The biochemical pathway of CO₂ fixation was discovered by feeding radioactively labelled CO₂ in the light to algae and then extracting the cells and examining which compounds accumulated radioactivity. Figure 2.1(b) shows a typical labelling ‘pattern’ for a C₃ plant. Here, a short burst of labelled CO₂ was given to the plants, then the label was ‘chased’ through the photosynthetic pathway by flushing with unlabelled air. Atmospheric CO₂ is initially incorporated into a five-carbon sugar phosphate (ribulose-1,5-bisphosphate or RuBP) to produce two molecules of the phosphorylated three-carbon compound 3-phosphoglycerate, often referred to as the acidic form 3-phosphoglyceric acid (3-PGA). Hence, plants which use Rubisco as their primary enzyme of CO₂ fixation from the air are called C₃ plants. Consequently, in C₃ plants, 3-PGA is the first labelled sugar phosphate detected after a pulse of ¹⁴CO₂ has been supplied (Figure 2.1b). In the PCR cycle, 3-PGA is phosphorylated by the ATP produced from thylakoid electron transport (see Chapter 1) and then reduced by NADPH to produce triose phosphate. Triose phosphates are the carbon backbones, produced by the PCR cycle, for the synthesis of critical carbohydrate for the maintenance of plant growth and the productive yield of stored carbohydrate in seed.

Newly synthesised triose phosphate faces three options. It can be (1) exported to the cytosol for sucrose synthesis and subsequent translocation to the rest of the plant, (2) recycled within the chloroplast to produce more RuBP or (3) diverted to produce starch (Figure 2.1a). This is shown by the time-course of the appearance of radioactivity in starch and sucrose after it has passed through 3-PGA (Figure 2.1b). The energy requirements of the PCR cycle are three ATP and two NADPH per CO₂ fixed, in the absence of any other energy-consuming processes.

Sucrose and starch synthesis

Most of the triose phosphate synthesised in chloroplasts is converted to either sucrose or starch. Starch accumulates in chloroplasts, but sucrose is synthesised in the surrounding cytosol, starting with the export of dihydroxyacetone phosphate and glyceraldehyde phosphate from the chloroplast. A condensation reaction, catalysed by aldolase, generates fructose-1,6-bisphosphate, and this is converted to fructose-6-phosphate after an hydrolysis reaction catalysed by fructose-1,6-phosphatase. Sucrose-6-phosphate synthase then generates sucrose-6-phosphate from the reaction of fructose-6-phosphate and UDP-glucose. The phosphate group is removed by the action of sucrose-6-phosphatase. This P_i is transported back into the chloroplast where it is available for ATP synthesis. For each molecule of triose phosphate exported from a chloroplast, one P_i is translocated inwards.

Sucrose synthesised within the cytosol of photosynthesising cells is then available for general distribution and is commonly translocated to other carbon-demanding centres via the phloem (see Chapter 5).

By contrast, starch synthesis occurs within chloroplasts. The first step is a condensation of glucose-1-phosphate with ATP. Starch synthase then transfers glucose residues from this molecule to the non-reducing end of a pre-existing molecule of starch. Starch consists of two types of glucose polymer, namely amylose and amylopectin. Amylose is a long, unbranched chain of D-glucose units connected via (α1–4) linkages. Amylopectin is a branched form, with (α1–6) linkages forming branches approximately every 24–30 glucose residues.

Ribulose bisphosphate (RuBP) is consumed in the carboxylating step of carbon fixation. If such fixation is to continue, RuBP must be regenerated, and in this case via the PCR cycle. The PCR cycle operates within the stroma of chloroplasts, and consists of a sequence of 11 steps where a three-carbon compound (3-phosphoglycerate) is phosphorylated, reduced to glyceraldehyde 3-phosphate and isomerised to dihydroxyacetone phosphate. Condensation of this three-carbon compound with glyceraldehyde 3-phosphate yields a six- carbon compound (fructose bisphosphate). Following a series of carbon shunts, involving four-, five- and seven-carbon compounds, RuBP is regenerated.

Important features of the PCR cycle include: (1) for every step of the cycle to occur once, three carboxylations must occur via ribulose bisphosphate carboxylase thus generating six moles of phosphoglycerate (18 carbons); (2) for one turn of the cycle, three molecules of RuBP participate (15 carbons) and thus a net gain of three carbons has occurred for the plant; (3) in regenerating three molecules of RuBP, nine ATP and six NADPH are consumed.

Photosynthetic carbon fixation in air is constrained by the kinetic properties of Rubisco. Form I Rubisco in higher plants is a large protein (approximately 550 kDa) comprised of eight large (approx. 50-55 kDa) and eight small subunits (approx. 13-18 kDa) to form an L₈S₈ hexadecamer. Rubisco synthesis and assembly in higher plants is a complex process whereby the large subunit gene (rbcL) is encoded in the chloroplast genome, while the small subunit genes (rbcS) are encoded as a multi-gene family in the nucleus. The Rubisco small subunits are translated as precursors in thc cytosol and are equipped with a transit-peptide to target them to the chloroplast. Upon import in the chloroplast the transit-peptide is cleaved by a stromal peptidase and the N-terminus modified by methylation of the n-terminal methionine. The large subunits  are synthesised within the chloroplast and also post-translationally modified through the removal of the the N-terminal methionine and serine amino acids and the subsequent acetylation of proline at the N-terminus and the methylation of lysine at position 14. The assembly of large and small subunits into functional hexadecameric Rubisco is reliant on the coordination of chloroplast-localised chaperones.

Despite selection pressure over evolutionary history, Rubisco remains an inefficient catalyst (Spreitzer and Salvucci 2002). Therefore, to achieve a productive maximum CO₂ assimilation rate (A_max), plants must compensate for catalytic inefficiency by investing large amounts of nitrogen in Rubisco. Consequently, Rubisco comprises more than 50% of leaf soluble protein in C₃ plants. On a global scale, this investment equates to around 10 kg of nitrogen per person!

More than 1000 million years of evolution has still not resulted in a ‘better’ Rubisco adapted for the current and future concentrations of CO₂. Such a highly conserved catalytic protein is an outcome of thermodynamic and mechanistic difficulties inherent to this reaction. Rubisco requires carbamylation of the absolutely-conserved residue K201 that is then stabilised by the binding of Mg²⁺. Without this activation step Rubisco is unable to function. The fixation of CO₂ to RuBP to form two molecules of 3-PGA is a five step catalytic process that produces highly reactive transition state intermediates that bind CO₂. The highly reactive transition states make Rubisco prone to generating misfiring products, which generate inhibitors within the active site. Therefore, Rubisco requires its own catalytic protection enzyme Rubisco activase. Plants devoid of this enzyme fail to grow properly in air as the activation and subsequent activity of Rubisco is impeded (Portis and Salvucci 2002). Rubisco activase is an ATP-dependent process that removes inhibitors from the active site of Rubisco allowing for activation and catalysis to proceed. Recently, the crystal structure of Rubisco activase has been solved, which will provide key insight into the molecular interaction between Rubisco and Rubisco activase (reviewed by Portis et al. 2008).

Rubisco first evolved when the earth’s atmosphere was rich in CO₂, but virtually devoid of O₂. With the advent of oxygen-producing photosynthesis by land plants, and the resulting increases in atmospheric O₂, one key deficiency of this enzyme became apparent. Rubisco would not only catalyse fixation of CO₂ but would also permit incorporation of O₂ into RuBP to produce, instead of two molecules of 3-PGA, just one molecule of 3-PGA with one molecule of a two-carbon compound, 2-phosphoglycolate (Section 2.3). Indeed, CO₂ and O₂ compete directly for access to the active sites of Rubisco. So feeble is Rubisco’s ability to distinguish between these two substrates that in air (20% O₂) approximately one molecule of O₂ is fixed for every three molecules of CO₂.

Fixation of O₂ and subsequent photorespiration (Section 2.3) is an energy-consuming process, due to competition between O₂ and CO₂ for RuBP, plus the energy cost of converting the phosphoglycolate product to a form which can be recycled in the PCR cycle. This energy cost is increased at higher temperatures because O₂ competes more effectively with CO₂ at the active site of Rubisco. Such sensitivity to temperature × O₂ explains why CO₂ enrichment, which reduces photorespiration, has a proportionally larger effect upon net carbon gain at higher temperatures than at lower temperatures (Section 13.3).

Notwithstanding a meagre catalytic effectiveness in present day Rubisco, more efficient variants would still have had a selective advantage, and especially during those times in the earth’s geological history when atmospheric CO₂ concentration was decreasing. Indeed there has been some improvement (Figure 2.3) such that specificity towards CO₂ as opposed to O₂ has improved significantly. Recently evolved angiosperms show a relative specificity almost twice that of 'older' organisms such as photosynthetic bacteria.

Despite such improvement, Rubisco remains seemingly maladapted to its cardinal role in global carbon uptake, and in response to selection pressure for more efficient variants of CO₂ assimilation, vascular plants have evolved with photosynthetic mechanisms that alleviate an inefficient Rubisco. One key feature of such devices is a mechanism to increase CO₂ concentration at active sites within photosynthetic tissues. Some of these photosynthetic pathways are dealt with below.

One hundred million years ago (Mid-Cretaceous), atmospheric CO₂ was between 1500 and 3000 µL L^–1, or four to ten times post-industrial levels. Atmospheric CO₂ declined during the Oligocene (20-30 million years ago) from the high Tertiary levels (>1000 µL L^-1), and oscillated between 180 and 300 µL L^-1 for the last 1-3 million years. The Oligocene was also a time when the Earth was dry and the tropics were relatively hot. The earliest origins of C₄ photosynthesis date back to this period. Curiously, C₄ plants remained in low abundance for a long period of time. According to stable carbon isotopic data, a worldwide expansion of C₄ grasslands and savannas occurred during the Late Miocene and Pliocene (3 to 8 million years ago), most probably through the displacement of C₃ vegetation (Edwards et al. 2010).

Under the early high concentration of CO₂, photorespiration of C₃ plants was inhibited (Section 2.3) so that photosynthetic efficiency was higher than it is now. In addition, maximum photosynthetic rates were double twentieth century values, and the energy cost of photosynthesis would have been around three ATP and two NADPH per molecule of CO₂ fixed. As atmospheric CO₂ concentrations declined to approximately 250–300 µL L^–1, photosynthetic rates were halved, photorespiration increased substantially, photosynthetic efficiency declined and the energetic costs of photosynthesis increased to approximately five ATP and 3.2 NADPH per CO₂ molecule fixed. Such events would have generated a strong selection pressure for genetic variants with increased carboxylation efficiency and increased photosynthetic rates.

Angiosperms have a higher relative specificity of Rubisco for CO₂ than ferns and mosses (see examples of other less evolutionarily advanced species in Figure 2.3). Such differences imply minor evolution in this highly conserved molecule of Rubisco and there is little variation between species of vascular plants. Consequently, alteration of Rubisco in response to a changing atmospheric CO₂ concentration has not been an option.

By contrast, evolution of a new photosynthetic pathway (C₄) has occurred independently and on many occasions in diverse taxa over 25 to 30 million years as CO₂ levels declined. Despite its complexity, C₄ photosynthesis evolved more than 60 independent times in 19 distantly related flowering families. About 50% of C₄ species are grasses (Poaceae) with ~18 distinct origins distributed over 370 genera and ~4600 species (Sage et al. 2011). The oldest identifiable fossils with pronounced bundle sheath layers are seven million years old, although necessary metabolic pathways could have evolved earlier, prior to this adaptation in anatomy. C₄ plants are known to differ from C₃ plants in their discrimination against atmospheric ¹³CO₂, and shifts in the stable carbon isotope signature of soil carbonate layers that reflect emergence of C₄ plants have been dated at 7.5 million years bp. Modern evidence from molecular phylogeny places the origin of the main C₄ taxa at 25-30 million years ago (Christin et al. 2009). By inference, C₄ photosynthesis evolved in response to a significant decline in atmospheric CO₂ concentration, from 1500–3000 µL L^–1 to about 300 µL L^–1. By evolving a CO₂-concentrating mechanism, C₄ plants presented their Rubisco with an elevated partial pressure of CO₂ despite lower atmospheric CO₂. As a consequence, photorespiration was inhibited, maximum photosynthetic rates increased and energetic costs reduced.

The C₄ pathway (Figure 2.3) is ‘a unique blend of modified biochemistry, anatomy and ultra-structure’ (Hatch 1987). The classical C₄ syndrome in most terrestrial plants consists of two photosynthetic cycles (C₃ (or PCR) and C₄) operating across two photosynthetic cell types (mesophyll and bundle sheath), which are arranged in concentric layers around the vascular bundle, also known as the kranz anatomy.

Initial and rapid fixation of CO₂ within mesophyll cells results in the formation of a four-carbon compound which is then pumped to bundle sheath cells for decarboxylation and subsequent incorporation into the PCR cycle in that tissue. This neat division of labour hinges on specialised anatomy and has even resulted in evolution of distinct classes of chloroplasts in mesophyll compared with bundle sheath cells. Three biochemical variants of C₄ photosynthesis (termed subtypes) are known to have evolved from C₃ progenitor and in all cases with a recurring theme where the C₄ cycle of mesophyll cells is complemented by a PCR cycle in bundle sheath cells, where Rubisco is exclusively localised. In effect, a biochemical ‘pump’ concentrates CO₂ at Rubisco sites in bundle sheath cells thereby sustaining faster net rates of CO₂ incorporation and virtually eliminating photorespiration. For this overall mechanism to have evolved, a complex combination of cell specialisation and differential gene expression was necessary. Figure 2.3a shows a low-magnification electron micrograph of a C₄ leaf related to a generalised scheme for the C₄ pathway.

By analogy with Calvin’s biochemical definition of the C₃ pathway at Berkeley in the 1950s, the C₄ pathway was also delineated with radioactively labelled CO₂ (see Feature Essay 2.1). Significantly, and unlike C₃ plants, 3-PGA is not the first compound to be labelled after a ¹⁴C pulse (Figure 2.3b). Specialised mesophyll cells carry out the initial steps of CO₂ fixation utilising the enzyme phosphoenolpyruvate (PEP) carboxylase. The product of CO₂ fixation, oxaloacetate, is a four-carbon organic acid, hence the designation ‘C₄’ photosynthesis (or colloquially, C₄ plant). A form of this four-carbon acid, either malate or aspartate depending on the C₄ subtype, migrates to the bundle sheath cells which contain Rubisco and the PCR cycle. In the bundle sheath cells, CO₂ is removed from the four-carbon acid by a specific decarboxylase and a three-carbon product returns to the mesophyll to be recycled to PEP for the carboxylation reaction. Thus, label first appears in the four-carbon acid after ¹⁴C feeding, followed by 3-PGA and, finally, in sucrose and starch (Figure 2.1b).

A physical barrier to CO₂ diffusion exists in the thickened walls of the bundle sheath cells (lined with suberised in some C₄ species), preventing CO₂ diffusion back to the mesophyll and allowing CO₂ build up to levels at least 10 times those of ambient air. Build up of CO₂ in the bundle sheath is also facilitated by the higher activity ratio (2-4 times) of PEP carboxylase relative to Rubisco in C₄ plants. Rubisco is thus exposed to a saturating concentration of CO₂ which both enhances carboxylation due to increased substrate supply, and forestalls oxygenation of RuBP (hence photorespiration) by outcompeting O₂ for CO₂ binding sites on Rubisco (Figure 2.5).

In leaves of C₃ plants, the PCR cycle operates in all mesophyll chloroplasts, but in C₄ plants the PCR cycle is restricted to bundle sheath cells (Figure 2.3). Rubisco is pivotal in this cycle, and can be used as a marker for sites of photosynthetic carbon reduction. Rubisco was visualised by localising this photosynthetic enzyme with antibodies via indirect immunofluorescent labelling (Hattersley et al. 1977; Figure 2.6). In this pioneering method, ‘primary’ rabbit anti-Rubisco serum (from rabbits injected with purified Rubisco) is first applied to fixed transverse sections of leaves. Rabbit antibodies to Rubisco bind to the enzyme in situ. Then ‘secondary’ sheep anti-rabbit immunoglobulin tagged with a fluorochrome (fluorescein isothiocyanate) is applied to the preparation. This fluorochrome binds specifically to the rabbit antibodies and fluoresces bright yellow wherever Rubisco is located (blue light excitation using an epifluorescence light microscope).

In the C₃ grass Microlaena stipoides (Figure 2.6a), all chloroplasts are fluorescing bright yellow and this indicates wide distribution of Rubisco throughout mesophyll tissue. By contrast, only bundle sheath cells are equipped with Rubisco in the C₄ grass Digitaria brownii (Figure 2.6b). These two native Australian grasses co-occur in the ACT but contrast in relative abundance. M. stipoides (weeping grass) is common in dry sclerophyll woodlands throughout southeast temperate Australia, whereas D. brownii (cotton panic grass) in the ACT is at the southern end of its distribution, being far more abundant in subtropical Australia and, in keeping with its C₄ physiology, especially prevalent in semi-arid regions.

One disadvantage of the C₄ pathway is that an energy cost is incurred by C₄ plants to run the CO₂ ‘pump’. This is due to the ATP required for recycling PEP from pyruvate by the chloroplastic enzyme pyruvate, Pi dikinase in the mesophyll cells (Figure 2.4 and Hatch 1987). Under ideal conditions, five ATP and two NADPH are required for every CO₂ fixed in C₄ photosynthesis (two ATP are required to run the CO₂ pump, i.e., regenerate PEP). In addition, a proportion (20-30%) of CO₂ fixed by PEP carboxylase in the mesophyll is not fixed by Rubisco in the bundle sheath, and subsequently leaks back to the mesophyll. This leaked (or overcycled) CO₂ represents an additional, inherent energetic cost of the C₄ pathway.

From the previous section, the C₄ pathway is obviously energetically more expensive than the C₃ pathway in the absence of photorespiration. However, at higher temperatures the ratio of RuBP oxygenation to carboxylation is increased and the energy requirements of C₃ photosynthesis can rise to more than five ATP and three NADPH per CO₂ fixed in air (for these calculations see Hatch 1987).

Representative light response curves for photosynthesis in C₃ cf. C₄ plants (Figure 2.7) can be used to demonstrate some of these inherent differences in photosynthetic attributes. At low temperature (10°C in Figure 2.7) a C₃ leaf shows a steeper initial slope as well as a higher value for light-saturated photosynthesis. By implication, quantum yield is higher and photosynthetic capacity is greater under cool conditions. In terms of carbon gain and hence competitive ability, C₃ plants will thus have an advantage over C₄ plants at low temperature and especially under low light.

By contrast, under warm conditions (35°C, upper curves in Figure 2.7) C₄ photosynthesis in full sun greatly exceeds that of C₃, while quantum yield (inferred from initial slopes) remains unaffected by temperature. Significantly, C₃ plants show a reduction in quantum yield under warm conditions (compare 10°C and 35°C curves; right side of Figure 2.7). At 35°C C₃ plants also show lower rates of light-saturated assimilation compared with C₄ plants. Increased photorespiratory losses from C₃ leaves at high temperature are responsible (Section 2.3). C₄ plants will thus have a competitive advantage over C₃ plants under warm conditions at both high and low irradiance.

C₄ photosynthesis calls for metabolic compartmentation which is in turn linked to specialised anatomy (Figure 2.4). Three biochemical subtypes of C₄ photosynthesis have evolved which probably derive from subtle differences in the original physiology and leaf anatomy of their C₃ progenitors.

CO₂ assimilation by all three C₄ subtypes (Figure 2.8) involves five stages:

1. carboxylation of PEP in mesophyll cells, thereby generating four-carbon acids (malate and/or aspartate);
2. transport of four-carbon acids to bundle sheath cells;
3. decarboxylation of four-carbon acids to liberate CO₂;
4. re-fixation of this CO₂ via Rubisco within the bundle sheath, using the C₃ pathway;
5. transport of three-carbon acid products following decarboxylation back to mesophyll cells to enable synthesis of more PEP.

Recognising some systematic distinctions in whether malate or aspartate was transported to bundle sheath cells, C₄ plants were further subdivided into three subtypes according to their four-carbon acid decarboxylating systems and ultrastructural features (Hatch et al. 1975). Members of each subtype contain high levels of either NADP-malic enzyme (NADP-ME), phosphoenolpyruvate carboxykinase (PCK) or NAD-malic enzyme (NAD-ME) (so designated in Figure 2.8). High NADP-malic enzyme activity is always associated with higher NADP-malate dehydrogenase activity, while those species featuring high activities of either of the other two decarboxylases always contain high levels of aminotransferase and alanine aminotransferase activities. As a further distinction, each of the decarboxylating enzymes is located in bundle sheath cells; NAD-malic enzyme is located in mitochondria but PEP carboxykinase is not.

In all three subtypes, the primary carboxylation event occurs in mesophyll cytoplasm with PEP carboxylase acting on HCO₃^– to form oxaloacetate. However, the fate of this oxaloacetate varies according to subtype (Table 2.1; Figure 2.8). In NADP-ME species, oxaloacetate is quickly reduced to malate in mesophyll chloroplasts using NADPH. By contrast, in NAD-ME and PCK species, oxaloacetate is transaminated in the cytoplasm, with glutamate donating the amino group, to generate aspartate. Thus, malate is transferred to bundle sheath cells in NADP-ME species and aspartate is transferred in NAD-ME and PCK species. The chemical identity of three-carbon acids returned to mesophyll cells varies accordingly.

In NADP-ME species, only chloroplasts are involved in decarboxylation and subsequent carboxylation via the PCR cycle (Figure 2.8). By contrast, in NAD-ME and PCK species, chloroplasts, cytoplasm and mitochondria are all involved in moving carbon to the PCR cycle of bundle sheath chloroplasts. In NAD-ME and PCK species, aspartate arriving in bundle sheath cells is reconverted to oxaloacetate in either mitochondria (NAD-ME) or cytoplasm (PCK) (Table 2.2). Reduction and decarboxylation of oxaloacetate occurs in mitochondria of NAD-ME species and CO₂ is thereby released for fixation by chloroplasts of bundle sheath cells. In PCK species, oxaloacetate in the cytoplasm is decarboxylated by PCK, thereby releasing CO₂ for fixation in bundle sheath chloroplasts (Figure 2.8).

Transport of metabolites to bundle sheath cells

A rapid transfer of malate and aspartate to bundle sheath cells from mesophyll cells is required if the CO₂ concentration in bundle sheath cells is to stay high. A very high density of plasmodesmata linking bundle sheath cells to mesophyll cells facilitates this traffic. Consequently, the permeability coefficient of C₄ bundle sheath cells to small metabolites such as four-carbon acids is about 10 times larger than that of C₃ mesophyll cells (Table 2.2). However, coupled with this need for a high permeability to metabolites moving into bundle sheath cells is a low permeability to CO₂ molecules so that CO₂ released through decarboxylation in the bundle sheath does not diffuse rapidly into mesophyll air spaces. For some species, a layer of suberin in the cell wall of bundle sheath–mesophyll junctions (suberin lamella) significantly reduces CO₂ efflux (Table 2.2).

Centrifugal versus centripetal chloroplasts

Not all species contain a suberin layer, but all C₄ plants have a need to prevent CO₂ from diffusing quickly out of bundle sheath cells, so that the location of chloroplasts of bundle sheath cells becomes critical in those species lacking a suberin layer (Figure 2.9). Where species have a suberin layer, chloroplasts are located in a centrifugal position, that is, on the wall furtherest away from the centre of the vascular bundle lying in the middle of the bundle sheath (Figure 2.9E, F). In those C₄ species lacking a suberin layer, chloroplasts are located centripetally, that is, on the wall closest to the centre of the vascular bundle lying within the bundle sheath (Figure 2.9A, B). Such a location would help restrict CO₂ diffusion from bundle sheath to mesophyll cells.

Regulation of C₄ photosynthesis

Fixation of CO₂ by C₄ plants involves the coordinated activity of two cycles in separate anatomical compartments (Figure 2.8). The first cycle is C₄ (carboxylation by PEP carboxylase), the second is C₃ (carboxylation by Rubisco). Given this biochemical and anatomical complexity, close regulation of enzyme activities is a prerequisite for efficient coordination.

PEP carboxylase, NADP-malate dehydrogenase and pyruvate orthophosphate dikinase are all light-regulated and their activities vary according to irradiance. NADP-malate dehydrogenase is regulated indirectly by light via the thioredoxin system.

PEP carboxylase in C₄ plants exists in the same homo-tetramer in light- and dark-acclimated leaves. This is in marked contrast to CAM species where different forms exist in light- and dark-acclimated leaves. In C₄ plants, PEP carboxylase has extremely low activity at night, thus preventing uncontrolled consumption of PEP. Such complete loss of activity in darkness is mediated via divalent metal ions, pH plus allosteric activators and inhibitors. As a consequence, and over a period of days, C₄ plants can increase or decrease PEP carboxylase in response to light regime.

Rubisco is characterised by its low affinity for its productive substrate, CO₂ and slow catalytic turnover rate (i.e., 1-3 cycles per sec). Importantly, Rubisco reacts with O₂ (photorespiration), and this culminates in loss of CO₂ and energy. In C₃ plants, photorespiration can drain more than 25% of fixed CO₂ under non-stressful conditions. The ratio of photorespiration to photosynthesis increases with increasing temperature and decreasing intercellular CO₂ such as occurs when stomatal conductance is reduced under water stress. C₃ plants compensate for Rubisco’s inefficiencies by (i) opening their stomata to increase CO₂ diffusion into chloroplasts, which increases water loss and lowers leaf-level water use efficiency, WUE; and (ii) investing up to 50% of leaf nitrogen in Rubisco, which lowers their leaf-level nitrogen use efficiency, NUE.

The C₄ pathway supercharges photosynthesis and suppresses photorespiration by operating a CO₂ concentrating mechanism which elevates CO₂ around Rubisco. Although C₄ photosynthesis incurs additional energy, the energy cost of photorespiration exceeds that of the CO₂ concentrating mechanism above 25^oC. Hence, higher radiation use efficiencies (i.e., efficiency of converting absorbed radiation into biomass) have been recorded for C₄ than C₃ crops. High bundle sheath CO₂ concentration saturates C₄ photosynthesis at relatively low intercellular CO₂, allowing C₄ plants to operate with lower stomatal conductance. Thus, leaf-level WUE is usually higher in C₄ than C₃ plants. Relative to C₃ plants, Rubisco of C₄ plants is faster (higher turnover rate) and operates under saturating CO₂. Thus, C₄ plants typically achieve higher photosynthetic rates with about 50% less Rubisco and less leaf nitrogen. Hence, photosynthetic NUE is higher in C₄ than C₃ plants. Accordingly, C₄ plants are advantaged relative to C₃ plants in hot and nitrogen-poor environments with short growing seasons, hence their great abundance in wet/dry tropics such as Northern Territory savannas.

As mentioned earlier, more than 50% of C₄ plants are grasses. C₄ grasses are confined to low latitudes and altitudes, whereas C₃ species dominate at higher latitudes and altitudes. Generally, C₄ species frequently occur in regions of strong irradiance. Ehleringer and colleagues (Ehleringer et al. 1997) proposed that these distribution patterns are best explained by the different responses of photosynthetic quantum yields to temperature between C₃ and C₄ plants.

C₄ photosynthesis suppresses photorespiration by operating a CO₂ concentrating mechanism that comes at additional energetic cost. This cost is independent of ambient CO₂ and temperatures. In contrast, photorespiration (and its associated energy cost) increases steeply with temperature in C₃ plants and is highly dependent on CO₂ concentrations. Under saturating irradiance and current ambient atmospheric CO₂ concentration, the threshold temperature where the cost of photorespiration in C₃ plants exceeds that of the CO₂ concentrating mechanism in C₄ plants is estimated around 25^oC. This model provides a physiological basis for understanding today’s contrasting geographic distribution between C₃ and C₄ grasses.

As an example, the C₄ grasses of the northern Australian savannas are relatively un-shaded because of the low tree density and sparse canopy. Light is abundant and since the CO₂ concentration inside C₄ leaves is high, a potentially high rate of light-saturated assimilation can be exploited. Most C₃ species reach light saturation in the range of one-eight to one-half full sunlight (Figure 2.7). In C₄ species, canopy assimilation might not become light saturated even in full sunlight. C₄ plants thus maintain a competitive advantage over C₃ plants in tropical locations, where average daily light receipt is much larger than in temperate zones, and associated with warmer conditions that also favour C₄ photosynthesis (Figure 2.6). Given strong sunlight, warmth and seasonally abundant water, biomass production by C₄ plants is commonly double the rate for C₃ plants. Typically, C₃ plants produce 15–25 t ha^–1 but C₄ plants easily produce 35–45 t ha^–1.

Physiological characteristics of the C₄ subtypes

As outlined in previous sections, characteristic biochemical, anatomical and physiological traits are associated with each of the three “classical” C₄ subtypes (Table 2.3). However, it should be noted that many C₄ plants have leaf structures that fall outside the “classical” subtype division (eg, NADP-ME tribes, Arundinelleae and Neurachneae). As many as 11 anatomical-biochemical suites have been identified in C₄ grasses. A curious aspect about the subtypes of C₄ grasses is their biogeography. In Australia and elsewhere, NADP-ME grasses are more frequent at higher rainfall, NAD-ME grasses predominate at lower rainfall, while the distribution of PCK grasses is even across rainfall gradients (Hattersley 1992).

The fundamental paradigm underpinning the efficiency of C₄ photosynthesis in terrestrial plants is the ‘division of labour’ between the initial fixation of CO₂ into C₄ acids, and their subsequent utilisation to generate high concentrations of CO₂ for ultimate fixation by Rubisco. The basic model for C₄ plants with classical kranz anatomy consists of two photosynthetic cycles (C₃ and C₄) operating across two photosynthetic cell types (mesophyll and bundle sheath), with strict cell- and organelle-specific localisation of key enzymes and with sufficient resistance to CO₂ back-diffusion. Indeed, the discovery of the kranz anatomy by Haberlandt preceded that of C₄ biochemistry by a century. The prevailing consensus has been that efficient C₄ photosynthesis necessitates the collaboration of two cell types.

Recently, this notion has been challenged by the discovery of non-kranz or single-cell C₄ photosynthesis in shrubs (Borszczowia aralocaspica and Bienertia cycloptera; Chenopodiaceae family) found in the salt deserts of Central Asia (Voznesenskaya et al. 2002, 2003). These plants show CO₂ and O₂ responses typical of C₄ photosynthesis but lack the kranz anatomy. They perform C₄ photosynthesis through the spatial localisation of dimorphic chloroplasts (as well as other organelles and photosynthetic enzymes) in distinct positions within a single chlorenchyma cell. Yet, the details of the partitioning differ between the two species (Edwards et al. 2004).

In Bienertia, the central cytoplasmic compartment of the chlorenchyma cell plays the role of bundle sheath cells in kranz-type C₄ (NAD-ME) plants; it is filled with mitochondria surrounded by chloroplasts. The peripheral cytoplasm lacks mitochondria and plays the role of the mesophyll cell in kranz-type C₄ plants. Accordingly, chloroplastic Rubisco and mitochondrial NAD-ME and glycine decarboxylase are restricted to the central compartment; chloroplastic pyruvate, Pi dikinase is restricted to the peripheral compartment, which is highly enriched with cytosolic PEP carboxylase. In Borszczowia, the compartmentation occurs at the distal (mesophyll equivalent) and proximal (bundle sheath equivalent) ends of the elongated, cylindrical chlorenchyma cell. The inter-connecting cytoplasm between the two intra-cellular compartments provides a liquid diffusion path, thus replacing the role of the bundle sheath cell wall in kranz-type C₄ plants (Edwards et al. 2004).

A low conductance for CO₂ diffusion out of the bundle sheath cells (or its equivalent cellular compartment) is critical for the efficient operation of C₄ photosynthesis. The total diffusive resistance to CO₂ has multiple components with different levels of contribution. These components include bundle sheath walls, membranes, bundle sheath chloroplast position, the site of C₄ acid decarboxylation, and the liquid-phase diffusion path. For kranz-type C₄ plants, calculated total bundle sheath resistance on a leaf area basis can range from 50 to 150 m² s^-1 mol^-1 (von Caemmerer and Furbank 2003). Evidently, single-cell C₄ plants have sufficient resistance to CO₂ back-diffusion which is essentially made of the cytoplasmic liquid phase and the special localisation of the (Rubisco-containing) chloroplasts surrounding the mitochondria (site of C₄ acid decarboxylation).

Thus, single-cell C₄ plants have efficient photosynthesis which is not inhibited by O₂, and their carbon isotope values are similar to kranz-type C₄ plants. Although, single-cell C₄ photosynthesis breaks away from the classical kranz anatomy, it remains within the general ‘division of labour’ paradigm.

More than 40 eudicot and monocot species distributed over 21 lineages have been reported to possess intermediate C₃ and C₄ photosynthetic characteristics and CO₂ compensation points. These intermediate species are likely remnants of the complex processes that led to the evolution of C₄ plants from C₃ ancestors, although reversions from the C₄ condition have also been suggested. Moreover, a number of identified C₃-C₄ species occur in taxa that are not closely related to any C₄ lineage, raising the possibility that the C₃-C₄ photosynthetic pathway may be a distinct adaptation. This, in addition to the small number of intermediate species found so far cast doubts over their physiological and ecological fitness, and whether they represent living fossils of evolutionary paths or evolutionary dead-ends (Rawsthorne 1992; Sage et al. 2011).

Leaves of all C₃-C₄ intermediates have partial or full kranz anatomy, with prominent bundle sheath cells containing chloroplasts and other organelles, and intermediate interveinal distances. Bundle sheath chloroplasts contain Rubisco and functional PCR cycle in both mesophyll and bundle sheath cells. Intermediate leaves also have CO₂ compensation points that are lower than what is observed for C₃ leaves and can be indistinguishable from C₄ leaves, due to reduced photorespiration. Biochemically, C₃-C₄ intermediates differ in the level of activity of the C₄ cycle and the extent to which CO₂ is concentrated in bundle sheath cells.

C₃-C₄ intermediate plants reduce photorespiration (and hence, CO₂ compensation point) using a ‘photorespiratory pump’ based on modified localisation of the mitochondrial photorespiratory enzyme, glycine decarboxylase (Figure 2.10). In these plants, glycine decarboxylase activity is restricted to bundle sheath cells and excluded from mesophyll cells. Consequently, photorespired CO₂ is released in the bundle sheath where it is largely refixed by Rubisco and the bundle sheath PCR before it diffuses back to the mesophyll. Such a system may weakly elevate CO₂ in bundle sheath cells. Intermediate species that rely on the ‘photorespiratory pump’ are termed C₃-like or Type I intermediates (e.g., Panicum milioides, Flaveria pubescens) and have intermediate CO₂ compensation points and negligible C₄ cycle activity.

In Type II and C₄-like intermediates (e.g., Flaveria brownii), up to 70% of atmospheric CO₂ may be first fixed into C₄ acids. These plants have C₄-like CO₂ compensation points but are not classified as C₄ plants because they lack the strict localisation of photosynthetic enzymes (e.g., Rubisco is present in mesophyll cells) and their bundle sheath cell walls have high CO₂ permeability, resulting in only a partial CO₂ concentrating mechanism (Brown 1980; Ku et al. 1991; Rawthorne 1992; Vogan and Sage 2011).

The physiological advantages of the intermediate photosynthetic pathway in all its naturally occurring forms remain unclear. It may be hypothesised that lowered photorespiration may lead to reduced CO₂ limitation of photosynthesis, and thus allow C₃-C₄ plants to operate with lower stomatal conductance, thus conferring higher water use efficiency relative to C₃ counterparts. Moreover, increased nitrogen cost associated with ‘building’ another set of photosynthetic cells (bundle sheath) may reduce nitrogen use efficiency if the gains in CO₂ uptake are not substantial.

Work conducted with C₃-C₄ species yielded inconclusive evidence on the likely advantages of C₃-C₄ photosynthesis relative to the ancestral C₃ mode. Generally, these studies demonstrated that, short of substantial C₄ cycle activity and advanced cell-specific localisation of C₃ and C₄ cycle enzymes between the mesophyll and bundle sheath cells, C₃-C₄ photosynthesis does not improve photosynthetic efficiency (Bolton and Brown 1980; Pinto et al. 2011, Vogan and Sage 2011). Therefore, partial recycling of photorespired CO₂ or a partial CO₂ concentrating mechanism reduce photorespiratory loss normally associated with C₃ photosynthesis, without leading to significant gains in plant fitness or productivity.

Joseph Holtum¹, Klaus Winter² and Barry Osmond³

¹Centre for Tropical Biodiversity and Climate Change, James Cook University, Australia; ²Smithsonian Tropical Research Institute, Balboa, Ancón, Republic of Panama; ³School of Biological Sciences, University of Wollongong, and Research School of Biology, Australian National University, Australia

Crassulacean acid metabolism (CAM) is a water-conserving mode of photosynthesis that, like C₄ photosynthesis, is a modification of the C₃ photosynthetic pathway fitted with a CO₂ concentrating mechanism (CCM) that can increase the [CO₂] around ribulose bisphosphate carboxylase/oxygenase (Rubisco) by more than 10-fold and suppress photorespiration. The overall energy demand of the CAM pathway is only about 10% more than that of C₃ photosynthesis, as costs of the CCM machinery are partially offset by reducing photorespiration.

In C₄ plants, as explained earlier in Section 2.2.2, this CCM is most commonly achieved by an “in-line turbocharger” based on initial CO₂ fixation by phosphoenolpyruvate carboxylase (PEPC) into C₄ acids in the cytoplasm of outer mesophyll cells. These acids diffuse rapidly to adjacent relatively CO₂-tight bundle-sheath cells (Figure 2.31 right) where CO₂ is released again. High [CO₂] builds up in this spatially separated compartment where it is refixed by Rubisco.

In CAM plants enzyme systems analogous to those in C₄ plants achieve the same result through a “battery-like” dark accumulation of CO₂ into the 2^nd carboxyl group of malic acid (acidification phase) in the vacuole of large mesophyll cells (Figure 2.31 left). Malic acid can accumulate to very high concentrations, attaining concentrations of greater than 1 mole acid per litre in mesophyll cells of tropical tree-CAM plants (Clusia spp.). Indeed one can sometimes taste the acid, with acid taste-testing for the presence of CAM being possibly first recorded in Aloe sp. by Nehemiah Grew in 1682 and in field reports from India by Benjamin Heyne in 1815.

In the light, malic acid returns to the cytoplasm where it is rapidly decarboxylated (deacidification phase). The CO₂ released, which accumulates to high internal [CO₂] as stomata close, is refixed by Rubisco in chloroplasts of the same mesophyll cell where it is further assimilated by the photosynthetic carbon reduction (PCR) cycle (Figure 2.32).  

Ultimately three of the four carbons recovered from the malic acid must be stored as starch and/or sugars in order to provide to PEPC the C₃ substrate required for CO₂ uptake during the following night. The fourth carbon, effectively that obtained from the atmosphere, is available for growth. Deacidification may generate high [CO₂] behind closed stomata but photorespiration is not completely abolished (Lüttge 2002) since photosynthesis also generates high internal [O₂]. While exploring Lake Valencia in Venezuela in 1800, Alexander von Humboldt measured elevated [O₂] in bubbles streaming from the cut base of presumably CAM Clusia leaves standing in water in the light.

Of course by closing stomata in the light CAM plants minimize water loss when evaporative demand is highest (von Caemmerer and Griffiths 2009). The biomass production per unit water utilized in CAM was 6 times higher than for C₃ plants and 2 times higher than for C₄ plants when plants exhibiting all three photosynthetic pathways were grown together in a garden outdoors (Winter et al. 2005). The attribute of water-use efficiency undoubtedly contributes significantly to the success of CAM photosynthesis in nature, with CAM species outnumbering C₄ species by about two to one. The paradoxes of CAM, a mode of photosynthesis that involves stomatal opening and CO₂ uptake during the dark, continue to inform many aspects of plant biochemistry, physiology, ecology and evolution. This article draws heavily on two recent reviews (Borland et al. 2011; Winter et al. 2015).

Vascular plants often inhabit regions subject to tidal submergence while others carry out their entire life cycle under water. Examples of common submerged aquatic macrophytes are pond weeds and seagrasses. Once again, an evolutionary selective pressure for these plants has been the availability of CO₂. Low levels of dissolved CO₂ are common in both inland and marine waters, particularly at more alkaline pH. In more productive inland lakes, CO₂ content can vary enormously, requiring considerable flexibility in the actual mode of carbon acquisition. At high pH, HCO₃^– becomes the more abundant form of inorganic carbon, whereas dissolved CO₂ will predominate at low pH (Section 18.2). Consequently, when SAM plants evolved from their C₃ progenitors on land, there was some adaptive advantage in devices for CO₂ accumulation because CO₂ rather than HCO₃^– is substrate for Rubisco. The nature of this ‘CO₂ pump’ and the energetics of carbon assimilation are not fully characterised in SAM plants but considerable CO₂ concentrations do build up within leaves, enhancing assimilation and suppressing photorespiration.

In summary:

Regardless of photosynthetic mode, and despite catalytic limitations, Rubisco is ubiquitous and remains pivotal to carbon gain in our biosphere. As a corollary, carbon loss via photorespiration is an equally universal feature of C₃ leaves, and the evolution of devices that overcome such losses have conferred significant adaptive advantages to C₄, CAM and SAM plants.

 

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Brown DA (1980). Photosynthesis of grass species differing in carbon dioxide fixation pathways. IV. Analysis of reduced oxygen response in Panicum milioides and Panicum schenckii. Plant Physiol 65: 346-349

Christin PA, Samaritani E, Petitpierre B et al. (2009) Evolutionary insights on C₄ photosynthetic subtypes in grasses from genomics and phylogenetics. Genom Biol Evol 1: 221–230

Edwards GE, Franceschi VR, Voznesenskaya EV (2004) Single-cell C₄ photosynthesis versus the dual-cell (Kranz) paradigm. Annu Rev Plant Biol 55: 173–196

Ehleringer JR, Cerling TE, Helliker BR (1997) C₄ photosynthesis, atmospheric CO₂, and climate. Oecologia 112: 285-299

Ghannoum O, von Caemmerer S, Conroy JP (2002) The effect of drought on plant water use efficiency of nine NAD–ME and nine NADP–ME Australian C₄ grasses. Funct Plant Biol 29: 1337-1348

Ghannoum O, Evans JR, Chow WS et al. (2005) Faster rubisco is the key to superior nitrogen-use efficiency in NADP-malic enzyme relative to NAD-malic enzyme C₄ grasses. Plant Physiology 137: 638-650

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(a) Historical evidence for photorespiration

The first line of evidence for photorespiration came from the different compensation points of C3 and C4 plants. When air is recirculated over an illuminated leaf in a closed system, photosynthesis will reduce CO₂ concentration to a low level where fixation of CO₂ by photosynthesis is just offset by release from respiration. For many C₃ plants this compensation point is around 50 µL L^–1 but is markedly affected by oxygen, photon irradiance and leaf temperature (Tregunna et al. 1966; Zelitch 1966). In low concentrations (1–2% O₂) the CO₂ compensation point of C3 plants is near zero. Significantly, early researchers in this area had already noted that some tropical grass species appeared to have a compensation point at or close to zero CO₂, even in normal air (20% O₂). This was first reported for corn (Zea mays) (Meidner 1962) and raised a very perplexing question as to whether these species even respired in light. However, we now know that C₄ photosynthesis is responsible for the low evolution of CO₂ (Section 2.2) and that C₄ plants have a CO₂ concentrating mechanism that forestalls photorespiration, resulting in a CO₂ compensation point close to zero.

A second line of evidence for leaf respiration in light was provided by a transient increase in release of CO₂ when leaves are transferred from light to dark. This ‘post-illumination CO₂ burst’ was studied extensively during the early 1960s by Gleb Krotkov and colleagues at Queens University (Kingston, Ontario). The intensity of this burst increased with the photon irradiance during the preceding period of photosynthesis. Understandably, the Queens group regarded this post-illumination burst as a ‘remnant’ of respiratory processes in light, and coined the term ‘photorespiration’. A functional link with the CO₂ compensation point was inferred, because the burst was abolished in low O₂ (Figure 2.12). A competitive inhibition by O₂ on CO₂ assimilation was suspected and was subsequently proved to be particularly relevant in defining Rubisco’s properties. Nevertheless, for many years a biochemical explanation for interaction between these two gases remained elusive.

(b) Biochemistry

Significant progress came when Ludwig and Krotkov designed an open gas exchange system in which ¹⁴CO₂ was used to separate the fixing (photosynthetic) and evolving (respiratory) fluxes of CO₂ for an illuminated leaf (Ludwig and Canvin 1971). Results using this steady-rate labelling technique were particularly revealing and provided the first direct evidence that respiratory processes in light were qualitatively different from those in darkness. They were able to show that CO₂ evolved during normal high rates of photosynthesis by an attached sunflower leaf was derived from currently fixed carbon. The specific activity of evolved CO₂ (ratio of ¹⁴C to ¹²C) was essentially the same as that of the CO₂ being fixed, indicating that photorespiratory substrates were closely related to the initial products of fixation. Ludwig and Krotkov concluded that ¹⁴CO₂ supplied to a photosynthesising leaf was being re-evolved within 28–45 s! Furthermore the rate of CO₂ evolution in light was as much as three times the rate in darkness, and while early fixed products of photosynthesis (intermediates of the PCR cycle) were respired in light, this was not the case in darkness.

The radiolabelling method of Ludwig and Krotkov had, for the first time, provided measurements of what could be regarded as a true estimate of light-driven respiration which was not complicated by transient effects (as the post-illumination burst had been), or by changes in CO₂ concentration (as was the case for measurements in closed gas exchange systems or in CO₂-free air) or by difficulties associated with detached organs. Ludwig and Canvin (1971) subsequently concluded that processes underlying photorespiration re-evolved 25% of the CO₂ which was being fixed concurrently by photosynthesis. Such a rate of CO₂ loss was not a trivial process so a biochemical basis for its operation had to be established, and particularly when photorespiration seemed to be quite different from known mechanisms of dark (mitochondrial) respiration.

The search for the substrates of ‘photorespiration’ occupied many laboratories worldwide for many years. Much work centred on synthesis and oxidation of the two-carbon acid glycolate because as early as 1920 Warburg had reported that CO₂ fixation by illuminated Chlorella was inhibited by O₂, and under these conditions the alga excreted massive amounts of glycolic acid (Warburg and Krippahl 1960). Numerous reports on the nature of the ¹⁴C-labelled products of photosynthesis showed that glycolate was a prominent early-labelled product. A very wide variety of research with algae and leaves of many higher plants established two significant features of glycolate synthesis: formation was enhanced in either low CO₂ or high O₂. Both of these features had been predicted from Ludwig’s physiological gas exchange work and eventually proved a key to understanding the biochemistry of photorespiration.

(c) Source of glycolate production

The photosynthetic carbon reduction (PCR) cycle for CO₂ fixation (Section 2.1) involves an initial carboxylation of ribulose-1,5-bisphosphate (RuBP) to form 3-PGA, but makes no provision for glycolate synthesis. However, Wang and Waygood (1962) had described the ‘glycolate pathway’, namely a series of reactions in which glycolate is oxidised to glyoxylate and aminated, first to form glycine and subsequently the three-carbon amino acid serine. The intracellular location of this pathway in leaves was established in a series of elegant studies by Tolbert and his colleagues who also established that leaf microbodies (peroxisomes) were responsible for glycolate oxidation and the synthesis of glycine. Kisaki and Tolbert (1969) suggested that the yield of CO₂ from the condensation of two molecules of glycine to form serine could account for the CO₂ evolved in photorespiration. This idea was incorporated in later formulations of the pathway.

What remained elusive was the source of photosynthetically produced glycolate. Many studies had suggested that the sugar bisphosphates of the PCR cycle could yield a two-carbon fragment which, on the basis of short-term ¹⁴CO₂ fixation, would have its two carbon atoms uniformly labelled (if the two carbons were to be derived directly from 3-PGA this would not be the case as PGA was asymmetrically labelled in the carboxyl group). The mechanism was likened to the release of the active ‘glycolaldehyde’ transferred in the thiamine pyrophosphate (TPP)-linked transketolase-catalysed reactions of the PCR cycle. In some cases significant glycolate synthesis from the sugar bisphosphate intermediates of the cycle were demonstrated in vitro; however, the rates were typically too low to constitute a viable mechanism for glycolate synthesis in vivo.

A more dynamic approach to carbon fixation was needed to resolve this impasse. In particular, the biochemical fate of early products would have to be traced, and using a development of the open gas exchange system at Queens, Atkins et al. (1971) supplied ¹⁴CO₂ in pulse–chase experiments to sunflower leaf tissue under conditions in which photorespiration was operating at high rates (21% O₂) or in which it was absent (1% O₂). A series of kinetic experiments showed that synthesis of ¹⁴C-glycine and ¹⁴C-serine was inhibited in low O₂ and that the ¹⁴C precursor for their synthesis was derived from sugar bisphosphates of the PCR cycle, especially RuBP. Indeed RuBP was the obvious source of glycine carbon atoms and the kinetics of glycine turnover closely matched those of RuBP. As these authors concluded, ‘we can no longer view this (glycolate) pathway as an adjunct to the Calvin cycle but must incorporate it completely into the carbon fixation scheme for photosynthesis’ (Atkins et al. 1971).

The question was finally and most elegantly resolved by Ogren and Bowes (1971) who demonstrated that the carboxylating enzyme of the PCR cycle, RuBP carboxylase, was both an oxygenase and a carboxylase! During normal photosynthesis in air, this enzyme thus catalysed formation of both P-glycolate (the precursor of glycolate) and 3-PGA from the oxygenation of RuBP as well as two molecules of PGA from carboxylation of RuBP. In effect, CO₂ and O₂ compete with each other for the same active sites for this oxygenation/carboxylation of RuBP, at last providing a biochemical mechanism which had confused and perplexed photosynthesis researchers since the 1920s. This primary carboxylating enzyme of the PCR cycle, which had hitherto rejoiced in a variety of names (carboxydismutase, fraction 1 protein, RuDP carboxylase and RuBP carboxylase), was renamed Rubisco (ribulose-1,5-bisphosphate carboxylase/oxygenase) to reflect its dual activity.

A scheme for the PCR cycle and photosynthetic carbon oxidation (PCO) pathways then represents the synthesis of almost 70 years of research effort, and integrates the metabolism of P-glycolate with the PCR cycle. This is shown in the following section. 

 

A scheme for photorespiration involving the photosynthetic carbon reduction (PCR) cycle and the photosynthetic carbon oxidation (PCO) pathways represents the synthesis of almost 70 years of research. It integrates the metabolism of P-glycolate with the PCR cycle. This scheme is shown in Figure 2.13.

Specialised reactions within three classes of organelles in leaf cells are required, namely chloroplasts, peroxisomes (originally called microbodies) and mitochondria. Their close proximity in leaf cells (Figure 2.14 below) plus specific membrane transporters facilitate the exchange of metabolites.

Within choroplasts, oxygenase activity by Rubisco results in formation of phosphoglycolate which then enters a PCO cycle, and is responsible for loss of some of the CO₂ just fixed in photosynthesis.

Within peroxisomes, O₂ is consumed in converting glycolate to glyoxylate, and aminated to form glycine.

Within mitochondria, CO₂ is released during conversion of glycine to serine. Subsequently, serine is recovered by peroxisomes where it is further metabolised, re-entering the PCR cycle of chloroplasts as glycerate. About 75% of carbon skeletons channelled into photorespiration are eventually recovered as carbohydrate.

Transport of glycerate and glycolate across the inner membrane of chloroplasts may involve separate translocators as shown in Figure 2.13, or it may involve a single translocator that exchanges two glycolate molecules for one molecule of glycerate. Transport of metabolites across the peroxisomal membrane most likely occurs through unspecific channel proteins, similar to those in the outer membranes of mitochondria and chloroplasts. These outer membranes are not included in this diagram. Mitochondria take up two molecules of glycine and release one molecule of serine. A specific translocator most probably mediates the exchange of these amino acids.

Not only does the photosynthetic oxidation pathway consume O₂ and release CO₂ but ammonia is also produced by mitochondria during synthesis of serine from glycine (Figure 2.13). This ammonia would be extremely toxic if it were not metabolised by either cytosolic or chloroplastic glutamine synthetase. A very effective herbicide that blocks glutamine synthetase has been developed (phosphinothricin, also known as glufosinate or Basta), and when it is applied to (or expressed in) actively growing plants they are killed by their photorespiratory ammonia release.

In summary, participation of photorespiration in leaf gas exchange, and thus dry matter accumulation by plants, reflects kinetic properties of Rubisco, and in particular a relatively high affinity for CO₂ (K_m = 12 µM) compared with a much lower affinity for O₂ (K_m = 250 µM). That contrast in affinity is, however, somewhat offset by the relative abundance of the two gases at catalytic sites of the enzyme where the ratio of O₂:CO₂ partial pressures approaches 1000:1! Thus, CO₂ assimilation always prevails over CO₂ loss in photorespiration. 

At around the same time as the nature of photorespiration was becoming clearer Hatch and Slack (1966) demonstrated that in tropical grasses (initially sugar cane) the first-formed products of photosynthetic CO₂ fixation were the four-carbon acids oxalacetate, malate and aspartate, rather than the 3-PGA formed in the PCR cycle. Furthermore, the carboxylation reaction involved PEP carboxylase and carbon was subsequently transferred to PCR cycle intermediates. As noted earlier (Section 2.2) C₄ plants show no apparent CO₂ release in light. The explanation lies in their anatomy and multiple carboxylation reactions rather than in the absence of the pathway of photorespiration. Bundle sheath cells are equipped with a CO₂-concentrating mechanism that favours carboxylation over oxygenation reactions due to increased partial pressure of CO₂, while photorespiratory release of CO₂ is further prevented through the activity of PEP carboxylase which refixes any respired CO₂ formed from the oxygenase function of Rubisco.

Unicellular green algae also posed a problem for the simple extrapolation of early models for photorespiratory metabolism in C₃ leaves. Although organisms such as Chlorella had been used to establish the PCR cycle, and indeed provided much early evidence for effects of O₂ on photosynthesis and formation of glycolate in light, they also appeared to lack CO₂ evolution in light (Lloyd et al. 1977). In this case the explanation lies in a CO₂-concentrating mechanism which effectively increases the internal pool of inorganic carbon (CO₂ and HCO₃^–) thereby favouring the carboxylase function of Rubisco over its oxygenase function.

Such a substantial loss of carbon concurrent with CO₂ fixation raises the question of whether eliminating or minimising photorespiration in C₃ plants could enhance their yield, and specifically that of major crop plants such as rice, wheat, grain legumes, oil seeds and trees, none of which are C₄ species. Faced with an expanding world human population and an increasing demand for food and animal feed, enhanced agricultural productivity is a global necessity. In its most obvious form a scenario which alters or removes the oxygenase function of Rubisco could achieve such a goal. In an early review of the process of photorespiration in plants, Ogren (1984) noted that ‘the sequence of reactions constituting the photorespiratory pathway in C₃ plants appears to be firmly established’ and he went on to suggest that, although reducing the loss of fixed carbon as CO₂ in the process may be a valid goal to improve the yield of crop plants, it is not clear whether or not this can be achieved by specific changes to the kinetic and catalytic properties of Rubisco alone.

Photorespiration may be loosely considered as a wasteful process because previously fixed carbon is lost and energy is dissipated. Ideal destinations for photoassimilates include synthetic pathways leading to fixed biomass and respiratory pathways for re-release of fixed energy in a controlled sequence of reactions leading to ATP and NAD(P)H for use in other synthetic events.

However, situations do exist where energy dissipation via photorespiration can be beneficial. For example, photo-oxidative damage can be alleviated in shade-adapted plants that experience strong irradiance if photorespiratory processes are allowed to proceed. Depriving such plants of an external oxygen supply, and hence preventing photosynthetic carbon oxidation, will exacerbate chloroplast lesions due to strong irradiance. Photosynthetic variants which obviate photo-respiratory loss, and most notably C₄ plants, integrate structure and function in a way that forestalls photo-oxidative damage and leads to their outstanding performance under warm conditions. Environmental factors that constitute selection pressure for such photosynthetic adaptation are reviewed by Sage et al (2012). 

Atkins CA, Canvin DT, Foch H (1971) Intermediary metabolism of photosynthesis in relation to carbon dioxide evolution in sunflower. In MD Hatch, CB Osmond, RO Slatyer, eds, Photosynthesis and Photorespiration. John Wiley, New York, pp 497-505

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Starch is the principal storage carbohydrate in plants and this carbon reserve plays a number of important roles in plants. It is composed of two polymers of glucose, amylose and amylopectin and is stored in the plastid (chloroplast in leaves, amyloplasts in non-photosynthetic tissues) as insoluble, semi-crystalline granules. Starch is accumulated during rapid growth in the day and is almost completely degraded at night to mostly glucose and maltose, which is exported from the chloroplast and metabolised in the cytosol (Figure 2.19). Starch degradation is initiated by the addition of phosphate groups at the C6-position and C3-position of individual glucosyl residues that act to disrupt the packing of the glucans at the granule surface. These phosphate additions are catalysed by two enzymes, glucan water dikinase (GWD) and phosphoglucan water dikinase (PWD) respectively. The hydrolysis of the resulting glucan and phosphoglucan chains is carried out by a suite of enzymes including the phosphoglucan phosphatases (SEX4/LSF2), β-amylases (BAM1/BAM3), debranching enzymes (DBE; ISA3/LDA), α-amylase (AMY3), α-glucan phosphorylase and the disproportionating enzyme 1 (D-enzyme 1; an α-1,4-glucanotransferase).). The resulting maltose and glucose are exported to the cytosol by the glucose transporter (pGlcT) and maltose transporter (MEX1) and glucose-1-phosphate is thought to be exported by a similar but yet unknown mechanism. Once in the cytosol the maltose and glucose are converted to substrates for either sucrose synthesis, glycolysis or the oxidative pentose phosphate pathway by a number of enzymes including the disproportionating enzyme 2 (D-enzyme 2; an α-1,4-glucanotransferase), α-glucan phosphorylase, hexokinase and phosphoglucomutase.

Sucrose is the world’s most abundant disaccharide, it is only produced by photosynthetic organisms and serves a role as a transportable carbohydrate and sometimes as a storage compound. The reactions in plant tissues leading to degradation of sucrose to hexose monophosphates are outlined in Figure 2.20.

The first step is cleavage of the glycosidic bond by either invertase (Equation 2.1) or sucrose synthase (Equation 2.2).

\[\text{Sucrose + H}_2\text{O} \rightarrow \text{D-Glucose + D-Fructose}\tag{2.1}\]

\[\text{Sucrose + UDP} \rightarrow \text{UDP-Glucose + D-Fructose}\tag{2.2}\]

Plant tissues contain distinct invertases located in the vacuole, cell wall (acid invertases) cytosol, mitochondria, nucleus, and cholorplast (neutral/alkaline invertases) which hydrolyse sucrose to glucose and fructose in an irreversible reaction. The invertases are differentially regulated by a number of mechanisms including pH to allow them to function in cell expansion, supply of carbon skeletons and energy metabolism. Multiple isoforms of sucrose synthase are located in the cytosol or cytosolic membranes that catalyse a thermodynamically reversible reaction, but this reaction probably acts only to breakdown sucrose in vivo. Their activity is developmentally regulated and they have functions in the supply of activated glucose for starch and cellulose biosynthesis. While both invertase and sucrose synthase can both breakdown sucrose, research using knockouts of multiple isoforms of both enzymes has shown that sucrose synthase is not required for normal growth in Arabidopsis, whereas invertase is indispensable. However this does not rule out the requirement of sucrose synthase in certain tissues of crop plant tubers, seeds and fruits where it has been shown to be crucial. Glucose and fructose are metabolised further by phosphorylation to the corresponding hexose-6-P by hexokinase. Hexokinase in plant tissues is associated with the outer surface of mitochondria.

The glycolytic pathway involves the oxidation of the hexoses and hexose phosphates molecules produced from the breakdown of starch or sucrose to generate ATP, reductants and pyruvate (Figure 2.21).

Regulation of PFK and PPi-PFK is achieved by a combination of mechanisms, including pH, the concentration of substrates and effector metabolites and changes in subunit association. Phosphoenolpyruvate (PEP) is a potent inhibitor of both of PFK and PPi-PFK, inhibiting at µM concentrations and Pi can activate the cytoplasmic PFK, whereas the plastidic form is slightly inhibited by Pi. A number of other effectors of PFK have been identified including ADP, 3-phosphoglycerate and phosphoglycolate as well as it ability to accept ribonucleoside triphosphates other than ATP as the phosphate donor. PP_i-PFK, has a catalytic potential higher than that of PFK and is strongly activated by Fructose-2,6-bisphosphate, but has no effect on PFK.

Fructose-1,6-bisphosphate is cleaved by fructose bisphosphate aldolase to form glyceraldehyde-3-phosphate and dihydroxyacetone phosphate, and these triose phosphates can be interconverted in a reaction catalysed by triose phosphate isomerase. Glyceraldehyde-3-phosphate is oxidised to 1,3-bisphosphoglycerate by a nicotinamide adenine dinucleotide (NAD⁺)-dependent glyceraldehyde 3-P dehydrogenase. Glyceraldehyde 3-P dehydrogenase is sensitive to inhibition by the reduced pyridine nucleotide cofactor (NADH), which must be reoxidised to maintain the flux through the glycolytic pathway. A phosphate group is then transferred from 1,3-bisphosphoglycerate to ADP forming ATP and 3-phosphogylcerate by phosphoglycerate kinase. In the cytosol a bypass is present that can convert glyceraldehyde-3-phosphate directly to 3-phosphoglycerate without phosphorylation by a non-phosphorylating NADP dependent glyceraldehyde 3-phosphate dehydrogenase. The resulting 3-phosphoglycerate is then converted to phosphenolpyruvate (PEP) by the action of phosphoglycerate mutase and then enolase.  

The end-products of glycolytic reactions in the cytosol are determined by the relative activities of the two enzymes that can utilise PEP as a substrate: pyruvate kinase, which forms pyruvate and a molecule of ATP, and PEP carboxylase, which forms oxaloacetate (Figure 2.21). Both of these reactions are essentially irreversible and there are fine controls that regulate the partitioning of PEP between these reactions. Pyruvate kinase is controlled post translationally by a partial C-terminal truncation which may yield altered regulatory properties and a phosphorylation and ubiquitin conjugation that targets the protein to the 26S proteasome for complete degradation, it is also inhibited by ATP. Whereas PEP carboxylase is inhibited by malate and thus its regulation is independent of cell energy status. The sensitivity of PEP carboxylase to malate is regulated by phosphorylation of a N-terminal serine of the enzyme, with the phosphorylated form less sensitive to malate inhibition. Oxaloacetate is then reduced by malate dehydrogenase to malate which, along with pyruvate, can be taken up into mitochondria and metabolised further in the TCA cycle (see below). The reduction of oxaloacetate in the cytosol could provide a cytosolic mechanism for oxidising NADH formed by glyceraldehyde 3-P dehydrogenase (Figure 2.21).

Another level of regulation of components of glycolysis is their physical location within the plant cell. Under conditions of high respiratory activity, a greater proportion of the cytosolic enzymes of glycolysis are present on the surface of mitochondria. In contrast when respiration is experimentally inhibited, a decrease in the association of glycolytic enzymes with the mitochondria is observed. It is likely that the glycolytic enzymes associate dynamically with mitochondria to support respiration and that this association restricts the use of glycolytic intermediates by competing metabolic pathways.

An alternative route for the breakdown of glucose-6-phosphate is provided by the oxidative pentose phosphate pathway (OPP) (Figure 2.22). This pathway functions mainly to generate reductant (i.e. NADPH) for biosynthetic processes including the assimilation of inorganic nitrogen and fatty acid biosynthesis and to maintain redox potential to protect against oxidative stress. In addition, the reversible oxidative section of the pathway is the source of carbon skeletons for the synthesis of a number of compounds. For example ribose-5-phosphate provides the ribosyl moiety of nucleotides and is a precursor for the biosynthesis of purine skeletons and erythrose-4-phosphate, which is the precursor for the biosynthesis of aromatic amino acids by the shikimic acid pathway.

 

The pathway begins with the dehydrogenation of glucose-6-phosphate catalyzed by glucose-6-phosphate dehydrogenase to produce 6-phosphoglucolactone and is the first step of the oxidative phase of the pathway. The 6-phosphoglucolactone is then hydrolysed to 6-phosphogluconate by 6-phosphogluconolactonase and then undergoes oxidative decarboxylation by 6-phosphogluconate dehydrogenase to produce ribulose-5-phosphate in the final step of the oxidative phase. Overall this phase of the pathway produces two molecules of NADPH from the conversion of glucose-6-phosphate to ribulose-5-phosphate. The non-oxidative phase begins with the reaction of ribulose-5-phosphate with either ribulose-5-phosphate isomerase or ribulose-5-phosphate epimerase followed by a series of reactions catalyzed by transaldolase and transketolase. These reactions result in the production of two molecules of fructose-6-phosphate and one glyceraldehyde 3-phosphate. The glyceraldehyde 3-phosphate and fructose-6-phosphate in the oxidative pentose phosphate pathway may be exchanged with enzymes of glycolysis.

As with glycolysis, reactions of the pentose phosphate pathway are catalysed by different isoforms of the enzymes that occur either in the cytosol or in plastids. Although transketolase and transaldolase may be absent from the cytosol of some species, the activity is maintained by phosphate translocator proteins on the plastid inner-envelope membrane that have the capacity to translocate sugar phosphates.

Organic acids such as pyruvate and malate produced in the cytosol by processes described above are further oxidised in mitochondria by the tricarboxylic acid (TCA) cycle and subsequent respiratory chain. Energy released by this oxidation is used to synthesise ATP which is then exported to the cytosol for use in biosynthesis and growth.

(a)  Mitochondrial structure

Plant mitochondria (Figure 2.23) are typically double-membrane organelles where the inner membrane is invaginated to form folds known as cristae to increase the surface area of the membrane. The outer membrane contains relatively few proteins (<100) and is permeable to most small compounds (< Mr=5 kDa) due to the presence of the pore-forming protein VDAC (voltage dependent anion channel) which is a member of the porin family of ion channels. The inner membrane is the main permeability barrier of the organelle and controls the movement of molecules by means of a series of carrier proteins many of which are members of mitochondrial substrate carrier family (MSCF). The inner membrane also houses the large complexes that carry out oxidative phosphorylation and encloses the soluble matrix which contains the enzymes of the TCA cycle and many other soluble proteins involved in a myriad of mitochondrial functions.

Mitochondria are semi-autonomous organelles with their own DNA and protein synthesis machinery. However, the mitochondrial genome encodes only a small portion of the proteins which make up the mitochondrion; the rest are encoded on nuclear genes and synthesised in the cytosol. These proteins are the transported into the mitochondrion by the protein import machinery and assembled with the mitochondrially synthesised subunits to form the large respiratory complexes. The number of mitochondria per cell varies with tissue type (from a few hundred in mature differentiated tissue to some thousands in specialised cells). Understandably, more active cells, with high energy demands, such as those in growing meristems are generally equipped with larger numbers of mitochondria per unit cell volume, and consequently show faster respiration rates.

(b)  Mitochondrial substrates

Two substrates are produced from glycolytic PEP for oxidation in mitochondria: malate and pyruvate (Figure 2.21). These compounds are thought to be the most abundant mitochondrial substrates in vivo. However, amino acids may also serve as substrates for mitochondrial respiration in some tissues, particularly in seeds rich in stored protein or under conditions of sugar depletion such as extended darkness, shading and senescence. β-oxidation of fatty acids typically does not occur in plant mitochondria, this oxidation is principally carried out in peroxisomes in plants.

(c) Carbon metabolism in mitochondria

Malate and pyruvate enter the mitochondrial matrix across the inner membrane via separate carriers. Malate is then oxidised by either malate dehydrogenase (a separate enzyme isoform from that in the cytosol), which yields oxaloacetate (OAA) and reduced nicotinamide adenine dinucleotide (NADH), or NAD⁺-linked malic enzyme, which yields pyruvate and NADH and releases CO₂ (Figure 2.24). Cytosolic pyruvate carboxylase is an alternative means of providing substrate to mitochondria by combining pyruvate with HCO₃ to yield OAA that can then be imported into mitochondria.

 

Pyruvate formed either from malate and malic enzyme or transported directly from the cytosol is oxidised inside mitochondria by the pyruvate dehydrogenase complex (PDC) to form CO₂, acetyl-CoA and NADH. This enzyme, which requires coenzyme A, thiamine pyrophosphate and lipoic acid as cofactors, effectively links the TCA cycle to glycolysis. PDC comprises three enzymes E1 (2-oxo acid dehydrogenase), E2 (acyltransferase) and E3 (lipoamide dehydrogenase). This complex is regulated by phosphorylation of the E1 subunit, lowering PDC activity in the day and increasing PDC activity at night. Pyruvate dehydrogenase is also subject to feedback inhibition from acetyl-CoA and NADH.

(d) Tricarboxylic acid (TCA) cycle

The TCA cycle begins with the condensation of acetyl-CoA and OAA, to form the six-carbon molecule citrate and the release coenzyme A (CoA) (Figure 2.24) in a reaction catalysed by citrate synthase. Aconitase catalyses the next step, converting citrate to isocitrate in a two-step reaction (dehydration/hydration) with cis-aconitate as an intermediate.

NAD-linked isocitrate dehydrogenase then oxidatively decarboxylates isocitrate to form CO₂ and 2-oxoglutarate, and reduce NAD⁺ to NADH. The 2-oxoglutarate formed is also oxidatively decarboxylated to succinyl-CoA in a reaction catalysed by the enzyme 2-oxoglutarate dehydrogenase. This enzyme complex has similarities to pyruvate dehydrogenase and its reaction is analogous to the formation of acetyl-CoA from pyruvate by pyruvate dehydrogenase. The reaction mechanisms are also very similar, with 3 subunit enzymes, but 2-oxoglutarate dehydrogenase is not subject to the phosphorylation control that regulates pyruvate dehydrogenase. Succinyl-CoA synthase then catalyses the conversion of succinyl-CoA to succinate, with the concomitant phosphorylation of ADP to ATP, the only substrate-level phosphorylation step in the mitochondrion. This enzyme in plants differs from its mammalian counterpart in that it is specific for ADP rather than GDP.

Succinate dehydrogenase (SDH, Complex II), which catalyses the oxidation of succinate to fumarate, is the only membrane-bound enzyme of the TCA cycle and is part of the respiratory electron transfer chain (Figure 2.25). SDH is a large complex consisting of four core subunits, as well as number other associated subunits.

Fumarase catalyses the hydration of fumarate to malate followed by malate dehydrogenase that catalyses the final step of the TCA cycle, oxidising malate to OAA and producing NADH. The reaction is freely reversible, although the equilibrium constant strongly favours the reduction of OAA, necessitating rapid turnover of OAA and NADH to maintain this reaction in a forward direction.

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

Regulation of carbon flux through the TCA cycle probably occurs via phosphorylation/dephosphorylation of pyruvate dehydrogenase, which will depend in turn on mitochondrial energy status and feedback inhibition of various enzymes by NADH and acetyl-CoA. The rate of cycle turnover thus depends on the rate of electron flow through the respiratory chain (to reoxidise NADH) and the utilisation of ATP in the cell to provide ADP for substrate level and oxidative phoshorylation. TCA cycle turnover will also depend on the rate of substrate provision by reactions in chloroplasts and cytosol. A number of studies of TCA cycle mutants have demonstrated the wide impact these enzymes have not simply on TCA cycle function but as steps for the delivery of organic acids for other processes in plant cells such a photosynthetic performance, plant biomass, root growth, photorespiration, nitrogen assimilation, amino acid metabolism, and stomatal function.

The respiratory electron transfer chain (ETC) of mitochondria consists of a series of large membrane-bound protein complexes (Complexes I, II, III, IV) which together with a small lipid ubiquinone (UQ) and the small protein cytochrome c catalyse the transfer of electrons from NADH and succinate to O₂, forming H₂O (Figure 2.25). Electron flow from NADH and succinate to oxygen is coupled to proton translocation out of the matrix to the intermembrane space which establishes a proton electrochemical gradient (Dµ_H⁺) across the inner membrane that is used to drive phosphorylation of ADP to form ATP by the F₁F_O ATP synthase (Complex V, Figure 2.25).

(a) Complex I (CI)

 

NADH-UQ oxidoreductase, is responsible for the oxidation of matrix NADH and reduction of ubiquinone (UQ) in the inner mitochondrial membrane (Figure 2.25). In plants it is a large multi-subunit complex composed of 49 subunits, up to ten of which are synthesised in the mitochondrion whilst the others are imported from the cytosol. One of the subunits, a 50 kDa protein, contains flavin mononucleotide as a cofactor and is the dehydrogenase which oxidises NADH and passes electrons to iron-sulphur containing subunits of the complex, and eventually to ubiquinone. The passage of electrons through the complex is accompanied by H⁺ translocation across the membrane. Complex I is inhibited specifically by the flavonoid rotenone and its analogues. The NADH-binding site is exposed to the matrix and the complex oxidises NADH produced by the TCA cycle and other NAD-linked enzymes (Figure 2.25). Studies of mutations of CI subunits have shown that plants can survive without CI due to the activity of alternative NAD(P)H dehydrogenases (see below). Such mutants have a variety of interesting phenotypes including viral infection tolerance, prolonged hydration under water-deficient conditions and altered organic and amino acid concentrations. Using a series of Arabidopsis CI subunit knockout mutants, a number membrane arm subcomplexes (of 200, 400, 450 and 650 kDa) have been identified using BN-PAGE and antibodies. It is proposed that at least some of these subcomplexes may be assembly intermediates during CI formation, and these are seen to accumulate when specific subunits are absent.

(b) Complex II (CII)

Succinate dehydrogenase, is an enzyme of both the respiratory ETC and the TCA cycle (see above) (Figure 2.25). It is composed of four core subunits: a flavoprotein (SDHI), an iron-sulphur subunit (SDH2) and two membrane anchor subunits (SDH3 and SDH4) in most organisms. In plants, the purification of the complex has revealed the common core subunits, but also additional proteins of unknown function that co-migrate with the complex. All SDH subunits are encoded in the nuclear genome in Arabidopsis. SDH contains FeS and flavin centres which participate in electron transfer from succinate to ubiquinone. Unlike complex I, complex II does not pump H⁺ and succinate oxidation is therefore linked to the synthesis of less ATP per O₂ reduced (see below). Malonate, an analogue of succinate, is a strong competitive inhibitor of succinate dehydrogenase. Knockout mutants of the SDH1 gene have been shown to be embryo lethal, but knockdown of SDH1 and SDH2 leads a array of phenotypes including altered stomatal aperture, mitochondrial ROS production and nitrogen use efficiency. SDHAF1 and SDHAF2 assist in CII assembly in plants and knockdown of the SDHAF2 homolog lowers SDH assembly and reduces root growth.

(c) Complex III (CIII)

Ubiquinone-cytochrome c oxidoreductase or the cytochrome b/c₁ complex as it is sometimes known, contains 10 subunits including a number of bifunctional core proteins. These proteins act both in CIII function and as a matrix processing peptidase, removing targeting presequences from imported matrix synthesized proteins (Figure 2.25). A single subunit of this complex, cytochrome b, is encoded by the plant mitochondrial genome, whilst the others are encoded by the nuclear genome. It contains two b-type cytochromes, b₅₆₆ and b₅₆₂, cytochrome c₁, an iron-sulpher protein named the Rieske iron–sulphur protein and several other polypeptides. Electron flow from ubiquinol to cytochrome c is accompanied by the translocation H⁺ across the membrane, via the so-called Q cycle. Various inhibitors of complex III have been discovered, with antimycin A and myxothiazol most widely used in research. The assembly of CIII is modular and includes an early core subcomplex, a late core subcomplex and the final dimeric CIII. Approximately 13 assembly factors implicated in aiding one or more of the different stages of CIII assembly in yeast, however little is known about CIII assembly or functional assembly factors in plants.

(d) Complex IV (CIV)

Cytochrome c oxidase is the final step of electron transfer of the classical ETC. As the name implies, cytochrome c oxidase accepts electrons from cytochrome c and transfers them to O₂ which is reduced to form H₂O. Purification of CIV in plants has identified a complex containing 14 protein subunits (Figure 2.25). Eight of these proteins are homologous to known CIV subunits from other organisms, together with a further six proteins that are probably plant specific. Two cytochrome haem centres, a and a₃, and two copper atoms make up its redox active components and like complex I, cytochrome oxidase is a proton pump. Cytochrome c oxidase is sensitive to a number of inhibitors, the best known of which are carbon monoxide and cyanide. Plants, however, show resistance to both carbon monoxide and cyanide because they are equipped with an alternative oxidase (see below). Studies of human and yeast CIV has shown an assembly pathway that involves the sequential incorporation of CIV subunits, initiated by subunit 1 and assisted by over 40 assembly factors. Research investigating the plant homolog of the yeast assembly factor COX19 has found it is capable of complementing the yeast cox19 null mutant. This suggests it might play a role in the biogenesis of plant cytochrome c oxidase or in the replacement of damaged forms of the enzyme. However, our knowledge of the assembly of CIV in plants is still incomplete.

(e) Ubiquinone and Cytochrome c

These large multi-subunit complexes (I, II, III, IV) of the respiratory ETC chain are embedded in the inner mitochondrial membrane by virtue of their hydrophobic subunits, and interact with one another via two smaller molecules: ubiquinone and cytochrome c. The lipid-soluble ubiquinone also known as coenzyme Q₁₀ is a small mobile electron carrier which moves rapidly along and across the membrane, and participates in H⁺ transport across the membrane via the Q-cycle as well as shuttling electrons from complexes I and II to complex III. Cytochrome c is a small haem-containing protein located on the outer surface of the inner membrane, which shuttles electrons between complexes III and IV. In this respect, the respiratory chain is similar in layout to the photosynthetic electron transport chain: three large complexes which communicate by a quinone and a small mobile protein (Cyt c or plastocyanin). However, orientation of components in the membrane is inverted and the net reaction catalysed is opposite to that in chloroplasts (Figure 1.12).

When electrons are transferred from NADH to O₂, a large release of redox energy enables ATP formation in complex V of the respiratory chain (Figure 2.24). Energy release associated with electron transport is conserved by H⁺ translocation across the membrane to form a proton electrochemical gradient (Δµ_H⁺) that has both an electrical membrane potential (Δψ) and a pH component (Δµ_H⁺ = Δψ + ΔpH). This is known as the chemiosmotic theory and was originally proposed by Peter Mitchell in 1960s. In plant mitochondria, Δµ_H⁺ exists mainly as a Δψ of ~150–200 mV, with a pH gradient (ΔpH) of ~0.2–0.5 units. ATP synthesis occurs as H⁺ move from a compartment of high potential (the intermembrane space) to one of low potential (the mitochondrial matrix) through the ATP synthase complex. Oxidation of NADH via the cytochrome pathway has three associated H⁺ translocation sites and is linked to synthesis of up to three ATP molecules for each molecule of NADH oxidized. By contrast, both succinate and alternative NADH oxidation by the rotenone-insensitive NADH dehydrogenases (see below) are linked to the synthesis of only two ATP molecules per NADH or succinate, as these events are associated with only two H⁺ pumping sites.

 

(a) Complex V (CV)

ATP synthase is a membrane-bound F₁F₀ type H⁺-ATP synthase that harnesses the Δµ_H⁺ generated by the ETC to produce ATP. It is composed of a hydrophobic F₀ component which channels protons through the inner mitochondrial membrane and also anchors the complex to the membrane and a hydrophilic F₁ component which catalyses ATP formation and protrudes into the matrix. The core subunits of the enzyme are highly conserved in both prokaryotic and eukaryotic organisms. In plants, the majority of mitochondrial F₁ subunits are encoded in the nucleus and translated in the cytosol before being imported into the mitochondria (including  α, β, γ and ε subunits), while most of the F₀ subunits are encoded in the plant mitochondrial genome and translated in the mitochondrial matrix (including a, b, c and A6L subunits).  The reaction mechanism of the ATP synthase is known as the three-site alternating binding site mechanism. According to this model, F₁ has three nucleotide-binding sites which can exist in three configurations: one with loosely bound nucleotides, one with tightly bound nucleotides and the third in a nucleotide-free state. H⁺ movement through F₀ results in rotation of F₁, causing a conformational change during which the site with loosely bound ADP and P_i is converted to one which binds them tightly in a hydrophobic pocket in which ATP synthesis occurs. Further H⁺ movement then causes another rotation of F₁ and the ATP binding site is exposed and releases the nucleotide. In the meantime, the other nucleotide-binding sites are undergoing similar changes, with ADP and P_i being bound and converted to ATP. Thus H⁺ translocation drives the three sites through three different configurations and the main expenditure of energy is in the induction of a conformational change that releases tightly bound ATP, rather than in ATP synthesis itself. The F₀ complex also contains a protein known as the oligomycin-sensitivity-conferring protein (OSCP) because it binds the antibiotic oligomycin that prevents H⁺ translocation through F₀ and inhibits ATP synthesis. Therefore, adding oligomycin to mitochondria oxidising a substrate in the presence of ADP restricts O₂ uptake. Generally knockouts of ATP synthase core subunits are lethal in plants, however inducible knockdowns have enabled investigations into the tissue-specific phenotypes incurred by slowing the rates of mitochondrial ADP:ATP cycling at a number of different developmental stages. It has been proposed that the assembly of plant CV comprises of three steps, the first being the formation of a rapidly turned over F1 subcomplex in the matrix, followed by an intermediate stage where F1 associates with the inner membrane and still turns over at a fast rate, and then finally a union of F₁ with F_O to form functional CV. A number of assembly factors (Atp10, Atp11, Atp12, Atp22, Atp23 and Fmc1) have been discovered for yeast ATP synthase, however, a detailed study of the presence and conservation of CV assembly factors in plants has not been undertaken.

(b) Respiratory control

Electron transport through the respiratory chain, and therefore rate of O₂ uptake, is controlled by availability of ADP and P_i, a phenomenon described as ‘respiratory control’. In the absence of ADP or P_i, the proton pore of ATP synthase is blocked and a Δµ_H⁺ builds up to a point where it restricts further H⁺ translocation across the inner membrane. Since electron transport is functionally linked to H⁺ translocation, this elevated Δµ_H⁺ will also restrict O₂ consumption. That outcome is easily seen with isolated mitochondria (Figure 2.26) where O₂ uptake is stimulated by adding ADP (‘State 3’ respiration). When all of the added ADP has been consumed, O₂ uptake decreases again (‘State 4’). In steady state, the rate of electron flow is determined by the rate of flow of H⁺ back across the membrane: when ADP and P_i are available the backflow is rapid and occurs via ATP synthase; in the absence of these compounds, backflow is by slow diffusion through the membrane.

The ratio of State 3 to State 4 (the respiratory control ratio) is thus an indication of coupling between ADP phosphorylation and electron transport. Larger values represent tighter coupling. The proton leak can be dramatically stimulated by some compounds which act as protonophores or proton channels; these compounds collapse the Δµ_H⁺ and increase O₂ uptake up to the State 3 rate (Figure 2.26). However, no ATP is formed and these compounds are called uncouplers because they uncouple the linked processes of electron transport and phosphorylation.

Plant mitochondria have a respiratory chain which is more complicated than that of animals and contains alternative NADH dehydrogenases, alternative oxidases which catalyse cyanide-insensitive O₂ consumption and uncoupling proteins that acts to dissipate the Δµ_H⁺. The alternative NADH dehydrogenases and alternative oxidase do not translocate protons and therefore are not linked to ATP synthesis; they are often referred to as the non-phosphorylating bypasses of the plant respiratory chain. These pathways were initially identified in plant mitochondria as they are able to continue to respire in the presence of the CIV inhibitor, cyanide and the CI inhibitor, rotenone and by their ability to exhibit natively uncoupled respiration in the absence of an ADP source.

(a) Alternative oxidase

Cyanide-insensitive respiration is catalysed by the alternative oxidase (AOX). This alternative terminal oxidase is a diiron quinol oxidase that branches from the classical respiratory chain at UQ and reduces oxygen to water without an associated proton translocation. The oxidase exists in mitochondria as a dimer which can be inactivated by covalent linkage via disulphide bonds. The reduced enzyme is stimulated allosterically by pyruvate and some other 2-oxo acids (such as glyoxylate), which interact directly with the oxidase. The exact role of AOX continues to be debated but it appears to play an antioxidant role in plant mitochondria. Research has shown it is actively induced by oxidative stress and the different genes for the oxidase have been shown to be both development- and tissue-specific. Knockout of AOX leads to reactive oxygen species and anthocyanin accumulation in the leaves exposed to a combination of high light and drought stress. AOX can be inhibited by hydroxamic acids such as n-propylgallate (nPG) and salicyl hydroxamic acid (SHAM).

(b) Alternative NAD(P)H dehydrogenases

Alternative NAD(P)H dehydrogenases have been shown to be present on both sides of the inner mitochondrial membrane. These type II NAD(P)H dehydrogenases oxidise external or cytosolic and matrix NADH and NADPH and are insensitive to the classical CI inhibitor rotenone. As with AOX, these enzymes do not translocate protons and therefore are not linked to ATP synthesis. The Arabidopsis genome contains seven genes encoding NAD(P)H dehydrogenases, although it appears that some of these isoforms are present in multiple subcellular compartments in addition to mitochondria.

(c) Uncoupling proteins

Uncoupling proteins (UCPs) are members of the mitochondrial carrier family of proteins. They act to dissipate the Δµ_H⁺ built up the ETC by transporting H⁺ back across the inner membrane uncoupling proton and electron transport. The reactive oxygen species superoxide activates UCPs and this suggests a possible mechanism for the engagement of this enzyme in vivo. Analysis of knockouts of UCP (AtUCP1) showed that its absence led to localized oxidative stress but did not impair the ability of the plant to withstand a wide range of abiotic stresses. However, knockout of UCP1 did limit the photorespiration rate of plants and led to a reduction in photosynthetic carbon assimilation. This suggests that the main role of UCP1 in leaves is to maintain the redox poise of the mitochondrial ETC to facilitate photosynthesis.

(a)  Efficiency

Respiration represents a substantial loss of carbon from a plant, and under adverse conditions can be as high as two-thirds of the carbon fixed daily in photosynthesis. Both the rate and the efficiency of respiration will therefore affect plant growth significantly. The overall process of respiration results in the release of a substantial amount of energy which may be harnessed for metabolic work. In theory, the energy released from the complete oxidation of one molecule of glucose to CO₂ and H₂O in respiratory reactions leads to the synthesis of 36 molecules of ATP. However, in plants, because there are alternative routes for respiration, this yield can be greatly reduced. Mechanisms for regulating respiration rates in whole plants remain unclear. Convention has it that the rate of respiration is matched to the energy demands of the cell through feed-back regulation of glycolysis and electron transport by cytosolic ATP/ADP. However, since plants have non-phosphorylating bypasses in their respiratory chain that are insensitive to ATP levels, and since PEP carboxylase and PFP might be involved in sucrose degradation, the situation in vivo is not so simple. For example, the rotenone-insensitive alternative NADH dehydrogenases requires high concentrations of NADH in the matrix before it can operate and seems to be active only when substrate is plentiful and electron flow through complex I is restricted by lack of ADP. Alternative oxidase activity also depends on carbon and ADP availability and its flux is very dependent on the degree of environmental stress of the plant. In other words, non-phosphorylating pathways act as carbon or reductant ‘overflows’ of the main respiratory pathway and will only be active in vivo when sugar levels are high and the glycolytic flux rapid, when the cytochrome chain is inhibited, or when the bypasses have been induced significantly during stress. In glycolysis, the interaction between environmental signals and key regulatory enzymes, as well as the role of PFP and its activator fructose-2,6-P₂, will be important.

(b)  Allocation of respiratory energy to process physiology

One way of viewing respiratory cost for plant growth and survival is by subdividing measured respiration into two components associated with (1) growth and (2) maintenance. This distinction is somewhat arbitrary, and these categories of process physiology must not be regarded as discrete sets of biochemical events. Such energy-dependent processes are all interconnected because ATP represents a universal energy currency for both, while a common pool of substrates is drawn upon in sustaining production of that ATP. Nevertheless, cells do vary in their respiratory efficiency, while genotype × environment interactions are also evident in both generation and utilisation of products from oxidative metabolism. The benefit of a high respiration rate is that more ATP is produced, which provides vital energy for growth of new tissue and defence processes, such as antioxidant activation, metabolite transport or production of resistant protein isoforms. However, the cost of high respiration rates is that carbon is expended on respiration instead of being allocated to synthesis of new tissue, therefore limiting growth capacity. Variation in respiration rate has implications for growth and resource use efficiency in plants during drought, temperature and salinity responses of plants.

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