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Carbon fixation in C 4 plants

Carbon fixation via crassulacean acid metabolism (cam), differences in carbon fixation pathways, the molecular biology of photosynthesis.

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Certain plants—including the important crops sugarcane and corn (maize), as well as other diverse species that are thought to have expanded their geographic ranges into tropical areas—have developed a special mechanism of carbon fixation that largely prevents photorespiration. The leaves of these plants have special anatomy and biochemistry. In particular, photosynthetic functions are divided between mesophyll and bundle-sheath leaf cells. The carbon-fixation pathway begins in the mesophyll cells, where carbon dioxide is converted into bicarbonate, which is then added to the three-carbon acid phosphoenolpyruvate (PEP) by an enzyme called phosphoenolpyruvate carboxylase. The product of this reaction is the four-carbon acid oxaloacetate , which is reduced to malate , another four-carbon acid, in one form of the C 4 pathway. Malate then is transported to bundle-sheath cells, which are located near the vascular system of the leaf. There, malate enters the chloroplasts and is oxidized and decarboxylated (i.e., loses CO 2 ) by malic enzyme. This yields high concentrations of carbon dioxide, which is fed into the Calvin-Benson cycle of the bundle sheath cells, and pyruvate , a three-carbon acid that is translocated back to the mesophyll cells. In the mesophyll chloroplasts, the enzyme pyruvate orthophosphate dikinase (PPDK) uses ATP and P i to convert pyruvate back to PEP, completing the C 4 cycle. There are several variations of this pathway in different species. For example, the amino acids aspartate and alanine can substitute for malate and pyruvate in some species.

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The C 4 pathway acts as a mechanism to build up high concentrations of carbon dioxide in the chloroplasts of the bundle sheath cells. The resulting higher level of internal carbon dioxide in these chloroplasts serves to increase the ratio of carboxylation to oxygenation, thus minimizing photorespiration. Although the plant must expend extra energy to drive this mechanism, the energy loss is more than compensated by the near elimination of photorespiration under conditions where it would otherwise occur. Sugarcane and certain other plants that employ this pathway have the highest annual yields of biomass of all species. In cool climates, where photorespiration is insignificant, C 4 plants are rare. Carbon dioxide is also used efficiently in carbohydrate synthesis in the bundle sheath.

PEP carboxylase, which is located in the mesophyll cells, is an essential enzyme in C 4 plants. In hot and dry environments , carbon dioxide concentrations inside the leaf fall when the plant closes or partially closes its stomata to reduce water loss from the leaves. Under these conditions, photorespiration is likely to occur in plants that use Rubisco as the primary carboxylating enzyme, since Rubisco adds oxygen to RuBP when carbon dioxide concentrations are low. PEP carboxylase, however, does not use oxygen as a substrate, and it has a greater affinity for carbon dioxide than Rubisco does. Thus, it has the ability to fix carbon dioxide in reduced carbon dioxide conditions, such as when the stomata on the leaves are only partially open. As a consequence, at similar rates of photosynthesis, C 4 plants lose less water when compared with C 3 plants. This explains why C 4 plants are favored in dry and warm environments.

In addition to C 3 and C 4 species, there are many succulent plants that make use of a third photosynthetic pathway: crassulacean acid metabolism (CAM). This pathway is named after the Crassulaceae , a family in which many species display this type of metabolism, but it also occurs commonly in other families, such as the Cactaceae , the Euphorbiaceae , the Orchidaceae , and the Bromeliaceae . CAM species number more than 20,000 and span 34 families. Almost all CAM plants are angiosperms ; however, quillworts and ferns also use the CAM pathway. In addition, some scientists note that CAM might be used by Welwitschia , a gymnosperm . CAM plants are often characterized by their succulence, but this quality is not pronounced in epiphytes that use the CAM pathway.

CAM plants are known for their capacity to fix carbon dioxide at night, using PEP carboxylase as the primary carboxylating enzyme and the accumulation of malate (which is made by the enzyme malate dehydrogenase) in the large vacuoles of their cells. Deacidification occurs during the day, when carbon dioxide is released from malate and fixed in the Calvin-Benson cycle, using Rubisco. During daylight hours, the stomata are closed to prevent water loss. The stomata are open at night when the air is cooler and more humid, and this setting allows the leaves of the plant to assimilate carbon dioxide. Since their stomata are closed during the day, CAM plants require considerably less water than both C 3 and C 4 plants that fix the same amount of carbon dioxide in photosynthesis.

The productivity of most CAM plants is fairly low, however. This is not an inherent trait of CAM species, because some cultivated CAM plants (e.g., Agave mapisaga and A. salmiana ) can achieve a high aboveground productivity. In fact, some cultivated species that are irrigated, fertilized, and carefully pruned are highly productive. For example, prickly pear ( Opuntia ficus-indica ) and its thornless variety, O. amyclea , produce 4.6 kg per square meter (0.9 pound per square foot) of new growth per year. Such productivity is among the highest of any plant species. Thus, the rates of photosynthesis of CAM plants may be as high as those of C 3 plants, if morphologically similar plants adapted to the similar habitats are compared.

The unusual capacity of CAM plants to fix carbon dioxide into organic acids in the dark, causing nocturnal acidification, with deacidification occurring during the day, has been known to science since the 19th century. (There is evidence, however, that the Romans noticed the difference between the morning acid taste of some of the house plants they cultivated.) On the other hand, the C 4 pathway was discovered during the middle of the 20th century. A full appreciation of CAM as a photosynthetic pathway was greatly stimulated by analogies with C 4 species.

A comparison of the differences between the various carbon pathways is provided in the table.

Oxygenic photosynthesis occurs in the prokaryotic cells called cyanobacteria and in eukaryotic plant cells ( algae and higher plants ). In eukaryotic plant cells , which contain chloroplasts and a nucleus , the genetic information needed for the reproduction of the photosynthetic apparatus is contained partly in the chloroplast chromosome and partly in chromosomes of the nucleus. For example, the carboxylation enzyme ribulose 1,5-bisphosphate carboxylase is a large protein molecule comprising a complex of eight large polypeptide subunits and eight small polypeptide subunits. The gene for the large subunits is located in the chloroplast chromosome, whereas the gene for the small subunits is in the nucleus. Transcription of the DNA of the nuclear gene yields messenger RNA (mRNA) that encodes the information for the synthesis of the small polypeptides. During this synthesis, which occurs on the cytosolic ribosomes , some extra amino acid residues are added to form a recognition leader on the end of the polypeptide chain. This leader is recognized by special receptor sites on the outer chloroplast membrane; these receptor sites then allow the polypeptide to penetrate the membrane and enter the chloroplast. The leader is removed, and the small subunits combine with the large subunits, which have been synthesized on chloroplast ribosomes according to mRNA transcribed from the chloroplast DNA. The expression of nuclear genes that code for proteins needed in the chloroplasts appears to be under control of events in the chloroplasts in some cases; for example, the synthesis of some nuclear-encoded chloroplast enzymes may occur only when light is absorbed by chloroplasts.

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Article Contents

Introduction, development, biochemistry, regulation of gene function, c 4 systems, future perspectives, acknowledgments.

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C 4 Cycles: Past, Present, and Future Research on C 4 Photosynthesis

Address correspondence to [email protected] .

www.plantcell.org/cgi/doi/10.1105/tpc.111.092098

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Jane A. Langdale, C 4 Cycles: Past, Present, and Future Research on C 4 Photosynthesis, The Plant Cell , Volume 23, Issue 11, November 2011, Pages 3879–3892, https://doi.org/10.1105/tpc.111.092098

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In the late 1960s, a vibrant new research field was ignited by the discovery that instead of fixing CO 2 into a C 3 compound, some plants initially fix CO 2 into a four-carbon (C 4 ) compound. The term C 4 photosynthesis was born. In the 20 years that followed, physiologists, biochemists, and molecular and developmental biologists grappled to understand how the C 4 photosynthetic pathway was partitioned between two morphologically distinct cell types in the leaf. By the early 1990s, much was known about C 4 biochemistry, the types of leaf anatomy that facilitated the pathway, and the patterns of gene expression that underpinned the biochemistry. However, virtually nothing was known about how the pathway was regulated. It should have been an exciting time, but many of the original researchers were approaching retirement, C 4 plants were proving recalcitrant to genetic manipulation, and whole-genome sequences were not even a dream. In combination, these factors led to reduced funding and the failure to attract young people into the field; the endgame seemed to be underway. But over the last 5 years, there has been a resurgence of interest and funding, not least because of ambitious multinational projects that aim to increase crop yields by introducing C 4 traits into C 3 plants. Combined with new technologies, this renewed interest has resulted in the development of more sophisticated approaches toward understanding how the C 4 pathway evolved, how it is regulated, and how it might be manipulated. The extent of this resurgence is manifest by the publication in 2011 of more than 650 pages of reviews on different aspects of C 4 . Here, I provide an overview of our current understanding, the questions that are being addressed, and the issues that lie ahead.

The Discovery

In 1956, the pathway through which plants fix CO 2 into organic acids was elucidated ( Bassham et al., 1956 ). The subsequently named Calvin-Benson or C 3 cycle uses the enzyme ribulose-1,5-bis-phosphate carboxylase/oxygenase (Rubisco) to fix CO 2 into the three-carbon compound 3-phosphoglycerate ( Figure 1A ). At the time, it was generally assumed that the Calvin-Benson cycle accounted for CO 2 assimilation in all plants. However, further 14 CO 2 labeling experiments revealed that in maize ( Zea mays ) and sugarcane ( Saccharum officinarum ), the four-carbon compounds malate and Asp were among the earliest labeled products ( Karpilov, 1960 ; Kortschak et al., 1965 ). The significance of these findings was not fully understood until M.D. Hatch and C.R. Slack proposed a model for the C 4 dicarboxylic acid pathway, wherein CO 2 is initially fixed into a four-carbon compound, subsequently decarboxylated, and then refixed into a three-carbon compound ( Hatch and Slack, 1966 ; Hatch, 2002 ). These three steps define the canonical C 4 photosynthetic pathway.

Schematic of C 3 Calvin-Benson and NADP-ME C 4 Cycles.

Calvin-Benson (A) and NADP-ME C 4 (B) cycles. The green box represents the chloroplast. Blue dots represent active transport steps.

Variations on a Theme

Variants of C 4 biochemistry have been found in a marine macroalga ( Udotea flabellum ) ( Reiskind and Bowes, 1991 ), a diatom ( Thalassiosira weissflogii ) ( Roberts et al., 2007 ), and in both aquatic (reviewed in Bowes, 2011 ) and terrestrial angiosperms. Some of these variants operate in the context of a single cell, but in most cases, the C 4 pathway is partitioned between two morphologically distinct cell types known as bundle sheath (BS) and mesophyll (M) cells (reviewed in Edwards et al., 2004 ). In C 4 plants, these BS and M cells surround the leaf veins in concentric circles, leading to a wreath-like appearance. This specialized arrangement was named Kranz anatomy (from the German word for wreath) many years before its association with the C 4 pathway was elucidated ( Haberlandt, 1896 ), but the link is now very well established, and as with the biochemistry, many variations on the Kranz theme exist ( Brown, 1975 ; reviewed in Edwards and Voznesenskaya, 2011 ).

In the context of the two-cell C 4 pathway, three biochemical subtypes have been defined that differ in the subcellular localization and type of C 4 acid decarboxylase used in the BS cells (reviewed in Drincovich et al., 2011 ). The first to be discovered was the NADP-malic enzyme (ME) type, in which the decarboxylation step is performed in BS chloroplasts by NADP-dependent ME ( Figure 1B ). In this pathway, CO 2 enters the M cell cytoplasm where it is first converted to bicarbonate ions by carbonic anhydrase (CA) and is then fixed by phospho enol pyruvate carboxylase (PEPCase) to form oxaloacetate (OAA). OAA is subsequently transported from the M cytoplasm to the M chloroplast where it is converted to malate by NADP-malate dehydrogenase. Malate is then transported out of the M chloroplast and into the BS chloroplast, a process that requires transport across the chloroplast and plasma membranes of both cell types. In the BS cell chloroplast, malate is decarboxylated by NADP-ME, and the released CO 2 is refixed by Rubisco in the Calvin-Benson (C 3 ) cycle. The pyruvate generated by the decarboxylation reaction is transported back from the BS chloroplast to the M chloroplast where it acts as a substrate for pyruvate orthophosphate dikinase (PPdK) to regenerate phosphoenolpyruvate (PEP). The cycle is restarted when PEP is transported from the M chloroplast to the M cell cytoplasm to combine once again with CO 2 .

The key features of the NADP-ME subtype are movement of malate and pyruvate between M and BS cells and decarboxylation of malate in the BS chloroplasts. By contrast, the NAD-ME and phosphoenolpyruvate carboxykinase (PEP-CK) subtypes both move Asp and Ala between M and BS cells. Asp is converted to either malate or OAA, and then malate is decarboxylated by NAD-ME in the BS cell mitochondria or OAA is decarboxylated by PEP-CK in the BS cell cytoplasm. Notably, the NAD-ME and PEP-CK pathways have higher energy requirements than the NADP-ME pathway, and both have more intracellular transport steps. In the PEP-CK subtype, PEP-CK and NAD-ME decarboxylases can operate in parallel, placing an even greater energetic load on the process ( Burnell and Hatch, 1988 ). PEP-CK activity has also been detected in maize, which is classically considered as an NADP-ME subtype, raising the question of whether the subtype classification is actually robust ( Furbank, 2011 ).

The energetic cost of the C 4 pathway is offset by the fact that all forms of the pathway act to concentrate CO 2 at the site of Rubisco. This carbon-concentrating mechanism prevents oxygen from competing for the active site of Rubisco and thus reduces the energetically wasteful process of photorespiration, which in C 3 plants can reduce photosynthetic output by up to 40% ( Ehleringer et al., 1991 ). However, these recognized gains demand the development of specialized leaf anatomy and the compartmentalization of biochemical reactions. This in turn requires sophisticated regulatory processes to operate at all levels of gene expression and protein function.

Phylogenetic Diversity

In land plants, the C 4 pathway is found only in angiosperms. In this group, there are 62 C 4 taxa that comprise 36 eudicots, 6 sedges, 18 grasses, and 2 aquatic lineages in the genera Hydrilla and Egeria ( Sage et al., 2011 ). While the evolutionary independence of all of these lineages is not clear, it is indisputable that the C 4 pathway arose multiple independent times from the ancestral C 3 pathway ( Christin et al., 2010 ). In most cases (58 lineages), the pathway evolved in association with Kranz anatomy, but in the aquatic lineages and in two Chenopod lineages ( Binertia and Suaeda ), the pathway operates in a single cell. In the aquatic species, CO 2 is concentrated from the cytoplasm to the chloroplast ( Bowes, 2011 ), whereas in the Chenopods, CO 2 is concentrated from an outer to an inner region of the cell ( Edwards and Voznesenskaya, 2011 ). In total, there are ~7500 C 4 species, most of which use the NADP-ME pathway and most of which (~4600 species) are grasses ( Sage et al., 2011 ).

The phylogenetic distribution of C 4 grasses is notable in that they all occur in the so-called PACMAD clade ( Christin et al., 2009a ). This group comprises the six subfamilies Panicoideae, Arundinoideae, Chloridoideae, Micrairoideae, Aristidoideae, and Danthonioideae and thus includes the agronomically important crop plants maize, sorghum ( Sorghum bicolor ), and sugarcane. With at least 17 independent origins of C 4 proposed in this clade ( Christin et al., 2007 , 2008a ), and none in the other seven grass families, it is tempting to speculate that a preconditioning event occurred in the last common PACMAD ancestor. In this regard, it may not be a coincidence that low levels of CA, the first enzyme of the C 4 shuttle, are a characteristic of the entire clade ( Edwards et al., 2007 ).

Ecological Drivers

Given the multiple independent origins of C 4 , it is not easy to identify the evolutionary drivers. However, because the C 4 pathway concentrates CO 2 at the site of Rubisco and because it is only energetically favorable in warm arid climates, three paleoclimatic drivers have been proposed: declines in CO 2 , increases in temperature, and periods of drought. Notably, C 3 photosynthesis evolved in a CO 2 -rich atmosphere of well over 1000 ppm, but atmospheric CO 2 levels dropped around 32 to 25 million years ago in the Oligocene, to ~500 ppm ( Pagani et al., 2005 ). Molecular dating of the C 4 grass lineages suggests that the first transition from C 3 to C 4 occurred around 30 million years ago, coincident with this reduction in atmospheric CO 2 levels ( Christin et al., 2008a ; Vicentini et al., 2008 ). However, C 4 lineages continued to appear over the subsequent 20 million years ( Christin et al., 2008a ; Vicentini et al., 2008 ) and the ecological dominance of C 4 grasslands did not occur until 8 to 6 million years ago ( Cerling et al., 1997 ). Thus, while declining CO 2 levels may have facilitated C 4 evolution, other factors influenced its expansion.

Biogeographical and phylogenetic studies have attempted to characterize the emergence and ultimate dominance of C 4 plants (particularly grasses) in certain environmental niches. Crucially, the level of atmospheric CO 2 at which C 4 outcompetes C 3 is dependent on temperature. C 4 is favored at 550 ppm CO 2 at 35°C, 450 ppm at 30°C, and 350 ppm at 25°C ( Ehleringer et al., 1997 ). Given this interdependence, it might be predicted that C 4 plants evolved first in the tropics and only moved north as atmospheric CO 2 levels dropped to levels of ~250 ppm in the Miocene. However, although most C 4 species are found growing in high-temperature climates, the analysis of a 1200-taxon grass phylogeny alongside climate data for each of the species failed to correlate C 4 with any of a number of temperature parameters ( Edwards and Smith, 2010 ) ( Figure 2 ). Instead, there was compelling evidence to suggest that 18 of the 20 C 4 origins examined were correlated with marked reductions in annual rainfall.

Comparison between Photosynthetic Pathway and Mean Annual Temperature for 1200 Grass Species Representing 20 Origins of C 4 .

(Reprinted with permission from Edwards and Smith [2010] , Figure 2.)

Despite the inference that the evolution of C 4 was influenced by reduced water availability ( Edwards and Smith, 2010 ), the issue remains far from resolved. Other reports suggest that although extant C 4 species are preferentially localized in arid environments, drought was not a driver for C 4 evolution. Instead, it is suggested that once the pathway had evolved, C 4 as opposed to C 3 grasses were more likely to make the transition into arid habitats ( Osborne and Freckleton, 2009 ). The Miocene-Pliocene expansion of C 4 grasslands, to the point where 3% of vascular plant species account for 25% of terrestrial photosynthesis, is further proposed to have resulted from combinations of coevolution with grazing mammals ( Bouchenak-Khelladi et al., 2009 ), increased temperature, increased summer rainfall, and more frequent occurrence of fire (discussed in Osborne, 2011 ).

The difficulty of trying to understand the complex interplay between paleoclimatic factors that favored C 4 versus C 3 physiology is illustrated by the results of a long-term elevated CO 2 experiment. Although elevated CO 2 is predicted to favor productivity in C 3 plants, when combined with an increase in temperature, the opposing effects of CO 2 and temperature on soil water content led to enhanced productivity in C 4 rather than C 3 prairie grasses ( Morgan et al., 2011 ). Enhanced photosynthetic activity was also seen in C 4 maize plants when exposed to elevated CO 2 levels in the field ( Leakey et al., 2004 ). In a similar paradox, despite the fact that C 4 species generally occupy drier habitats than C 3 species, a comparison of physiological properties in a range of grass species demonstrated that the performance advantages of C 4 photosynthesis are actually reduced by drought ( Taylor et al., 2011 ). In light of such apparent contradictions, it seems that we may have overestimated our ability to identify the ecological drivers for C 4 evolution and to predict how the pathway will respond to future climate change.

Developmental Innovations

The evolution of C 4 photosynthesis required the modification of leaf development programs. In single-cell C 4 systems, intracellular partitioning mechanisms evolved, while in two-cell systems, specialized Kranz leaf anatomy developed. Insight into how these developmental pathways may have evolved has been obtained from comparisons between development in extant C 3 and C 4 species and by the examination of species that develop traits intermediate between C 3 and C 4 . Such intermediates have been identified in a number of genera, most of which are eudicots (reviewed in Sage et al., 2011 ). In families such as Flaveria , C 3 , C 3 -C 4 intermediate, C 4 -like, and C 4 species have all been identified ( Ku et al., 1983 ). Intermediate Flaveria species may thus represent a transitional phase of C 4 evolution. However, other intermediates, such as Moricandia arvensis ( Holaday et al., 1981 ), occur in families with no known C 4 species. Although it is possible that C 4 species have yet to evolve in these families, it is perhaps more likely that such intermediates define a distinct developmental state.

In addition to obligate C 3 -C 4 intermediates, there are a number of examples where C 4 development is induced by environmental cues. These facultative C 4 systems provide an opportunity to examine the developmental transition from C 3 to C 4 in the context of individual plants. Examples of this type include Eloecharis vivipara , an aquatic species that develops C 3 anatomy in submerged leaves and C 4 anatomy in aerial leaves ( Ueno et al., 1988 ). Interestingly, in this system, the transition from C 3 to C 4 can also be induced by abscisic acid ( Ueno, 1998 ). Another well-studied example is Flaveria brownii , in which the extent of C 4 induction is correlated with light intensity; plants grown in higher light intensities are more C 4 -like than those grown at lower intensities ( Monson et al., 1987 ; Cheng et al., 1989 ).

C 3 Development Is Default

The single-cell C 4 pathway operates in aquatic C 4 species and in the terrestrial chenopods Binertia and Suaeda . Leaf development in these species is quite remarkable in that chlorenchyma cells are organized into two distinct cytoplasmic compartments that are maintained by an organized network of actin filaments and microtubules ( Chuong et al., 2006 ). In Binertia , there is a centrally located compartment surrounded by a more peripheral compartment ( Voznesenskaya et al., 2002 ; Offermann et al., 2011 ), whereas in Suaeda , the two compartments are distal (toward the outside of the leaf) and proximal in the cell ( Voznesenskaya et al., 2001 ). In each of the two compartments, chloroplasts accumulate a distinct complement of photosynthetic enzymes with the peripheral/distal chloroplasts analogous to M cell chloroplasts of the Kranz system and the central/proximal chloroplasts analogous to the BS cell chloroplasts. Crucially, this dimorphism is not apparent early in development in that a monomorphic C 3 chloroplast state develops by default and the C 4 pattern is induced by later developmental cues ( Voznesenskaya et al., 2005 ; Lara et al., 2008 ).

The development of a default C 3 state in C 4 plants is not confined to species with single-cell systems. A similar situation occurs in both the monocot maize and the eudicot amaranth, where Rubisco accumulates in both BS and M cell chloroplasts unless light and/or developmental cues restrict accumulation to BS cells ( Langdale et al., 1988 ; Wang et al., 1993 ). In maize, it has been concluded that the C 4 -inducing signals are only perceived in cells that are within a two-cell radius of a vein ( Langdale and Nelson, 1991 ). This deduction is based on the observation that in leaf-like organs, such as the husk leaf sheath, where up to 20 cells separate vein pairs, dimorphic chloroplast development is only observed in cells immediately surrounding the vasculature ( Langdale et al., 1988 ; Pengelly et al., 2011 ).

Veins Act as Organizing Centers

It is perhaps not surprising that veins play a key role in the differentiation of C 4 leaf anatomy since one of the most obvious differences between leaf morphology in C 3 and C 4 plants is leaf venation pattern. Measurements of vein density in a range of C 3 and C 4 species demonstrated that veins are consistently more closely spaced in C 4 species ( Crookston and Moss, 1974 ). Furthermore, quantitative measurements of BS-to-M cell ratios in C 3 and C 4 leaves showed that in C 4 plants the ratio approaches 1:1 ( Hattersley and Watson, 1975 ; Dengler et al., 1994 ; Muhaidat et al., 2007 ). This ratio equates to veins (V) being separated by only four photosynthetic cells in C 4 leaves as opposed to up to 20 cells in C 3 leaves ( Figure 3 ). As such, the repeating V-BS-M-M-BS-V unit of Kranz anatomy is generated. One notable exception to this repeating pattern is found in Arundinella hirta , a C 4 grass that exhibits an atypical anatomy where wreaths of so-called distinctive (D) cells are found between V-BS-M-M-BS-V units ( Crookston and Moss, 1973 ; Dengler and Dengler, 1990 ). The D cells carry out the same function as BS cells but are not themselves associated with veins ( Reger and Yates, 1979 ; Dengler et al., 1996 ; Wakayama et al., 2006 ). Notably, if the number of BS and D cells is combined, the 1:1 ratio is also observed in A. hirta .

Transverse Leaf Sections and Corresponding Schematics of C 3 Rice and C 4 Maize.

Rice (left) and maize (right). Bars = 30 μm.

A comparison of vascular development in C 3 and C 4 Flaveria species showed that both the major and minor veins were initiated at comparable stages in development but that a greater number of minor veins were initiated in the C 4 species ( McKown and Dengler, 2009 ). A first step in the evolution of Kranz anatomy may thus have been the acquisition of a mechanism to induce procambium at more regular intervals across the leaf. Given the established role of auxin in vascular development, it is likely that such a mechanism was adapted from existing auxin pathways. A study that compared anatomical and biochemical differences between 16 Flaveria species that encompassed C 3 , C 3 -C 4 intermediate, C 4 -like, and C 4 types further supported the suggestion that altered vein patterning was an early modification in the evolution of C 4 . Based on the phylogeny of Flaveria , it was concluded that C 4 vein pattern traits were acquired prior to either intermediate or C 4 -like biochemistry ( McKown and Dengler, 2007 ). Because the presence of extra veins leads to an effective increase in BS cell area and a decrease in M cell area, it is likely that these traits also preceded biochemical changes.

Metabolic Modifications

Most of the enzymes involved in the C 4 pathway play housekeeping roles in C 3 plants (reviewed in Aubry et al., 2011 ). For example, chloroplast-localized CA ensures a supply of CO 2 into the Calvin-Benson cycle ( Price et al., 1994 ), and PEPCase generates malate as a photosynthetic product ( Ting and Osmond, 1973 ). PEPCase-generated malate is used to provide carbon skeletons to the TCA cycle ( Miyao and Fukayama, 2003 ) and for ammonium assimilation ( Masumoto et al., 2010 ). In addition, PEPCase activity contributes to the extension of fibers in cotton ( Gossypium hirsutum ; Li et al., 2010b ) and to salt and drought responses in wheat ( Triticum aestivum ; González et al., 2003 ) and Arabidopsis thaliana ( Sánchez et al., 2006 ). The decarboxylase PEP-CK has similarly diverse roles in C 3 plants. These roles include mobilization of sugars from lipids in seeds during germination ( Leegood and ap Rees, 1978 ), provision of PEP to the shikimate pathway, and metabolism of nitrogenous compounds ( Walker et al., 1999 ). PPdK-generated PEP has also been shown to contribute to seed metabolism ( Kang et al., 2005 ), the shikimate pathway ( Hibberd and Quick, 2002 ), and nitrogen remobilization ( Lin and Wu, 2004 ). C 4 biochemistry thus evolved through modification of existing functions rather than de novo. This conclusion is supported by the fact that C 3 remains the default developmental state in C 4 plants (discussed above) and that biochemical characteristics of C 4 photosynthesis are found around the vascular bundles of C 3 plant stems ( Hibberd and Quick, 2002 ; Brown et al., 2010 ).

One of the main advantages of the C 4 pathway is a reduction in photorespiration because O 2 cannot effectively compete for the active site of Rubisco in the CO 2 -enriched environment of the BS cells. However, it is a misconception that C 4 plants eliminate the photorespiratory pathway entirely. Maize mutants that are deficient in glycolate oxidase, a key enzyme in the pathway, are seedling lethal at ambient CO 2 ( Zelitch et al., 2009 ). When grown at higher CO 2 levels that inhibit photorespiration, however, the seedlings survive. This suggests that the early stages of the pathway are functional in the mutant and that a buildup of glycolate is toxic for the plant. Most of the photorespiratory pathway is localized to the BS cells of C 4 plants ( Majeran et al., 2005 ), and as a consequence, the released CO 2 further enriches the environment for Rubisco. The use of the photorespiratory pathway as a shuttle to enrich CO 2 in the BS cells is also found in C 3 -C 4 intermediates, where the final step of the pathway is restricted to BS cells (reviewed in Bauwe, 2011 ). This step is catalyzed by Gly decarboxylase and as such it has been proposed that one of the first steps in the evolution of C 4 metabolism was the localization of Gly decarboxylase to the BS ( Sage, 2004 ; Gowik and Westhoff, 2011 ). This would have enriched the BS environment with CO 2 and may have acted as a driver to induce the Calvin-Benson cycle in this cell type.

Metabolite Transport

Increased photosynthetic efficiency in C 4 plants results from the CO 2 -enriched BS cell environment in which Rubisco operates. This environment can only be maintained if the CO 2 that is generated by the BS-localized decarboxylation reaction cannot diffuse back out of the cell. It is generally assumed that the suberized BS cell wall prevents CO 2 leakage. However, the situation cannot be that simple, not least because NAD-ME C 4 species do not have suberized BS cell walls and the different C 4 subtypes carry out the decarboxylation reaction in different sub-cellular compartments. It is thus likely that the diffusion kinetics are also affected by chloroplast and mitochondrial position in the cell and by the distance of the decarboxylation site from the BS-M cell interface ( von Caemmerer and Furbank, 2003 ). A role for porins in CO 2 movement across intracellular membranes has been discussed, but their importance in C 4 plants is far from clear ( Weber and von Caemmerer, 2010 ). Regardless of the exact mechanism, mathematical modeling has shown that the efficiency of the C 4 pathway can only be maintained (through development or in different environmental conditions) if BS cell resistance to leakage increases as the amount of C 4 acid that is decarboxylated decreases (and vice versa) ( von Caemmerer and Furbank, 2003 ). The mechanisms that regulate this dynamic process are far from clear.

Although CO 2 must be prevented from moving between BS and M cells, many metabolic intermediates of the pathway must diffuse between the two cell types and must be actively transported between compartments in individual cells. In C 3 plants, one transport process has to occur across the chloroplast envelope for every three CO 2 molecules assimilated into triose phosphate (TP). By contrast, 30 transport steps are required per TP generated in NADP-ME C 4 plants (reviewed in Weber and von Caemmerer, 2010 ). This difference has implications in terms of the energetic cost of photosynthesis, the establishment of plasmodesmatal connections between the two cell types, and the proteins that had to be modified during C 4 evolution. Until recently, the identities of the transporter proteins were not known, and even now there remain big gaps in our knowledge (reviewed in Majeran and van Wijk, 2009 ). The only two transporter proteins that have been unambiguously identified are the TP transporter, which moves TP from the M chloroplast to M cytoplasm and from the BS cytoplasm to the BS chloroplast, and the PEP/phosphate translocator, which moves PEP from the M chloroplast into the M cytoplasm ( Bräutigam et al., 2008 ). Candidates for the M cell malate/OAA antiporter (dicarboxylate transporter) ( Taniguchi et al., 2004 ; Majeran et al., 2008 ) and for a sodium-dependent pyruvate transporter (bile acid:sodium symporter family protein 2) have also been identified ( Furumoto et al., 2011 ). Other transporters (including all of the BS cell–specific transporters) remain to be identified. Given the quantitative and cell-specific proteomic data available, however, it is presumably only a matter of time before functional assays ( Nozawa et al., 2007 ) of potential candidates ( Bräutigam et al., 2008 ; Majeran et al., 2008 ) provide insight.

Physiological Efficiencies

Although the transition from C 3 to C 4 can be considered at the level of individual genes and proteins (see below), C 4 is in effect a complex trait. In addition to the modified photosynthetic pathway, aspects of nitrogen and sulfur metabolic pathways are also altered or localized in specific cell types ( Friso et al., 2010 ; Bräutigam et al., 2011 ). Key physiological enhancements include greater radiation, nitrogen, and water use efficiencies (RUE, NUE, and WUE) than C 3 plants. For example, measured at 30°C and 380 ppm CO 2 , estimates for the maximum conversion efficiency of solar energy to biomass is 4.6% for C 3 plants and 6% for C 4 plants ( Zhu et al., 2008 ). The relatively higher CO 2 assimilation rates in C 4 plants result from increased efficiency of Rubisco, and this in turn means that only 8% of leaf N needs to be allocated to the enzyme. This contrasts with a >20% allocation to Rubisco in some C 3 plants, leading to a much higher proportion of N required per CO 2 fixed (reviewed in Ghannoum et al., 2011 ). Increased WUE has also been proposed to result from increased CO 2 assimilation rates ( Wong et al., 1985 ), although decreased stomatal conductance has also been implicated ( Taylor et al., 2010 ).

Gene Families

A comparison of photosynthetic gene expression patterns in independently evolved C 4 grass lineages demonstrated that the only patterns common to all origins were an upregulation of PEPCase and a downregulation of Rubisco in M cells ( Sinha and Kellogg, 1996 ). All other gene expression patterns varied between different lineages and different C 4 subtypes. The recruitment of PEPCase into an M cell–specific photosynthetic role was thus a key step in the evolution of the C 4 pathway. PEPCase genes are members of a multigene family that encodes multiple isoforms of the enzyme, only one of which is involved in the C 4 pathway ( Lepiniec et al., 1994 ). Phylogenetic analyses of these gene families in the grasses have shown that the C 4 gene evolved eight independent times from the same non-C 4 gene ( Christin et al., 2007 ). During this transition, 21 amino acids evolved under positive selection and converged to similar or identical amino acids. In some amino acid positions, identical changes have also been recorded in non-grass C 4 species ( Bläsing et al., 2000 ; Gowik et al., 2006 ; Christin et al., 2007 , 2011 ). At some sites, such convergence appears to reflect the need for a specific amino acid for C 4 function, whereas at other sites, there appears to be a requirement for loss of the C 3 -associated amino acid.

In addition to PEPCase, examples of positive selection and gene convergence during the evolution of C 4 have also been reported for genes encoding Rubisco and PEP-CK ( Christin et al., 2008b , 2009b ). In the case of PEP-CK, there is evidence for initial acquisition of the C 4 gene followed by recurrent losses and at least three independent reacquisitions. All of these examples point to gene duplication in C 3 ancestors being a prerequisite for C 4 evolution. Neofunctionalization then presumably occurred either in the context of the C 3 ancestor or, at least in the case of PEP-CK, within the C 4 lineages (for a discussion, see Monson, 2003 ). Support for this evolutionary trajectory has been provided by a comparative analysis of C 3 (rice [ Oryza sativa ]) and C 4 (maize and sorghum) genomes ( Wang et al., 2009 ).

While in some cases, the recruitment to C 4 involved changes in protein function, in other cases, protein targeting mechanisms were altered. For example, there are three genes encoding chloroplast-localized CA in the C 3 species Flaveria pringlei . In the C 4 species Flaveria bidentis , two genes also encode chloroplast-localized proteins, as in F. pringlei , whereas the third has lost the chloroplast-targeting signal, facilitating CA function in the M cell cytoplasm ( Tanz et al., 2009 ).

Cis - and Trans -Regulators of Transcription

Over the last 25 years, considerable effort has been invested into understanding how the cell-specific and light-induced regulation of C 4 enzymes is achieved. These studies have examined the activity of cis - and trans -regulatory factors through the use of biochemical assays, transient expression assays in protoplasts, transgenic manipulation of gene expression in both C 3 plants and C 4 plants, and mutant analysis. Two substantial reviews, written 10 years apart, cover the detailed information for each gene and by comparison illustrate how the field has advanced in recent years ( Sheen, 1999 ; Hibberd and Covshoff, 2010 ). A few key points emerge from a synthesis of the data, and they can be grouped according to level of gene regulation ( Wang et al., 2011 ).

At the epigenetic level, both nucleotide and histone methylation have been associated with the M cell–specific regulation of genes encoding PEPCase ( Ngernprasirtsiri et al., 1989 ; Langdale et al., 1991 ; Offermann et al., 2006 ; Danker et al., 2008 ) and histone methylation with BS cell–specific regulation of NADP-ME ( Danker et al., 2008 ). However, such examples are limited both with respect to the generality across C 4 species and in terms of how epigenetic mechanisms interact with other levels of gene regulation. Information about the epigenetic regulation of other C 4 genes is similarly lacking.

In terms of transcriptional control, cis -regulatory elements that direct M or BS cell–specific expression have been identified for a number of genes (reviewed in Hibberd and Covshoff, 2010 ). In the case of any individual C 4 gene, however, the identified elements differ between C 4 species in terms of both sequence composition and position within the gene (particularly between monocots and eudicots). With few exceptions, these cis -regulators of transcription have yet to be proven sufficient for cell-specific expression. One exception is the 41-bp mesophyll expression module 1 (MEM1) element from the C 4 species Flaveria trinervia ppcA gene promoter. MEM1 is both necessary and sufficient to drive M cell–specific accumulation of ppcA gene transcripts in both C 4 and C 3 Flaveria species ( Gowik et al., 2004 ; Akyildiz et al., 2007 ). Two other exceptions have been reported for genes of the NAD-ME C 4 species Cleome gynandra . The 5′ and 3′ untranslated region sequences from the C. gynandra genes encoding PPdK and CA have been shown to be sufficient for M cell–specific expression in transient assays ( Kajala et al., 2011 ). Interestingly, these sequences are conserved in the orthologous genes of the C 3 species Arabidopsis . This observation suggests that cell specificity in the C 4 species evolved through changes in trans -regulatory mechanisms.

A similar scenario of altered trans -regulators in C 4 species relates to the gene encoding NAD-ME. In this case, a novel mechanism of gene regulation has been revealed. Specifically, a 240-bp sequence of the coding region of the gene encoding NAD-ME, which must be transcribed to be functional, is necessary and sufficient to direct BS cell–specific expression ( Brown et al., 2011 ). As with the PPdK and CA examples discussed above, this sequence is also present in the C 3 orthologs of Arabidopsis , where expression is not cell specific.

Putative trans -regulators of cell-specific gene expression in maize have been identified by gel retardation assays with 5′ promoter sequences of genes encoding PEPCase ( Taniguchi et al., 2000 ), Rubisco small subunit ( Xu et al., 2001 ), and PPdK ( Matsuoka and Numazawa, 1991 ). However, the context in which these proteins act is not understood, and the properties of the proteins are not known. The only known transcription factors that have been proposed to play a role in C 4 regulation are members of the DNA binding with one finger (DoF) and Golden2 -like ( GLK ) gene families. DoF1 is a zinc finger DNA binding protein that was shown to bind to the promoter of the maize PEPC gene and was proposed to play a role in regulating cell-specific gene expression ( Yanagisawa and Sheen, 1998 ). While this may be the case, subsequent analyses showed that DoF proteins also perform a more general role in the transcriptional activation of non-photosynthesis-related genes in maize ( Yanagisawa, 2000 ). Golden2 ( G2 ) is a GARP transcription factor that was initially identified by mutant analysis in maize, where loss of function led to impaired BS cell development ( Hall et al., 1998a ; Rossini et al., 2001 ). The first mutant that was isolated exhibited rudimentary chloroplast development and reduced accumulation of transcripts for C 4 enzymes in the BS cells, leading to the suggestion that G2 was a global regulator of C 4 development in BS cells ( Langdale and Kidner, 1994 ). However, subsequent analysis of an allelic series of g2 mutations determined that the effects on C 4 gene expression were a secondary consequence of perturbed plastid development and thus showed that G2 is not a direct regulator of genes encoding C 4 enzymes ( Cribb et al., 2001 ).

Although G2 is not a direct regulator of C 4 gene expression, it nevertheless functions specifically in maize BS cells, whereas its paralog Zm- Glk1 functions specifically in M cells. By contrast, GLK gene pairs in C 3 plants act redundantly in a single photosynthetic cell type ( Rossini et al., 2001 ; Fitter et al., 2002 ; Yasumura et al., 2005 ). It is now known that in the C 3 plant Arabidopsis , GLK proteins act cell autonomously to directly regulate the expression of a suite of genes encoding chlorophyll biosynthesis enzymes, light harvesting, and electron transport components ( Waters et al., 2008 , 2009 ). As such, they are proposed to synchronize photosynthetic gene expression in response to environmental and developmental cues. This suggestion is supported by the number of pathways in which GLK proteins have been shown to play a role ( Savitch et al., 2007 ; Gutiérrez et al., 2008 ; Yu et al., 2011 ). Importantly, overexpression of GLK1 in the C 3 plant rice leads to the light-induced development of chloroplasts in most cell types ( Nakamura et al., 2009 ). However, a similar response is not seen in Arabidopsis ( Waters et al., 2008 ). As such, GLK proteins are sufficient to induce the proplastid-to-chloroplast transition, but only in certain developmental contexts. Given the cell-autonomous action of GLK proteins and cell-specific accumulation of G2 and Glk1 transcripts in maize ( Rossini et al., 2001 ) and sorghum (unpublished transcriptome data; U. Gowik and P. Westhoff, personal communication), it remains possible that the compartmentalization of GLK function played a critical role in the evolutionary transition to C 4 photosynthetic development.

Posttranscriptional Regulation

Mechanisms that posttranscriptionally regulate gene expression can be divided into those that regulate transcript turnover, translation, or posttranslational activation. Genes encoding the large ( rbcL ) and small ( RbcS ) subunits of Rubisco are regulated at all of these levels. This observation is perhaps not surprising given that Rubisco function in C 4 plants requires the integration of nuclear and chloroplast gene expression programs in addition to BS cell–specific regulation of subunit assembly. The DNAJ-like chaperone BUNDLE SHEATH DEFECTIVE2 (BSD2) has been shown to bind polysome-associated rbcL RNA and is thought to mediate Rubisco assembly and stability in maize ( Brutnell et al., 1999 ). Loss of BSD2 function leads to absence of Rubisco protein and to ectopic accumulation of rbcL transcripts in M cells ( Roth et al., 1996 ). Although cell-specific posttranscriptional mRNA turnover has been implicated for both rbcL and RbcS genes, it is not understood how the failure to assemble Rubisco in BS cells of the bsd2 mutant leads to a failure to repress rbcL transcript accumulation in the M cells. Similarly, there is nothing known about the mechanism of mRNA turnover that operates during normal development. In this regard, it is somewhat surprising that there have been no reports of C 4 gene regulation by noncoding RNAs, given that such RNAs are regulatory components of so many developmental processes ( Vaucheret, 2006 ).

Because research into C 4 photosynthesis was founded in biochemistry, it has been known for many years that posttranslational mechanisms play a key role in the regulation of at least two enzymes of the pathway. PEPCase is posttranslationally and diurnally regulated by the enzyme PEPCase kinase (PEPCk) (not to be confused with PEP-CK, which is the decarboxylase PEP carboxykinase) ( Nimmo et al., 1987 ; Saze et al., 2001 ). The relatively rapid activation and inactivation that is demanded for the diurnal activity of PEPCase is facilitated by the rapid turnover and degradation of PEPCk by the ubiquitin-proteosome pathway ( Agetsuma et al., 2005 ). PPdK is also reversibly light activated by a protein kinase, but in this case, rapid deactivation is facilitated by the same protein. PPdK regulatory protein is a bifunctional Ser/Thr kinase phosphatase that catalyzes both the ADP-dependent inactivation and Pi-dependent activation of PPdK ( Burnell and Hatch, 1985 ; Burnell and Chastain, 2006 ).

Although maize has been a model genetic organism for almost a century, genetic approaches to understand C 4 have yielded limited information. Screens for maize mutants with perturbed vein spacing patterns were unsuccessful (J. Langdale, unpublished data), while those for disrupted BS or M cell development led to the identification of only a handful of examples ( Langdale and Kidner, 1994 ; Roth et al., 1996 ; Hall et al., 1998b ; Covshoff et al., 2008 ). Of those that were characterized in depth, bsd2 and high chlorophyll fluorescence136 were shown to be perturbed in the assembly and/or stabilization of BS (Rubisco) and M (photosystem II) cell–specific proteins, respectively ( Brutnell et al., 1999 ; Covshoff et al., 2008 ), whereas the bsd1 mutant phenotype resulted from loss of G2 transcription factor activity ( Hall et al., 1998a ) (see above). In Panicum maximum , a potential vein spacing mutant was identified in an ethyl methanesulfonate–mutagenized population, but the pleiotropic nature of the phenotype led to lethality and the line was lost ( Fladung, 1994 ).

Other approaches to identify genetic regulators of C 4 include the generation of hybrids between C 3 and C 3 -C 4 Flaveria species and the characterization of oat ( Avena sativa ) lines with single maize chromosomes added. While the Flaveria experiments provided some insight into whether aspects of C 4 were dominant or recessive in F1 hybrids, the sterility of the hybrids precluded quantitative trait loci analysis for C 4 traits ( Brown et al., 1986 , 1993 ; Cameron and Bassett, 1988 ; Holaday et al., 1988 ; Cameron et al., 1989 ). Similarly, the oat-maize addition lines provided insight into certain aspects of C 4 regulation but failed to reveal global regulators of the pathway. In particular, oat-maize addition lines that contained maize chromosomes 6 and 9 were shown to accumulate maize PEPCase and PPdK ( Kowles et al., 2008 ). Notably, both enzymes were active, suggesting that oat PEPCk and PPdK regulatory protein can phosphorylate the maize proteins. However, even in lines with both chromosomes present, photosynthesis was more C 3 -like than C 4 .

The introduction of Setaria viridis as a new model organism for studying the C 4 pathway in monocots provides hope that future genetic analyses will be informative because the plant is relatively small and the generation time is short ( Brutnell et al., 2010 ). This will allow mutant screens to be performed on a much larger scale than has been possible in maize and other C 4 large plants. That said, the dearth of insight thus far provided by genetic approaches may simply be a reflection of the quantitative nature of C 4 traits, and it will be some time before molecular and genetic tools are sufficiently advanced to make substantive progress in S. viridis .

Over the last few years, a number of approaches have been taken to assess C 4 at a systems level. These include proteome comparisons between isolated BS and M cell chloroplasts ( Majeran et al., 2005 ; Friso et al., 2010 ), microarray analysis of BS and M cell transcriptomes ( Sawers et al., 2007 ), transcriptome profiling of mature sugarcane leaves ( Calsa and Figueira, 2007 ), transcriptome and proteome profiling in a single-cell C 4 species ( Park et al., 2010 ) and across a developmental gradient in the maize leaf ( Li et al., 2010a ; Majeran et al., 2010 ), comparative transcriptomics between closely related C 4 and C 3 species ( Bräutigam et al., 2011 ; Gowik et al., 2011 ), and genome-scale models of flux distribution between BS and M cells ( Dal’Molin et al., 2010 ). All of these studies have generated a substantial amount of data, and more is on the way.

For now, we can say that 64% of maize genes are differentially expressed along the developing leaf gradient and that 21% (i.e., 3441 genes) are differentially expressed between BS and M cells ( Li et al., 2010a ). Included in the 21% are members of 180 transcription factor families. Proteomes of a similar developmental gradient elucidate key metabolic and structural transitions along five phases of leaf development (phase 1 being the youngest basal leaf section and phase 5 being the oldest tip section) ( Majeran et al., 2010 ). Three key features emerge from this analysis. First, BS cells (with associated vascular strands) can be isolated from whole-leaf tissue at all phases along the gradient. Second, distinct BS and M cell plastids are observed at phase 2. Third, distinct proteome specialization only becomes apparent in the regions of the leaf that are autotrophic (i.e., phases 3 to 5). In combination, these observations demonstrate that BS and M cell identity is determined early in development and that photosynthetic/metabolic distinctions are mapped onto this anatomical template much later in development.

Transcriptome comparisons between closely related species are harder to analyze for C 4 -specific signatures because of background species differences. However, as more pairwise comparisons are added to the data set, the signal-to-noise ratio will increase. Thus far, a comparison between fully expanded Cleome spinosa (C 3 ) and C. gynandra (C 4 ) leaves has shown that 603 transcripts (2.8% of those identified) are more abundant in C 4 leaves ( Bräutigam et al., 2011 ). These include genes encoding transport proteins, putative plasmodesmata-related proteins, cell wall–modifying enzymes, and 17 transcription factors. At a pathway level, the C 4 species had lower levels of transcripts associated with one-carbon metabolism, the shikimate pathway, amino acid metabolism, the Calvin-Benson cycle, photorespiration, and protein synthesis (both cytoplasmic and plastidic). By contrast, starch metabolism, cofactor synthesis, and nitrogen metabolism–associated transcripts were elevated in the C 4 species. Similar observations were made when comparing five Flaveria species with C 3 , intermediate, or NADP-ME C 4 photosynthesis ( Gowik et al., 2011 ). In this study, the authors placed an upper limit of 3582 expression changes required for the transition to C 4 . Of course the key will be to determine which of those changes are necessary and sufficient for the transition.

The renewed interest in C 4 biology results from increased global awareness of the difficulty we face in trying to provide food and fuel for a growing population. One way to increase yields while simultaneously improving WUE and NUE could be to introduce C 4 traits into C 3 crops. This idea was first proposed in the late 1990s when transgenic experiments to understand C 4 gene function were initiated (reviewed in Matsuoka et al., 2001 ) and Japan Tobacco was granted a U.S. patent on the generation of PEP-CK type C 4 cycle in rice ( Arai et al., 2003 ).

One of the most promising reports at the time showed that introducing the intact maize Ppc gene into rice led to high levels of transgene expression, PEPCase enzyme activity two- to threefold of that found in maize, and reduced O 2 inhibition of photosynthesis ( Ku et al., 1999 ). Introduction of the maize PPdk gene also produced increases in enzyme activity (as much as 40-fold in some lines) and that activity was light/dark regulated as normal ( Fukayama et al., 2001 ). Similarly, introduction of the sorghum NADP-ME gene led to elevated transcript and protein levels and a 1.7-fold increase in enzyme activity ( Chi et al., 2004 ). However, neither the PPdK nor NADP-ME transgenics showed changes in carbon assimilation, and in the case of the PEPCase transgenics, subsequent reports went on to show that the reduced O 2 inhibition was due to reduced rates of photosynthesis. This reduction was in part because of Pi limitation ( Agarie et al., 2002 ) but also because the enzyme was phosphorylated in the dark (in the same way as the endogenous rice enzyme) instead of in the light (like the maize enzyme) ( Fukayama et al., 2003 ). These findings highlighted the complexity of trying to alter the activity of just one enzyme, and when subsets of the different transgenes were combined, the picture became even more complicated ( Taniguchi et al., 2008 ). In no case was a CO 2 concentrating mechanism generated, and in the case of PEPCase and NADP-ME, overexpression led to stunted growth that was only slightly mitigated by overexpression of NADP-malate dehydrogenase.

So why would the more recently formed C 4 rice consortium ( http://irri.org/c4rice ) and its funders, The Bill and Melinda Gates Foundation, once again consider introducing C 4 traits into rice? The rationale is straightforward: C 4 plants have higher RUE than C 3 plants, and yield increases in C 3 cereal crops are becoming limited by RUE ( Hibberd et al., 2008 ). In addition, technology has improved significantly over the last few years. Phenotypes can now be assessed at the whole-plant level ( Furbank et al., 2009 ), gene interactions can be diagnosed at a systems level ( Zhu et al., 2010 ; Wang et al., 2011 ), and mutated genes associated with specific phenotypes can be identified through whole-genome sequencing. Even with these advances, however, the project remains a grand challenge.

Another driver of the current C 4 research agenda is the global focus on biofuels. Two of the current major biofuel crops, sugarcane and maize, are both C 4 species. Whereas the future of sugarcane as a fuel crop is almost certain, the use of maize can only be defended in a future where lignocellulosic fermentation means that grain is not used to produce ethanol. However, another C 4 species may hold the key to biofuel demands, at least in the US. The perennial grass Miscanthus × giganteus is capable of producing higher biomass than maize, primarily because it can photosynthesize efficiently for a longer period during the growing season. This increased efficiency is achieved in two ways. First, Miscanthus can photosynthesize at cooler temperatures than maize as a consequence of cold-tolerant PPdK activity ( Wang et al., 2008 ). Second, its perennial habit means that it is able to capture more light early in the season because at that time the canopy is bigger than that of annual crops, such as maize ( Dohleman and Long, 2009 ). Current estimates suggest that 9.7 million hectares of Miscanthus would provide enough biomass to meet the annual U.S. energy mandate ( Somerville et al., 2010 ). Given that long-term field trials have shown that Miscanthus yields highly even on poor soils and that 14 million hectares of land dropped out of agricultural use in the US between 1997 and 2007 ( http://www.ers.usda.gov/statefacts/us.htm ), this C 4 perennial could resolve the food versus fuel dilemma in the US for the foreseeable future.

I thank all of the postdocs and students who have worked with me over the years on the chloroplast project. For many of them, their time came when the field was very unfashionable, but they shared my passion. I apologize to all C 4 researchers whose work has not been cited here; in providing a general overview for non-C 4 aficionados, I have had to be selective, and in some cases, that selection was a randomly chosen example. I thank all of my colleagues in the C 4 rice consortium and the EU 3to4 Project for sharing their vision, ideas, and data. In particular, I thank Jim Fouracre, Peng Wang, Mark Waters, and Julian Hibberd for comments on the manuscript. In the past, my C 4 research has been funded by the Biotechnology and Biological Science Research Council and The Gatsby Charitable Foundation. It is now funded by the Bill and Melinda Gates Foundation, EUFP7, and the Oxford Martin School.

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How Light Reactions of Photosynthesis in C4 Plants Are Optimized and Protected under High Light Conditions

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Most C4 plants that naturally occur in tropical or subtropical climates, in high light environments, had to evolve a series of adaptations of photosynthesis that allowed them to grow under these conditions. In this review, we summarize mechanisms that ensure the balancing of energy distribution, counteract photoinhibition, and allow the dissipation of excess light energy. They secure effective electron transport in light reactions of photosynthesis, which will lead to the production of NADPH and ATP. Furthermore, a higher content of the cyclic electron transport components and an increase in ATP production are observed, which is necessary for the metabolism of C4 for effective assimilation of CO 2 . Most of the data are provided by studies of the genus Flaveria , where species belonging to different metabolic subtypes and intermediate forms between C3 and C4 are present. All described mechanisms that function in mesophyll and bundle sheath chloroplasts, into which photosynthetic reactions are divided, may differ in metabolic subtypes as a result of the different organization of thylakoid membranes, as well as the different demand for ATP and NADPH. This indicates that C4 plants have plasticity in the utilization of pathways in which efficient use and dissipation of excitation energy are realized.

1. Diversity of C4 Photosynthesis

Photosynthesis is a complex metabolic process in which solar energy is utilized to convert atmospheric carbon dioxide (CO 2 ) into organic compounds. Traditionally, it is divided into two phases: (1) the light energy captured by the pigment–protein complexes is exchanged into energy-rich bonds of molecules such as ATP and NADPH, and (2) when ATP and NADPH drive the fixation of CO 2 in carbohydrates. Approximately 85% of all higher plants use the C3 photosynthetic pathway [ 1 ]. In this process, the first step of CO 2 fixation is catalyzed by ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) and two three-carbon molecules (3-phosphoglyceric acid) are produced by carboxylation of ribulose-1,5-bisphosphate. However, RuBisCO also shows oxygenase activity, which leads to carbon loss by photorespiration. Plants have evolved mechanisms that allow them to concentrate CO 2 at the RuBisCO site and thus eliminate or reduce the oxygenase activity of RuBisCO. One of them is C4 photosynthesis, a pathway described in the 1960s [ 2 , 3 , 4 ]. In this type of photosynthesis ( Figure 1 A), the assimilation and reduction of CO 2 are spatially separated and catalyzed by two different enzymes. Carbon dioxide assimilation occurs in the cytoplasm of mesophyll (M) cells, where CO 2 is initially converted to HCO 3 − , which is then incorporated into phosphoenolpyruvate (PEP) to form oxaloacetate (OAA) by phosphoenolpyruvate carboxylase (PEPC). Then, OAA is converted to malate or aspartate, and these carboxylic acids are transported to bundle sheath (BS) cells, where they are decarboxylated. The CO 2 released is then assimilated by RuBisCO (present only in the chloroplasts of BS cells) through the Calvin–Benson cycle [ 5 ]. Due to differences in a type of carboxylic acid transferred to BS cells, and the main enzyme responsible for the decarboxylation and regeneration of the CO 2 acceptor in M cells, three biochemical subtypes of C4 photosynthesis are recognized: NADP-ME, NAD-ME, and PEPCK or PCK [ 5 , 6 , 7 ]. In the most common NADP-ME subtype, malate is transferred from the mesophyll to bundle sheath cells and CO 2 is released into the chloroplasts of BS by the NADP-dependent malic enzyme (NADP-ME). However, in the NAD-ME subtype, aspartate is the main carboxylic acid that is transported from M to BS cells, and decarboxylation is catalyzed by mitochondrial NAD-dependent malic enzyme (NAD-ME). In the third PEPCK subtype, both aspartate and malate are transferred to BS cells, where aspartate is converted to OAA in the cytosol. Then, it is decarboxylated by PEP carboxykinase (PEPCK or PCK). Moreover, malate synthesized in the mesophyll chloroplast is transported into BS mitochondria, where decarboxylation catalyzed by NAD-ME occurs. Furthermore, although C4 photosynthesis has been classified into three subtypes, accumulating evidence indicates that many C4 plants use a combination of organic acids and decarboxylases during CO 2 fixation, and the C4-type categories are not rigid [ 8 ]. The C4 metabolic cycle consumes more energy than C3 photosynthesis. Fixation of 1 CO 2 requires 5 ATP and 2 NADPH for plants driving the NADP-ME and NAD-ME photosynthesis subtypes and 3.6 ATP and 2.3 NADPH for the PEPCK subtype ( Figure 1 B), whereas for the C3 cycle this energy demand is 3 ATP and 2 NADPH [ 9 , 10 ].

( A ) Schematic describing the general metabolic pathway of C4 photosynthesis. ( B ) The requirement for ATP and NADPH per fixed CO 2 in mesophyll and bundle sheath cells of plants representing the NADP-ME, NAD-ME, and PEPCK subtypes of C4 photosynthesis.ATP and NADPH requirements estimated by Edwards and Voznesenskaya [ 9 ]. CA: carbonic anhydrase; Chl: chloroplast; DC: decarboxsylase; OAA: oxaloacetate; PCR cycle: photosynthetic carbon reduction cycle; PEP: phosphoenolpyruvate; PEPC: phosphoenolpyruvate carboxylase; PPDK: pyruvate, phosphate dikinase; RuBisCO: ribulose-1,5-bisphosphate carboxylase/oxygenase.

Separation of assimilation and CO 2 reduction into two cell types involves several adaptations in the anatomy and ultrastructure of the leaf [ 5 ]. This special leaf anatomy in the C4 plant is called Kranz and characterizes thick-walled (containing suberin) large parenchyma cells (BS cells) that tightly surround vascular bundles, while mesophyll cells, with much thinner walls, are located between the leaf epidermis and BS cells [ 11 ]. In addition, mesophyll and BS cells are connected by a dense network of plasmodesmata through which an intensive transport of metabolites takes place [ 5 ]. Furthermore, the M and BS chloroplasts in species belonging to different subtypes of C4 photosynthesis exhibit structural dimorphism. In the NADP-ME subtype, chloroplasts localized in M cells have grana, whereas BS chloroplasts are deficient in these structures or grana stacks are rare [ 12 , 13 ]. In contrast, in the NAD-ME type, mesophyll chloroplasts are more deficient in grana than the BS chloroplast [ 14 , 15 ]. In the PEPCK subtype, M and BS chloroplasts demonstrate a similar pattern of granal development [ 13 , 16 ]. This dimorphism is related to differences in the need for NADPH and ATP between the chloroplasts of the mesophyll and bundle sheath cells to support C4 photosynthesis [ 9 ].

This spatial separation of CO 2 assimilation and reduction allows for up to a tenfold increase in the CO 2 concentration in BS cells compared to the natural concentration of this gas in the air [ 17 ]. The maintenance of the high concentration of CO 2 at the RuBisCO site is also related in part to the lack or low levels of carbonic anhydrase. This prevents a rapid conversion of carbon dioxide into a carbonate anion, which is not the substrate for RuBisCO [ 18 ]. The mechanism of CO 2 concentration, the higher CO 2 assimilation capacity, and the reduction of stomatal conductance help C4 plants maintain higher rates of carbon gain compared to C3 plants [ 19 , 20 ], 50–300% higher water use efficiency [ 20 ], and higher nitrogen use efficiency [ 21 , 22 ]. These advantages allow C4 plants to grow at high light intensities, high temperatures, and under arid conditions, where the photorespiration process can reduce C3 photosynthesis by up to 30% [ 23 ].

Photosynthesis of C4 is a relatively recent innovation that has evolved independently more than sixty times during the last 30 million years of land plant evolution [ 24 ]. The significant differences in anatomy, biochemistry, and physiology between different C4 species are based on various genetic changes that include simple modifications of the molecular sequence of genes involved in photosynthesis, alterations of regulatory elements, gene duplications, or subcellular retargeting of proteins, and lateral transfer of genetic material. A good example of simple molecular sequence modification is PEP carboxylase, in which 21 different amino acid positions have been elucidated in grasses [ 25 ]. Larger changes in DNA sequences are also frequent and include duplication of entire genomes or genes involved in the C4 cycle [ 25 , 26 , 27 ]. Interestingly, it is currently thought that the duplication of genes encoding proteins of the core C4 cycle generally occurred before the origin of C4 photosynthesis and this may have been a contributing factor that allowed the evolution of C4 plants through the acquisition of new functions by duplicated genes. However, genetic modifications related to the evolution of C4 photosynthesis are not fully understood and are still intensively studied.

2. Ways of Light Energy Utilization: Balanced Distribution between Photosystems and Emission of Excess Energy as a Heat

Environmental factors, especially light intensity and quality during growth, can cause changes in pigment–protein complexes involved in the light reactions of photosynthesis. In some cases, light can cause photoinhibition, which is associated with the overproduction of reactive oxygen species (ROS) and damage of thylakoid membrane components [ 28 ]. The plant response may be more complex when there are several environmental factors, which in the case of C4 plants is quite natural due to their ability to grow in hot, and dry conditions with high light intensity [ 29 , 30 ].

The light energy that has been absorbed by chlorophyll molecules can be used in three ways: it can be used for photochemical reactions, its excess can be dissipated as heat, or it can be transferred as light, i.e., fluorescence ( Figure 2 ). These processes are interdependent, so limiting photochemical reactions will result in an increase in the other two parameters [ 31 ]. As can be seen in the figure below, only a small part of the absorbed light energy is used for the course of photochemical reactions. Most of it is dissipated as heat and only 3–5% of the absorbed energy is emitted by fluorescence, but these values may change depending on environmental conditions and stress factors, including different intensities or qualities of light. Measurements of chlorophyll a fluorescence are commonly performed to investigate the influence of environmental factors on the photosynthetic apparatus. They provide a lot of valuable information on how much light energy is used in photochemical reactions and how much is lost (dissipated) as heat.

The scheme presents the use of absorbed light energy for photochemical reactions, heat dissipation, and radiation by fluorescence. PAR: photosynthetically active radiation.

In this review, we focus mainly on the mechanisms that allow the adjustment of light reactions of photosynthesis to changing light conditions, including high light intensities. Unfortunately, the response of C4 photosynthetic apparatus to changing light intensities has been barely studied at the level of gene expression, and thus this review focused mainly on the role of post-translational modifications.

Changes in light intensity under natural conditions occur rapidly, and the amount of light absorbed often exceeds the efficiency of photosynthetic reactions. It should be noted that photosynthetic light reactions play a crucial role due to the products of linear (LET) and cyclic electron transport (CET), that is, the supply of ATP and NADPH, necessary for the assimilation of CO 2 in the Calvin cycle [ 32 ]. Mechanisms that function as safety valves in chloroplasts can be divided into those universal and concern not only the protection of C4 plants, but also C3, such as the xanthophyll cycle, heat dissipation, or state transitions, and those more characteristic for C4, such as promoting cyclic electron transport and alternative CET pathways, routes to increase ATP production [ 32 ]. It should be noted that in C4 plants the response to environmental factors can be more complicated due to the existence of different metabolic subtypes (NADP-ME, NAD-ME, PEPCK), different structures of M and BS chloroplasts within these subtypes, and different requirements for ATP and NADPH.

2.1. Elevation of Cyclic Electron Transport Components

C4 plants have higher cyclic electron transport activity compared to C3, and this increase may have been most important for the production of the additional ATP pool required for C4 metabolism. Studies on the genus Flaveria showed an increase in the content of cyclic electron transport components, such as PGR5 (proton gradient regulation 5), PGRL1 (proton gradient regulation like 1) proteins, and the NDH (NADH dehydrogenase-like) complex in the chloroplast of BS cells during the evolution of C4 photosynthesis ( Figure 3 and Figure 4 ). Furthermore, the reduction in the amount of grana or the presence of agranal thylakoid organization, as in the maize BS chloroplasts, was associated with a higher content of components such as PGR5, PGRL1, and NDH. In C4 plants, two extra ATP molecules are required to assimilate each CO 2 molecule in the Calvin cycle. As mentioned above ( Figure 1 ), in the NADP-ME subtype, the demand for ATP is higher in chloroplasts of BS than in mesophyll chloroplasts. It is indicated that additional ATP molecules are produced by the cyclic electron transport, which dominates in this type of chloroplast and ensures the generation of a pH gradient across the membrane thylakoid without NADPH production. Moreover, a reduction in the number of grana is associated with a lower PSII content and activity, and this down-regulation of PSII and the promotion of cyclic electron transport are related to C4 metabolism. The higher activity and content of the cyclic electron transport components were confirmed, among others, by electrophoretic methods, immunodetection, fluorescence, as well as thermoluminescence [ 33 ].

The scheme of alterations that take place in bundle sheath chloroplasts (BS) during the evolutionary process from C3 to C4 in the Flaveria genus. As described by Munekage and Taniguchi [ 38 ], in species representing C4 photosynthesis, the content of complexes involved in cyclic electron transport increased and the allocation of LHCII trimers as the PSI antenna occurred due to changes in the organization of thylakoid membranes.

The ways of electron transport in the thylakoid membrane under high light intensities and over-reduction of the plastoquinone pool (PQ). The scheme shows linear electron transport (LET), cyclic electron transport (CET), which takes place with the participation of the proteins PGR5 and PGRL1 or/and the NDH complex, and the activity of PTOX in the chlororespiration process. The content of complexes and the activity of the electron transport pathway will differ in the M and BS chloroplasts in particular metabolic subtypes and depending on environmental conditions.

There are two key pathways in the cyclic electron transport in plants ( Figure 3 ). One of them is classic transport with the participation of PGR5-PGRL1 proteins, the other is an alternative route involving the components of the NDH complex. The PGR5–PGRL1 complex, together with the cytochrome b 6 f complex, is involved in electron transport from ferredoxin to plastoquinone (PQ) [ 34 ]. The NDH complex, which consists of at least 29 subunits, also donates electrons from ferredoxin to plastoquinone. Using different electrophoretic methods, Darie et al. [ 35 ] showed that the molecular mass of the NDH complex is 550 kDa, in both M and BS maize chloroplasts. Additionally, it may exist in monomeric (550 kDa) or dimeric form (1000–1100 kDa). Moreover, they indicate that if the structure of the NDH complex is the same in two different types of maize chloroplasts, it will be very similar in all other cases [ 35 ], and only its content in thylakoid membranes may be different.

An increase in the number of individual subunits of the NDH complex is observed in BS chloroplasts of NADP-ME C4 plants, including Zea mays , Sorghum bicolor, and Flaveria C4 species [ 33 , 36 , 37 , 38 ] ( Figure 4 ). In C4 plants, the content of the NDH complex has been shown to be four times higher in the M chloroplasts and even fourteen times higher in the BS chloroplasts compared to C3 plants [ 33 ]. Furthermore, in C3 plants, the amount of NDH complex is low and can consist of approximately 4% of total PSI, whereas in BS chloroplasts of C4 plants it can even be 40% [ 39 ]. In maize, a higher content of such subunits as NdhH, NdhK, NdhJ, and NdhE was observed, and the proportion of the NDH complex in BS and M chloroplasts was 3:1 [ 35 ].

When comparing various species representing different subtypes of C4 photosynthesis, a higher NDH content was observed in M chloroplasts in the NAD-ME subtype and in BS chloroplasts in the NADP-ME subtype, i.e., in these chloroplasts that have a higher demand for ATP. This confirms the hypothesis that NDH-dependent cyclic electron transfer plays a key role in providing the ATP needed to drive the CO 2 concentrating mechanism [ 39 ]. It is speculated that the increase in the concentration of the NDH complex was caused by the accumulation of NADPH, resulting from a higher need for ATP than NADPH during C4 photosynthesis. In the NADP-ME subtype, NADPH is released during decarboxylation by the NADP-malic enzyme, leading to an increase in its amount in the BS chloroplast [ 33 ]. Furthermore, it has been shown that the NDH complex can participate in the dark reduction of the plastoquinone pool. Reduced plastoquinone is a signal that activates Stn7 kinase, which phosphorylates LHCII antenna systems. Presumably, the NDH complex and its functioning can maintain a certain pool of antenna phosphorylated under all conditions, allowing an even distribution of incoming light energy, efficient cyclic transport of electrons, and ATP production [ 40 ], as it occurs in chloroplasts of the maize bundle sheath cells with permanent state 2 (explained below) [ 41 ].

ATP synthase, which is responsible for the synthesis of ATP, has a localization similar to that of the complexes involved in the cyclic electron transport, in the marginal regions and stroma lamellae. The elevation in the content of the CET components is correlated with the amount of ATP synthase, especially under conditions of high light intensities, as demonstrated by Romanowska et al. [ 42 ]. The authors investigated the effect of light on the level of the CF1-α subunit in the chloroplast of M and BS cells in maize ( Zea mays , type NADP-ME), millet ( Panicum miliaceum , type NAD-ME), and guinea grass ( Megathyrsus maximus , type PEPCK). They showed that light intensity had an impact on the amount of α subunit of ATP synthase, and the accumulation of additional isoform ά might have a protective role for C4 plants under high light to prevent the thylakoid lumen from overprotonation and from PSII photooxidative damage [ 42 ].

2.2. Function of PTOX Protein and Chlororespiration

In addition to transmembrane complexes, such as PSI, cytochrome b 6 f , or PSII, there are also additional components that are important in electron transport. These include the NADH chloroplast dehydrogenase complex (described above) and the plastid terminal oxidase (PTOX) [ 43 ] ( Figure 3 ). The approximately 40 kDa PTOX protein is similar to mitochondrial alternative oxidase (AOX) and is a plastoquinone:O 2 oxidoreductase, commonly found in plants. It is located on the stromal side of the thylakoid membrane between PSII and the cytochrome b 6 f complex [ 43 ]. It oxidizes the plastoquinone pool and, by donating electrons to oxygen, leads to the formation of a water molecule. This protects the plastoquinone against excessive reduction [ 44 ], for example, under conditions where the consumption of the reduced form is limited by the dark reactions of photosynthesis. PTOX accounts for 1% of the PSII content in Arabidopsis thaliana , and it is located in the non-appressed regions of thylakoids where PSI is dominant [ 45 ].

Exposing plants to high light intensities leads to an excessive reduction of the plastoquinone pool, which can cause photoinhibition. PTOX content is indicated to increase under environmental stress conditions, as shown in the C3 Ranunculus glacialis plant [ 46 ] exposed to full sunlight. At high light intensity, when the pH of the stroma increases, PTOX can bind to the membrane and access the substrate plastoquinol, but under nonsaturating light, the protein detaches from the membrane and is inactive [ 44 ].

It is noted that maize (NADP-ME subtype) has more PTOX in BS chloroplasts. This may be due to the agranal structure of chloroplasts and the presence of non-appressed regions. Moreover, the RuBisCO enzyme is present only in BS chloroplasts. The presence of PTOX can reduce oxygen concentration at the RuBisCO site and increase the efficiency of CO 2 assimilation in the Calvin cycle [ 47 ]. This observation could indicate that in all metabolic subtypes of C4 plants, a higher PTOX content should be expected in chloroplasts of BS cells due to the exclusion of the oxygenase activity of RuBisCO.

There is little data on PTOX activity in individual C4 subtypes in response to high light intensities. This protein has been shown to be involved in tolerance to salt stress in a representative of PEPCK Spartina alterniflora , which is a halophyte. PTOX may provide an alternative pathway for the protection of this species against over-reduction and minimize or avoid damage to both photosystems (PSI and PSII) [ 48 ]. The authors suggest that Spartina alterniflora gained increased salt tolerance because of increased electron flow through PTOX, which can be a major sink of electrons in salt stress and functions as a safety valve to protect photosystems from over-reduction, in contrast to the response observed in the glycophyte Setaria viridis (NADP-ME). These experiments were carried out on whole leaves, so they do not provide information on the content and activity of PTOX individually in the M and BS chloroplasts.

The final product in the reaction catalyzed by the PTOX is water. Due to the close location of the NDH complex and the PTOX protein in the thylakoid stroma membranes, they participate in a pathway called chlororespiration, which is important in protecting PSI from over-reduction, protecting against reactive oxygen species, and preventing PSI photoinhibition [ 49 ].

2.3. Changes in the Amount of Thylakoid Complexes and Rearrangement of Super- and Megacomplexes

The LHCII antenna system is one of the complexes in thylakoid membranes whose content changes under conditions of varying light intensities [ 50 ]. Adaptation to high light intensities in maize is a tightly coordinated regulation of the components/activity of the light reaction in both mesophyll and bundle sheath chloroplasts [ 51 ]. Under different light intensities, both the content of individual proteins and the arrangement of the complexes can change. Low light intensity promotes the development of antenna systems to capture as much light energy as possible, in contrast to high light when LHC systems decrease. When the intensity of light increases during growth, the levels of the PSII and PSI reaction centers, as well as the cytochrome b 6 f complex, increase [ 51 ]. It has also been shown that the content of LHCII antenna systems increases under conditions of low light, even in maize BS chloroplasts where the PSII content is reduced.

Light, both its quantity and quality, as well as other environmental factors (e.g., temperature, CO 2 and O 2 concentration, drought, and also phosphate availability) affect the expression of chloroplast genes, which is dependent on the redox state of the chloroplasts. The PQH 2 /PQ ratio influences the transcription genes that encode the proteins of the PSI and PSII reaction centers [ 52 ], allowing the photosynthetic apparatus to be adjusted to the actual conditions. Maintaining the oxidized pool of plastoquinone (PQ) by exposure to PSI excitation light or by inhibiting electron transport in PSII was found to activate transcription of the psbA gene encoding D1—the core protein of PSII. In contrast, when the pool of plastoquinone is in a reduced state (PQH 2 ), during exposure to PSII excitation light, the transcription of the psaA and psaB genes encoding the PSI reaction center proteins is activated. Variable light conditions also influence the rate of the Calvin cycle. Therefore, if the demand for ATP and NADPH and their production in light reactions of photosynthesis changes, the degree of reduction of the plastoquinone pool will also change. This has an impact on the expression of genes encoding core proteins of the photosystems, and this, in turn, may change the proportion of cyclic or linear electron transport under given conditions.

In acclimation to changing light conditions, the organization of complexes in thylakoid membranes is also important. Urban et al. [ 53 ] reported the presence of PSI–LHCI–LHCII–Lhcb4 supercomplexes and PSI–LHCI–PSII–LHCII megacomplexes in the stroma lamellae and grana margins of maize mesophyll chloroplasts. These complexes contained various LHCII trimers and monomers in combination with PSI. They were formed under both low light and high light conditions, but their exact composition differed. It was shown that exposure of plants growing in low light intensity to far-red (FR) light caused dissociation of the PSI-LHCI–LHCII–Lhcb4 supercomplex into free LHCII–Lhcb4 and PSI–LHCI complexes and which then associated with the PSII monomer. The process was different in plants grown in high light. Exposure to FR light causes dissociation of both PSI–LHCI–LHCII–Lhcb4 supercomplexes and PSI–PSII megacomplexes. These results suggest that the reorganization of the super- and megacomplex has a different function than balancing light absorption between the two photosystems under light stress. Such changes may have an influence on energy quenching and the PSII turnover cycle [ 53 ]. These studies were innovative and data on the formation of these complexes are not available for maize BS chloroplasts and for the chloroplast of other C4 subtypes.

2.4. Photoinhibition and Role of D1 Protein Phosphorylation

Photoinhibition is a phenomenon that leads to a decrease in photosynthetic activity and a reduction in CO 2 assimilation. It is defined as the light-induced inhibition of photosystem II activity [ 54 ] when photosystem II degradation dominates over its repair [ 55 ]. The classic model of photoinhibition assumed the generation of reactive oxygen species by excessive reduction of the plastoquinone pool. The formed reactive oxygen species are responsible for the damage to the PSII reaction center. Currently, many authors indicate that PSII repair processes are more sensitive to environmental stresses [ 28 , 56 ]. Photosystem II is considered to be the most damage-sensitive complex of the thylakoid membrane, which does not mean that PSI is not affected by photoinhibition as well. PSI photoinhibition occurs when the supply of electrons from PSII exceeds the acceptor capacity of PSI [ 57 ], but PSI is effectively protected against damage, for example, by photoinhibition of a certain pool of PSII. Tikkanen et al. [ 58 ] showed that PSII photoinhibition, which reduces electron transport, prevents ROS formation and PSI damage. Furthermore, Ballotari et al. [ 59 ] observed that zeaxanthin can bind to PSI and participate in the dissipation of excess energy from PSI. It should be emphasized that the state transition process described later is also the one that protects photosystems from overexcitation. Furthermore, the results of Lima-Melo et al. [ 60 ] suggest that rapid activation of PSI photoinhibition under severe photosynthetic imbalance protects the chloroplast from over-reduction and excess ROS formation.

Repair of damaged PSII reaction centers requires the degradation of the D1 protein destroyed during photoinhibition, its de novo synthesis, and reconstruction of the PSII complex. D1 degradation is a multistage process regulated by protein phosphorylation and dephosphorylation, and also by the level of ATP in chloroplasts. The D1 protein, one of the most easily degraded, is phosphorylated under the influence of medium and high intensity of light in the granum of thylakoids by membrane-bound serine threonine kinase. This modification affects both intact and damaged reaction centers, protects against proteolytic degradation, and has no effect on the electron transport rate in PSII [ 61 ]. When irreversible damage of D1 is caused by photoinhibition, this D1 is directed to the thylakoid stroma, where it is dephosphorylated and then degraded [ 62 ].

Pokorska et al. [ 61 ] showed that the rate of degradation of the D1 protein in maize (NADP-ME plant) BS chloroplasts during photoinhibition is faster compared to mesophyll chloroplasts, and the content of Deg1 protease, which is one of the enzymes that degrades the D1 protein, was higher. Furthermore, individual steps of D1 turnover such as dephosphorylation, degradation, and de novo synthesis of PSII subunits are known to take place in stroma exposed regions, and enzymes responsible for D1 proteolysis are also present there [ 63 ]. The organization of thylakoid membranes as an agranal system in maize BS chloroplasts makes processes related to the D1 protein turnover cycle much faster.

There is no data in the literature on photoinhibition in other metabolic subtypes of C4 plants. Differences in the metabolic reaction and structure between mesophyll and bundle sheath chloroplasts may have an influence on the rate of D1 turnover.

2.5. State Transitions and Phosphorylation of LHCII

At different light intensities, the migration of LHCII between photosystems is observed in the process called state transitions. The LHCII antenna, and especially the Lhcb2 protein, undergoes reversible phosphorylation, which is crucial for the switching of LHCII between photosystems. The levels of LHCII phosphorylation are lower at high light compared with those under low light conditions. State 1 is traditionally defined as the condition when PSI is preferentially excited and all LHCIIs become associated with PSII. Illumination conditions, which lead to an excess excitation of photosystem II (PSII), compared to photosystem I (PSI), induce a transition to state 2, in which the more absorbed excitation energy is diverted to PSI because the phosphorylated LHCII antennas are associated with PSI [ 64 ]. State transitions act as a mechanism to balance the excitation of the two photosystems under changing light regimes [ 41 ].

The state transition has been described mainly in C3 plants, including A. thaliana , in which this process has been studied using mutants, e.g., in the stn7 gene, which encodes the key enzyme, and STN7 kinase, which phosphorylates LHCII. Little is known about this process in C4 plants. In C4 plants where there are differences in the organization of thylakoid membranes in the M and BS chloroplasts, the process may be quite different. Thylakoid membranes are heterogeneous, and while PSII with the LHCII antenna is located in the stacked regions of the grana, the PSI occurs in the stroma lamellae and marginal grana regions. Thus, the number of grana in a given chloroplast type in each metabolic subtype will determine the LHCII content and the amount of PSI to which these antennas can potentially be attached.

Nakamura et al. [ 33 ] showed that with changes in the Flaveria genus, which led to the formation of C4 species, such as Flaveria trinervia (NADP-ME), increased mobility within the thylakoid membranes and increased LHCII functioning as a PSI antenna was observed in the chloroplasts of BS cells ( Figure 4 ). It has also been shown that in maize belonging to the NADP-ME subtype, a different regulation of antenna switching is present in the two types of chloroplasts. In M chloroplasts, the state transition occurs in a classical way, and depending on the light conditions, the LHCII systems are attached to PSII or PSI. In BS chloroplasts, a permanent state 2 was observed ( Figure 5 ), in which a certain pool of LHCII antennas remains attached to the PSI, regardless of light conditions, even at high intensities of far red, which preferentially excites PSI [ 41 ]. This can be crucial in regulating cyclic electron transport and making it work more efficiently, allowing the production of higher amounts of ATP. It should be remembered that ATP drives the energy for CO 2 assimilation in the Calvin cycle, which operates only in BS chloroplasts.

Models of state 1 state 2 transitions in two types of chloroplasts in three subtypes of C4 plants. In maize (NADP-ME) mesophyll chloroplasts, the typical transition from state 1 to state 2 occurs, while in bundle sheath chloroplasts it is permanent state 2 where some pool of LHCII antennas are bound to PSI [ 41 ]. For the NAD-ME and PEPCK subtypes, in which the regulation of antenna migration is unknown, the shown model of state transitions is based on the organization of thylakoid membranes and the demand for ATP.

The regulation of antenna phosphorylation in the remaining C4 subtypes may be quite different due to the organization of the thylakoid membrane in the chloroplasts and the demand for ATP. Based on this demand and the organization of the thylakoid membranes in various types of chloroplasts, we propose a possible state transition scheme ( Figure 5 ).

2.6. Xanthophyll Cycle and Heat Dissipation

Among several mechanisms in chloroplasts that allow them to function under stress conditions, preventing the generation of ROS is extremely important. One of the protection mechanisms is the quenching of excess energy as a thermal dissipation. This process involves the xanthophyll cycle, related to the conversion of violaxanthin to zeaxanthin [ 65 ] and the protonation of the PsbS protein. Generally, when plants are exposed to high light intensities, violaxanthin is oxidized by violaxanthin de-epoxidase (VDE). This leads to the formation of antheraxanthin, followed by zeaxanthin. Zeaxanthin creates a barrier that prevents overexcitation of the PSII reaction center. The energy from LHCII is dissipated and is not directed to the reaction center. At low light intensities, zeaxanthin epoxidation catalyzed by zeaxanthin epoxidase (ZE) occurs. The content of carotenoids, including xanthophylls participating in the xanthophyll cycle, in M and BS chloroplasts was investigated by the group of Romanowska et al. [ 66 ]. Studies were carried out on three species of C4 plants belonging to the NADP-ME subtype: Zea mays , Echinochloa crus galli , and Digitaria sanguinalis, characterized by different tolerances to high light intensity. The total content of carotenoids, including zeaxanthin and antheraxanthin, a final and an intermediate product of the xanthophyll cycle, was measured. It was shown that after exposure to high light intensity, the amount of zeaxanthin in the chloroplast of barnyardgrass ( Echinochloa crus galli ) was higher than in other investigated species. Barnyardgrass is a plant highly resistant to various environmental stresses, and the functioning of the xanthophyll cycle may be one of the crucial mechanisms that allows growth under stress conditions. An increase in zeaxanthin content was also observed in Sorghum bicolor (NADP-ME plant), after exposure to the same light intensity as described above. The amount of zeaxanthin, antheraxanthin, and violaxanthin participating in the xanthophyll cycle was twice as high compared to the control [ 67 ], which may indicate that regardless of the species, the functioning of the xanthophyll cycle is an important element of protection of the photosynthetic apparatus and dissipation of excess energy.

Moreover, under conditions of excess light, a transthylakoidal proton gradient (ΔpH) is generated. At a low pH of the thylakoid lumen, the PsbS protein, which is involved in energy dissipation, is protonated. The dimeric forms of PsbS are associated with the PSII core, whereas the PsbS monomers are associated with the LHCII antenna [ 68 ]. Conversion of PsbS homodimers to monomers occurs at low pH in a thylakoid lumen at high light intensity [ 68 ]. The energy quenching in the form of qE is higher when there are more quenching sites, so it should depend on the PsbS protein content.

The participation of the LHCII antenna in energy quenching was confirmed in the mesophyll chloroplasts of maize (NADP-ME), where after the high intensity of far-red light, the LHCII were dephosphorylated, detached from the PSI in the stroma lamellae, moved to the grana, and either bound to PSII or formed aggregates which in consequence, lead to induction of the qE parameter [ 41 ]. In M chloroplasts, light is not a factor that limits the production of ATP and NADPH, so the excess light energy is dissipated as heat. In the agranal BS chloroplasts, there was a slight dephosphorylation of the LHCII connected to PSI, but a high dephosphorylation of free and aggregated LHCII. The dephosphorylated antenna could connect to PSII and caused an energy transfer redirecting it to this photosystem. Because the proportion of LHCII aggregates decreases, therefore, the amount of energy dissipated is also reduced [ 41 ]. The authors suggest that because of the leaf structure, less light energy can reach the BS chloroplasts, thereby decreasing the pH gradient and lowering the qE parameter.

It can be assumed that in plants that perform C4 photosynthesis (excluding those with single-cell C4 photosynthesis), the mechanisms of quenching and dissipation of excess excitation energy should be more efficient in M chloroplasts due to the specific anatomical structure of the leaves. This causes much more light energy to reach the chloroplasts localized in M cells than those in BS cells.

Mechanisms that allow C4 plants to adapt the light reaction of the photosynthesis function under changing light conditions, particularly in high light intensity, are summarized in Table 1 . Many of them are universal and are also found in C3 plants. However, some are modified in C4 plants to provide more efficient CO 2 assimilation. The close relationship between the light phase of photosynthesis and the enzymatic reactions in chloroplasts, and the associated demand for ATP and NADPH, results that in C4 plants the linear and cyclic electron transport operate in a different ratio in the chloroplasts of M and BS cells. In addition, differences in the intensity of light reaching M and BS chloroplasts and in the thylakoid structure (granal and agranal) will affect the processes of the redistribution of excitation energy between photosystems and the dissipation of its excess. Therefore, it can be assumed that, in the M chloroplasts, because of increased incoming light energy, the mechanisms related to the dissipation of excess energy must function better than in BS chloroplasts to prevent photosystems from photoinhibition and, in consequence, from a decrease in ATP and NADPH. On the other hand, BS chloroplasts, which receive less light energy, must have better functioning mechanisms that allow for its efficient use. More research is needed on other subtypes of C4 plants to help explain the importance of chloroplast structure in the processes involved in the use and dissipation of excitation energy.

Summary of the chloroplast processes involved in the adaptation/acclimatization of C4 plants to high light intensities, compared to C3 plants, described in the article.

Acknowledgments

The authors are grateful to Elzbieta Romanowska (Faculty of Biology, UW) for critical reading of the manuscript.

Abbreviations

AOX: mitochondrial alternative oxidase; ATP: adenosine triphosphate; BS: bundle sheath; CA: carbonic anhydrase; CET cyclic electron transport; DC: decarboxylase; FR: far red; LET: linear electron transport; LHC: light-harvesting complex; LHCI: light-harvesting complex I; LHCII: light-harvesting complex II; M: mesophyll; NAD-ME: NAD-dependent malic enzyme: NADPH: nicotinamide adenine dinucleotide phosphate; NADP-ME: NADP-dependent malic enzyme; NDH: NADH dehydrogenase-like; OAA: oxaloacetate; PAR: photosynthetically active radiation; PCK: PEP carboxykinase; PCR cycle: photosynthetic carbon reduction cycle; PEP: phosphoenolpyruvate; PEPC: phosphoenolpyruvate carboxylase; PEPCK: PEP carboxykinase; PGRL1: proton gradient regulation like 1; PGR5: proton gradient regulation 5; PPDK: pyruvate, phosphate dikinase; PQ: plastoquinone; PQH 2 : reduced plastoquinone; PSI: photosystem I; PSII: photosystem II; PTOX: plastid terminal oxidase; ROS: reactive oxygen species; RuBisCO: ribulose-1,5-bisphosphate carboxylase/oxygenase; VDE: violaxanthin de-epoxidase; ZE: zeaxanthin epoxidase.

Author Contributions

Conceptualization, W.W.-D. and A.D.; writing manuscript, W.W.-D., A.D. and M.Z.; figures and table preparation, W.W.-D. and A.D. All authors have read and agreed to the published version of the manuscript.

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Data availability statement, conflicts of interest.

The authors declare no conflict of interest.

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