In the Calvin Cycle, How Many Atp Molecules Are Required to Regenerate Rubp From Five G3p Molecules?
C
In Dictionary of Energy (Second Edition), 2015
Calvin cycle Biological Energetics. the complete route that carbon travels through a plant during photosynthesis. [Named for U.S. biochemist Melvin Calvin, 1911–1977.] Also called the Calvin-Benson cycle.
See next column.
Calvin cycle The metabolic pathway by which carbon dioxide (CO2) is incorporated into carbohydrate. Nobel Laureate Melvin Calvin had a major role in elucidating this cyclic series of enzyme-catalyzed reactions. The enzyme RuBisCO (ribulose bisphosphate carboxylase/oxygenase) catalyzes the initial reaction of CO2 with a five-carbon compound ribulose-1, 5-bisphosphate (RuBP). The resulting 6-carbon intermediate splits into two copies of a 3-carbon compound that is converted in subsequent steps to the carbohydrate glyceraldehyde-3-phosphate. Some of this product exits the pathway to be used for synthesis of more complex carbohydrates or other carbon compounds. The rest is converted back to RuBP (the substrate for the initial CO2 fixation reaction), completing the cycle. Most carbon compounds in the biosphere are derived from the carbohydrate product of the Calvin Cycle. The abbreviated structure of a typical carbohydrate is (H-C-OH)n. Due to unequal sharing of electrons in a C-O bond, the carbon atom in CO2 is electron deficient relative to a carbon atom in a carbohydrate, that bonds with only one oxygen atom. Carbon in CO2 is thus said to be more oxidized, while carbon in a carbohydrate is more reduced. The Calvin Cycle does not directly utilize light energy, but is part of the process of photosynthesis. Some Calvin cycle reactions require ATP (adenosine triphosphate), a compound that functions in energy transfer, and NADPH (reduced nicotinamide adenine dinucleotide phosphate), a source of hydrogen atoms for reduction reactions. ATP and NADPH are formed during light-energized reactions of photosynthesis.
Joyce Diwan
Rensselaer Polytechnic Institute
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CO2 capture by bacteria and their enzymes
Alessandro Senatore , ... Angelo Basile , in Advances in Carbon Capture, 2020
18.2.1 Calvin cycle
Calvin cycle also known as Calvin-Benson-Bassham or reductive pentose pathway has been the first CO 2 fixation cycle discovered by Calvin, Benson and Bassham in plants and after reported in many other microorganisms [24–26]. In this cycle represented in Fig. 18.1, ribulose-1,5-biphosphate carboxylase oxygenase (RuBisCO), a key enzyme of this process, sequesters a CO2 molecule from the environment, to convert carboxylate ribulose-1,5-biphosphate (RuBP) to form 3-phosphoglycerate (PGA) with subsequent generation of glyceraldehyde-3-phosphate and a series of other reactions at the end of which RuBP is regenerated by the enzyme phosphoribulokinase (PRK).
Fig. 18.1. Calvin cycle. The key enzymes of this process are shown in red.
From R. Saini, R. Kapoor, R. Kumar, T.O. Siddiqi, A. Kumar, CO2 utilizing microbes—a comprehensive review, Biotechnol. Adv. 29 (6) (2011) 949–960.Read full chapter
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Biofixation of carbon dioxide (CO2) by microorganisms
B. Wang , C.Q. Lan , in Developments and Innovation in Carbon Dioxide (CO2) Capture and Storage Technology, 2010
15.2.1 Calvin cycle
The Calvin cycle, which is also called the reductive pentose phosphate cycle, is the most widespread CO 2 biofixation pathway among autotrophs. It exists in plants and microalgae, as well as photoautotrophic and chemoautotrophic bacteria. As shown in Fig. 15.1, the key step of the Calvin cycle is catalyzed by the enzyme ribulose bisphosphate carboxylase, which fixes a CO2 molecule onto a molecule of ribulose-1,5-diphosphate (RuBP), resulting in two molecules of glyceric acid-3-phosphate (3PG). These 3PG molecules are then converted into two glyceraldehyde-3-phosphate (G3P, aka phosphoglyceraldehyde, PGAL) molecules by adding a high-energy phosphate group from ATP to each molecule. The two 3PG molecules are then converted to a RuBP molecule, which stays in the cycle for another round of CO2, and an organic carbon unit [C]. Three rounds of Calvin cycle lead to fixation of 3 CO2 molecules and the production of a G3P molecule, which can be further converted to glucose, lipids and other cell materials.
Figure 15.1. The Calvin cycle: three molecules of CO2 fixed give a net yield of one molecule of glyceraldehyde 3-phosphate at the net cost of nine molecules of ATP and six molecules of NADPH.
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Resource Use Efficiency
D.W. Sheriff , ... P.B. Reich , in Resource Physiology of Conifers, 1995
b Partitioning of Nutrient within Assimilatory Pathway
Within the photosynthetic pathway, nitrogen is principally in Calvin cycle proteins, mainly ribulose bisphosphate carboxylase, and in thylakoid membranes of chloroplasts. The proportion of foliar nitrogen in each of these components varies both with species and with a leaf's microenvironment. A major factor of the microenvironment that influences this is the light regime under which a leaf develops ( Evans, 1989). Usually, there is a negative relationship between the "average" PPFD incident on a leaf and the proportion of nitrogen partitioned to the thylakoids in its chloroplasts; at lower light more thylakoids are needed for light capture. There is a corresponding positive relationship between PPFD and the proportion of nitrogen partitioned to ribulose bisphosphate carboxylase: at high light levels more is needed to utilize the captured energy. When nitrogen is partitioned to maximize carbon assimilation in a leaf's usual light environment, leaf ANnUE will be improved compared to that resulting from a fixed partitioning of nitrogen. Thus, PANnUEs of leaves from different light environments need to be measured under light regimes similar to those they normally experience. In low light environments foliar nitrogen concentrations do not usually affect carbon assimilation, so under these conditions values of PANnUE or of ANnUE have little meaning (e.g., Gulmon and Chu, 1981). Changes in assimilatory capacity not caused by nutrition, and changes in foliar nutrient concentration, will usually both affect a leaf's ANUE. The general effects of these changes are shown in Table I.
Plants that live in an environment normally deficient in a nutrient may optimize use of the nutrient partly by maximizing its ANUE. However, this can have a deleterious effect on the efficiency of water use (see below).
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Metabolic engineering approaches for high-yield hydrocarbon biofuels
Kalyan Gayen , in Hydrocarbon Biorefinery, 2022
9.6 Bioconversion of CO2 to hydrocarbons
Oxygenic photosynthesizers, like cyanobacteria, can consume naturally abundant CO2 and convert it to several organic compounds via Calvin cycle [69]. Cyanobacteria is a suitable candidate for converting CO2 to hydrocarbons using metabolic engineering approaches. Moreover, the photosynthetic biosynthesis pathway of cyanobacteria can be introduced to heterotrophic bacteria like E. coli for the production of hydrocarbon biofuels. The biosynthesis of hydrocarbons from CO2 is advantageous compared to lignocellulosic materials because of no requirement of feedstock pretreatment, which is a cost- and time-expensive step for lignocellulosic materials.
Production of two hydrocarbons, namely alkanes and isoprenoids, has been examined significantly in cyanobacteria cells [70, 71]. Cyanobacteria take a common route, like other alkane-producing organisms, through decarbonylation of fatty aldehydes [45]. The involvement of enzymes such as fatty aldehyde decarbonylase and fatty acyl-ACP reductase and related gene sequences was found in the decarbonylation pathway [72]. Cyanobacteria can also be employed to synthesize fatty acids of preferred chain lengths such as C8, C10, and C12 to compensate the jet fuel and gasoline. To increase such specificity of hydrocarbons, different heterologous genes encoding different acyl-ACP thioesterases such as FatB, FatB2, and FatB1 from Arabidopsis, Cuphea hookeriana, and Umbellularia californica, respectively, can be introduced to cyanobacteria metabolism. In addition to the insertion of an adequate acyl-ACP thioesterase, a high synthesis of free fatty acid in the cells will be needed to optimize a high yield of hydrocarbons from cell factories. At this end, increasing the activity of the gene like ACC can play a crucial role in enhancing the production of free fatty acids because the product of ACC gene catalyzes a critical rate-limiting step in fatty acid ACP biosynthesis. ACC gene contains many subunits such as AccA, AccB, AccC, and AccD, and not all cyanobacteria genomes include all these genes. Another strategy can be to insert heterologous genes from other organisms to enhance the synthesis of free fatty acids in cells. Along with the insertion of ACC, inactivation of AAS can reduce the re-thioesterification of free fatty acids, hence will enhance the net synthesis of free fatty acids [73].
Several studies illustrated the capabilities of cyanobacteria to produce a wide range of linear, branched, and cyclic alkanes. Biosynthesis of some of alkanes such as methyl and ethylalkanes is exclusively present in cyanobacterial cells. For instance, cyanobacterial strain, Anabaena cylindrica can produce C9–C16 n-alkanes under high salt (NaCl) concentration in the media [74]. Moreover, Microcoleus vaginatus was demonstrated to produce several n-alkanes and branched alkanes. More exploration efforts to identify and investigate the different cyanobacterial strains will lead us to an increasing variety of alkanes in terms of chain lengths and structures.
Isoprenoids have a higher-octane rating compared to n-alkanes, hence, they can be employed for high-performance gasoline engines. Cyanobacteria can naturally produce carotenoids such as geranyl pyrophosphate, geranylgeranyl pyrophosphate, and farnesyl pyrophosphate. These carotenoids can be employed directly as biofuel after some processing; however, carotenoids can be transformed to isoprenoids for more efficient use. Above carotenoids can be used as precursors for monoterpenes, sesqui- and triterpenes, and di- and tetraterpenes [75]. Although the production of isoprene was not reported naturally in cyanobacteria, however, insertion of isoprene synthase especially from plants demonstrated the biosynthesis of isoprene hydrocarbons in cyanobacteria cells [76]. The engineered carotenoid pathway in cyanobacteria, for producing isoprene, monoterpene, or sesquiterpene, can be established by moderate efforts through the insertion of a single isoprene synthase gene or different terpene synthase genes. However, manipulating carotenoid pathway for synthesizing pinene and farnesene will be more beneficial as these isoprenoids are considered as precursors of jet fuels and diesel fuels.
The capability of cyanobacteria to cultivate in low to high concentrations of CO2 makes them a suitable candidate for producing biofuels by consuming the CO2 produced from coal-based industries. Despite several advantages, considering cyanobacteria as biofuel producers brings many challenges. For example, (i) the dependency on the light will affect the biofuel production in the dark; (ii) harvesting light for growth is a slow process in the photosynthetic organism that will affect the productivity of biofuel production; and (iii) harvesting and extracting hydrocarbons can be a costly process as the current microalgae-based biofuel industry is facing this challenge. The above challenges should be addressed for establishing the economically feasible hydrocarbon-based biofuels from CO2.
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Challenges in industrialization of biological CO2 capture
Saeid Samipour , ... Payam Setoodeh , in Advances in Carbon Capture, 2020
19.2.1.1 Reductive pentose pathway or Calvin-Benson-Bassham (CBB) pathway or Calvin cycle
Not only the photosynthetic eukaryotic organisms, but also a number of prokaryotic microorganisms have been known to benefit from the Calvin cycle, and many more have been shown to at least harbor RuBisCO (see Fig. 19.2A) [15]. Many types of bacteria are considered to be responsible for the operation of the Calvin cycle, including purple nonsulfur bacteria (Rhodopseudomonas, Rhodospirillum, and Rhodobacter) [19], and purple sulfur bacteria (Chromatium), cyanobacteria (Anabaena, Anacystis, and Synechococcus) [20], along with hydrogen bacteria (Hydrogenovibrio and Ralstonia) [21], and other chemoautotrophs like Thiobacillus [22]. This cycle requires 13 enzymatic reactions. Among all, the RuBisCO-mediated reactions are responsible for CO2 biofixation [15].
Fig. 19.2. Natural CO2 biofixation pathways: (A) Calvin cycle, (B) reductive tricarboxylic acid cycle, (C) Wood-Ljungdahl pathway, and (D) 3-hydroxypropionate 4-hydroxybutyrate cycle [15, 16].
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Agricultural and Related Biotechnologies
E.D. Leonardos , B. Grodzinski , in Comprehensive Biotechnology (Second Edition), 2011
4.14.2.1 Photosynthetic Variants in Vascular Plants
Vascular plants are divided into at least three different photosynthetic types depending on the way they concentrate atmospheric CO2 at the site of the primary fixation step of the Calvin cycle catalyzed by RuBisCO [2, 35, 38, 39] (see Chapter 4.13). Each photosynthetic type possesses a unique set of anatomical ( Figure 1 ), biochemical, and physiological features ( Figure 2 ), which have developed through evolution, allowing for adaptation to different environmental conditions in which H2O losses and CO2 uptake are balanced.
Figure 1. Anatomy of source leaves of Flaveria species with different photosynthetic pathways. Cross sections are (a) a C3 species, F. robusta; (b) a C3–C4 intermediate species, F. floridana; and (c) a C4 species, F. trinervia.
Figure 2. A schematic of the major photosynthetic pathways (C3, CAM, C4, and C3–C4 subtypes) showing the intracellular flow of C from the atmosphere to the phloem in source leaves of vascular plants.
The three major biochemical variants of photosynthesis are: C3 photosynthesis, the first product of CO2 fixation being a three-C compound (3-phosphoglycerate, PGA); C4 photosynthesis, the first product being a four-C compound (e.g., oxaloacetic acid, OAA); and crassulacean acid metabolism (CAM), the first product being OAA during nighttime and PGA during daylight depending on prevailing plant or environmental factors. It is important to note that more than 90% of vascular plants are classified as C3, about 7% are CAM, and approximately 1.5% are only C4 plants [35, 38, 39].
In the C3 pathway, atmospheric CO2 enters the leaf via the stomata, diffuses as a gas to the chloroplast and is assimilated there directly through carboxylation using ribulose-1,5-bisphosphate (RuBP) as the acceptor by RuBisCO, in the photosynthetic C-reduction cycle also known as the Calvin cycle ( Figures 1(a) and 2 ). The same enzyme (RuBisCO) also catalyzes the oxidation of RuBP by O2, in the photorespiration cycle, resulting in released CO2. Thus, due to RuBisCO's low affinity for CO2 and its oxygenase activity, the major limitation of C3 photosynthesis is the fact that at present atmospheric CO2 concentrations, the net photosynthetic capacity of the Calvin cycle is reduced from that that can operate at elevated levels of CO2 (see Chapter 4.13 and references therein).
In C4 metabolism, inorganic CO2 in the form of HCO3 − is initially fixed by phosphoenolpyruvate carboxylase (PEPCase) in mesophyll cells (MCs) to form oxaloacetate (OAA) which is converted to malate or aspartate ( Figure 2 ). These four-C compounds (thus the name, 'C4') can be transferred to the bundle sheath cells (BSCs), neighboring the vascular conduits ( Figures 1 and 2 ). The concentric arrangement of BSCs around the vasculature is a feature of Kranz anatomy classically associated with the C4 metabolism ( Figure 1 (c)). A major feature of a C4 is that upon decarboxylation in the BSC, the CO2 released is refixed by RuBisCO in the Calvin cycle. The C4 species are classified into three subtypes according to the decarboxylation enzyme: Nicotinamide adenine dinucleotide phosphate (NADP)-dependant malic enzyme (NADP-ME subtype); NAD-dependant malic enzyme (NAD-ME subtype); and phosphoenolpyruvate carboxykinase (PEP-CK subtype) [17]. It suffices here to note that it appears that overall the C4 variants have improved the operation of RuBisCO by concentrating CO2 at the site of the Calvin cycle or supplying it when H2O losses through evaporation are minimized [35, 38, 39].
In addition, a small number of species which cannot be classified as C3, C4, or CAM have been found to have some aspects of both C3 and C4 photosynthesis and are named C3–C4 intermediate species [7]. Photorespiration in C3–C4 intermediate species may be suppressed by different means. According to the mechanism by which they reduce photorespiration, they can be classified into type I or type II C3–C4 intermediates. Type I C3–C4 intermediates have no C4 metabolism present, whereas type II intermediates have a limited but functional C4 metabolism ( Figure 2 ).
Two factors contribute to the reduction of photorespiration in C4 and C3–C4 intermediates. First, the availability of CO2 for RuBisCO provided by the decarboxylation of a transient metabolite favors carboxylation rather than oxidation by RuBisCO. Second, any CO2 generated internally through photorespiration can be refixed. Kranz anatomy requires many intercellular movements of intermediates between the MCs and BSCs. It is not clear yet to what extent a full and robust C4-type CCM requires all features of the Kranz anatomy for full functionality, but the proximity of the BSC to the vascular tissue especially for efficient phloem loading cannot be overlooked [15, 24, 29, 37, 40].
Most of the C4 types have what is known as Kranz anatomy ( Figure 1 ), although C4-like metabolism can occur in single cells [8, 39] – an observation that supports the theory that it may be possible with new genetic tools to engineer a C4-type CCM system intracellularly in a C3 plant, thus suppressing photorespiration. Understanding how such a spatial arrangement of enzymes is accomplished intracellularly versus intercellulary and maintained is very important in developing strategies to genetically engineer a functional C4 pathway in selected C3 plants (see Chapter 4.13).
Plants with CAM also have C4 biochemical features to concentrate the CO2 at the site of RuBisCO; however, the operation of the CAM-specific CCM involves a temporal separation of activities, can proceed intracellularly, and seems to be related to stomatal closure due to drought and heat stress during the day. The main difference between CAM and C4 fixation is that whereas C4 metabolism is based on synergistic operation in the light with spatial separation of the PEPase and RuBisCO, in CAM, the CCM is based on temporal separation of the synthesis and decarboxylation of a C4 intermediate and refixation of the CO2 by the action of RuBisCO in the light ( Figure 2 ). In CAM, the C4 intermediate compound is made in low light or darkness when the environment is cooler and there is less loss of H2O from the leaf. With the discovery of single-cell C4-like traits, it is evident that the difference between CAM and C4 in terms of evolution and gene regulation might be closer than once thought [38] (see Chapter 4.13). In a discussion of adapting crops to warm arid climates, it is noteworthy that with a little irrigation, many slower growing CAM crops can be as productive as their C4 relatives [35].
In defining leaf source strength, its photosynthesis rate is only one parameter of interest. Except perhaps for a crop like a leafy lettuce where the leaves are harvested (harvest index above 0.80), source strength in vascular plants is usually defined in terms of how much reduced C is exported from the source leaf to developing sinks.
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"Bioplastics from microalgae"—Polyhydroxyalkanoate production by cyanobacteria
Martin Koller , in Handbook of Microalgae-Based Processes and Products, 2020
22.4.3 Other storage compounds produced by cyanobacteria in parallel with PHA biosynthesis
In addition to PHA, glycogen and other polysaccharides are storage compounds frequently found in cyanobacteria; similar to PHA production, fixation of the required carbon starts with the Calvin cycle. Stal et al. (1990) investigated the different functions of these two reserve materials (glycogen and PHA) in the metabolic network of the marine cyanobacterium Oscillatoria limosa. These authors detected that the intracellular PHA fraction was growing during the late exponential phase of growth, and reached its maximum value during the stationary phase of growth. If transferred into carbon-limited fresh medium, O. limosa cultures rich in PHA rapidly started to degrade these PHA reserves under illumination, while in dark reaction, PHA degradation was not observed, whether with or without aeration. In contrast, polysaccharide (glycogen) reserves were degraded during the dark reaction in order to maintain the cell's metabolism (Stal et al., 1990). Further studies by Stal (1992) with dark reaction cultures of PHA-rich Gloeothece sp. PCC 6909 substantiated this finding; this strain also did not degrade its PHA reserves after stopping the light supply.
These results evidenced that PHA primarily acts as carbon reserve in cyanobacteria, while, for energy generation, cyanobacteria preferably degrade their glycogen storage. Further, PHA's role in regulating the intracellular redox state in phototrophic organisms like cyanobacteria was evidenced by the studies of De Philippis et al. (1992), who described the competition between nitrogenase and PHA synthesis enzymes for reducing equivalents in a photoheterotrophic purple non-sulfur bacterium; in this context, regeneration of the oxidized state of reduction equivalents by PHA biosynthesis is referred to as "pseudofermentation" due to its biochemical similarities to ethanol production by yeast as a response to oxygen limitation. The concomitant production of lipids, glycogen, and PHA in Synechocystis sp. PCC 6803 was studied by Monshupanee and Incharoensakdi (2014). Under photoautotrophic cultivation conditions, the sum of accumulation products peaked in the mid-stationary growth phase with about 0.4 g accumulation products (0.227 g glycogen, 0.141 g lipids, 0.024 g PHA) per gram of biomass. The mass fraction of accumulated products was increased to 0.615 g (0.368 g glycogen, 0.112 g lipids, 0.135 g PHA) per gram of biomass under nitrogen-limited conditions, which outperformed storage compounds production under phosphate-, sulfur-, iron-, or calcium-limited conditions investigated in parallel.
More recently, parallel intracellular production of carbohydrates and PHA was also reported for mixed microbial cultures enriched by cyanobacteria under different photoperiods and nutrient availability (Arias et al., 2018a), and under feast-and-famine feeding conditions (Arias et al., 2018b). For both storage compounds, sufficient carbon supply and parallel limitation of nitrogen (favoring PHA biosynthesis) or phosphate (favoring carbohydrate biosynthesis) were reported. In addition, Troschl et al. (2018) reported the cyclic nature of PHA and glycogen accumulation and utilization by Synechocytis sp. CCALA192 in pilot-scale experiments; as a new insight into the cyclic nature of storage compound metabolism in cyanobacteria, these authors emphasized the intracellular conversion of glycogen to PHA by older cells of this cyanobacterium.
De Philippis et al. (1992) studied Spirulina maxima under different cultivation conditions in order to shed further light on the function of PHA in the metabolism of cyanobacteria. In photoautotrophic batch cultures, grown either under balanced conditions or under nitrogen or phosphate deprivation, PHA fractions did not exceed 0.00005 wt.-% (50 μg PHA per g biomass). After long nitrogen starvation, the PHA fraction increased to 0.007 wt.-%, and 0.012 wt.-% after depletion of intra- and extracellular phosphate reserves; this was the first study reporting enhanced PHA production by cyanobacteria under phosphate-limited conditions. When suddenly increasing the photosynthesis activity by increasing illumination, or by shifting the cultivation temperature to 18°C, PHA biosynthesis was stopped completely. Under all these photoautotrophic growth conditions, considerably more glycogen than PHA was produced (De Philippis et al., 1992).
Production of an extracellular polysaccharide (EPS) was determined for A. cylindrica in parallel to PHA biosynthesis (Lama et al., 1996), which is analogous to reports for the bacteria Azotobacter beijerininckii (Pal et al., 1999) and Azotobacter vinelandii (Brivonese and Sutherland, 1989), or the haloarchaeon Hfx. mediterranei (Koller et al., 2015; Parolis et al., 1996). Moreover, the characterization of an EPS produced by Spirulina strain LEB 18 revealed the presence of six different pentoses and hexoses plus pending sulfate groups (Martins et al., 2014).
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Analysis, fate, and toxicity of engineered nanomaterials in plants
Wenjuan Tan , ... Jorge L. Gardea-Torresdey , in Comprehensive Analytical Chemistry, 2019
3 The effects of ENMs on photosynthesis
Plants rely on the photosynthesis process to obtain energy by two steps: (1) to convert carbon dioxide and water into glucose from sunlight in thylakoids; (2) in the Calvin cycle, energy (adenosine triphosphate, ATP) and oxygen (O 2) are released when decomposing glucose into pyruvate in stroma [14]. These two steps are vital for the circulation of carbon and oxygen in the ecosystem. Photosynthetic parameters including morphology of photosynthetic tissues or structures (chloroplast, stomata, and trichrome), photosynthetic pigments (chlorophyll a, b, and total chlorophyll) contents, gas exchange parameters [photosynthesis rate (Pn), transpiration rate (E), and stomata conductance (Gs)], as well as metabolite compounds (e.g. phytol), are essential indicators of plants to respond their interaction with ENMs in the environment.
Schematic diagram of the effects of ENMs exposure on photosynthesis in plant systems is shown in Fig. 1. Besides, a summary of the recent findings from studies on photosynthesis in vascular plants is provided in Table 2. As can be seen, several researchers reported that the root-to-leave or leave-to-root uptake of ENMs resulted in deformation of photosynthetic tissues [41]. For example, it was found that swollen chloroplasts with squeezed nuclei and thylakoids were shown in the wheat exposed to 100 and 400 mg kg− 1 nCeO2 [8]. Moreover, Lalau et al. [29] reported that grana in the chloroplast were displaced by distensible stroma in the duckweed exposed to 10 mg kg− 1 nCuO. Besides, Rajput et al. [41] found that a decrease in the number and size of stomata and trichomes was shown in spring barley exposed to 10,000 mg kg− 1 nCuO. These structural changes may be correlated with a disruption of chlorophyll synthesis genes. In a rice study of nZnO, by performing qRT-PRC technique, Chen et al. [17] found that several chlorophyll synthesis genes, such as CAO, CHLG, HEMA and HEMG, were significantly up-regulated at 25, 50, and 100 mg kg− 1, while CHLD and CHLM genes were down-regulated at 25 and 50 mg kg− 1.
Fig. 1. Schematic diagram of the effects of ENMs exposure on photosynthesis in plant systems.
Table 2. Summary of the findings from studies on photosynthesis in vascular plants.
| ENMs | Concentration (mg kg− 1) | Plant | Exposure way | Exposure time | Detection method | Findings | References |
|---|---|---|---|---|---|---|---|
| nAg | 4 or 40 mg/plant | Cucumber (Cucumis sativus) | Foliar | 7 days | GC-MS |
| [22] |
| nAg | 1, 10, 100 | Lettuce (Lactuca sativa) | Foliar | 7 days | UV-vis |
| [24] |
| nAg | 1.0, 2.5 | Arabidopsis thaliana | Root | 3 days 45 days | UV-vis |
| [25] |
| nCeO2 | 100, 400 | Wheat (Triticum aestivum) | Root | 7 months | TEM, UV-vis |
| [8] |
| nCeO2, nCuO | 50, 100, 200 | Cucumber (Cucumis sativus) | Foliar | 74 days | Gas exchange system |
| [15] |
| nCeO2 (uncoated and coated with citric acid), bCeO2 | 62.5, 125, 250, 500 | Tomato (Solanum lycopersicum) | Root | 210 days | UV-vis |
| [39] |
| nCu, bCu | 50, 100, 200 | Oregano (Origanum vulgare) | Root | 60 days | UV-vis |
| [40] |
| nCuO | 10,000 | Spring barley (Hordeum sativum distichum) | Root | 4 weeks | SEM, fluorometer |
| [41] |
| nCuO | 0.1, 1.0, 10.0 | Duckweed (Landoltia punctata) | Foliar | 7 days | UV-vis, TEM |
| [29] |
| nCuO, nZnO | 500 | Wheat (Triticum aestivum) | Root | 14 days | UV-vis |
| [42] |
| nCu(OH)2 | 100, 1000 | Cucumber (Cucumis sativus) | Foliar | 7 days | UV-vis |
| [43] |
| nCu(OH)2 | 100, 1000 | Corn (Zea mays) | Foliar | 7 days | GC-TOF-MS |
| [44] |
| nTiO2 | 1, 2, 4 | Long raceme elm (Ulmus elongata) | Foliar | 2 days | Gas exchange system |
| [45] |
| nTiO2 | 0.125, 1.25, 12.5 mmol L− 1 | Lettuce (Lactuca sativa) | Foliar | 7 days | UV-vis |
| [33] |
| nTiO2 | 0.025%, 0.05%, 0.1%, 0.15%, 0.2%, 0.25%, 0.4%, 0.6% | Spinach | Seed | 2 days | Portable photosynthesis detector |
| [46] |
| nTiO2 (unmodified, hydrophobic, hydrophilic) | 125, 250, 500, 750 | Basil (Ocimum basilicum) | Root | 65 days | UV-vis |
| [47] |
| nZnO | 20,225, 250, 900 | Bean (Phaseolus vulgaris), maize (Zea mays), pea (Pisum sativum), wheat (Triticum aestivum), Beet (Beta vulgaris) | Root | 35 days | UV-vis |
| [31] |
| nZnO | 25, 50, 100 | Rice (Oryza sativa) | Root | 7 days | UV-vis, qRT-PRC |
| [17] |
Furthermore, by using GC-MS or GC-TOF-MS methods, it has been shown that the production of the photosynthetic secondary metabolite, phytol, was interfered. Significantly higher phytol content was found in cucumber leaves exposed to 4 or 40 mg/plant nAg [22]. In the contrast, Zhao et al. [44] found that the exposure of 1000 mg kg− 1 nCu(OH)2 resulted in statistically lower phytol content in corn leaves. Further study is required to discover the mechanism of the modification of phytol levels in plant systems by different ENMs.
The above findings suggest the chlorophyll degradation would be impacted. Moreover, as can be seen in Table 2, the presence of ENMs affected the synthesis of chlorophyll contents. A few researchers found no significant effects on chlorophyll contents in lettuce exposed to 1, 10, and 100 mg kg− 1 nAg [24] or 10, 100, and 1000 mg kg− 1 paint-coated (hydrophobic) nTiO2 [33], and oregano exposed to 50, 100, and 200 mg kg− 1 nCu [40]. However, more studies have shown significant reductions or increases in chlorophyll productions. In a study of wheat exposed to 500 mg kg− 1 nCuO and nZnO, total chlorophyll contents were statistically decreased by 38% and 37%, respectively [42]. Tan et al. [47] studied photosynthetic responses of nTiO2 with different surface coatings in basil. They found that significant reductions in relative chlorophyll contents were shown in plants exposed to hydrophilic nTiO2 at 250 (24%) and 750 mg kg− 1 (18%). In a 3 days' nAg exposure study of Arabidopsis thaliana, Ke et al. [25] reported that significantly decreases in chlorophyll a, b, and total chlorophyll contents by 25% and 35% at 1.0 and 2.5 mg kg− 1. Similar results were found in the duckweed study exposed to 0.1 and 1 mg kg− 1 nCuO [29] and rice treated with 25, 50, and 100 mg kg− 1 nZnO [17]. Differently, Barrios et al. [39] reported that nCeO2 coated with citric acid (CA + nCeO2) resulted in significantly higher chlorophyll a, b, and total chlorophyll contents at 250 mg kg− 1. Zhao et al. [43] found similar results were shown in cucumber leaves exposed to 1000 mg kg− 1 nCu(OH)2. Recently, Garcia-Gomez et al. [31] found that the effects of ENMs on photosynthesis vary with soil type and plant species. In the calcareous soil, significant increases in chlorophyll a and b contents were only shown in beet. In the acidic soil, significant decreases in chlorophyll a and b were shown in maize, wheat, bean, and pea.
The changes in chlorophyll contents might be associated with the disturbance of gas exchange procedure (Table 2). Researchers found that in cucumber, the foliar exposure of 200 mg kg− 1 nCeO2 or nCuO resulted in significantly lower photosynthesis rate and transpiration rate [15]. In the spinach study of rutile nTiO2, statistically higher chlorophyll formation and the photosynthetic rate were determined [46]. Gao et al. [45] reported in the leaves of long raceme elm exposed to 1, 2, and 4 mg kg− 1 nTiO2 resulted in significantly higher stomatal conductance, intercellular carbon concentration, and transpiration rate. Later on, in another study, a statistically higher maximal quantum yield of photosystem II was shown in spring barley treated with 10,000 mg kg− 1 nCuO [41]. The changes in gas exchange parameters might be explained by the disruption of electron flows between photosystem I and II in the Hill reaction and Rubisco activities in the Calvin cycle [48].
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Agricultural and Related Biotechnologies
H. Ashida , A. Yokota , in Comprehensive Biotechnology (Second Edition), 2011
4.13.3.2 Enhancement of RuBP Regenerative Capacity
Flux control analyses in antisense plants revealed that the activities of sedoheptulose-1,7-bisphosphatase (SBPase), transketolase, and aldolase in the RuBP-regeneration phase contribute to the flux control of the Calvin cycle [32]. The extent to which each enzyme exerts control over flux through the cycle is indicated by its flux control coefficient [33]. The flux control coefficient varies from zero, for an enzyme that has no contribution to control, to one, for an enzyme that exerts total control. SBPase shows high flux control coefficient values, 0.35–0.7, indicating that its activity is a major determinant of flux through the Calvin cycle [32]. In fact, small decreases in SBPase activity reduce the CO2 assimilation rate in antisense plants. These experimental findings suggest that an increase in SBPase activity could enhance photosynthesis, and several studies have confirmed this. Miyagawa et al. [34] used Agrobacterium-mediated transformation to produce transgenic tobacco lines expressing the cyanobacterial gene for the bifunctional enzyme, fructose-1,6-bisphosphatase/SBPase (FBP/SBPase). Transformants showed approximately twofold increases in FBPase and SBPase activities compared with those of the wild type, and showed increases in CO2 assimilation rate and dry matter (124% and 150%, respectively, compared with the wild type) [34]. In the transformants, the RuBP level and the activation ratio of RuBisCO were increased by 1.8-fold compared with those of the wild type, despite the fact that there were no changes in total activities or amounts of other enzymes in the Calvin cycle. These data clearly demonstrated that enhancement of the CO2 assimilation rate in transgenic tobacco was due to an increase in the activation level of RuBisCO. Activation of RuBisCO is strongly dependent on the RuBP concentration [3], and RCA requires a mM-level of RuBP [28]. Thus, the upregulation of the activation state of RuBisCO is probably induced by activation of RCA via an increase in the RuBP level due to overexpression of FBP/SBPase. This result has been reproduced in transplastomic tobacco overexpressing the same enzyme [35]. These transplastomic plants showed 1.7- and 1.8-fold increases in CO2 assimilation rate and dry matter, respectively, relative to the wild type. These resulted from increases of FBPase and SBPase activities to levels 69 and 33 times greater than those of the wild type, respectively. Several transplastomic tobacco lines with different expression levels of FBP/SBPase were produced using various promoters. Analyses of these transformants showed that the effect of introducing FBP/SBPase on photosynthesis was saturated with two- to threefold increases in FBP/SBPase activity. Thus, there may be another step that limits photosynthesis in transgenic plants with excessive expression of FBP/SBPase [35]. Tobacco with increased SBPase activity, which was achieved by introducing the SBPase from Arabidopsis, also showed increased photosynthesis and productivity [36].
These results suggested that the reaction step catalyzed by SBPase is a crucial target to improve photosynthesis, and that the CO2 assimilation rate can be enhanced by promoting RuBP regeneration and, as a consequence, activating RCA.
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In the Calvin Cycle, How Many Atp Molecules Are Required to Regenerate Rubp From Five G3p Molecules?
Source: https://www.sciencedirect.com/topics/engineering/calvin-cycle