experiment limitations photosynthesis

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Recent advances in understanding and improving photosynthesis

Alicia v perera-castro.

1 Department of Biology, Universitat de les Illes Balears, INAGEA, Palma de Mallorca, Spain

Jaume Flexas

Since 1893, when the word “photosynthesis” was first coined by Charles Reid Barnes and Conway MacMillan, our understanding of the elements and regulation of this complex process is far from being entirely understood. We aim to review the most relevant advances in photosynthesis research from the last few years and to provide a perspective on the forthcoming research in this field. Recent discoveries related to light sensing, harvesting, and dissipation; kinetics of CO 2 fixation; components and regulators of CO 2 diffusion through stomata and mesophyll; and genetic engineering for improving photosynthetic and production capacities of crops are addressed.

Introduction

Photosynthesis is the chemical reaction that sustains most life on Earth. Since the description of the Hill reaction and the Calvin-Benson cycle 1 – 3 , knowledge about their components, regulation, and limitations experienced a vertiginous increase. It is widely known that plants have important handicaps related to photosynthesis. First, the photosynthetic apparatus that harvests and transforms light energy into electron transport for the generation of ATP and NADPH 2 must cope with the generation of dangerous reactive oxygen species (ROS) 4 and most of the energy must be “wasted” in dynamic heat dissipation mechanisms 5 . Second, the enzyme that catalyzes CO 2 fixation in the Calvin-Benson cycle—ribulose 1,5-bisphosphate carboxylase oxidase or rubisco—is inefficient owing to several intrinsic characteristics, the most notable being the competitiveness between carboxylation and oxidation processes, since the oxidation of D-ribulose-1,5-bisphosphate results in the energetically expensive but perhaps convenient photorespiratory pathway 6 . And, third, the diffusion of CO 2 from the atmosphere surrounding leaves through stomata and the leaf tissues to the carboxylation sites in the chloroplast stroma, where rubisco is located, is a dynamic pathway that is full of barriers and includes gaseous, lepidic, and aqueous phases, the latter with a small solubility and diffusivity for CO 2 .

In the last few years, researchers have tried to determine the limitations and components of the processes described above. Engineering photosynthesis targeting different aspects of photosynthesis and its regulation has also advanced. The aim of this review is to compile and organize these advances in photosynthesis from the last few years and suggest a next horizon for plant physiologists, ecologists, and geneticists.

Light harvesting and use

Light energy is absorbed and transferred to the photosystem II (PSII) core by the light-harvesting complex II (LHCII). The way this absorption is regulated is relevant, since excessive and/or unbalanced exposure to light can lead to the generation of ROS and, in the long term, to the initiation of senescence processes 7 . Some isoforms of LHCII upregulate its transcription and translation as a response to high irradiance 8 , 9 , and their interaction with PsbS—a protein that plays a special role in photoprotection—has been described in detail 10 . Furthermore, Janil et al . 11 discussed the enhanced dimerization of LHCII under strong light conditions as a photoprotective response partially responsible for the dissipation of excess excitation. In line with this, Albanese et al . 12 recently described how the organization of PSII–LHCII supercomplexes changed with the diversification of land plants, contributing to their adaptability to different light environments. However, photoprotective processes and their ecophysiological implications remain far from fully characterized 5 . At the extreme opposite to excess light, shaded leaves within the canopy exhibit lower photosynthesis rates and slower activation of rubisco, stomata opening, and relaxation of photoprotection states. These delays, especially in rubisco activation, have been estimated to decrease wheat assimilation by 21% in shade to sun transitions 13 . Indeed, the fact that light is often in excess in the most illuminated leaves while limited in the shaded leaves within the canopy has led to the suggestion that lowering chlorophyll content may result not only in negligible effects on leaf-level photosynthesis rates but also in a higher distribution of light harvesting through the canopy, hence potentially enhancing whole plant photosynthesis rates and yield 14 , 15 . On the other hand, alterations of the canopy structure have also been suggested as a mechanism to improve light interception and canopy assimilation (see the recent review by Morales et al . 16 and references therein), mainly through long-term breeding but also through hormonal and/or genetic means 17 .

Besides studies on the photosynthetic management of light amount , the effect of light quality on photosynthesis-related issues has also been addressed. It is widely known that growing under blue light conditions induces lower photosynthetic rates, increases the synthesis of carotenoids and anthocyanins and the photoprotection capacity, and decreases stomata size while increasing their density 18 . Light quality also affects the level of ROS and the expression of antioxidant enzymes 19 . Recently, Górecka et al . 20 demonstrated that PsbS is not only a compulsory protein for enhancing dissipation of the excess of light energy as heat but also relevant for the red/blue light-associated enhancement of tolerance to UV-C and chloroplast signaling for light memory. A recent study has also described a species-specific response of photosynthesis to the quality of light independent of its intensity 21 . These interspecific differences in light response represent an opportunity to deeply understand the elements of light harvesting and their adaptation to different light environments.

Rubisco kinetics and CO 2 -concentrating mechanisms

Interspecific variation of rubisco kinetics has also been a focus over the last several years. In two almost simultaneously published works, Hermida-Carrera et al . 22 and Orr et al . 23 assessed rubisco kinetics, their temperature dependency, and the aminoacidic replacements in the large subunit of rubisco in many crop species. Orr et al . 23 extended their study to include 75 angiosperm species and found that some undomesticated plants presented inherently better rubisco kinetics, being thus a potential source for crop photosynthesis improvement. Iñiguez et al . 24 and Flamholz et al . 25 extended the analysis of differences in rubisco catalysis across the phylogeny and correlated them with the incidence of CO 2 concentration mechanisms (CCMs), showing that organisms that had evolved CCMs tended to have faster rubiscos yet with lower affinity and specificity for CO 2 . Hermida-Carrera et al . 26 found similar results when comparing rubisco catalytic traits of orchids and bromeliads with and without CCMs. These results suggest that equipping C 3 crops with CCMs could be another strategy for fueling their photosynthetic capacity.

C 4 photosynthesis is often envisaged as an efficient CCM and thus converting typical C 3 crops into C 4 has been a long-standing goal, resulting in the development of large-scale projects like the ongoing C4Rice ( https://c4rice.com/ ), yet the goal has not been fully accomplished yet 27 . Furthermore, transitioning from mostly C 3 to mostly C 4 crops may be an efficient way to enhance productivity in a world exhibiting increased global aridity 28 , 29 , as it has been shown that in some cases C 4 plants performed better under drought than did C 3 species 30 . In the same vein, introducing crassulacean acid metabolism (CAM) into C 3 crops has been suggested as a strategy to increase water use efficiency, i.e. to maximize CO 2 fixation with minimum water loss through transpiration 31 , 32 . On the other hand, other CCMs like those found in algae and other aquatic organisms (e.g. pyrenoids and carboxysomes) have been reported to concentrate more CO 2 around rubisco than C 4 photosynthesis. Hence, while the C 4 mechanism allows CO 2 concentrations around rubisco of at least 10-times higher than those of the surrounding atmosphere 33 , eukaryotic algae like Chlamydomonas containing pyrenoids can concentrate CO 2 40-times 34 and prokaryotic cyanobacteria possessing carboxysomes 100-times 35 higher than the surrounding atmosphere. Consequently, the potential expression of cyanobacterial and algal CCMs in crop plants has been proposed as an opportunity to improve their photosynthesis 36 .

Despite the inefficiencies of light harvesting and rubisco, photochemical and/or biochemical limitations to photosynthesis are not larger than the diffusional limitations related to both stomatal and mesophyll resistances to CO 2 in most of the studied species 37 – 45 . Gago et al . 46 recently presented a compilation of photosynthetic limitations across land plants’ phylogenies, in which angiosperms showed a well-balanced distribution among biochemical, stomatal, and mesophyll limitations; photosynthesis in gymnosperms and ferns was co-limited mostly by stomatal and mesophyll limitations; and in bryophytes and lycophytes the mesophyll limitation largely predominated.

Mesophyll conductance components

Mesophyll conductance to CO 2 ( g m ) depends on several leaf structures that comprise the pathway from sub-stomatal cavities to carboxylation sites of rubisco. Intercellular air spaces, cell walls, plasma membranes, cytosol, double chloroplast membranes, and stroma offer resistance to CO 2 diffusion. Values of g m vary strongly among species, and short-term changes in g m have been reported in response to many different environmental variables 46 – 49 , although a part of them could reflect methodological errors or uncertainties 50 – 52 . While interspecific differences are largely explained by anatomical traits 37 – 39 , 53 , 54 , short-term changes cannot be explained either by variable leaf anatomy or by the temperature coefficient reported for CO 2 diffusion 55 – 57 . Consequently, it has been suggested that a biochemically facilitated CO 2 diffusion must contribute to g m instead of solely physical diffusion 56 , 58 – 60 . Short-term chloroplast movement, aquaporins, and carbonic anhydrases have been indicated as candidates 53 , 56 , 61 , although their actual involvement is far away from being conclusive.

For instance, despite the fact that chloroplast surface area facing intercellular airspaces per unit leaf area ( S c / S ) is one of the anatomical parameters more correlated with g m 37 , 53 , 54 , no evidence for an association between short-term changes of g m and chloroplast movement or leaf anatomy has been found 57 , 62 , 63 , with the exception of Arabidopsis mutants with phytochrome-mediated impairment of the chloroplast avoidance response 64 . In a similar way, the contribution of carbonic anhydrases to g m variations remains elusive and is a matter of ongoing debate 65 . The most recent studies showed that latitudinal variation of g m correlates with variations in carbonic anhydrase activity 66 , 67 and that a coupled inhibition of both g m and carbonic anhydrases is obtained with treatment with mercuric chloride 68 . Han et al . 69 also reported a decrease in the expression of carbonic anhydrase ( CA1 ) during drought. On the contrary, Kolbe and Cousins 70 did not find any variation in g m in five lines of maize despite their differences in carbonic anhydrase activity.

The role of aquaporins as enhancers of CO 2 diffusion across membranes has been widely reported 48 , 71 . Changes in g m had been induced by inhibitors of aquaporins 68 , 72 in transgenics 73 – 76 and in mutants 77 – 80 . Direct measurement of the CO 2 permeability of chloroplasts also revealed a 50% reduction in chloroplasts of an Arabidopsis aquaporin mutant as compared to the wild-type 81 . Despite these findings, Kromdijk et al . 82 recently reported null differences in g m among several knockout aquaporin mutants and wild-type, probably due to functional redundancy of aquaporin isoforms.

Additionally, the relative importance of these biochemical processes and anatomical traits in regulating g m remains unknown. Furthermore, recent studies showed uncertainty about estimating some relevant anatomical parameters from microscopic images of 2D cross-sections compared to 3D microscopy, especially the mesophyll surface area exposed to air-filled spaces 83 and chloroplast volume 84 . This could partially explain the differences in the g m calculated from chlorophyll fluorescence and/or gas exchange and g m calculated so far from anatomical models 38 , 39 , 53 , 85 , 86 . Earles et al . 87 have emphasized the need to improve 3D techniques and models to properly characterize leaf-level photosynthesis in its whole complexity.

Within the anatomical components, S c / S and cell wall thickness ( T cw ) have been recognized as especially determinant for g m 46 . Besides the effect of T cw , an effect of cell wall composition and porosity in short- and long-term variations of g m has been suggested 88 , 89 , and recently the first empirical evidence was provided. Thus, a reduction of g m was observed by Ellsworth et al . 90 in mutants with disrupted β-glucosyl polysaccharides of the cell wall. More recent studies have shown that the decrease of g m provoked by drought, salinity, and low temperatures is coupled with variations in the relative levels of cellulose, hemicelluloses, and pectins 91 , 92 . More evidence is needed to understand how cell wall composition affects porosity and CO 2 diffusion.

Stomatal conductance

As mentioned above, an additional important limiting factor of photosynthesis is the stomatal conductance ( g s ). Several internal and environmental factors are widely known to affect g s . Stomatal shape, size, density, and clustering influence g s and therefore photosynthesis 93 . These traits are established during leaf development and regulated by several phytohormones, especially abscisic acid (ABA) 94 . Light, CO 2 , and water supply also affect g s 95 , 96 .

The speed of g s responses to light and CO 2 has been recently compared among phylogenetic plant groups. Although fern and lycophyte stomata are not insensitive to light and CO 2 , their response is lower and slower than that observed in angiosperms 97 – 100 . Furthermore, unlike angiosperms, fern and lycophyte stomata do not respond to endogenous levels of ABA 97 , 98 and their closure is based on a passive response of guard cells to dehydration 101 . The mechanism that explains this different response remains unclear, although it is likely related to differences in the molecular mechanisms operating in the guard cells along the phylogeny. Among other factors affecting g s (kinases, anion channels, etc.), it is known that carbonic anhydrases can be involved in the biochemical mechanism by which guard cells of angiosperms sense CO 2 (see the review by Engineer et al . 95 ), although details of signal transduction and the identity of the second messengers (bicarbonate, protons) are still debated. Furthermore, a higher CO 2 assimilation related to phosphoenolpyruvate carboxylase activity followed by gluconeogenesis and maybe sucrose synthesis has been described for guard cells in comparison to those of mesophyll cells of C 3 plants 102 .

In addition, recent studies suggest that stomata movement is regulated by mesophyll-derived signals. Sucrose has been identified as an important metabolite for the regulation of stomatal opening and closure 100 , 103 , 104 . Wang et al . 105 reported that the maize mutant cst1 —with an impaired membrane glucose transporter CST1 located in the subsidiary cell membrane—presented lower g s , lower photosynthesis, and earlier senescence than the wild-type. In line with this, Fujita et al . 106 demonstrated that stomatal responses are disrupted when a membrane excluding molecules of 100–500 Da is transplanted between mesophyll and guard cells, which would avoid the transport of sucrose, malate, and ABA. In a study of ABA-regulated genes in Arabidopsis , Yoshida et al . 107 found highly expressed genes in guard cells related to the tricarboxylic acid cycle and sucrose and hexose transport and metabolism. These studies support the hypothesis of stomatal regulation driven by carbohydrate/hormone-related mesophyll signals. However, the differences in the mechanism of mesophyll cell signaling and in guard cell metabolism among fern, lycophytes, and angiosperms—both anisohydric and isohydric species—remain unknown.

Even in angiosperms, the predominance of hormonal vs . hydraulic stomatal regulation is currently under debate 108 – 110 . Traditionally, stomatal closure has been understood as a safety valve to prevent cavitation (see Hochberg et al . 111 and references therein). However, a detailed chronological description of the drought response of g s and hydraulic conductance ( K leaf ) in rice revealed that the decline in K leaf preceded and probably triggered the decline of g s and g m 108 . Nadal et al . 112 suggested that both types of drought response are not necessarily incompatible and can be related to the spectrum of the iso-anisohydric response of angiosperms.

Engineering photosynthesis

While there are some opposing views 113 , improving photosynthesis is often envisaged as an important goal for improving crop yields 114 – 117 , including the cultivation of photosynthetic microorganisms, which constitutes a huge and important branch of bioengineering for bioenergy production 118 , 119 . Regarding land plant bioengineering, optimizing production with a minimum investment of resources (water, land, and nutrients) is the aim of ongoing large-scale projects, such as the already mentioned C4rice or the RIPE project ( https://ripe.illinois.edu/ ). Several targets for manipulation—including all those mentioned in the above sections—have been proposed with the aim of improving photosynthesis and crop yield 120 , 121 . Neglecting which are the main limitations for photosynthesis when targeting genes for improving photosynthesis is an example of the mutual disregard that ecophysiologists and biotechnologists have had for each other in the last few decades 122 , i.e. biotechnologists attempting to improve photosynthetic targets that ecophysiologists were showing to be non-limiting for photosynthesis. Using a model approach, Flexas 116 showed that only modest improvements of photosynthesis can be expected from relaxing only one limiting factor, since photosynthetic limitations are generally well-balanced in angiosperms 46 . Nevertheless, even with this relatively modest approach, increases of yield of >40% have been reported in some successful attempts 117 .

Rubisco kinetics have been among the most common targets for improving photosynthesis. All the advances in rubisco engineering have implied important improvements in our understanding of rubisco regulation and assembly but unsuccessfully improved the catalytic performance of rubisco 123 , 124 or photosynthesis 125 . While faster rubisco from cyanobacteria have been successfully engineered in transplastomic tobacco 126 , post-transcriptional assembly of functional rubisco in large enough quantities remains a limiting factor, likely due to the inability of local chaperones to deal with foreign rubisco fragments (see Whitney et al . 127 and their attempt to solve this problem by the use of ancillary chaperone genes). For this reason, this is a very active area of ongoing research 127 , 128 . Rubisco activase is another potential limiting factor, as Fukuyama et al . 129 also showed how increased expression of rubisco activase resulted in a negative correlation with rubisco content.

Besides achieving more efficient rubiscos, an alternative strategy has been to increase CO 2 concentration by either introducing elements of algal CCMs or bypassing photorespiration by different processes. While theoretically CCMs should increase photosynthesis 130 , introducing CCMs into either tobacco 131 or Arabidopsis failed to increase photosynthesis 132 , 133 , probably because of insufficient encapsulation of local rubisco in the foreign carboxysomes, which can be improved by simultaneously replacing the native large subunit of rubisco 134 . Additional elements might also be essential for a proper assemblage of fully functional carboxysome–rubisco CCMs, as recently demonstrated for bestrophin-like proteins 135 .

More successful results have been obtained when the photorespiration pathway has been manipulated in Arabidopsis and tobacco 136 , 137 . While photosynthesis increases 136 , biomass production has been shown to vary from decreasing through unaffected to increasing by 10–50% 117 , 138 . Recently, South et al . 137 obtained a 24% maximum increase of biomass when glycolate byproducts of photorespiration are processed by foreign malate synthase and a green algal glycolate dehydrogenase, substituting the native pathway. Tissue-specific overexpression of one of the subunits forming in the glycine dehydrogenase system also increased biomass yield by 13–38% in tobacco 139 . This is a very promising approach for improving grain crop yields in the near future.

Also, modifications of the Calvin-Benson cycle have resulted in improved photosynthesis and yield. Overexpression or transgenic insertion of several enzymes involved in the cycle (mostly sucrose bis-phosphatase—SBPase—and fructose bis-phosphatase—FBPase—but also FBPaldolase) has also resulted in increased photosynthesis and dry weights, although generally not in improved yield. However, Driever et al . 140 showed an up to 40% increase in grain yield in wheat, and Simkin et al . 141 a 35–53% increase in seed yield in Arabidopsis . Furthermore, overexpression of FBP/SBPase has been recently combined with an improved electron transport by the addition of the algae cytochrome C 6 , which also resulted in up to 53% of increase of biomass 142 . These results open up the possibility of using this approach for improving crop yields in the very near future.

Few attempts have focused on modifying CO 2 diffusive characteristics of leaves. Altered stomatal density in epidermal patterning factor (EPF) mutants of Arabidopsis 143 and wheat 144 resulted in an increased photosynthetic water-use efficiency (WUE) but not increased photosynthesis itself. Similarly, Yang et al . 145 showed that overexpression of the ABA receptors RCAR6/PYL12 increases the sensitivity of the stomata in Arabidopsis lines, reducing g s even in the absence of water stress without affecting photosynthesis, thus also enhancing WUE. As described in previous sections, g s was also enhanced by overexpression of glucose transporters in subsidiary cell membranes 105 .

Generally speaking, increasing stomatal conductance does not result in enhanced photosynthesis because stomatal limitations are generally minor in the absence of stress. However, during leaf development, the presence of well-developed and functional stomata appears to be the main driver of the development of mesophyll porosity, which is an essential anatomical trade favoring g m and hence photosynthesis 146 . This finding is remarkable as it implies that, while it is likely that a mesophyll signal is involved in stomata regulation (see above sections), stomata define the developmental set-up of the mesophyll structure, hence establishing a very intricate co-dependency between g s and g m limitations at different time scales that deserves further study. In line with this, Lehmeier et al . 147 showed that it is possible to genetically modify cell density and the arrangement of the air channels with an overall decreased path tortuosity in the palisade air spaces in a way that facilitates g m without affecting g s . Similarly, alteration of leaf mesophyll anatomy of Eucalyptus has been attempted by the overexpression of the transcription factor EcHB1 , which is involved in multiple genes related to cell wall biosynthesis and cell growth, increasing the number of chloroplasts per unit leaf area and therefore enhancing CO 2 diffusion into chloroplasts and photosynthesis 148 . These results offer new possibilities in improving photosynthesis by reducing CO 2 diffusion limitations. Advances in the understanding of cell wall composition determinants of g m may open complementary doors in the near future.

While significant and important in some cases, the above-described manipulations aimed to improve maximum photosynthesis rates, i.e. light-saturated photosynthesis in the absence of abiotic and biotic stresses. However, photosynthesis in nature occurs in largely variable conditions, e.g. in fluctuating light. For instance, De Souza et al . 43 showed in cassava that, while under steady-state high-light conditions, g m and biochemical limitations accounted for up to 84% of the total photosynthetic limitation and, under non-steady state conditions during shade to sun transition, g s became the most dominant limitation. Thus, in recent years, research has focused on improving photosynthesis and efficiency under non-steady-state conditions by decreasing the excess absorption of light 15 , 149 or increasing the relaxing velocity of photoprotection 150 – 152 . More surprisingly, overexpressing PsbS in transgenic tobacco resulted in enhanced WUE by reducing g s , not increasing photosynthesis, again pointing to potential mesophyll signals in stomata regulation 153 . Recently, Papanatsiou et al . 154 used an optogenetic approach to improve photosynthesis, WUE, and growth in Arabidopsis . They expressed a synthetic light-gated K + channel in stomatal guard cells (BLINK1), which improved the speed of stomata kinetics in response to varying light. Increased velocity of stomata opening from a dark-to-light transition and closing from a light-to-dark transition resulted in increased plant growth and WUE by approximately 30% 154 .

Light sensing, photoprotection, CO 2 diffusion, and its fixation involve numerous and complex processes that are far from fully understood. In the last few years, new insights have been obtained into how interaction and conformation of light-harvesting complexes and photosystems affect photoprotection and heat dissipation. Advances have been made also in the understanding of the variability in rubisco kinetics and photosynthetic limitations at steady state along the plant’s phylogeny, of the genetics and mechanistic aspects of carbon-concentrating mechanisms, and of the major anatomical determinants of g m and the metabolic determinants of stomatal conductance and kinetics. Important links between mesophyll and stomatal cells have been revealed, although the signaling between mesophyll cells and guard cells that regulates g s requires further research, as does understanding the chemical and biochemical determinants of g m .

Nevertheless, owing to the new knowledge acquired, engineering efforts for improving photosynthesis and photosynthetic WUE have been attempted, some of them with significant success, which open up the opportunity for photosynthesis-mediated improvement of crop productivity in the forthcoming years. To achieve this goal, a close collaboration among plant physiologists, molecular biologists, geneticists, and agronomists might be essential for generating multiple new photosynthetic genotypes and evaluating them under realistic conditions, both under steady- and non-steady-state conditions, from a photosynthetic limitations perspective to a yield and WUE perspective 122 . Technical advances in analytical tools, like the recently implemented rapid CO 2 response curves of gas exchange 155 – 159 , would be crucial to allow in-depth phenotyping of photosynthesis in record times.

Funding Statement

Alicia V. Perera-Castro and Jaume Flexas’s research was supported by the project EREMITA (PGC-2018-093824-B-C41) from the Ministerio de Economía y Competitividad (MINECO, Spain) and the ERDF (FEDER). The Ministerio de Educación, Cultura y Deporte (MECD, Spain) supported a pre-doctoral fellowship (FPU-02054) awarded to Alicia V. Perera-Castro. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

The peer reviewers who approve this article are:

• Asaph B. Cousins , School of Biological Sciences, Washington State University, WA, USA No competing interests were disclosed.
• Esa Tyystjärvi , Department of Biochemistry/Molecular Plant Biology, University of Turku, Turku, Finland No competing interests were disclosed.

Practical Biology

A collection of experiments that demonstrate biological concepts and processes.

Observing earthworm locomotion

Practical Work for Learning

Published experiments

Investigating the light dependent reaction in photosynthesis.

It is fairly easy to show that plants produce oxygen and starch in photosynthesis . At age 14–16 students may have collected the gas given off by pond weed (for example Elodea ) and tested leaves for starch.

It is not quite so easy to demonstrate the other reactions in photosynthesis. For the reduction of carbon dioxide to carbohydrate there must be a source of electrons . In the cell, NADP is the electron acceptor which is reduced in the light-dependent reactions, and which provides electrons and hydrogen for the light-independent reactions.

In this investigation, DCPIP (2,6-dichlorophenol-indophenol), a blue dye, acts as an electron acceptor and becomes colourless when reduced, allowing any reducing agent produced by the chloroplasts to be detected.

Lesson organisation

This investigation depends on working quickly and keeping everything cool. Your students will need to understand all the instructions in advance to be sure that they know what they are doing.

Apparatus and Chemicals

Per student or group of students:.

Centrifuge – with RCF between 1500 and 1800g

Centrifuge tubes

Fresh green spinach, lettuce or cabbage, 3 leaves (discard the midribs)

Cold pestle and mortar (or blender or food mixer) which has been kept in a freezer compartment for 15–30 minutes (if left too long the extract will freeze)

Muslin or fine nylon mesh

Filter funnel

Ice-water-salt bath

Glass rod or Pasteur pipette

Measuring cylinder, 20 cm 3

Beaker, 100 cm 3

Pipettes, 5 cm 3 and 1 cm 3

Bench lamp with 100 W bulb

Test tubes, 5

Boiling tube

Pipette for 5 cm 3

Pipette for 0.5 cm 3

Pipette filler

Waterproof pen to label tubes

Colorimeter and tubes or light sensor and data logger

0.05 M phosphate buffer solution, pH 7.0: Store in a refrigerator at 0–4 °C ( Note 1 ).

Isolation medium (sucrose and KCl in phosphate buffer): Store in a refrigerator at 0–4 °C ( Note 2 ).

Potassium chloride (Low Hazard) ( Note 3 ).

DCPIP solution (Low Hazard): (1 x 10 - 4 M approx.) ( Note 4 )

Health & Safety and Technical notes

Although DCPIP presents minimal hazard apart from staining, it is best to avoid skin contact in case prolonged contact with the dye causes sensitisation. Do not handle electric light bulbs with wet hands. All solutions used are low hazard – refer to relevant CLEAPSS Hazcards and Recipe cards for more information.

Read our standard health & safety guidance

1 0.05 M phosphate buffer solution, pH 7.0. Na 2 HPO 4 .12H 2 O, 4.48 g (0.025 M) KH 2 PO 4 , 1.70 g (0.025 M). Make up to 500 cm 3 with distilled water and store in a refrigerator at 0–4 °C. Low hazard – refer to CLEAPSS Hazcard 72.

2 Isolation medium. Sucrose 34.23 g (0.4 M) KCl 0.19 g (0.01 M). Dissolve in phosphate buffer solution (pH 7.0) at room temperature and make up to 250 cm 3 with the buffer solution. Store in a refrigerator at 0–4 °C. Low hazard – refer to CLEAPSS Hazcard 40C.

3 Potassium chloride 0.05 M. Dissolve 0.93 g in phosphate buffer solution at room temperature and make up to 250 cm 3 . Store in a refrigerator at 0–4 °C. Use at room temperature.(Note that Potassium chloride is a cofactor for the Hill reaction.) Refer to CLEAPSS Hazcard 47B and Recipe card 51.

4 DCPIP solution DCPIP 0.007–0.01 g, made up to 100 cm 3 with phosphate buffer. Refer to CLEAPSS Hazcard 32 and Recipe card 46.

Keep solutions and apparatus cold during the extraction procedure, steps 1–8, to preserve enzyme activity. Carry out the extraction as quickly as possible.

Preparation

a Cut three small green spinach, lettuce or cabbage leaves into small pieces with scissors, but discard the tough midribs and leaf stalks. Place in a cold mortar or blender containing 20 cm 3 of cold isolation medium. (Scale up quantities for blender if necessary.)

b Grind vigorously and rapidly (or blend for about 10 seconds).

c Place four layers of muslin or nylon in a funnel and wet with cold isolation medium.

d Filter the mixture through the funnel into the beaker and pour the filtrate into pre-cooled centrifuge tubes supported in an ice-water-salt bath. Gather the edges of the muslin, wring thoroughly into the beaker, and add filtrate to the centrifuge tubes.

e Check that each centrifuge tube contains about the same volume of filtrate.

f Centrifuge the tubes for sufficient time to get a small pellet of chloroplasts. (10 minutes at high speed should be sufficient.)

g Pour off the liquid (supernatant) into a boiling tube being careful not to lose the pellet. Re-suspend the pellet with about 2 cm 3 of isolation medium, using a glass rod. Squirting in and out of a Pasteur pipette five or six times gives a uniform suspension.

h Store this leaf extract in an ice-water-salt bath and use as soon as possible.

Investigation using the chloroplasts

Read all the instructions before you start. Use the DCPIP solution at room temperature.

i Set up 5 labelled tubes as follows.

j When the DCPIP is added to the extract, shake the tube and note the time. Place tubes 1, 2 and 4 about 12–15 cm from a bright light (100 W). Place tube 3 in darkness.

k Time how long it takes to decolourise the DCPIP in each tube. If the extract is so active that it decolourises within seconds of mixing, dilute it 1:5 with isolation medium and try again.

Teaching notes

Traditionally the production of oxygen and starch are used as evidence for photosynthesis. The light-dependent reactions produce a reducing agent. This normally reduces NADP, but in this experiment the electrons are accepted by the blue dye DCPIP. Reduced DCPIP is colourless. The loss of colour in the DCPIP is due to reducing agent produced by light-dependent reactions in the extracted chloroplasts.

Students must develop a clear understanding of the link between the light-dependent and light-independent reactions to be able to interpret the results. Robert Hill originally completed this investigation in 1938; he concluded that water had been split into hydrogen and oxygen. This is now known as the Hill reaction.

You can examine a drop of the sediment extract with a microscope under high power to see chloroplasts. There will be fewer chloroplasts in the supernatant – which decolourises the DCPIP more slowly, reinforcing the idea that the reduction is the result of chloroplast activity.

Sample results

Using a bench centrifuge

The experimental procedure was followed. A standard lab centrifuge was used to spin down the chloroplasts (Clifton NE 010GT/I) at 2650 RPM, 95 X g for 10 minutes.

The experiment was started within 5 minutes of preparing the chloroplasts. The reaction was followed using an EEL colorimeter with a red filter – readings taken every minute.

Tube 3 (incubated in the dark) gave a reading of 5.4 absorption units after 20 minutes. Tube 2 (DCPIP with no leaf extract) was 6.2 absorption units.

Using a micro-centrifuge

The experiment was repeated using a micro-centrifuge.

Tube 3 (incubated in the dark) gave a reading of 4.9 absorption units after 10 minutes.

Tube 2 (DCPIP with no leaf extract) was 6.4 absorption Units.

The relative activity of the pellet was higher than when the bench centrifuge was used. The micro-centrifuge tubes were only 1.5 cm 3 capacity – not ideal for this practical. A higher speed bench centrifuge would be better.

In order to check for loss of chloroplast activity, the experiment was repeated using the same chloroplast suspension 1 and 2 hours after preparation. Chloroplast suspension was kept in a salt-ice bath. There was no loss of activity when the extract was kept in ice for up to 2 hours.

Student questions

1 Describe and explain the changes observed in the five tubes. Compare the results and make some concluding comments about what they show.

2 The rate of photosynthesis in intact leaves can be limited by several factors including light, temperature and carbon dioxide. Which of these factors will have little effect on the reducing capacity of the leaf extract?

3 Describe how you might extend this practical to investigate the effect of light intensity on the light-dependent reactions of photosynthesis.

1 Colour change and inferences that can made from the results: Tube 1 (leaf extract + DCPIP) colour changes until it is the same colour as tube 4 (leaf extract + distilled water). Tube 2 (isolation medium + DCPIP) no colour change. This shows that the DCPIP does not decolourise when exposed to light. Tube 3 (leaf extract + DCPIP in the dark) no colour change. It can therefore be inferred that the loss of colour in tube 1 is due to the effect of light on the extract. Tube 4 (leaf extract + distilled water) no colour change. This shows that the extract does not change colour in the light. It acts as a colour standard for the extract without DCPIP. Tube 5 (supernatant + DCPIP) no colour change if the supernatant is clear; if it is slightly green there may be some decolouring. The results should indicate that the light-dependent reactions of photosynthesis are restricted to the chloroplasts that have been extracted.

2 Carbon dioxide will have no effect, because it is not involved in the light-dependent reactions.

3 Students should describe a procedure in which light intensity is varied but temperature is controlled.

Health and safety checked, September 2008

Related experiment

Investigating photosynthesis using immobilised algae

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Biology Teaching Resources

Photosynthesis Virtual Lab

This lab was created to replace the popular waterweed simulator which no longer functions because it is flash-based. In this virtual photosynthesis lab , students can manipulate the light intensity, light color, and distance from the light source.

A plant is shown in a beaker and test tube which bubbles to indicate the rate of photosynthesis. Students can measure the rate over time. There is an included data table for students to type into the simulator, but I prefer to give them their own handout ,

The handout is a paper version for students to write on as the work with the simulator. The document is made with google docs so that it can be shared with remote students.

There are several experiments that can be done in the lab that would complement this virtual experiment. For example, students can use elodea and measure the number of bubbles released when the plant is under a bright light. Algae beads can also be used to measure changes in pH as the plants consume carbon dioxide.

In experiment 2, students specifically look at light color to determine which wavelength of light increases the rate of photosynthesis. Students should discover that green light has a very slow rate. Their collected data is then compared to a graph of the absorption spectrum of light.

Shannan Muskopf

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• Published: 23 February 2016

Differences on photosynthetic limitations between leaf margins and leaf centers under potassium deficiency for Brassica napus L.

• Zhifeng Lu 1 , 2 ,
• Tao Ren 1 , 2 ,
• Yonghui Pan 1 , 2 ,
• Xiaokun Li 1 , 2 ,
• Rihuan Cong 1 , 2 &
• Jianwei Lu 1 , 2  

Scientific Reports volume  6 , Article number:  21725 ( 2016 ) Cite this article

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• Plant physiology

Analyzing the proportions of stomatal (S L ), mesophyll conductance (MC L ) and biochemical limitations (B L ) imposed by potassium (K) deficit and evaluating their relationships to leaf K status will be helpful to understand the mechanism underlying the inhibition of K deficiency on photosynthesis ( A ). A quantitative limitation analysis of K deficiency on photosynthesis was performed on leaf margins and centers under K deficiency and sufficient K supply treatments of Brassica napus L. Potassium deficiency decreased A , stomatal ( g s ) and mesophyll conductance ( g m ) of margins, S L , MC L and B L accounted for 23.9%, 33.0% and 43.1% of the total limitations. While for leaf centers, relatively low limitations occurred. Nonlinear curve fitting analysis indicated that each limiting factor generated at same leaf K status (1.07%). Although MC L was the main component of limitations when A began to fall, B L replaced it at a leaf K concentration below 0.78%. Up-regulated MC L was related to lower surface area of chloroplasts exposed to intercellular airspaces ( S c / S ) and larger cytosol diffusion resistance but not the cell wall thickness. Our results highlighted that photosynthetic limitations appear simultaneously under K deficiency and vary with increasing K deficiency intensity.

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Introduction.

Potassium (K), one of the macronutrients essential for plant growth and development, is involved in many physiological processes, such as photosynthesis, enzyme activation, water relations, assimilate transport and protein synthesis 1 , 2 . K deficiency profoundly decreased crop yield 3 , 4 , thus strategies of survival and improvement would be important for plant growing under adverse conditions. It is a truism that most of the dry matter is formed by leaf photosynthesis ( A ) which is intimately connected with K status. Multifarious studies have come to a nearly consistent conclusion that leaf A decreases in K-starved plants 3 , 5 , 6 , therefore an even deep comprehending of mechanism underlying the inhibition of K deficiency on A is necessary 2 .

During photosynthesis, CO 2 moves from external atmosphere to the internal leaf air spaces through the stomata and from there to carboxylation sites inside the chloroplasts 7 . It is established that stomatal closure is the foremost limitation to CO 2 assimilation due to the vital role of K in stomatal aperture 8 , 9 . As a major osmotica, K + accumulation in vacuole is essential for stomatal opening, which had been verified to be initially dropped under K deficiency 10 . And because of this, Bednarz et al. 5 stated that the most limiting resistance to A of Gossypium hirsutum L. came from stomata 5 . In contrast, K starvation caused a decreased A and stomatal conductance ( g s ), but an increased intercellular CO 2 concentration ( C i ) of Carya cathayensis leaves, suggesting that, in addition to g s , mesophyll conductance ( g m ) and biochemical limitations might be involved in the depression of photosynthesis in K deficient conditions 6 . Numerous studies have shown that g m is relatively low, leading to great draw-down of chloroplastic CO 2 concentration from C i and changed along the variation of water status, nitrogen nutrient, irradiance, temperature and CO 2 concentration 11 , 12 , 13 , 14 . Moreover, leaf structures, specific aquaporins, plasma membrane etc. are involved in the determinations of g m 15 . K starvation might have reduced aquaporin activity 16 and increased leaf dry mass per unit area ( M A ) 1 , 17 , therefore, causing a stronger mesophyll diffusion resistance to CO 2 delivery 18 . Besides, K nutrition also known to increase the leaf intercellular air space to enhance g m 1 .

Additionally, biochemical processes may restrain photosynthesis, particularly under severe and/or long-time K starvation 1 , 5 , 6 . It was reported that Rubisco (ribulose-1,5-bisphosphate carboxylase/oxygenase, EC 4.1.1.39) activity was decreased under K deficiency, becoming a major limiting factor for photosynthesis in Oryza sativa leaves 3 . Chlorophyll synthesis was observed to be significantly impaired under K deficiency in Eucalyptus grandis leaves 1 . Moreover, K starvation up-regulated the fraction of electron transport to O 2 , resulting in an increased reactive oxygen species (ROS) 19 . Carbohydrate accumulation which may feedback regulation of leaf photosynthesis is more easily observed in K starved leaves 20 , 21 . Indeed, the relative contributions of these three limiting processes to photosynthesis under K deficiency and the underlying mechanisms have not been fully explored, due to the complicated physiological processes and variation of dominant limiting factors under differ K deficiencies 2 , 5 . No matter what the primary cause of decrease A , the discrepancy between researches was believed to be derived from differ physiological K deficiency severities. For this reason, a comprehensive consideration of whole limiting factors and their relationships with leaf K status seems to be important.

In 2005, Grassi and Magnani proposed a method to accurately quantify photosynthetic limitations by separating the relative controls on A resulting from S L , mesophyll conductance (MC L ) and biochemical limitations (B L ) 22 . This method has been successfully applied for evaluating the relative control of leaf A under water stress and during their recovery processes, among inter- and intra-species 13 , 23 , 24 , 25 . It showed not only great potential for elucidating the magnitude changes of limitations and their dominance in photosynthetic restraints with increasing severity of K deficiency, but also revealing the corresponding critical K concentrations for their transformation.

Winter oilseed rape ( Brassica napus L.), a model-plant of winter cover crops who needs substantial amount of potassium to growth was used for a deeply aggregate analysis of K deficiency on photosynthetic limitations 26 . However, malfunction of physiological processes like photosynthesis is hard to be affected when K concentration above the threshold value (1.5% in dry matter, or less) 10 . On consideration of the fact that potassium deficiency symptoms, characterized by a chlorosis and even scorch around the periphery can be obviously observed when leaf K concentration below 1.0% in most species 27 . And the withdrawal K initially occurred at the edge of leaf tip, as tip cells are initially proliferated and oldest 28 , resulting in different K levels as well as visible distinctions between centers and margins. These natural K gradients are therefore precious for photosynthetic limitation analysis, from which we may seek out the main limiting factors under variable leaf K status and the corresponding threshold values. This phenomenon occurred more frequently under a complex biological and abiological environment system during a long-time and low-temperature wintertide, which may conducive to generate a physiology K deficiency in a K-deficient soil, i.e., it may bring K function into full play 29 , 30 . Accordingly, the objectives of the present study were to: (1) estimate the differences of contributions for three limiting factors to photosynthesis between leaf margins and leaf centers, (2) uncover the relationships between photosynthetic limitations and diminishing leaf K status and therefore the critical K concentration for the predominate restraint transformation, (3) reveal the mechanism underlying the K-induced variation of limiting factors. It is hoped that this research will facilitate a better understanding of the photosynthetic physiological mechanism by which potassium deficiency leads to growth retardation in oilseed rape.

Plant performance, leaf K concentration and net photosynthesis

The total dry matter of the –K treatment decreased significantly by 29.9% on average versus the +K treatment ( Table 1 ). The leaf expansion was also restrained, with a 22.1% and 18.0% decline in the individual leaf dry matter and leaf area, respectively. Leaf K concentration was dramatically influenced by potassium supply and leaf position, which was significantly lower in the –K treatment than in the +K treatment. Meanwhile, within an individual leaf, K concentration was remarkably lower in margins than in centers. The mean net photosynthesis ( A ) in the leaf margins of the –K treatment was 56.9% that of the +K treatment. However, there was no significant difference between leaf margins and centers under the –K treatment, as well as the two positions under the +K treatment.

Stomatal conductance

Potassium deficiency led to a significant decline of the mean stomatal conductance ( g s ) in leaf margins, which was 63.6% that of the +K treatment. However, the mean g s value of the leaf centers was not influenced by K nutrient ( Table 2 ). There was a significantly lower g s in leaf margins than in leaf centers under the –K treatment, whilst the g s values of these two positions were the same under the +K treatment. Despite a decrease of the g s value in leaf margins of the –K treatment, the intercellular CO 2 concentrations ( C i ) value was raised and the mean C i were similar to those of other groups.

Potassium supply and leaf position had no effects on stomatal frequency and stomatal length ( Table 2 ). However, stomatal width was significantly decreased in the –K treatment, especially in the leaf margins where the width decreased by 20.9% as compared with the +K treatment. Stomatal pore area was therefore considerably decreased due to K deficiency, particularly in the leaf margins with a 28.0% decline in the single stomatal pore area. Nevertheless, the stomatal length and width as well as the stomatal pore area showed no significant difference in these two positions under the +K treatment.

Mesophyll conductance

Despite a dramatic decrease in the mean mesophyll conductance ( g m ) in the leaf margins of the –K treatment, the mean chloroplastic CO 2 concentrations ( C c ) was 9.4% higher than that of the +K treatment ( Table 3 ). The mean g m and C c values were similar in the leaf centers of different K treatments, as well as between the two positions in the +K treatment. Potassium niutrient and leaf position did not affect mean intercellular CO 2 compensation point ( C i * ) and mitochondrial respiration rate in the light ( R d ), however, the mean chloroplastic CO 2 compensation point (Γ*) was significantly increased in leaf margins of the –K treatment, but showed no statistical differences among the other three groups.

Biochemical characteristics

The mean maximum rate of electron transport ( J max ) and maximum rate of carboxylation ( V c,max ) in the leaf margins of the –K treatment were the lowest and minor changes were observed among the other three treatments ( Table 4 ). However, the mean J max / V c,max in leaf margins of the –K treatment was dramatically increased compared with the mean values of the other three groups in the range from 1.42 to 1.46. The variation of photosynthetic parameters was verified by chemical analyses ( Table 4 ). A significant decline of leaf chlorophyll concentration was found in the –K treated leaves, especially in the leaf margins, with a 31.1% decrease. Furthermore, Rubisco activity was dramatically decreased in leaf margins of the –K treatment, but it was the same in the leaf centers of the –K treatment and the two positions of the +K treatment. Potassium deficiency caused severe ROS production in leaf margins where O 2 .− generation rate increased by 22.8% and meanwhile, POD activity increased by 25.5%.

The relationship between relative A , g s and g m with leaf K concentration

A significant curvilinear relationship between relative A , g s or g m and leaf K concentrations is shown in Fig. 1 . The relative values increased with increasing leaf K concentration and remained stable when the leaf K concentration was beyond a certain concentration. Here a photosynthesis-based concentration threshold with the relative values reaching 95.0% of the maxima was defined. The relative A values increased rapidly with increasing leaf K concentration when it was less than 1.07% ( Fig. 1a ) and varied little when the leaf K concentration was above 1.07%. Therefore, the K concentration (1.07%) was used to evaluate the relative g s and g m ( Fig. 1b,c ) and the calculated result (93.7% and 94.3%) was close to 95.0%, indicating that this threshold value was acceptable for g s and g m .

Relationship between relative photosynthetic parameters and leaf K concentration.

Relative ( a ) photosynthesis rate ( A ), ( b ) stomatal conductance ( g s ), ( c ) mesophyll conductance ( g m ). The values were the relative proportion of measured values over the mean values of K-sufficient leaf centers. Each point represents one leaf measurement. Open triangles and closed triangles represent the values of leaf margin and leaf center under the –K treatment, while open circles and closed circles represent those of the +K treatment. Equations, regression coefficients and significance are shown when P  ≤ 0.05 (* P  ≤ 0.05; ** P  ≤ 0.01).

Quantitative limitation analysis

The restrictions of A max in the –K leaves posed by stomatal (S L ), mesophyll conductance (MC L ) and biochemical limitations (B L ) are presented in Fig. 2a . In symptomatic margins, total limitations reached a value of 46.9% and the contribution of S L , MC L and B L represented 23.9%, 33.0% and 43.1% of total limitations, respectively. By contrast, despite the relatively low limitation (4.8%) in the leaf center, MC L contributed a primary limitation to A max . Accordingly, the dominant limitations changed from symptomatic leaf margins to centers. The relationship between relative limitations and leaf K concentrations were further analyzed ( Fig. 2b ). All the limitations declined precipitously with the leaf K concentration increased from 0.6 to 1.07% (according to the K-based concentration threshold), particularly the B L with the maximum slope of the fitted curve, but they gradually decrease as leaf K concentration continues to increase. Their relative contribution also varied with the change of the leaf K status. While MC L largely predominated at the leaf K concentration of less than 1.07%, B L replaced it when the K concentration was below 0.78% (leaf K concentration of the intersection point between B L and MC L fitted curves).

Photosynthetic limitations and their response to leaf K concentration.

( a ) Quantitative limitation analysis of photosynthetic CO 2 assimilation in leaf margins and centers under the –K treatment. Values are mean ± SE of four replicates per position. The open, gray and dark gray bars represent the percentages of stomatal (S L ), mesophyll conductance (MC L ) and biochemical (B L ) limitations, respectively. ( b ) Relationships between limitations and leaf K concentration. Each point with the same shape represents a single leaf (n = 16). The symbols are as follows: S L , closed squares; MC L , open circles; B L , closed triangles. Solid, dash and dot lines are regression curves of S L , MC L and B L , respectively. Equations, regression coefficients and significance are shown when P  ≤ 0.05 (* P  ≤ 0.05; ** P  ≤ 0.01).

Limitations imposed by K deficiency occur at the same time

In the present study, A in leaf margins were weakened by K deficiency. Generally, the declining A is considered to be limited by stomatal and mesophyll resistances to CO 2 diffusion and biochemical obstacles 13 , 22 . Here we demonstrated that g s , g m and biochemical activities were profoundly restricted as A down-regulated. Stomatal conductance which determine the vital step of CO 2 diffuse from the atmosphere to the interior of leaf was markedly decreased in –K leaf margins, as reported for Eucalyptus grandis 1 , Gossypium hirsutum 17 and Oryza sativa 3 . This is mainly because the lack of vacuole K to keep stomatal aperture by providing driving force to promote water inpour into the guard cell vacuole 31 . The declined A , to a certain extent, revealed that the K in cytoplasm identified as biochemical functional component was below the critical value 10 . Therefore, malfunction of physiological process could come with limited A .

Likewise, g m was decreased in parallel with A . Indeed, g m might be down-regulated by increasing leaf dry mass per area ( M A ) 7 , 23 , however, in the present study, there was no remarkable difference in M A between the –K and +K leaves ( Table 5 ; Supplementary Fig. S1 ). Cell wall thickness ( T cell-wall ) and surface area of chloroplasts exposed to intercellular airspaces ( S c / S ) are reported to be the most substantial anatomical traits in determining g m 23 , 32 . However, significant differences in mesophyll cell wall surface area exposed to intercellular airspace per leaf area ( S m / S ) and S c / S , but not T cell-wall between leaf margins of two K treatments were observed ( Table 5 ). Besides, chloroplast size 33 has also been proved to influence g m . In the present study, though the chloroplast length ( L chl ) decreased under lowest K status, the thickness ( T chl ), surface area ( S chl ) and volume ( V chl ) of chloroplast were largely increased, however, the S chl / V chl was smaller ( Fig. 3 ). The chloroplast enlarging under lowest K concentration was not completely same to that discovered under low nitrogen conditions 33 , 34 . The increase of T chl was more likely to be based on the sacrifice of length owing to roughly circular envelope ( Fig. 3a,b ; Supplementary Fig. S2 ). Mathematically, ellipsoidal chloroplasts, combining with an increscent chloroplast number (see Supplementary Fig. S3 ) were more probably to have longer length of chloroplasts facing the cell wall than swollen even sphere envelopes. Furthermore, the resistance along diffusion pathway length in cytoplasm (distance of chloroplast from cell wall, D chl-cw ) and stroma (taken as half of the chloroplast thickness) account for 10–50% of g m limitation 23 , which however, reported only up to 22% of liquid phase resistance ( r liq ) by Evans et al. in 1994 35 . Low K status strongly increased T chl and D chl-cw ( Fig. 3b,f ), accordingly, the corresponding resistance would be increased. It is therefore proved that the decreased g m is primary due to the reduced S c / S and larger cytosol diffusion resistance but not T cell-wall . More evidences may seek from the influence of K on plasma membrane and chloroplast envelope conductance 32 , carbonic anhydrase and aquaporins that participated in determination of g m 14 , 15 , 23 .

The relationship between chloroplast characteristics and leaf K concentration.

( a ) Chloroplast length ( L chl ), ( b ) thickness ( T chl ), ( c ) surface area ( S chl ), ( d ) volume ( V chl ), ( e ) S chl / V chl , ( f ) distance of chloroplast from the cell wall ( D chl-cw ). Values are mean ± SE of four replicates for K concentration and at least thirty replicates for microstructure parameters. Regression coefficients and significance are shown when P  ≤ 0.05 (* P  ≤ 0.05; ** P  ≤ 0.01).

It should be noted that A , g s or g m started to decline almost at the same time with an extremely similar leaf K status. By another way, the quantitative analysis of limitations indicated that three limiting factors coexist when K concentration below 1.07%. This was similar to the results reported by Grassi and Magnani 22 and Tezara et al. 36 in plants suffering from water stress. However, the investigation carried out by Galmés et al. 13 revealed that B L of Hypericum balearicum and Phlomis italica still remained zero under mild water stress even if the total limitation reached 20–30%. The present finding highlights that all photosynthetic limitations simultaneously occur when leaf is in a physiological K-deficiency state.

Limitations vary with increasing K deficiency intensity

The leaf K concentration threshold value observed in this study was 1.07%, in consistent with the range of 0.5 to 2.0% reported by Leigh and Wyn Jones 37 . Quantitative limitation analysis gives insight into the contributions of different photosynthetic limitations, revealing that the B L and MC L accounted for the majority of total limitations in K-starved leaf margins and centers, respectively. This is mainly ascribed to the discrepancy of relative severity of K deficiency 5 , 6 . As has been stated in the previous studies that some irreversible damages, such as impaired ATP synthesis, depressed Rubisco activity and cell damage occurred when the limiting A , for the most part, is attributed to B L 3 , 38 . Some of which were verified in the present study, such as degraded chloroplast, limited photoassimilate transportation (see Supplementary Fig. S2 ) and increased O 2 .− generation rate under severe K deficiency. The obstacle of these physiological processes alleviated as K deficient stress mitigating, however, the role of MC L on A began to stand out.

The relationship between relative limitations and leaf K concentration verified that, at a leaf K concentration of less than 1.07%, MC L represented the main component of limitations, but B L replaced it when leaf K concentration below 0.78%. This pattern, to a lesser extent, could be found in plants suffering from water stress which suggested that the variation of limitations depends on the stress intensity and duration 22 . Regrettably, the present study failed to reveal whether or not there is a critical concentration in the shifting process from S L predominance to MC L predominance. Further studies focusing on the photosynthetic limitations of rapeseed leaves subjected to a serial K gradient may help to elucidate this issue.

Study site and growth conditions

A field experiment was conducted in Wuxue county, Hubei province, central China (30° 06′46″N, 115° 36′9″E) during the 2013–2014 oilseed rape growing season. The mean temperature of the season was 13.8 °C and the average temperature during winter (from December 2013 to February 2014) was 5.9 °C. The total precipitation during oilseed rape cropping season was 660.7 mm, with wintertide accounting for 26.1% of the total. The soil was a sandy loam with pH 5.3, organic matter 30.5 g kg −1 , total N 1.7 g kg −1 , NH 4 OAc-K 42.5 mg kg −1 , Olsen-P 15.7 mg kg −1 and hot-water soluble B 0.78 mg kg −1 in the topsoil layer (0–20 cm). As stated by Zou, the soil belongs to a K-deficient type, which would cause yield reduction without K fertilizer addition 4 .

Experimental design

A complete randomized block design was set up with two treatments and four replicates. The treatments were: (1) Sufficient K supply treatment (+K), with a K fertilizer recommendation rate of 120 kg K 2 O ha –1 which was tested and well-proved to ensure the optimal growth and yield formation of oilseed rape based on field experiments in this region 39 . (2) K deficiency treatment (–K), with no K fertilizer applied throughout the growing season.

To ensure that nutrients other than K did not limit plant K uptake, 180 kg N ha −1 , 90 kg P 2 O 5 ha −1 and 1.6 kg B ha −1 were applied for these two treatments. The N, P, K, B fertilizers used in the experiment consisted of urea (46% N), superphosphate (12% P 2 O 5 ), potassium chloride (60% K 2 O) and borax (10.8% B). The N fertilizer was applied in three splits: 60% prior to transplanting, i.e., BBCH (Biologische Bundesantalt, Bundessortenamt and Chemische Industrie) 15–16 40 , 20% at the over-wintering stage (i.e., BBCH 29) and 20% at the initiation of stem elongation (i.e., BBCH 30). Besides, all the P, K, B fertilizers were applied as basal fertilizers. The experimental field was plowed and leveled with a rotary tiller and basal fertilizers were incorporated during the process. The plot measured 20 m 2 with a length of 10 m and a width of 2 m.

The oilseed rape cultivar was Zhongshuang 11, supplied by Oil Crops Research Institute of the Chinese Academy of Agricultural Sciences. Rapeseeds were sown in prepared seedbeds on 16 September 2013 and then, on 22 October, about 36 d after sowing, oilseed-rape seedlings with five to six leaves (i.e., BBCH 15–16, 3–4 g dry weight plant −1 ) were uniformly selected and transplanted by hand in double rows spaced approximately 0.3 m apart with 0.2–0.3 m between plants, corresponding to 112 500 plants ha –1 . The oilseed rape was grown under rain-fed conditions. Meanwhile, weeds, pests and disease stresses were controlled by spray herbicides, insecticide and fungicide so that no obvious weeds, insect pests and diseases infestation occurred during cropping season.

Plant and leaf tagging

There was an obvious phenotypic difference in plants between the –K and +K treatments 60 d after transplanting. The discrepancy was highlighted in the fifth to ninth fully expanded leaves (with a total average of 9 fully expanded leaves (i.e., BBCH 19) in both treatments) from apex downwards, specifically, obvious etiolation symptoms around the periphery in K-deficient leaves and asymptomatic leaves in the +K treatment. For each treatment, 24 fifth fully expanded leaves and six uniform plants were tagged in each of the four replicate plots for destructive and non-destructive analysis described later in the methods.

Leaf gas exchange and fluorescence measurements

Leaves under the –K and +K treatments.

The vertical dash lines divide leaf into two parts, the leaf apexes are separated into leaf margins and leaf centers by half-elliptic lines. White circles indicate the gas exchange measuring positions.

A / C i curves were measured on the two positions that had been previously acclimated to saturating light conditions for 20 min. The CO 2 concentration ( C a ) in the gas exchange chamber was reduced stepwise from 400 to 300, 250, 200, 150, 100, 50 μmolCO 2 mol −1 and then increased from 50 to 400, 600, 800, 1000, 1200, 1500, 1800 μmol CO 2 mol −1 at a constant PPFD of 1200 μmol m −2 s −1 at 25 ± 0.2 °C and 50–60% relative humidity. In all cases, the parameters were recorded after the gas exchange rate stabilized at the given C a . At least four leaves were performed in each treatment.

The actual photochemical efficiency of photosystem II (Φ PSII ) was then determined as follows 41 :

The electron transport rate ( J ) can be calculated as:

Where α is the leaf absorptance and β is the fraction of light distributed to PSII. As routinely assumed, α was taken as 0.85 42 , 43 and β was taken as 0.5 44 , 45 . A sensitivity analysis of J biases resulting from rough assumption of α and β on g m variations was also conducted (See Supplementary Table S5 ).

Mesophyll conductance was estimated according to Harley et al. from combined gas exchange and chlorophyll fluorescence measurements 46 .

where A , C i and J were determined as previously described for each treatment, mitochondrial respiration rate in the light ( R d ) and the intercellular CO 2 compensation point ( C i * ) were measured by Laisk method, as described by Brooks and Farquhar 47 . Briefly, the A / C i curves generated with PPFD values of 75, 150, 500 μmol m −2 s −1 , respectively, with each having five different C a in chamber (i.e. 50, 80, 100, 120 and 150 μmol CO 2 mol −1 ). A linear regression was then fitted to each A / C i curve. The x -axis and y -axis of intersection point of three A / C i curves were defined as C i * and R d 48 . The Γ * is the chloroplastic CO 2 photocompensation point calculated from C i * and R d as:

Therefore, A - C i curves were converted into A - C c curves. On the basis of C c , the maximum rate of Rubisco-catalysed carboxylation ( V c, max ) and the maximum rate of electron transport ( J max ) as defined by Farquhar et al. 49 , were calculated 11 , 50 .

Since variable J method is sensitive to many sources of errors, e.g. (1) Γ * and R d biases; (2) a wrong assumption of p 1 and p 2 ; (3) biases in the measurements of C i , A and J , a sensitivity analysis would be great values to improve the confidence in g m estimates and following limitation calculations 51 . Following the method of Harley et al. 46 , we used actual Γ*, R d and J values calculated in this study and a deviation from the measured values to analyze the effects of Γ * , R d and J on g m estimates (see Supplementary Table S1, S3, S5 ). RuBP regeneration can be limited by either insufficient NADPH or ATP, according to Farquhar model, A and J can be linked as follows:

For insufficient NADPH, p 1  = 4 and p 2  = 8; for insufficient ATP, p 1  = 4.5 and p 2  = 10.5 or p 1  = 4 and p 2  = 9.33. Finally, the sensitivity analysis for photosynthetic limitations was conducted basing on these calculated g m values (see Supplementary Table S2, S4, S6 ). The analysis showed that the g m was significantly affected by varying Γ * and R d (see Supplementary Table S1 ), p 1 and p 2 inputs (see Supplementary Table S3 ) and J biases (see Supplementary Table S5 ). However, the g m variation derived from Γ * , R d , J , p 1 and p 2 biases, did not cause profound effects on photosynthetic limitations (see Supplementary Table S2, S4, S6 ). In addition, g m appears to be strikingly affected by C i 12 , 14 , 51 , nevertheless, the similar C i in different treatments and positions here seems to have no impact on g m ( Table 3 ). Therefore, the results obtained was unlikely to be altered by these methodological artifacts.

Plant dry matter, leaf area and dry matter

Six tagged leaves and six tagged plants in each plot were used to determine the individual leaf area, dry matter and total dry matter. Each leaf was digitally scanned using an Epson ES-1200C scanner (Epson, Long Beach, CA, USA) and the area determined using ImageJ software (National Institutes of Health, Bethesda, Maryland, USA) 1 . Individual leaf dry matter and total dry matter were weighed after oven drying at 65 °C for 48 h.

Biochemical analysis

Twelve tagged leaves per plot were picked immediately after the determination of photosynthesis. They were divided into two parts along vertical dashed lines ( Fig. 4 ), followed by dissecting the leaf apexes into leaf margins and leaf centers and removing all the veins. A portion of segments were immersed in liquid N and then stored at −78 °C and the rest were used for leaf K concentration determination. There were four replications for biochemical determinations.

Leaf segments (2 g) were oven dried at 65 °C for 48 h. After that, about 0.15 g dried leaves were milled and digested with H 2 SO 4 -H 2 O 2 as described by Thomas et al. 52 and K concentration in digestion solution was measured by a flame photometer (M-410, Cole-Parmer, Chicago, IL, USA).

The Rubisco extracts were prepared according to Weng et al. with minor modifications 3 . Briefly, leave segments (0.2 g) were ground to a powder using a chilled mortar and pestle with liquid N 2 and a small amount of quarzsand, followed by homogenization with 4 mL pre-cooled extraction buffer containing 50 mM Tris-HCl (pH 7.5), 1 mM EDTA,10 mM MgCl 2 , 12.5% (v/v) glycerol, 10 mM (v/v) β-mercaptoethanol and 1% (w/v) PVP-40 (soluble PVP) at 0–4 °C. The homogenate was centrifuged for 15 min at 15 000  g at 4 °C and then the supernatant was immediately used to determine the activity of ribulose-1, 5-bisphosphate carboxylase/oxygenase (Rubisco, EC 4.1.1.39) by an enzyme-linked immunosorbent assay method with a RuBPcase ELISA kit (CK-E91697P, Shanghai jijin Chemistry and Technology Co., Ltd, China) according to the manufacturer’s instructions. The chlorophyll concentration was determined according to the method of Huang et al. 53 .

Superoxide radical O 2 .− production rate was measured by monitoring the nitrite formation from hydroxylamine in the presence of O 2 .− according to Elstner and Heupel 54 . A 0.5 g aliquot of leaf margins and centers was ground and homogenized in 5 mL of 65 mM pre-cooled phosphate buffer (pH 7.8), followed by centrifuging the homogenate at 10,000 g for 15 min at 4 °C and mixing 0.5 mL of the supernatant with phosphate buffer (0.5 mL) and 0.1 mL of 10 mM hydroxylamine hydrochloride. This mixture was incubated at 25 °C for 20 min, followed by the addition of 1 mL of 58 mM sulfanilic acid and 1 mL of α-naphthylamine and then another 20 min incubation at 25 °C. The as-prepared solution was shaken with equal volume of ether, followed by centrifuging the mixture at 10,000 g for 3 min and measuring the absorbance of the pink water phase at 530 nm. The activity of POD (EC 1.11.1.7) was determined using the guaiacol oxidation method 55 .

Anatomical analysis

Another six tagged leaves per treatment were collected and removed all the veins for anatomical analysis. The stomatal size and frequency were measured in six sub-samples either for leaf margin or center. The materials were prepared as described by Meng et al. 56 . Briefly, leaf samples (about 1cm in length and 1 cm in width) were fixed in 2.5% glutaraldehyde (v/v) at 4 °C for 2 h and washed twice in 0.1 M phosphate buffer (pH 6.8). Next, they were sequentially dehydrated in ethanol (30%, 50%, 70%, 80%, 90%, 95% and 100%) for 10 min at each gradient concentration, with 100% ethanol repeated twice. After further drying and spraying with gold, the as-treated leaf samples were observed and photographed with a scanning electron microscope (JSM-5310LV, Jeol Co, Tokyo, Japan). Images were taken of the lower leaf surface for five microscope fields per sub-sample at a magnification of ×500. The number of stomata was counted in each field (a total of 20 measurements of stomatal frequency for each position) as described by Battie-Laclau et al. 1 and the stomatal frequency was calculated by dividing the stomatal count by the area of the field of view 57 . Moreover, the length and width of ten stomata selected at random were measured in each field. Assuming the stomatal pore as an ellipse, the total stomatal pore area was calculated (stomatal frequency × π × 0.25 × stomatal length × stomatal width).

Leaf segments (1–2 mm 2 ) were cut from each part and fixed with 2.5% glutaraldehyde (v/v) in 0.1 M phosphate buffer (pH 7.4) for 4 h, followed by washing twice in the same buffer for 30 min and postfixing with 2% osmium tetroxide for 4 h at 4 °C. Next, the samples were dehydrated with an ethanol series (10–100%) and in propylene oxide, followed by embedding them in Epon 812 resin.

For the light microscope observation, they were cut into 1 μm transverse sections by LKB-5 ultramicrotome 359 (LKB Co., Ltd., Uppsala, Sweden) and stained with 0.5% toluidine blue. Micrographs were captured at a magnification of ×400 with a Nikon Eclipse E600 microscope equipped with a Nikon 5 MP digital microscope camera DS-Fi1 (Nikon Corporation, Kyoto, Japan). There were four samples per treatment. For each samples, three cross-sections were chosen to measure their thickness ( T leaf ), mesophyll cell wall surface area exposed to intercellular airspace per leaf area ( S m / S ) and surface area of chloroplasts exposed to intercellular airspaces ( S c / S ) according to Tosens et al. (2012) 32 .

Where L mes and L c are the length of mesophyll cell wall exposing to intercellular air space and chloroplast surface area touching the intercellular air space. W is the width of measured cross-section. F is the curvature correction factor which was obtained as the weight average of palisade and spongy mesophyll.

For the ultrastructural observations, ultrathin sections (90 nm) were examined with a transmission electron 360 microscope (H-7650, Hitachi, Japan) after staining with 2.0% uranyl acetate (w/v) and lead citrate. Cell wall thickness ( T cell-wall ), chloroplast length ( L chl ) and thickness ( T chl ) were measured from at least 30 chloroplasts. Chloroplasts were assumed as ellipsoids and chloroplast surface area ( S chl ) and volume ( V chl ) were calculated according Cesaro formula 58 :

The limitations (stomatal limitations, S L ; mesophyll conductance limitations, MC L ; biochemical limitations, B L ) imposed by K deficiency on photosynthesis were investigated by analyzing the leaf margins and centers under two treatments using the quantitative limitation analysis method proposed by Grassi and Magnai 22 . Relative changes in light-saturated assimilation is expressed in terms of relative changes in stomatal, mesophyll conductance and biochemical capacity as Equation (11) .

where l s , l mc and l b are the corresponding relative limitations calculated as Eqns from (12) to (14), g sc is stomatal conductance to CO 2 ( g s /1.6) and V c,max is maximum rate of carboxylation estimated from A - C i curve.

Then the relative change of A , g sc , g m and V c,max in Equation (11) can be approximated by Chen et al. 59 .

Statistical analysis

One-way analysis of variance ( ANOVA ) was caiculated usig SPSS 18.0 software (SPSS, Chicago, IL, USA). The mean values were compared using the least significant difference (LSD) test ( P  < 0.05). Graphics and regression analysis were performed using the OriginPro 8.5 software (OriginLab Corporation, Northampton, MA, USA).

Additional Information

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Acknowledgements

This work was supported by the earmarked fund for China Agriculture Research System (CARS-13); the Special Fund for Agro-scientific Research in the Public Interest (201203013); and the PhD Candidate Research Innovation Project of Huazhong Agricultural University (2014bs17).

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Zhifeng Lu, Tao Ren, Yonghui Pan, Xiaokun Li, Rihuan Cong & Jianwei Lu

Key Laboratory of Arable Land Conservation (Middle and Lower Reaches of Yangtze River) Ministry of Agriculture, Wuhan, 430070, China

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Z.L., T.R. and J.L. conceived and designed the experiment. Z.L. and Y.P. performed the experiments, Z.L. analyzed the data, wrote the main manuscript text and prepared all of the figures and Z.L., T.R., Y.P., X.L., R.C. and J.L. reviewed and approved the manuscript.

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Lu, Z., Ren, T., Pan, Y. et al. Differences on photosynthetic limitations between leaf margins and leaf centers under potassium deficiency for Brassica napus L.. Sci Rep 6 , 21725 (2016). https://doi.org/10.1038/srep21725

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Received : 16 September 2015

Accepted : 29 January 2016

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DOI : https://doi.org/10.1038/srep21725

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Investigating the Rate of Photosynthesis ( AQA A Level Biology )

Revision note.

Biology & Environmental Systems and Societies

Apparatus & Techniques: Investigating the Rate of Photosynthesis

• Investigations to determine the effects of light intensity, carbon dioxide concentration and temperature on the rate of photosynthesis can be carried out using aquatic plants , such as Elodea or Cabomba (types of pondweed )
• Light intensity – change the distance ( d ) of a light source from the plant (light intensity is proportional to 1/ d 2 )
• Carbon dioxide concentration – add different quantities of sodium hydrogencarbonate (NaHCO 3 ) to the water surrounding the plant, this dissolves to produce CO 2
• Temperature (of the solution surrounding the plant) – place the boiling tube containing the submerged plant in water baths of different temperatures
• For example, when investigating the effect of light intensity on the rate of photosynthesis, a glass tank should be placed in between the lamp and the boiling tube containing the pondweed to absorb heat from the lamp – this prevents the solution surrounding the plant from changing temperature
• Distilled water
• Aquatic plant, algae or algal beads
• Sodium hydrogen carbonate solution
• Thermometer
• Test tube plug
• This will ensure oxygen gas given off by the plant during the investigation form bubbles and do not dissolve in the water
• This will ensure that the plant contains all the enzymes required for photosynthesis and that any changes of rate are due to the independent variable
• Ensure the pondweed is submerged in sodium hydrogen carbonate solution (1%) – this ensures the pondweed has a controlled supply of carbon dioxide (a reactant in photosynthesis)
• Cut the stem of the pondweed cleanly just before placing into the boiling tube
• Measure the volume of gas collected in the gas-syringe in a set period of time (eg. 5 minutes)
• Change the independent variable (ie. change the light intensity, carbon dioxide concentration or temperature depending on which limiting factor you are investigating) and repeat step 5
• Record the results in a table and plot a graph of volume of oxygen produced per minute against the distance from the lamp (if investigating light intensity), carbon dioxide concentration, or temperature

The effect of light intensity on an aquatic plant is measured by the volume of oxygen produced

Results - Light Intensity

• The closer the lamp, the higher the light intensity (intensity ∝ 1/ d 2 )
• Therefore, the volume of oxygen produced should increase as the light intensity is increased
• This is when the light stops being the limiting factor and the temperature or concentration of carbon dioxide is limiting the rate of photosynthesis
• The effect of these variables could then be measured by increasing the temperature of water (by using a water bath) or increasing the concentration of sodium hydrogen carbonate respectively
• Rate of photosynthesis = volume of oxygen produced ÷ time elapsed

Limitations

• Immobilised algae beads are beads of jelly with a known surface area and volume that contain algae, therefore it is easier to ensure a standard quantity
• Immobilised algae beads are easy and cheap to grow, they are also easy to keep alive for several weeks and can be reused in different experiments
• The method is the same for algae beads though it is important to ensure sufficient light coverage for all beads

Light intensity – the distance of the light source from the plant (intensity ∝ 1/ d 2 )

Temperature - changing the temperature of the water bath the test tube sits in

Carbon dioxide - the amount of NaHCO 3 dissolved in the water the pondweed is in

Also remember that the variables not being tested (the control variables) must be kept constant.

Required Practical: Affecting the Rate of Dehydrogenase Activity

• The light-dependent reactions of photosynthesis take place in the thylakoid membrane and involve the release of high-energy electrons from chlorophyll a molecules
• These electrons are picked up by the electron acceptor NADP in a reaction catalysed by dehydrogenase
• However, if a redox indicator (such as DCPIP or methylene blue ) is present, the indicator takes up the electrons instead of NADP
• DCPIP: oxidised ( blue ) → accepts electrons → reduced ( colourless )
• Methylene blue: oxidised ( blue ) → accepts electrons → reduced ( colourless )
• The colour of the reduced solution may appear green because chlorophyll produces a green colour
• When light is at a higher intensity, or at more preferable light wavelengths, the rate of photoactivation of electrons is faster, therefore the rate of reduction of the indicator is faster

Light activates electrons from chlorophyll molecules during the light-dependent reaction. Redox indicators accept the excited electrons from the photosystem, becoming reduced and therefore changing colour.

• Isolation medium
• Pestel and mortar
• Aluminium Foil

Method - Measuring light as a limiting factor

• This produces a concentrated leaf extract that contains a suspension of intact and functional chloroplasts
• The medium must have the same water potential as the leaf cells so the chloroplasts don’t shrivel or burst and contain a buffer to keep the pH constant
• The medium should also be ice-cold (to avoid damaging the chloroplasts and to maintain membrane structure)
• The room should be at an adequate temperate for photosynthesis and maintained throughout, as should carbon dioxide concentration
• If different intensities of light are used, they must all be of the same wavelength (same colour of light) - light intensity is altered by changing the distance between the lamp and the test tube
• If different wavelengths of light are used, they must all be of the same light intensity - the lamp should be the same distance in all experiments
• DCPIP of methylene blue indicator is added to each tube, as well as a small volume of the leaf extract
• A control that is not exposed to light (wrapped in aluminium foil) should also be set up to ensure the affect on colour is due to the light
• This is a measure of the rate of photosynthesis
• A graph should be plotted of absorbance against time for each distance from the light
• This is because the lowered light intensity will slow the rate of photoionisation of the chlorophyll pigment, so the overall rate of the light dependent reaction will be slower
• This means that less electrons are released by the chlorophyll, hence the DCPIP accepts less electrons. This means that it will take longer to turn from blue to colourless
• A higher rate of decrease, shown by a steep gradient on the graph, indicates that the dehydrogenase is highly active.
• This experiment is not measuring the rate of dehydrogenase activity directly (through measuring the rate of substrate use or product made) but is instead predicting what the rate would be by measuring the rate of electron transfer from the photosystems
• It is therefore important to control the amount of leaf used to produce the chloroplast sample and also how much time is spent crushing the leaf to release the chloroplast
• It is also a good idea to measure a specific wavelength absorption by each sample on the colorimeter before and after the experiment so you can get a more accurate change in oxidised DCPIP concentration
• Results should also be repeated and the mean value calculated
• The time taken to go colourless is subjective to each person observing and therefore one person should be assigned the task of deciding when this is

In chemistry the acronym ‘OILRIG’ is used to remember if something is being oxidised or reduced. Oxidation Is Loss (of electrons) and Reduction Is Gain (of electrons). Therefore the oxidised state is when it hasn’t accepted electrons and the reduced state has accepted electrons.

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Author: Alistair

Alistair graduated from Oxford University with a degree in Biological Sciences. He has taught GCSE/IGCSE Biology, as well as Biology and Environmental Systems & Societies for the International Baccalaureate Diploma Programme. While teaching in Oxford, Alistair completed his MA Education as Head of Department for Environmental Systems & Societies. Alistair has continued to pursue his interests in ecology and environmental science, recently gaining an MSc in Wildlife Biology & Conservation with Edinburgh Napier University.

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Increasing global population and climate change uncertainties have compelled increased photosynthetic efficiency and yields to ensure food security over the coming decades. Potentially, genetic manipulation and minimization of carbon or energy losses can be ideal to boost photosynthetic efficiency or crop productivity. Despite significant efforts, limited success has been achieved. There is a need for thorough improvement in key photosynthetic limiting factors, such as stomatal conductance, mesophyll conductance, biochemical capacity combined with Rubisco, the Calvin–Benson cycle, thylakoid membrane electron transport, nonphotochemical quenching, and carbon metabolism or fixation pathways. In addition, the mechanistic basis for the enhancement in photosynthetic adaptation to environmental variables such as light intensity, temperature and elevated CO 2 requires further investigation. This review sheds light on strategies to improve plant photosynthesis by targeting these intrinsic photosynthetic limitations and external environmental factors.

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Acknowledgements

The research was supported by the National Natural Science Foundation of China (31871570) and Sichuan Innovation Team Project of National Modern Agricultural Industry Technology System (SCCXTD-2020-20) and VEGA-1-0589-19 & APVV-18-0465. And Collaborative Innovation Center for Modern Crop Production co-sponsored by Province and Ministry (CIC-MCP), the Science and Technology Department of Zhejiang Province (14th 5-year New Oil Crops Breeding).

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Sajad Hussain and Zaid Ulhassan have contributed equally to this work.

Authors and Affiliations

College of Agronomy, Sichuan Agricultural University, 211-Huimin Road, Wenjiang District, Chengdu, 611130, People’s Republic of China

Sajad Hussain, Wenyu Yang & Weiguo Liu

Key Laboratory of Crop Ecophysiology and Farming System in Southwest China (Ministry of Agriculture), Sichuan Engineering Research Center for Crop Strip Intercropping System, Sichuan Agricultural University, Chengdu, People’s Republic of China

Institute of Crop Science, Ministry of Agriculture and Rural Affairs Laboratory of Spectroscopy Sensing, Zhejiang University, Hangzhou, 310058, People’s Republic of China

Zaid Ulhassan &  Weijun Zhou

Department of Plant Physiology, Slovak University of Agriculture, 94976, Nitra, Slovakia

Marian Brestic & Marek Zivcak

К.A. Timiryazev Institute of Plant Physiology, Russian Academy of Sciences, Botanicheskaya St. 35, Moscow, Russia, 127276

Suleyman I. Allakhverdiev

Department of Plant Physiology, College of Life Sciences, Shandong Agricultural University, Daizong Road No. 61, 271018, Taian, People’s Republic of China

Xinghong Yang

College of Agriculture, University of Sargodha, Sargodha, Punjab, 40100, Pakistan

Muhammad Ehsan Safdar

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Hussain, S., Ulhassan, Z., Brestic, M. et al. Photosynthesis research under climate change. Photosynth Res 150 , 5–19 (2021). https://doi.org/10.1007/s11120-021-00861-z

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Accepted : 28 June 2021

Published : 07 July 2021

Issue Date : December 2021

DOI : https://doi.org/10.1007/s11120-021-00861-z

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• Science is LIT

Explore How Light Affects Photosynthesis

Algae are aquatic, plant-like organisms that can be found in oceans, lakes, ponds, rivers, and even in snow. But don’t worry, if you’re not near a waterway, it can easily be ordered from Amazon or Carolina Biological. Algae range from single-celled phytoplankton (microalgae) to large seaweeds (macroalgae). Phytoplanktons can be found drifting in water and are usually single-celled. They can also grow in colonies (group of single-cells) that are large enough to see with the naked eye. The specific types of algae that can be used in this experiment are  Scenedesmus, Chlamydomonas, or  Chlorella , all of which are phytoplanktons or microalgae. 

Experimental variables

• Color filter paper
• Table/desk lamp
• Light bulbs (varying intensities and colors)

Laboratory Supplies

• Transfer pipettes
• Vials with caps
• Freshwater Algae ( Scenedesmus , Chlorella , or Chlamydomonas )
• Small beakers or cups

Laboratory Solutions

• 2% Calcium Chloride
• 2% Sodium alginate
• Cresol red/thymol blue pH indicator solution

Solution Preparations

2% calcium chloride (cacl 2 ).

• 20 g of CaCl 2
• Fill to 1000 mL with water

2% CaCl 2 is stable at room temperature indefinitely.

2% Sodium alginate (prepared in advance)

• 2 g sodium alginate
• Fill to 100 mL with water

It takes a while for the alginate to go into solution. We recommend to dissolve by stirring using a magnetic stir bar overnight at room temperature. Store at 4 °C for up to 6 months or use immediately.

Cresol red/Thymol blue pH indicator solution (10x)

• 0.1 g cresol red
• 0.2 g thymol blue
• 0.85 g sodium bicarbonate (NaHCO 3 )
• 20 mL ethanol
• Fill to 1L with fresh boiled water

Measure indicators and mix with ethanol. Measure sodium bicarbonate and mix with warm/hot water. Mix the solutions together and fill with remaining freshly boiled water up to 1L final solution. The 10x stock solution is stable for at least a year.

In preparation for doing the experiment, prepare 1x indicator solution by diluting the 10x indicator solution with distilled water (e.g. 20 ml 10x into 200 mL final solution).

Experimental Bench Set-Up

• ~10 mL of 2% CaCl 2 in a cup or beaker
• ~3-5 mL of sodium alginate in cup or beaker
• Cup with ~10 mL of water
• Empty cup or beaker that holds a minimum of 30 mL

Preparing Algae for Experiment

• Prepare a concentrated suspension of algae. Without centrifuge : leave ~50 mL of algae suspension to settle (preferably overnight), then carefully pour off the supernatant to leave ~3-5 mL of concentrated algae. With centrifuge : Centrifuge ~50 mL of algae suspension at low speed for 10 minutes and then carefully pour off the supernatant, leaving behind ~3-5 mL of concentrated algae.
• In a small beaker, add equal volumes of sodium alginate and then add in the concentrated algae. Gently mix algae and sodium alginate together using a transfer pipette until its evenly distributed.
• Using the transfer pipette, carefully add single drops of the algae/sodium alginate mixture into the CaCl 2 to make little “algae balls”
• Once all of the “algae balls” are in the CaCl 2 solution, allow them to harden for 5 minutes
• Place the strainer over the empty cup or beaker, and pour over the entire solution of “algae balls” and CaCl 2 into the strainer allowing the CaCl 2 to pass through, leaving just the algae in the strainer
• Keeping the strainer over the container, pour the water over the “algae balls” to rinse the remain CaCl 2
• Transfer your newly made “algae balls” to a new cup or beaker

Setting up Photosynthesis Experiment

• Distance from light (using ruler) – group can set up vials different distances from one light source
• Different color lights (using color filter paper or different color light bulbs) – group can set up by covering the vials with different colored films and arrange them the same distance away from the light source or set up 1 vial in front of a different colored lamp same distance away.
• With or without light – group places 1 vial in front of an illuminated lamp and another has the vial or lamp covered with black paper the same distance away

• When starting your experiment, be sure to take note of the time that you placed your vial in front of the light source. Vials should be left for ~1-2 hours.

What would happen if the algae photosynthesizes (increase O2) in a solution that started at pH8.2?

Analyzing photosynthesis results

• After 1-2 hours, return to the experiment. Without disturbing the vials, analyze and take pictures of results. Have students write down the time that their experiment ended.
• Using the color chart above, determine which pH matches your sample the closest.
• Have students determine if they got what they expected and discuss amongst their group members.

Explain how the rate of photosynthesis is affected by their different variables.

What were your conclusions from this experiment? If you were to repeat the experiment, what would you change and why? What’s the relationship with O2 and CO2 during the process of photosynthesis? Is there a “best” source of light that allowed the algae to photosynthesize better?

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Experiment on photosynthesis is heading to the space station to explore effects of microgravity

by Tom Rickey, Pacific Northwest National Laboratory

An experiment aimed at learning more about how plants grow in space will be aboard a National Aeronautics and Space Administration launch in early August from the Cape Canaveral Space Force Station in Florida.

A Northrop Grumman Cygnus spacecraft perched atop a SpaceX Falcon 9 rocket will carry the plants to the orbiting laboratory, where astronauts will tend to them before the plants are returned to Earth.

The experiment created by scientists at the Department of Energy's Pacific Northwest National Laboratory will look at how two different types of grass grow on the space station. A PNNL team led by biologist Pubudu Handakumbura designed the experiment and will compare the results from space to identical plants being grown at the Kennedy Space Center.

The study focuses on photosynthesis—how plants take in light and then use it to grow, converting carbon dioxide to sugars and oxygen in the process. The two grass types under study, Brachypodium distachyon and Setaria viridis, use different carbon dioxide -concentrating mechanisms. Handakumbura's team will compare the two methods in a microgravity environment.

While most plants on Earth use a carbon-concentrating mechanism known as C3, there is some evidence that a method known as C4 holds more promise for plant growth in space.

"How will the plants respond in a microgravity environment?" said Handakumbura. "Plants naturally send their roots downward due to gravity. But how will they grow in microgravity? This is important for future deep space exploration, for growing food and supporting life."

The team will monitor three identical sets of plants as they grow for 32 days—two sets at Kennedy Space Center and one set on the space station. Altogether, the experiment includes 288 plants.

On the space station , astronauts will tend to the plants and record how efficiently they are carrying out photosynthesis. After the plants are returned to Earth on a subsequent mission, they will be sent to PNNL, where Handakumbura's team will spend several months analyzing the molecular activity that took place.

The experiments measuring proteins, metabolites and other molecules will be done at the Environmental Molecular Sciences Laboratory, a DOE Office of Science user facility.

Handakumbura's experiment is named Advanced Plant Experiment-09 or APEX-09 . PNNL colleagues Chaevien Clendinen, Summer Duckworth, Kim Hixson, Madeline Southworth and Kylee Tate are also working on the project.

Handakumbura will be on hand to watch the experiment, three years in the making, head into space as part of Northrop Grumman's 21st Commercial Resupply Services Mission.

"I look forward to the knowledge we will uncover from the team-driven science we are conducting with APEX 09," said Handakumbura. "And I am excited to contribute to the foundational research that will shape future plant system designs."

Provided by Pacific Northwest National Laboratory

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