase upon their import across the i.m., with the concomitant export of the product ATP to the cytosol. It will be of interest to assess whether a similar ATP synthasome supercomplex exists in the i.m. of plant mitochondria.

F.

Tissue and/or Developmental-Specific Isozymes of Plant Respiratory Enzymes Biochemical, genomic, and proteomic evidence indicate that tissue- or developmental-specific isozymes of plant respiratory enzymes are widespread, and that this could be important in contributing to cell-specific metabolism in vascular plants. For example, PEPC and its corresponding PEPC protein kinase are encoded by a small multigene family in which the expression of each member is controlled by exogenous and/or endogenous stimuli in a tissue-specific fashion (Chollet et al., 1996; Fontaine et al., 2002; Lepiniec et al., 2003; Marsh et al., 2003; Xu et al., 2003; Sullivan et al., 2004; Izui et al., 2004; Nimmo, 2005). Similarly, detailed biochemical studies of purified PKc s from expanding leaves, germinating endosperm and cotyledons, and developing endosperm of the castor oil plant clearly demonstrate that as with mammalian PKs, the vascular plant enzyme exists as tissue-specific isozymes that exhibit substantial differences in their various molecular and kinetic/regulatory properties (Plaxton, 1989; Podestá and Plaxton, 1991, 1992, 1993, 1994b; Hu and Plaxton, 1996; Turner et al., 2005). These differences are believed to contribute to tissue-specific respiratory metabolism within the different tissues of the castor plant (Turner et al., 2005). The surprising biochemical complexity of vascular plant PK has been corroborated by recent genome sequencing initiatives. Currently the NCBI-CDD (Marchler-Bauer et al., 2002) identifies 15 Arabidopsis and 8 rice genes that putatively encode different PK polypeptides. Likewise, various individual components of plant PDC, the TCA cycle and miETC (including AOX and UCP) are encoded by multigene families (Tovar-Mendez

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et al., 2003; Vance et al., 2003; Fernie et al., 2004; McDonald and Vanlerberghe, 2005). Evidently, a complete description of the organization and control of plant respiratory metabolism will necessitate determination of the tissue- and/or developmentalspecific expression of the various isozymes of glycolysis, PDC, the TCA cycle, and the miETC, coupled with thorough analyses of their individual biochemical and kinetic/regulatory characteristics. The proteomic approach now offers researchers the opportunity to undertake a systematic examination of which isozymes are expressed in different plant tissues or subcellular compartments (Sweetlove, 2005). For example, a recent proteomic study identified 53 proteins in rice anthers at the young microspore stage (Imin et al., 2001). Amongst these proteins were several enzymes associated with respiratory metabolism including triose-phosphate isomerase, phosphoglyceromutase, enolase, MDH (cytosolic), PDC, aconitase, and subunits of mitochondrial Complex I and ATP synthase. The identification of these enzymes of glycolysis and mitochondrial respiration as particularly abundant in the anther tissues indicates the importance of respiratory energy supply for pollen formation (Imin et al., 2001). This has been corroborated by a study demonstrating that antisense knockout of the mitochondrial PDC E1α subunit in tobacco anthers leads to mitochondrial male sterility, a phenocopy of the cytoplasmic male sterility (CMS) in sugar beet (Yui et al., 2003). Similarly, the CMSII tobacco mutant is impaired in Complex I function (Dutilleul et al., 2003), whereas antisense repression of mitochondrial citrate synthase led to a specific disintegration of the ovary tissues of the flowers (Landschütze et al., 1995). The most comprehensive study of tissue-specific proteomes of a single plant species was conducted in rice (Koller et al., 2002). Mass spectrometric analysis of rice root, seed, and leaf proteomes resulted in identification of over 2,500 unique proteins; 1,002 leaf-specific proteins, 1,350 root-specific proteins, and 877 seed-specific proteins. In accordance with the distribution of functional classes of proteins encoded by the rice genome, metabolic proteins represented the second most abundant class of proteins in the dataset. Within this dataset, the main metabolic pathways of primary metabolism were fully represented (glycolysis/gluconeogenesis, TCA cycle, oxidative pentose pentose-phosphate pathway and most pathways of amino acid biosynthesis) and were found to occur within each of the three tissues that were examined (Koller et al., 2002). However, there were distinct tissue-specific patterns of the distribution of the isozymes of these enzymes. Some plastidial and cytosolic isozymes of glycolysis were found to be present in all tissues (i.e., aldolase, triose-phosphate isomerase, GAPDH, 3-PGA kinase, and phosphoglyceromutase), but the majority of metabolic enzymes were found to be tissue-specific in their location (Koller et al., 2002). Recent mass spectrometry analyses of the Arabidopsis and rice mitochondrial proteomes, coupled with bioinformatic analyses of the corresponding genomes have been invaluable in this regard (Bykova et al., 2003; Heazlewood et al., 2003a, 2003b,

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2004; Herald et al., 2003; Millar et al., 2004a, 2004b, 2005; Sweetlove, 2005; Taylor et al., 2005). As the main function of mitochondria is the production of cellular ATP and biosynthetic precursors, it is not surprising that the enzymes associated with the TCA cycle and miETC dominate the mitochondrial proteome (Herald et al., 2003; Millar et al., 2004b, 2005; Sweetlove, 2005; Taylor et al., 2005). However, these studies have unexpectedly revealed new metabolic, regulation, and signalling pathways in plant mitochondria, providing key data for future experimental analyses of plant mitochondrial biogenesis and function. In a recent large-scale analysis of the Arabidopsis mitochondrial proteome, almost 20 percent of the 416 identified proteins were of unknown function (Heazlewood et al., 2004), suggesting a wealth of undiscovered mitochondrial functions in plants. Such proteins may have specific roles in respiration, or may exist as “moonlighting” proteins having additional functions that are not directly related to respiration. For example, a protease associated with Complex III of the plant miETC hydrolyzes the N-terminal transit peptide of mitochondrial-targeted proteins during their import from the cytosol across the i.m. (Braun et al., 1992). III. METABOLIC CONTROL OF PLANT RESPIRATION Because plant glycolysis, PDC/TCA cycle, and the miETC are tightly coupled processes, there must be mechanisms to integrate them so as to accommodate for short- or long-term changes in cellular demand for C-skeletons, reducing power, and ATP. A.

Coarse versus Fine Control of Plant Respiration It is possible to group mechanisms of metabolic control into two major classes on the basis of the relative lengths of time that they take to bring about a change in the velocity of a particular enzyme. These are coarse and fine metabolic control (Plaxton, 2004b). 1.

Coarse Control Coarse control is a longer-term energetically expensive response that is achieved through changes in the total cellular population of enzyme molecules. Thus, any alteration in the rates of gene expression (i.e., transcription, translation, mRNA processing or degradation), or proteolysis (turnover) can be considered as coarse metabolic control (Plaxton, 2004b). There are numerous reports outlining significant alterations in the amounts of plant respiratory and associated enzymes and/or their corresponding transcripts during plant development, over the diurnal cycle in leaves, and following longer-term environmental (adaptive) changes (Duff et al., 1989b; Wong et al., 1990; Blakeley et al., 1992; Sangwan et al., 1992; Podestá and Plaxton, 1994a; Theodorou et al., 1992; Theodorou and Plaxton, 1993, 1994; Gottlob-McHugh et al., 1995; Plaxton, 1996, 2004a, 2005; Givan, 1999; Golombek et al., 1999; Moraes and Plaxton, 2000; Scheible et al., 1997, 2000; Gonzàlez-Meler et al., 2001; Juszczuk et al., 2001; Mooney et al., 2002; Ruuska et al., 2002; Stitt et al., 2002; Vanlerberghe and Ordog, 2002; Vance et al.,

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2003; Daley et al., 2003; Hammond et al., 2003, 2004; Tang et al., 2003; Uhde-Stone et al., 2003; Wang et al., 2003; Wasaki et al., 2003; Wu et al., 2003; Fernie et al., 2004; Foyer et al., 2005; Hajduch et al., 2005; McDonald and Vanlerberghe, 2005; Misson et al., 2005; Turner et al., 2005). An understanding of the transcriptional and translational regulation of plant genes encoding respiratory enzymes is beginning to emerge (Stitt et al., 2002; Daley et al., 2003; Fernie et al., 2004; Thimm et al., 2005; Wasaki et al., 2003; Wu et al., 2003). It is important to note that an underlying assumption of many transcript profiling studies is that the expression of a gene at the mRNA level is a quantitative indicator of function of the encoded enzyme. Thus, an n-fold increase in transcript levels (detected via Northern blots or microarray screening) equates to n-fold more enzyme and hence n-fold more activity in vivo. However, it is becoming apparent that this assumption does not always reflect reality (Plaxton, 2004b; Gibon et al., 2004; Sweetlove and Fernie, 2005). For example, quantification of glycolytic fluxes in three parasitic protists found that these did not correlate proportionally with the concentration of the corresponding glycolytic enzymes; i.e., relative to various fine metabolic control mechanisms, gene expression alone exerts little control on glycolytic flux (ter Kuile and Westerhoff, 2001). Moreover, alterations in transcript abundance do not necessarily translate into a correlated change in protein amount, or vice versa (Gibon et al., 2004; Sweetlove and Fernie, 2005). It is thus crucial that transcript profiling of respiratory enzymes is paralleled by studies quantifying relative amounts of the encoded proteins and their enzymatic activities, and ideally in vivo flux studies through the relevant pathways (Schwender et al., 2004). In this regard, a high throughput enzyme assay platform developed by Mark Stitt’s group has been used to investigate the relationship between the abundance of a variety of respiratory enzymes and their corresponding transcripts in Arabidopsis leaves during the diurnal cycle and during prolonged darkness (Gibon et al., 2004). There was a poor quantitative correlation with transcript abundance and the corresponding enzyme protein amount. Generally, the changes in protein were much smaller than the respective changes in transcript abundance (Gibon et al., 2004). Moreover, there was a temporal delay between changes in transcript abundance and the corresponding enzymes. This reflects the fact that the processes of protein synthesis and turnover that regulate protein abundance are relatively slow in comparison to gene transcription. Relative to gene expression, far less is known about the mechanisms governing the proteolytic degradation of plant respiratory enzymes. Enzymes that coexist in the same subcellular compartment may exhibit vastly different turnover rates, ranging from several minutes to hundreds of hours (Plaxton, 2004b). In general, larger oligomeric proteins that display complex biological properties tend to show much shorter half-lives in vivo than do less complex monomeric proteins. It is becoming apparent that proteolysis of plant respiratory enzymes can be selectively targeted and may be initiated in response to specific stimuli. For example, phosphorylation of plant SuSy and PKc was reported

to target these enzymes for their ubiquination and subsequent degradation by the proteasome (Tang et al., 2003; Hardin et al., 2003, 2004). Vicia fabia PEPC and maize PEPC kinase both appear to be degraded via ubiquitin-dependent proteolysis (Schulz et al., 1993; Agetsuma et al., 2005), whereas an asparginyl endopeptidase has been suggested to play a role in the turnover of PEPC and leucoplast PK in maturing castor beans (Cornel and Plaxton, 1994; Negm et al., 1995, Crowley et al., 2005). Cotelle et al. (2000) reported the phosphorylation dependent binding of 14-3-3 proteins (see below) to a wide range of plant metabolic enzymes. In studies of sugar-starved Arabidopsis cell cultures, it appeared that a general influence of 14-3-3 binding was to protect the 14-3-3 bound proteins from proteolytic degradation (Cotelle et al., 2000). Thus, certain phosphorylation sites may constitute phosphodegrons that trigger proteolytic degradation. Nevertheless, this area of research is in its infancy and a very considerable effort will be required before the complex mechanisms that control the in vivo proteolysis of plant respiratory enzymes is fully understood. 2.

Fine Control By modulating the activity of pre-existing enzymes, fine controls function as “metabolic transducers” that sense the momentary metabolic requirements of the cell, and adjust the rate of metabolite flux through the various pathways accordingly (Plaxton, 2004b). Diverse fine control mechanisms have evolved to regulate the interplay of glycolysis with the TCA cycle and miETC, so as to coordinate glycolytic and TCA cycle flux with cellular needs for energy and anabolic precursors. The traditional view of fine metabolic control is that this is accomplished through altering the activity of at least one “pacemaker” enzyme (or “rate determining step”) of the pathway. Substantial efforts have been directed to identifying the pacemaker enzyme(s) of plant respiration, as well as the complex mechanisms that serve to modulate the activities of these key control enzymes. However, one problem with the pacemaker approach to metabolic control is that these studies are for the most part qualitative, rather than quantitative. Designation of an enzyme as a pacemaker is not based upon any direct measurement of the precise contribution of each enzyme in a pathway to the control of pathway flux, or how the degree of control exerted by each enzyme might vary under differing physiological circumstances. Furthermore, biological systems display regulatory properties that are not possessed by their isolated components. For instance, it would be impossible to understand how a mitochondrion functions in respiration by only studying purified mitochondrial enzymes and miETC proteins in isolation of each other and mitochondrial membranes. Thus, another important approach to the problem of metabolic control is to analyze the whole system. B.

Metabolic Control Analysis The metabolic control analysis (MCA) theory developed by Kacser and Burns (1973) attempts to provide a quantifiable

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mechanism for probing intact biological systems and interprets resultant data without preconceived notions as to which enzymes in a pathway are rate-determining steps or pacemakers (ap Rees and Hill, 1994; Thomas et al., 1997; Thomas and Fell, 1998; Cornish-Bowden, 1999; Giersch, 2000; Plaxton, 2004b). An important tenet of MCA theory is that metabolic control is shared among many if not all enzymes in a pathway. The various formulations and concepts of the mathematical models of MCA have given rise to considerable debate over the meaning and usefulness of MCA. For example, ATP-PFK has been traditionally considered as an important pacemaker enzyme of the glycolytic pathway in eukaryotes (Plaxton, 1996; Thomas et al., 1997; Thomas and Fell, 1998; Cornish-Bowden, 1999; Givan, 1999). Eukaryotic ATP-PFK catalyzes the first unique step of glycolysis, a non-equilibrium reaction in vivo, and shows a strong positive crossover concomitant with glycolytic stimulation (Plaxton, 1996, 2004b; Givan, 1999). Purified ATP-PFKs are multimeric enzymes that generally display sigmoidal substrate (Fru-6-P) saturation kinetics as well as complex and potent allosteric control by numerous effectors, the levels of which are controlled by the physiological and hormonal status of the tissue. For example, the role of the signal metabolite Fru-2,6-P2 as a potent allosteric activator of animal and yeast ATP-PFKs is well established (Stitt, 1990; Plaxton, 1996, 2004b; Givan, 1999; Thomas and Fell, 1998; Cornish-Bowden, 1999). However, the use of molecular genetic techniques for the selective overexpression of ATP-PFK in transgenic yeast, mammals, and vascular plants has failed to yield the significant increases in glycolytic flux or respiration that were expected (Stitt and Sonnewald, 1995; Thomas et al., 1997; Thomas and Fell, 1998; Cornish-Bowden, 1999; Plaxton, 2004b). It appears that the elevated ATP-PFK concentration in these transgenic organisms was somewhat compensated for in vivo by changes in the levels of ATP-PFK allosteric activators and inhibitors (Stitt and Sonnewald, 1995). Animal and plant ATP-PFK flux control coefficients were determined to be very small, leading to the surprising conclusion that this enzyme exerts very little or no control over glycolytic flux or respiration in vivo (Thomas et al., 1997; Thomas and Fell, 1998; Cornish-Bowden, 1999). Although some of the leading proponents of MCA have interpreted these findings to dispute the traditional concept of ATP-PFK as a pacemaker enzyme of glycolysis, several of the same MCA advocates have nevertheless agreed that there is little doubt that “. . . control of PFK activity plays a part in glycolytic flux control” (Thomas and Fell, 1998) and “. . . PFK makes an important contribution to the control of glycolysis in most cells (Cornish-Bowden, 1999). A possible explanation for this “PFK paradox,” is that MCA also indicated that significant flux control of glycolysis and respiration lies in the metabolism of key feedback inhibitors of ATP-PFK, namely ATP and citrate in yeast and mammalian cells, and PEP in plant cells (Thomas et al., 1997, Plaxton, 2004b). There is ample evidence that low flux control coefficient enzymes can play a major role in the control of metabolic pathways (Stitt and Sonnewald, 1995). This would suggest the somewhat contradictory (and con-

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fusing!) conclusion, that although the flux control coefficient for ATP-PFK may be low, it does indeed play a pivotal role in the in vivo control of carbohydrate catabolism in most cells. Most metabolic biochemists would likely agree that flux control of a metabolic pathway is generally dominated by a minority of its component enzymes (i.e., the pacemakers), although under different physiological conditions the degree of control exerted by the individual enzymes may vary. MCA of non-plant respiration shows that the contribution of the various enzymes to the overall flux control was highly dependent upon the organism, tissue, and physiological condition (Fother-Gillgilmore and Michels, 1993; Thomas and Fell, 1998; Cornish-Bowden, 1999). Changes in physiological conditions causes a redistribution of the control that each enzyme exerts on respiratory carbon flux, and this no doubt applies to plant respiration. However, the application of MCA to plant respiration has been greatly complicated by the inherent flexibility of plant respiratory metabolism and miETC. Nevertheless, as discussed below, MCA has provided some important contributions to our understanding of plant respiratory control.

C.

Key Control Enzymes of Plant Respiratory Carbon Metabolism Any explanation of the integration and control of plant respiration must include consideration of the factors that serve to regulate and coordinate the activities of PDC along with relevant glycolytic, TCA cycle, and miETC enzymes. The traditional (or qualitative) approach to metabolic control involves identifying the key pacemaker enzymes of a pathway, and the mechanisms whereby their activities are controlled in vivo. These reactions are often characterized by their strategic position in a pathway, by being greatly displaced from equilibrium in vivo, and were classically recognized by showing that their substrate(s): product(s) concentration ratio changed in the opposite direction to the flux when the latter was varied (Kubota and Ashihara, 1990; ap Rees and Hill, 1994; Plaxton, 1996, 2004b; Givan, 1999). Elucidation of the fine control of plant respiration is complicated by the alternative reactions that exist in the plant glycolysis, TCA cycle, and miETC. Flexibility in the organization of the plant respiratory metabolism implies flexibility in respiratory control and this will vary according to the specific tissue, its developmental stage, and the external environment. Nevertheless, various approaches have demonstrated that, as in non-plant systems, fine control of plant respiratory carbohydrate metabolism is primarily exerted by those enzymes that catalyze reactions involved in the conversion of hexose to hexose-6phosphate (hexokinase and fructokinase), Fru-6-P to Fru-1,6-P2 (ATP- and PPi-PFK), PEP to pyruvate or OAA (PKc and PEPC), pyruvate to acetyl-CoA (PDC), citrate to isocitrate (citrate synthase), isocitrate to 2-OG (isocitrate dehydrogenase), and 2OG to succinyl-CoA (2-OG dehydrogenase) (Figure 5) (Adams and Rowan, 1970; Kobr and Beevers, 1971; Paul et al., 1978; Dennis and Greyson, 1987; Beaudry et al., 1989; Kubota and

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FIG. 5. Regulation of plant respiration includes the fine or acute metabolic control of key cytosolic and mitochondrial enzymes by allosteric effectors and covalent modification. The central role of PEP in providing a “bottom up” type of control of glycolysis is highlighted. Also depicted are some of the most important control sites of mitochondrial respiration. Dotted arrows with a circled plus and minus sign indicate enzyme activation and inhibition, respectively, by allosteric effectors. Shaded arrows with a circled plus and minus sign denote enzyme activation and inhibition, respectively, by reversible covalent modification (phosphorylationdephosphorylation or disulfide-dithiol interconversion). Abbreviations are as defined in the text and as follows: DHAP, dihydroxyacetone-phosphate; PFK-2, bifunctional 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase Trred , reduced thioredoxin.

Ashihara, 1990; Klöck and Kreuzberg, 1991; Geigenberger and Stitt, 1991; Hatzfield and Stitt, 1991; Vanlerberghe et al., 1992; Gauthier and Turpin, 1994; Huppe and Turpin, 1994; Plaxton, 1996; Hill, 1997; Givan, 1999; Siedow and Day, 2000; Podestá, 2004; McDonald and Vanlerberghe, 2005). The partitioning of electrons between the phosphorylating versus non-energy conserving bypasses of miETC is also under significant fine control (Møller, 2001; Siedow and Day, 2000; Siedow and Umbach, 2000; Millenaar and Lambers, 2003; Rasmusson et al., 2004; McDonald and Vanlerberghe, 2005).

D.

Specific Mechanisms of Fine Metabolic Control as Applied to Plant Respiration Four mechanisms of fine control that can potentially modulate the activity of a preexisting enzyme are: variation in substrate(s) concentration, variation in pH, allosteric effectors, and covalent (post-translational) modification (Plaxton, 2004b). Examples of how each mechanism may apply to the control of plant respiration are briefly considered below. It has been argued that the first committed step of plant glycolysis is the conversion of Fru-6-P to Fru-1,6-P2 (Dennis and Greyson, 1987). Thus, our

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emphasis is the fine control of plant respiration beginning with Fru-6-P. 1.

Fine Control #1: Variation in Substrate Concentration Changes in substrate concentrations that normally occur in vivo, do not generally play a major role in metabolic control (Plaxton, 2004b). Nevertheless, control of several plant respiratory enzymes may be achieved, in part, through alterations in the in vivo level of a cosubstrate. For example, as discussed above the cellular pools of adenylates and Pi become severely depressed during long-term Pi deprivation (Ashihara et al., 1988; Duff et al., 1989b; Lauer et al., 1989; Dancer et al., 1990; Rychter and Mikulska, 1990; Rychter et al., 1992; Lee and Ratcliffe, 1993; Rao and Terry, 1995; Plaxton, 1996, 2004a; Plaxton and Carswell, 1999; Le Roux et al., 2005). This decline has been proposed to: (i) restrict the activities of the adenylateand Pi-dependent glycolytic enzymes, as well as the phosphorylating (cytochrome) pathway of mitochondrial respiration; and thereby (ii) promote the flux of hexose-phosphates through inducible adenylate- and Pi-independent cytosolic glycolytic bypasses, concomitant with an increased participation of the nonenergy conserving cyanide- and rotenone-insensitive pathways of miETC (Figure 2). Particularly relevant is the intimate link between the PDC and the TCA cycle, and the miETC, since the activities of the NAD+ -dependent PDC and several TCA cycle dehydrogenases are dependent upon the regeneration of their cosubstrate NAD+ by various NADH dehydrogenases of the miETC. Conversely, high NADH concentrations lead to increased engagement of the rotenone-insensitive NADH dehydrogenase bypasses to Complex I (Hill, 1997; Møller, 2001; Rasmusson et al., 2004). Fine Control #2: Variation in pH The pH dependence of enzyme activity may be an important aspect of the fine control of plant respiration since intracellular pH can change in response to a variety of environmental factors. For example, PEPC’s activity and response to metabolite effectors are well known to be sensitive to cytosolic pH changes that may occur in vivo. All plant PEPCs examined to date demonstrate a greater activity and weaker response to various metabolite inhibitors as assay pH is increased from about pH 7 to 8 (Chollet et al., 1996; Nimmo, 2005). Glu inhibition of PKc from endosperm of developing castor beans was positively correlated with pH increases in the physiological range (Turner et al., 2005), whereas with castor bean PEPC the converse was true (Blonde and Plaxton, 2003). Therefore, cytosolic pH has been suggested to be one factor that may help to coordinate the relative flux of PEP through PKc and PEPC in developing castor beans, with the balance moving from PEPC to PKc upon cytosolic acidification and vice versa (Turner et al., 2005). The pH-dependency of PEPC has given rise to the proposed “pHstat” function of this enzyme since its activity should increase in response to cytosolic alkalinization owing to direct [H+ ] ef-

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fects on its activity as well as the enzyme’s desensitization to inhibitory metabolites (Sakano, 1998). This view has been challenged in a recent paper that thoroughly analyzes ionic movements in the plant cell in response to N assimilation (Britto and Kronzucker, 2005). According to these authors, although pH may well have a role in controlling PEPC activity, the converse (i.e., that PEPC controls intracellular pH) appears to be invalid. The control of the PKc isozymes from endosperm and cotyledons of germinating castor beans may also arise from pHdependent alterations in the enzyme’s response to a variety of metabolite inhibitors (Podestá and Plaxton, 1991, 1992, 1994b). It was proposed that an enhancement in PKc activity of germinating castor bean: (i) endosperms occurs during anaerobiosis through concerted decreases in cytosolic pH and concentrations of several key inhibitors (Podestá and Plaxton, 1991, 1992, 1993); and (ii) cotyledons will arise from reduced cytosolic pH and ATP levels caused by operation of a plasmalemma H+ -symport which powers the uptake of endosperm-derived sucrose and amino acids from the apoplast (Podestá and Plaxton, 1994b). Similarly, binding of substrates and allosteric effectors to purified potato tuber PPi-PFK, as well as the corresponding PPi-PFK kinetic and allosteric properties were markedly influenced by pH in a manner that suggested an increased glycolytic role for PPi-PFK during cytosolic acidification that accompanies hypoxia stress (Podestá and Plaxton, 2003). Similarly, the allosteric properties of pineapple leaf and B. napus suspension cell PPi-PFKs were also significantly influenced by pH changes within the physiological range; in particular, markedly enhanced activation by Fru-2,6-P2 was observed at slightly acidic pH values (Theodorou and Plaxton, 1996; Tripodi and Podestá, 1997). In the case of pineapple leaf PPi-PFK, this effect was suggested to endow the CAM cell with a greater glycolytic capacity during the night (Tripodi and Podestá, 1997).

2.

3.

Fine Control #3: Allosteric Effectors Tissue-specific expression of kinetically distinct isozymes of key control enzymes such as PEPC, PKc , and PDC gives rise to important tissue-specific differences with respect to what metabolites are key effectors of plant respiratory enzymes (Plaxton, 1996; Turner et al., 2005). Nevertheless, there are two fundamental facets of this regulation that emerge. Firstly, adenine nucleosides (and hence energy charge) do not appear to play as prominent a role in the control of plant glycolysis, PDC, and the TCA cycle as they do in most non-plant systems. This perhaps arises because: (i) PPi can function as an autonomous energy donor in the plant cytosol and mitochondrial matrix; and (ii) respiration in many plant tissues has a cardinal function in supplying C-skeletons for biosynthesis and N-assimilation, rather than ATP production per se. An exception to this view may be in anoxia-tolerant tissues such as germinating seeds. For example, ATP has been demonstrated to be a key inhibitor of the ATP-PFK from germinating cucumber seeds (Botha et al., 1988). Similarly, the switch from gluconeogenesis to glycolysis that follows anoxia stress in the endosperm of germinating castor

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beans has been suggested to arise partially from the release of inhibition of the endosperm-specific PKc isoform by ATP (Podestá and Plaxton, 1991, 1992, 1993). Potent feedback inhibition of cytosolic aldolase (reaction 11, Figure 2) from germinated mung beans by ATP was suggested to help balance cellular ATP demands with the control of cytosolic glycolysis and respiration in this tissue (Lal et al., 2005). The addition of 2 mM ATP to the purified mung bean cytosolic aldolase: (i) reduced the enzyme’s Vmax by about 2-fold, (ii) increased its K m (Fru-1,6-P2 ) value by about 4-fold, and (iii) shifted its Fru-1,6-P2 saturation kinetic plot from hyperbolic to sigmoidal (Lal et al., 2005). Although aldolase catalyzes a readily reversible reaction and is not generally considered to be an important regulatory enzyme, under certain situations it may exert significant metabolic control in vivo since: (i) moderate reductions in plastid aldolase activity by antisense techniques markedly inhibited photosynthetic CO2 fixation and growth of transgenic potato plants (Haake et al., 1998), and (ii) the in vitro activity of several purified plant aldolases is significantly modulated by physiologically relevant concentrations of various metabolite effectors (Botha and O’Kennedy, 1989; Moorhead and Plaxton, 1990; Hodgson and Plaxton, 1998; Lal et al., 2005). A second notable attribute of the allosteric control of plant respiratory enzymes is the important roles played by PEP, Pi, Fru-2,6-P2 , pyruvate, TCA cycle intermediates, aspartate (Asp), Glu, NADPH, and NADH (Plaxton, 1996; Givan, 1999; Igamberdiev and Gardeström, 2003; Turner et al., 2005). Virtually all plant ATP-PFKs examined to date show potent allosteric inhibition by PEP, and this inhibition is relieved by the activator Pi (Garland and Dennis, 1980; Dennis and Greyson, 1987; Botha et al., 1988; Häusler et al., 1989; Lee and Copeland, 1996; Plaxton, 1996, 2004b; Givan, 1999; Turner and Plaxton, 2003). It is thus the cytosolic concentration ratio of Pi:PEP that is believed to be a critical factor that controls ATP-PFK activity in vivo. PEP also appears to feedback inhibit PPi-PFK, since the response of the banana fruit PPi-PFK to its allosteric activator Fru-2,6-P2 was markedly attenuated by PEP (Turner and Plaxton, 2003). In contrast to ATP-PFK, Pi is a potent inhibitor of PPi-PFK in the forward direction and of PEP phosphatase (Duff et al., 1989a; Plaxton, 1996; Theodorou and Plaxton, 1996; Givan, 1999; Tripodi and Podestá, 1997; Theodorou and Kruger, 2001; Podestá and Plaxton, 2003; Turner and Plaxton, 2003) (Figure 5). The large (up to 50-fold) reductions in cytoplasmic Pi levels that follow extended Pi starvation have, therefore, been proposed to promote the activity of PPi-PFK and PEP phosphatase while curtailing that of ATP-PFK (Figure 2) (Duff et al., 1989a, 1989b; Theodorou and Plaxton, 1996; Plaxton, 1996, 2004b; Fernie et al., 2002). Two reports that add to the complexity of PPi-PFK control pertain to the activating effect of Fru-1,6-P2 . Nielsen (1995) and later Podestá and Plaxton (2003) showed that Fru-1,6-P2 activates plant PPi-PFK by binding to its Fru2,6-P2 allosteric site. The difficulties inherent to characterizing Fru-1,6-P2 effects, which functions both as a PPi-PFK substrate/product and allosteric effector make the physiological im-

plications of Fru-1,6-P2 activation of PPi-PFK uncertain at this point. Around 1980, the cytosolic signal metabolite Fru-2,6-P2 was discovered to reciprocally control liver glycolysis and gluconeogenesis owing to its potent activation and inhibition of ATP-PFK and fructose-1,6-bisphosphatase (FBPase), respectively. Subsequent studies demonstrated that Fru-2,6-P2 potently activates and inhibits plant PPi-PFK and cytosolic FBPase, respectively, but exerts no effect on plant ATP-PFK (Figure 5) (Sabularse and Anderson, 1981; Ball and ap Rees, 1988; Botha et al., 1988; Stitt, 1990; Theodorou and Plaxton, 1996; Plaxton, 1996, 2004a; Hodgson et al., 1998; Theodorou and Kruger, 2001; Podestá and Plaxton, 2003; Turner and Plaxton, 2003; Nielsen et al., 2004; Podestá, 2004). Recent studies of transgenic plants containing altered levels of Fru-2,6-P2 have provided unequivocal evidence of the major impact that this signal metabolite has on carbohydrate metabolism and its control (Scott et al., 1995; Truesdale et al., 1999; Draborg et al 2001, Scott et al. 2000; Rung et al., 2004; Villadsen and Nielsen, 2001). It has been established that the diurnal variation in Fru-2,6-P2 content participates in the in vivo control of photosynthetic carbon partitioning between starch and sucrose in plant leaves (Truesdale et al., 1999; Kulma et al., 2004). Although the roles of Fru-2,6P2 in plants have not been fully resolved, the mode of action on carbon metabolism appears to differ between photosynthetic and sink tissues. A Fru-2,6-P2 -dependent coordination between starch and sucrose formation has been suggested to be promoted in photosynthetic tissue (Nielsen et al., 2004). Thus, a rise in the cytosolic concentration ratio of triose-phosphate:Pi triggered by photosynthesis will decrease Fru-2,6-P2 levels owing to the reciprocal allosteric effects of Pi (activator) and triosephosphate (inhibitor) on the bifunctional 6-phosphofructo-2kinase/fructose-2,6-bisphosphatase (Nielsen et al., 2004). Interestingly, PEP potently inhibits plant 6-phosphofructo-2-kinase activity (Markham and Kruger, 2002). So, in addition to directly inhibiting ATP-PFK and PPi-PFK (see above), high PEP concentrations will diminish Fru-2,6-P2 levels which will thus further reduce glycolytic flux due to decreased allosteric activation of PPi-PFK by Fru-2,6-P2 . It is also remarkable that pyruvate, an end-product of glycolysis, activates 6-phosphofructo-2-kinase and inhibits fructose2,6-bisphosphatase (Villadsen and Nielsen, 2001). Clearly, substantial work is needed to better comprehend the complex control of plant Fru-2,6-P2 metabolism, and to more fully elucidate the influence of Fru-2,6-P2 on the overall control and integration of plant carbohydrate partitioning and respiratory metabolism. Apart from its allosteric control of the bifunctional 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase (Villadsen and Nielsen, 2001), pyruvate is also a potent activator of AOX (Siedow and Umbach, 2000; Ordog and Vanlerberghe, 2002; Millenaar and Lambers, 2003; McDonald and Vanlerberghe, 2005). This is believed to help integrate the miETC with glycolysis such that a high rate of glycolytic flux will effectively enhance miETC capacity by activating AOX. It has been demonstrated

PLANT RESPIRATION

that pyruvate accumulates in Pi-deficient bean roots, and that this may provide a mechanism for AOX activation, thereby reducing oxidative stress imposed by Pi deficiency (Juszczuk and Rychter, 2002). By contrast, the TCA cycle intermediates citrate, 2-oxoglutarate, succinate, and/or malate are effective feedback inhibitors of many plant PKc s and PEPCs (Podestá and Plaxton, 1991, 1993; Law and Plaxton, 1995; Chollet et al., 1996; Plaxton, 1996; Smith et al., 2000; Turner and Plaxton, 2000; McCloud et al., 2001; Izui et al., 2004; Nimmo, 2005; Turner et al., 2005). This provides respiratory control of glycolysis, such that glycolytic flux will increase whenever TCA cycle intermediates are consumed via anabolism or respiration and vice versa. All plant PEPCs examined to date display varying degrees of inhibition by malate, and this is usually relieved by the activator glucose-6-phosphate (Glc-6-P) or through protein kinase-mediated phosphorylation (Chollet et al., 1996; Law and Plaxton, 1995, 1997; Moraes and Plaxton, 2000; Blonde and Plaxton, 2003; Nimmo, 2005; Tripodi et al., 2005). Feedforward activation of PEPC by Glc-6-P may help to balance sucrose availability with the flux of PEP carboxylation to dicarboxylic acids via PEPC. The amino acids Asp and Glu are important feedback effectors of PEPC and PKc in green algae and plant tissues active in N-assimilation, thus providing a link between N-assimilation and the control of respiratory metabolism (Figure 5) (Huppe and Turpin, 1994; Plaxton, 1996; Givan, 1999; Foyer et al., 2005; Nimmo, 2005). For example, potent inhibition by Asp and Glu has been reported for PEPCs from diverse plant sources including green algae (Rivoal et al., 1996, 1998, 2001), soybean root nodules (Schuller et al., 1990), B. napus suspension cells (Moraes and Plaxton, 2000), cotyledons of germinated castor beans (Podestá and Plaxton, 1994b), developing castor and V. faba seeds (Golombek et al., 1999; Blonde and Plaxton, 2003), and ripened banana fruit (Law and Plaxton, 1995, 1997). Likewise, Glu is a potent allosteric inhibitor of PKc from green algae (Lin et al., 1989; Wu and Turpin, 1992), cotyledons of germinated castor beans (Podestá and Plaxton, 1994b), spinach and castor leaves (Baysdorfer and Bassam, 1984; Hu and Plaxton, 1996), B. napus suspension cells (Smith et al., 2000), ripened banana fruit (Turner and Plaxton, 2000), and developing castor bean endosperm and soybean cotyledons (Tang et al., 2003; Turner et al., 2005), but not PKc from endosperm of germinating castor beans (Podestá and Plaxton, 1991). Feedback inhibition of some plant PEPCs and PKc s by Glu provides a rationale for the known activation of the two enzymes that occurs in vivo during periods of enhanced N-assimilation when cellular Glu concentrations are reduced (Paul et al., 1978; Hammel et al., 1979; Baysdorfer and Bassam, 1984; Huppe and Turpin, 1994; Plaxton, 1996; Givan, 1999; Murchie et al., 2000; Foyer et al., 2005). In contrast to PEPC, Asp functions as an allosteric activator of various plant PKc s by effectively relieving the enzyme’s Glu inhibition (Baysdorfer and Bassham, 1984; Lin et al., 1989; Hu and Plaxton, 1996; Plaxton, 1996; Smith et al., 2000; Turner and Plaxton, 2000; Turner et al., 2005). Reciprocal control of

183

PKc and PEPC by Asp in spinach and castor leaves, ripening banana fruit, B. napus cell cultures, and endosperm of developing castor beans was suggested to provide a mechanism for decreasing flux from PEP to Asp (via PEPC and Asp aminotransferase) while promoting PKc activity whenever cytosolic Asp levels become elevated (Baysdorfer and Bassham, 1984; Hu and Plaxton, 1996; Smith et al., 2000; Turner and Plaxton, 2000; Blonde and Plaxton, 2003; Turner et al., 2005). This would be expected to occur when the cell’s demands for N are satisfied, and the overall rate of protein synthesis becomes more dependent upon ATP availability, rather than the supply of amino acids. In this instance respiration (and hence PKc ) may assume a more significant role in terms of satisfying a large ATP demand, rather than the generation of biosynthetic precursors. The production of transgenic plants expressing allosteric mutants of PEPC and PKc will help to define the precise roles of Asp and Glu in coordinating respiratory carbon flux and PEP partitioning with N-assimilation in vascular plants. Mitochondrial PDC and TCA cycle dehydrogenases such as NAD-ICDH demonstrate potent product inhibition by NADH in vitro (Igamberdiev et al., 2003; Tovar-Mendez et al., 2003; McDonald and Vanlerberghe, 2005). Their activities are probably tightly controlled by the in vivo NADH/NAD+ ratio, which provides a mechanism to balance the rate of pyruvate oxidation by PDC and the TCA cycle with the rate of miETC and thus oxidative phosphorylation. Additionally, plant mitochondrial PDC is controlled by the acetyl-CoA:CoA ratio (Tovar-Méndez et al., 2003). 4.

Fine Control #4: Reversible Covalent Modification Enzyme control by reversible covalent modification plays a dominant role in fine metabolic control and is the major mechanism whereby extracellular stimuli such as hormones or light coordinate the control of intermediary metabolism (Cohen, 2002; Huber and Hardin, 2004; Plaxton, 2004b; Moorhead et al., 2005). The general model is that the covalently modified enzyme is interconverted between less active (or inactive) and more active forms owing to conformational-induced effects on the binding of its substrates and/or allosteric effectors. Reversible covalent modification of enzymes typically allows a very marked sensitivity (i.e., amplification) to signals, much greater than is possible for an enzyme responding to allosteric effectors, and indeed, often regulates enzymes in a virtual “on-off” manner (Cohen, 2002; Huber and Hardin, 2004; Plaxton, 2004b). There is convincing evidence for the crucial importance of reversible covalent modification as a mechanism to integrate glycolysis with mitochondrial metabolism in animals and plants. Recent advances in proteomics, particularly the development of novel and specific chemistries for detection of a diverse number of post-translational modifications, are rapidly expanding our awareness of respiratory enzymes whose functions are likely to be controlled by reversible covalent modification (Huber and Hardin, 2004). Phosphorylationdephosphorylation and disulfide-dithiol interconversions appear

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to be the most prevalent types of reversible covalent modification used in higher eukaryote, including plant, enzyme control (Cohen, 2002; Huber and Hardin, 2004; Plaxton, 2004b; MoraGarcia et al., 2005; Moorhead et al., 2005). 4.1. Phosphorylation-Dephosphorylation. Reversible protein phosphorylation is the most important post-translational modification in eukaryotic cells, as it pervades virtually all aspects of cell physiology, metabolism, and development (Cohen, 2002; Huber and Hardin, 2004; Plaxton, 2004b; Moorhead et al., 2005). Protein kinases and phosphatases catalyze the covalent incorporation or hydrolysis, respectively, of Pi groups on target proteins. Studies of mammalian cells have demonstrated that approximately one of every three proteins is modified by phosphorylation of either serine, threonine, or tyrosine residues, and this is believed to apply to other eukaryotes as well (Moorhead et al., 2005). The central role of phosphorylation-dephosphorylation in cell biology is illustrated by the fact that the largest known protein family consists of the eukaryotic protein kinase “superfamily” (Plaxton, 2004b; Moorhead et al., 2005). The protein phosphatase catalytic subunits responsible for the hydrolysis of Pi from proteins constitute a smaller group of genes compared to the protein kinases. However, protein phosphatase catalytic subunits (such as protein phosphatase type-1) appear to associate with a large number of other proteins to form a large variety of catalytic and regulatory subunit complexes (Moorhead et al., 2005). The first precedent for enzyme control by reversible covalent modification was provided in the mid-1950’s by Nobel Prize laureates Edwin Krebs and Eddy Fischer who were studying muscle glycogen phosphorylase, the enzyme that cleaves Glc-1-P units from the glycogen polymer (Cohen, 2002). Subsequent research revealed that protein phosphorylation-dephosphorylation plays a major role in the control of many enzymes involved in animal respiratory metabolism (Cohen, 2002; Plaxton, 2004b; Moorhead et al., 2005). Not surprisingly, at least 7 cytosolic or mitochondrial enzymes involved in plant respiration appear to be controlled, in part, by reversible phosphorylation (Table 1). For example, it is well established that the activity of most vascular plant PEPCs is enhanced by reversible phosphorylation catalyzed by an endogenous Ca2+ -independent PEPC protein kinase, and dephosphorylation by a protein phosphatase type 2A (PP2A) at a highly conserved N-terminal seryl residue (Chollet et al., 1996; Lepiniec et al., 2003; Izui et al., 2004; Nimmo, 2005). This typically results in reduced sensitivity of the enzyme to malate inhibition and increased sensitivity to activation by Glc-6-P. The post-translational control of plant PEPC by reversible phosphorylation has been best characterized in CAM and C4 leaves (Chollet et al., 1996; Lepiniec et al., 2003; Izui et al., 2004; Nimmo, 2005), and to a lesser extent in C3 leaves and soybean root nodules (Duff and Chollet, 1995; Zhang et al., 1995; Li et al., 1996; Wadham et al., 1996; Zhang and Chollet, 1997; Xu et al. 2003). However, regulatory PEPC phosphorylation in germinated wheat, sorghum and barley seeds, developing castor bean endosperm, ripening banana

fruit, and green algae has also been reported (Osuna et al., 1996, 1999; Law and Plaxton, 1997; Nhiri et al., 2000; Rivoal et al., 2002; Tripodi et al., 2005). The results from experiments in which pod excision or prolonged darkness pretreatments were imposed on castor plants indicate that similar to soybean root nodule PEPC (Zhang et al., 1995) the in vivo phosphorylation status of PEPC in developing castor beans is reversibly modulated in some manner by the supply of photosynthate recently translocated from the leaves (Tripodi et al., 2005). The in vivo regulatory phosphorylation of novel high and low molecular mass PEPC isoforms in developing castor bean endosperm was hypothesized to be one the regulatory components involved in photosynthate partitioning between storage lipids and storage proteins in this tissue (Tripodi et al., 2005). The phosphorylation of SuSy (reaction 1, Figure 2) by a calcium-dependent protein kinase (CDPK) occurs on two conserved Ser residues (Zhang et al., 1999; Winter and Huber, 2000; Hardin et al., 2003, 2004). In Zea mays, the phosphorylation of the major site (Ser15) has been linked to subsequent phosphorylation on its second site (Ser170) (Hardin et al., 2004). Phosphorylation of the major site may activate the cleavage activity of the enzyme by altering the structure of the amino terminus. In addition, Ser15 phosphorylation results in changes in the kinetic properties and subcellular localization of SuSy, as the enzyme’s partitioning between the soluble and membrane fractions is associated with the developmental stage of the organ examined and the phosphorylation status of Ser15. It is thought that membrane associated SuSy provides UDP-Glc to the membrane localized cellulose synthase complex (Winter and Huber, 2000; Hardin et al., 2004; Huber and Hardin, 2004). By contrast, phosphorylation of maize SuSy at Ser170 appears to target the enzyme for its proteasome-mediated degradation (Hardin et al., 2003). Furumoto and co-workers (2001) have demonstrated the in vivo phosphorylation of the bifunctional 6-phosphofructo2-kinase/fructose-2,6-bisphosphatase on serine and threonine residues in Arabidopsis leaves, although the identity of the specific phosphorylation sites has yet to be determined. The enzyme’s relative phosphorylation level varied diurnally, becoming maximal at the beginning of the night period. The physiological consequences of this phosphorylation remain to be clarified. However, apart from possible kinetic effects, 6-phosphofructo2-kinase/fructose-2,6-bisphosphatase phosphorylation in Arabidopsis leaves appears to generate a specific docking site for 14-3-3 proteins (discussed below) (Kulma et al., 2004). The mitochondrial PDC of plants and animals has a tightly associated PDC kinase and phospho-PDC phosphatase. Phosphorylation by PDC kinase almost completely inactivates PDC, whereas dephosphorylation by PDC phosphatase (a type 2C protein phosphatase) reactivates it (Hill, 1997; Siedow and Day, 2000; Mooney et al., 2002; Tovar-Méndez et al., 2003; McDonald and Vanlerberghe, 2005). PDC activity is therefore largely determined by the relative activities of the PDC kinase and phosphatase. Plant PDC kinase activity is dependent upon ATP, and is activated by NH+ 4 and inhibited by pyruvate and ADP

185

Activation (?)

Fru-6-P + ATP → Fru-2,6-P2 + ADP; Fru-2,6-P2 + H2 O → Fru-6-P + Pi

6-Phosphofructo-2Cytosol kinase/Fructose-2,6bisphosphatase (bifunctional enzyme) Non-phosphorylating Cytosol NADP-dependent glyceraldehyde-3-P dehydrogenase Pyruvate Kinase Cytosol

a

Mitochondrial ADP + Pi → ATP Inner Membrane

ATP Synthase

Remarks

Reference

Phosphorylation ‘tags’ the enzyme for its Tang et al. (2003) ubiquination and subsequent proteolytic degradation in maturing soybean seeds Phosphorylation of conserved seryl residue by a Chollet et al. (1996); Ca2+ -independent PEPC kinase typically Rivoal et al. (2002); relieves feedback inhibition of PEPC by Nimmo, (2005); malate and Glu. However, the novel high Tripodi et al. (2005) molecular mass PEPC2 isoform of green algae and developing castor seeds was recently reported to be less active when phosphorylated PDC phosphorylation status controlled by tightly Tovar-Méndez et al. bound PDC kinase and PDC phosphatase (2003); McDonald and Vanlerberghe (2005) Phospho-form inhibited by binding of a 14-3-3 Bunney et al., 2001; protein McDonald and Vanlerberghe (2005)

Phospho-activation may only apply to leaf form Zhang et al., 1999; of SuSy. Phosphorylation of root nodule SuSy Winter and Huber occurs in vivo, but has no known influence on (2000); Hardin et al., its kinetic properties in vitro. Phosphorylation 2003, 2004; Huber may also help to partition SuSy between the and Hardin, 2004. cytosol, plasma membrane, and cytoskeleton, as well as to target it for proteolytic turnover. Phospho-form of this bifunctional enzyme Furomoto et al. (2001); specifically binds 14-3-3 proteins. Kinetic Kulma et al. (2004) effect of this interaction is unclear, but phosphorylation in vivo was correlated with elevated Fru-2,6-P2 levels in Arabidopsis. Phospho-form in non-green wheat tissues is Bustos and Iglesias inhibited by binding of a 14-3-3 protein (2003)

Thermodynamically irreversible and reversible reactions are denoted by → and ↔, respectively.

Inhibition

Mitochondrial Pyruvate + CoA + Inactivation Matrix NAD+ → Acetyl-CoA + CO2 + NADH

Pyruvate Dehydrogenase Complex

Activation or Inhibition

Cytosol

PEP Carboxylase

PEP + HCO− 3 → OAA + Pi

Glyceraldeyde-3-P + Inhibition NADP+ → 3-P-glycerate + NADPH PEP + ADP → Pyruvate Inactivation + ATP

Activation

Sucrose Synthase

Sucrose + UDP ↔ Fructose + UDP-Glc

Reaction catalyzeda

Influence of phosphorylation on enzyme activity

Cytosol

Enzyme

Subcellular location

TABLE 1 Enzymes associated with plant respiration subject to protein kinase-mediated phosphorylation

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(Schuller et al., 1993; Tovar-Méndez et al., 2003). Upon illumination, pea leaf PDC is rapidly phosphorylated/inactivated. This has been suggested to be due to the stimulation of PDC kinase by NH+ 4 ions produced during photorespiratory glycine oxidation (Hill, 1997; Schuller et al., 1993; Tovar-Méndez et al., 2003). However, inhibition of mitochondrial oxidative phosphorylation by oligomycin disrupted photosynthesis similarly at both high and low CO2 levels, implying that reducing equivalents generated by the TCA cycle can replace those derived from glycine oxidation. Indeed, light dependent phospho-inactivation of leaf PDC does not appear to be common to all species (Hill, 1997; Tovar-Méndez et al., 2003). Antisense repression of PDC kinase in transgenic Arabidopsis plants resulted in elevated mitochondrial PDC activity as the enzyme mainly existed in its dephosphorylated, active form (Zhou and Taylor, 1998). These plants exhibited an altered phenotype with reduced accumulation of vegetative mass, earlier flower initiation, and a shortened generation time. Subsequent studies noted an increased respiratory rate in leaves and seeds of the PDC kinase antisense transgenics (Marilla et al., 2003). Moreover, these plants were also characterized by the enhanced contribution of carbon moieties from pyruvate to fatty acid biosynthesis and storage lipid accumulation in developing seeds, implicating a role for mitochondrial PDC in fatty acid biosynthesis in seeds (Marilla et al., 2003). Recent proteomic studies have revealed widespread phosphorylation of plant mitochondrial proteins. 32 P-labelling of intact potato tuber mitochondria yielded up to 30 radioactive polypeptides, whereas 2-dimensional gel electrophoresis of potato mitochondrial proteins followed by mass spectrometry identified 14 new phosphoproteins including several TCA cycle enzymes (i.e., NAD-ICDH, aconitase, NAD-MDH, and succinate dehydrogenase) and various components of the miETC (i.e., subunits of Complex I and III, and ATP synthase) (Bykova et al., 2003). Future work must determine what influence phosphorylation has on the functional properties of each of these mitochondrial proteins, as well as physiologically relevant situations and signalling processes that lead to alterations in their in vivo phosphorylation status. Identifying the in vivo mechanisms that serve to control the activities of the corresponding protein kinases and phosphoprotein phosphatases will be a daunting task. The pioneering studies of Zhou et al. (1999) and Marilla et al. (2003) involving genetic manipulation of PDC kinase in transgenic Arabidopsis plants indicate that protein kinases and phosphatases will become important targets for metabolic engineering of plant respiration. Since most of the early studies of protein phosphorylation were linked to metabolic enzymes, it was believed that the phosphorylation of proteins was primarily an event that caused conformational changes in enzymes that altered their ability to bind substrates, cofactors, and allosteric effectors. It is now known that protein phosphorylation may not only control enzymatic activity, but can also generate specific docking sites for other proteins, may control the shuttling of proteins within or between cellular compartments, and regulates pro-

teolytic degradation (Cohen, 2002; Hardin et al., 2003; Tang et al., 2003; Huber and Hardin, 2004; Moorhead et al., 2005). In fact, the generation of specific phosphorylation dependent docking or interaction sites may be the most common function of protein phosphorylation. Of particular interest in this regard are the 14-3-3s, a family of highly conserved and abundant proteins that play a central regulatory role in all eukaryotic cells (Winter and Huber, 2000; Huber and Hardin, 2004; Moorhead et al., 2005). The 14-3-3s bind to specific phosphorylated sites on diverse target proteins, thereby forcing conformational changes or influencing interactions between their targets and other molecules. In these ways, 14-3-3s “finish the job” when phosphorylation alone lacks the power to drive changes in the activities of intracellular enzymes. For example, the phospho-forms of nitrate reductase and sucrose-phosphate synthase, which are key enzymes in N- and C-metabolism, respectively, are both inhibited by 14-3-3 binding (Winter and Huber, 2000; Moorhead et al., 2005). Recent evidence suggests that 14-3-3 proteins also participate in the control of at least three enzymes involved in plant respiration. These are the aforementioned bifunctional 6-phosphofructo-2-kinase/fructose-2,6bisphosphatase, non-phosphorylating NADP-GAPDH, and ATP synthase (Table 1) (Cotelle et al., 2000; Bunney et al., 2001; Bustos and Iglesias, 2003; Kulma et al., 2004; Møller, 2001; McDonald and Vanlerberghe, 2005; Moorhead et al., 2005). The role of phosphorylation or 14-3-3 binding to Arabidopsis 6phosphofructo-2-kinase/fructose-2,6-bisphosphatase has yet to be determined, as a thorough kinetic examination did not find any changes in kinetic parameters or ratio of Fru-2,6-P2 synthesis to degradation activities (Kulma et al., 2004). Non phosphorylating NADP-GAPDH is a cytosolic enzyme that bypasses a Pi- and an ADP-dependent reaction of classical glycolysis (see reaction 14, Figure 2). 14-3-3 binding inhibits the phosphorylated form of the wheat endosperm enzyme (Bustos and Iglesias, 2003). 143-3 proteins have also been localized to the mitochondrial i.m. and shown to interact with the ATP synthase in a phosphorylation dependent manner (Bunney et al., 2001). The interaction was demonstrated to inhibit ATP synthase, and was suggested to provide a means to prevent ATP hydrolysis following stressinduced collapse of the electrochemical gradient across the i.m. (as would occur during anoxia stress) (Bunney et al., 2001). As noted above, a number of ATP synthase subunits appear to be phosphorylated in vivo (Bykova et al., 2003). Future research will undoubtedly uncover additional phosphoprotein targets for 14-3-3 binding control in metabolic pathways and miETC components associated with plant respiration. 4.2. Disulfide-Dithiol Interconversion. Covalent modification of cysteine (Cys) residues by disulfide-dithiol exchange links photosynthetic electron transport flow to the light regulation of several key enzymes of the chloroplast stroma via the thioredoxin system (Plaxton, 2004b; Mora-Garcia et al., 2005). This process is critical for the light-dependent activation and inhibition of the stromal-localized reductive and oxidativepentose phosphate pathways, respectively (Mora-Garc´ıa et al.,

PLANT RESPIRATION

2005). The discovery of plant cytosolic- and mitochondrialspecific thioredoxins indicated that this form of covalent modification may also participate in the control of plant respiration (Laloi et al., 2001; Balmer and Buchanan, 2002). That plant glycolytic and/or TCA cycle enzymes might be subject to disulfide-dithiol control in vivo was indicated by the observations that reduced thiol groups: (i) cause activation of plant cytosolic NAD-GAPDH and mitochondrial citrate synthase (Anderson et al., 1995; Stevens et al., 1997), and (ii) elicit maximal activation of tomato fruit or wheat flour PPi-PFK by Fru-2,6-P2 (Kiss et al., 1991). Plant PEPC kinase may be under thioredoxin control as the maize leaf enzyme was rapidly inactivated by incubation with oxidized glutathione and reactivated by subsequent incubation with thiol reducing agents, particularly in the presence of thioredoxin (Saze et al., 2001). Recent proteomics studies have assessed possible in vivo targets for thioredoxin action in the mitochondria and cytosol of Arabidopsis (Balmer et al., 2004; Marchand et al., 2004; Yamazaki et al., 2004). Cytosolic aldolase and NAD-GAPDH (reactions 11 and 13, Figure 2) were both determined to be likely targets for thioredoxinmediated disulfide to dithiol interconversion (Marchand et al., 2004; Yamazaki et al., 2004). A similar approach identified 50 probable thioredoxin-linked proteins in plant mitochondria that are involved in photorespiration, the TCA cycle, miETC, and ATP synthesis, among other processes (Balmer et al., 2004). Deduced amino acid sequences for most of these proteins contained a pair of conserved Cys residues consistent with these residues being thioredoxin targets. However, AOX provides a well documented role for thioredoxin mediated dithiol-disulfide interconversion in the control of plant respiration. AOX exists in the i.m. as either a non-covalently or covalently linked homodimer (Rhoads et al., 1998; Millenaar and Lambers, 2003; McDonald and Vanlerberghe, 2005). When covalently linked by an intersubunit disulfide bond AOX is inactive. When this bond is reduced by a specific form of thioredoxin h the AOX monomers become non-covalently associated, and increased activity results from an interaction of AOX’s regulatory thiols with its allosteric activator, pyruvate (Millenaar and Lambers, 2003; Gelhaye et al., 2005). Hence, AOX activity is modulated by both the redox state of the mitochondria (controlling AOX reduction) and glycolytic flux (defining levels of activating pyruvate). Both represent a feed-forward activation of this miETC component by upstream respiratory metabolism. AOX reduction to its active dithiol form is believed to be catalyzed by a mitochondrial thioredoxin, which itself may be reduced following the oxidation of specific TCA cycle substrates, namely isocitrate or malate (Møller, 2001). While it has been clearly demonstrated that AOX can be activated in vitro by reduced thioredoxin h (Gelhaye et al., 2005), it remains to be established whether this is the mechanism of AOX disulfide to dithiol interconversion that exists in vivo. 4.3. S-Nitrosylation. Nitric oxide (NO) has emerged as a key signalling molecule in animals and plants over the past few years, controlling the activity and expression of various

187

enzymes in response to various endogenous and exogenous stimuli (Mannick and Schonoff, 2002; Lamattina et al., 2003; Wendehenne et al., 2004). Analyses of NO-dependent events in animal systems have demonstrated that protein S-nitrosylation of Cys residues is an important control mechanism for many proteins (Mannick and Schonoff, 2002). The specificity of Snitrosylation is determined by the proximity of Cys residues to specific intracellular sources of NO, by the redox status of the microenvironment, and by the identity of residues that flank the Cys residue (Huber and Hardin, 2004). Leaves and roots of healthy plants produce and emit NO, and at least 100 possible targets of S-nitrosylation in plants were identified by searching the SwissProt database for the degenerate motif that is characteristic of S-nitrosylated proteins (Huber and Hardin, 2004). This in silico analysis has been corroborated by a recent proteomic analysis of Arabidopsis that identified 63 proteins from cell cultures and 53 proteins from leaves as likely candidates for S-nitrosylation (Lindermayr et al., 2005). Interestingly, aldolase, 3-PGA kinase, NAD-GAPDH, enolase, PEPC, and MDH were amongst the S-nitrosylated respiratory enzymes that were identified. At the enzymatic level, NO-dependent reversible inhibition of cytosolic NAD-GAPDH activity suggested that this enzyme might be controlled by S-nitrosylation in vivo (Lindermayr et al., 2005). The miETC appears to be a major source of NO emissions in plants (Planchet et al., 2005). The disruption of oxidative phosphorylation by NO has been suggested to occur via the direct inhibition of cytochrome oxidase by NO (Yamasaki et al., 2000). Although this disturbance of mitochondrial function by NO may be harmful to the plant cell (Caro and Puntarulo, 1999; Neill et al., 2003), a possibly greater deleterious impact of NO is avoided by the presence of AOX. Millar and Day (1996) noted that, unlike cytochrome oxidase, electron flow through AOX is insensitive to NO. Moreover, AOX expression is induced by NO in Arabidopsis (Huang et al., 2002). Far more research is required to fully assess the relationships between NO and plant respiration, including the extent and importance of reversible S-nitrosylation of key enzymes in respiratory control. E.

Overview of the Fine Control of Plant Respiratory Carbon Metabolism 1. Glycolysis In non-plant systems such as mammalian liver, primary control of glycolytic flux of hexose-phosphates to pyruvate is mediated by ATP-PFK, with secondary control at PK (Fothergill-Gilmore and Michels, 1993; Plaxton, 2004b). Activation of ATP-PFK increases the level of its product, Fru-1,6P2 , which is a potent feed-forward allosteric activator of the majority of non-plant PKs examined to date (Plaxton, 2004b). By contrast, quantification of changes in levels of glycolytic intermediates that occur following stimulation of respiration in many plant systems (including green algae, ripening fruit, aged storage root slices, germinating seeds, and suspension cell cultures) consistently demonstrate that plant glycolysis is controlled

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from the bottom up with primary and secondary regulation exerted at the levels of PEP and Fru-6-P utilization, respectively (Figure 5) (Adams and Rowan, 1970; Kobr and Beevers, 1971; Paul et al., 1978; Beaudry et al., 1989; Klöck and Kreuzberg, 1991; Vanlerberghe et al., 1992; Geigenberger and Stitt, 1991; Hatzfield and Stitt, 1991; Gauthier and Turpin, 1994; Huppe and Turpin, 1994; Plaxton, 1996; Givan, 1999). These findings are compatible with enzymological studies demonstrating that plant PKc s are not Fru-1,6-P2 activated (Podestá and Plaxton, 1991, 1992, 1994b; Hu and Plaxton, 1996; Smith et al., 2000; Turner and Plaxton, 2000; Turner et al., 2005), whereas plant ATP- and PPi-PFKs demonstrate potent allosteric inhibition by PEP (Garland and Dennis, 1980; Dennis and Greyson, 1987; Botha et al., 1988; Häusler et al., 1989; Lee and Copeland, 1996; Plaxton, 1996, 2004b; Givan, 1999; Turner and Plaxton, 2003). It follows that any enhancement in the activity of PK or PEPC will relieve the PEP inhibition of ATP-PFK and PPi-PFK, thereby elevating glycolytic flux from hexose-phosphate (Figure 5). Reduced cytosolic PEP levels also cause elevated Fru-2,6-P2 levels (and thus activation and inhibition of PPi-PFK and cytosolic FBPase, respectively) since a drop in PEP results in a fall in 3-PGA (these metabolites are at equilibrium in vivo), and PEP and 3-PGA are potent inhibitors of plant 6-phosphofructo-2kinase (Plaxton, 1996; Stitt, 1990; Markham and Kruger, 2002). Despite the obvious importance of PKc and PEPC in controlling plant respiration and cytosolic PEP partitioning, the two enzyme have thus far been subjected to concurrent biochemical characterization in only four systems; B. napus suspension cell cultures (Moraes and Plaxton, 2000; Smith et al., 2000), ripening banana fruit (Law and Plaxton, 1995, 1997; Turner and Plaxton, 2000), germinating castor bean cotyledons (Podestá and Plaxton, 1994), and soybean root nodules (Schuller et al., 1990; Zhang et al., 1995; McCloud et al., 2001). It has been suggested that a possible advantage of bottom-up regulation of glycolysis is that it permits plants to control net glycolytic flux to pyruvate independent of related plant-specific metabolic processes such as the Calvin-Benson cycle and sucrose-starch interconversion (Plaxton, 1996). Consistent with the view that plant glycolysis is controlled from the bottom up was the application of MCA to assess the distribution of respiratory flux control in aged disks from tubers of transgenic potato plants expressing different amounts of E. coli ATP-PFK (Thomas et al., 1997). It was determined that ATPPFK exerts a low flux control coefficient over both glycolysis and respiration, whereas far more flux control was exerted in the oxidative metabolism of PEP. The relatively low flux control coefficient of ATP-PFK was explained by MCA as a consequence of its potent feedback inhibition by PEP. In this way PEPC and/or PKc play a central role in the overall control of plant respiration, since the control of their activities in vivo will ultimately dictate the rate of mobilization of starch or sucrose for respiration, while simultaneously controling the provision of: (i) pyruvate for ATP production via oxidative phosphorylation, and (ii) TCA cycle C-skeletons needed for N-assimilation or as biosynthetic

precursors. It is obvious that PEPC and PKc represent promising targets for metabolic engineering of plant respiration and photosynthate partitioning. The abnormal growth, carbon partitioning and dark respiration rate of transgenic tobacco lacking leaf PKc (Knowles et al. 1998; Grodzinski et al. 1999) highlighted the importance of this enzyme in the control and integration of plant carbon and energy metabolism. Biotechnological approaches involving PEPC have also been implemented. PEPC overexpression in transgenic plants initially aimed to improve CO2 assimilation by suppressing photorespiration in C3 leaves (Jeanneau et al., 2002). Transgenic bean plants were subsequently generated that expressed a bacterial, malate-insensitive, PEPC in a seed-specific manner (Rolletschek et al., 2004). Analysis of the transgenic seeds indicated enhanced PEP partitioning through the PEPC anaplerotic pathway, resulting in a shift of photosynthate partitioning from sugars/starch into organic acids and amino acids. Consequently, the transgenic seeds accumulated up to 20 percent more storage protein (Rolletschek et al., 2004). 2.

PDC and TCA Cycle There are many well characterized regulatory properties of PDC and TCA cycle enzymes, but it is important that the significance of these properties is eventually evaluated in the context of metabolite concentrations that occur in vivo and the flux control coefficient of the enzyme concerned. Based upon in vitro regulatory properties, the most likely sites for controlling flux through the TCA cycle are PDC, and control of ICDH and 2-OG dehydrogenase by the NADH:NAD+ ratio (Chen and Gadal, 1990; Hill, 1997; Millar et al., 1999; Siedow and Day, 2000; Igamberdiev and Gardeström, 2003; McDonald and Vanlerberghe, 2005) (Figure 5). The available evidence is consistent with a tight but flexible control of flux through PDC, the TCA cycle and various components of the miETC, which is perhaps not surprising considering the range of metabolic functions performed by respiration in plants. IV.

CONCLUDING REMARKS From the preceeding discussion, it is clear that remarkable progress has been made over the past several decades in our understanding of plant respiration. The ever-growing collections of plant genes and the implementation of high-throughput transcriptomic and proteomic studies have dramatically increased the inventory of respiration related proteins in plants, while simultaneously raising numerous questions that remain to be answered. New insights have been provided by the production of transgenic plants having modified levels of respiratory enzymes, as well as enzymes such as protein kinases that mediate their reversible covalent modification in response to intra- or extracellular signals. However, there are still large gaps in our understanding of this complex process that is at the core of primary plant metabolism. Further refinement and integration of enzymology, genomics/bioinformatics, proteomics, transcriptomics, MCA, metabolomics and associated non-aqueous subcellular

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fractionation techniques, cell biology, and in vivo flux analyses via non-invasive NMR, gas exchange, and mass spectrometry technologies will undoubtedly help to further enhance our understanding of plant respiratory metabolism. Only with the coordinated efforts of plant physiologists, biochemists and molecular biologists will the tissue-specific organization and control of plant respiration be fully comprehended. Fundamental information is still lacking on how respiration and the processes supporting it are physiologically controlled. In particular, more information is needed on the in vivo cell-specific mechanisms by which partitioning of electrons between the phosphorylating versus non-energy conserving pathways of miETC reacts to changes in glycolytic and TCA cycle flux, and in response to changing environments and different catabolic and anabolic cellular demands. A better understanding of how flux through alternative glycolytic, TCA cycle, and miETC enzymes is controlled and the extent to which this influences plant stress tolerance is of significant practical interest. Recent theoretical and experimental studies indicate that the majority of metabolic control occurs at the post-transcriptional level (ter Kuile and Westerhoff, 2001; Sweetlove and Fernie, 2005). Thus, a thorough understanding of the organization and control of plant respiratory metabolism will ultimately require a systematic analysis not only of transcript and protein abundance changes in various tissues and cell types under different environmental and physiological regimes, but also of: (i) covalent modifications of enzymes in response to different stimuli, (ii) protein-protein interactions, (iii) the kinetic and allostric properties of the relevant enzymes, (iv) in vivo flux analyses, (v) cytosolic versus mitochondrial metabolite levels, and (vi) the location and kinetic properties of membrane transporter proteins that selectively transport specific metabolites between subcellular compartments. Given the central role that respiration plays in crop productivity, this fundamental knowledge will lead to important rational approaches to crop improvement via the process of metabolic engineering. At the ecosystem level, plant respiration has a profound impact on the CO2 concentration in the atmosphere, and is therefore a key component influencing global climate change (Atkin and Tjoelker, 2003; Gifford, 2003; Atkin et al., 2005). The role of temperature in mediating a potential large increase in respiratory CO2 release from plants to the atmosphere is of concern for its effects on global atmospheric CO2 levels and contribution to further greenhouse warming. Predicting plant responses to global climate change will necessitate a thorough assessment of the influence of temperature and CO2 on plant respiration.

ACKNOWLEDGMENTS Research in our laboratories is supported by grants from the Natural Sciences and Engineering Research Council of Canada (W.C.P.), Fundación Antorchas and CONICET (F.E.P.). We are greatly indebted to past and present members of our laboratories who have examined biochemical aspects of plant glycolysis and its control. We also thank Profs. Greg Moorhead and Greg Van-

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