Plant Science

The flux in photosynthesis can be studied by performing ¹³CO₂ pulse labelling and analysing the temporal labelling kinetics of metabolic intermediates using gas or liquid chromatography linked to mass spectrometry. Metabolic flux analysis (MFA) is the primary approach for analysing metabolic network function and quantifying intracellular metabolic fluxes. Different MFA approaches differ based on the metabolic state (steady vs. non-steady state) and the use of stable isotope tracers. The main methodology used to investigate metabolic systems is metabolite steady state associated with stable isotope labelling experiments. Specifically, in biological systems like photoautotrophic organisms, isotopic non-stationary ¹³C metabolic flux analysis at metabolic steady state with transient isotopic labelling (¹³C-INST-MFA) is required. The common requirement for metabolic steady state, alongside its very short half-timed reactions, complicates robust MFA of photosynthetic metabolism. While custom gas chambers design has addressed these challenges in various model plants, no similar tools were developed for liquid photosynthetic cultures (e.g., algae, cyanobacteria), where diffusion and equilibration of inorganic carbon species in the medium entails a new dimension of complexity. Recently, a novel tailor-made microfluidics labelling system has been introduced, supplying short ¹³CO₂ pulses at steady state, and resolving fluxes across most photosynthetic metabolic pathways in algae. The system involves injecting algal cultures and medium containing pre-equilibrated inorganic ¹³C into a microfluidic mixer, followed by rapid metabolic quenching, enabling precise seconds-level label pulses. This was complemented by a ¹³CO₂-bubbling-based open labelling system (photobioreactor), allowing long pulses (minutes–hours) required for investigating fluxes into central C metabolism and major products. This combined labelling procedure provides a comprehensive fluxome cover for most algal photosynthetic and central C metabolism pathways, thus allowing comparative flux analyses across algae and plants.

Metals are essential in many biological processes, including oxygenic photosynthesis. Here we described a method to measure the metal pool in whole cells and thylakoids, including the bioactive pool in intact photosynthetic protein complexes in the model oxygenic cyanobacterium Synechocystis PCC6803. In the first part of the protocol, whole cells and thylakoid membranes are carefully prepared, in which the total metal concentrations are measured by inductively coupled plasma triple-quadrupole mass spectrometry (ICP-QQQ-MS). In the second part of the protocol, isolated thylakoids are solubilized to release the integral membrane proteins and the metal binding protein complexes. These intact photosynthetic protein complexes are subjected to size exclusion chromatography (SEC) and metal binding in the size separated complexes is analyzed by hyphenation with ICP-QQQ-MS.

Chlorophyll fluorescence measurements have been widely used to monitor the condition of photosynthesis. Furthermore, chlorophyll fluorescence from cyanobacteria reflects the condition of respiration, since cyanobacterial photosynthesis shares several components of electron transport chain with respiration. This protocol presents the method to monitor the condition of both photosynthesis and respiration in cyanobacteria simply by measuring chlorophyll fluorescence in the dark and in the light with pulse amplitude modulation (PAM) chlorophyll fluorometer.

Phloem loading and transport of photoassimilate from photoautotrophic source leaves to heterotrophic sink organs are essential physiological processes that help the disparate organs of a plant function as a single, unified organism. We present three protocols we routinely use in combination with each other to assess (1) the relative rates of sucrose (Suc) loading into the phloem vascular system of mature leaves (this protocol), (2) the relative rates of carbon loading and transport through the phloem (Yadav et al., 2017a), and (3) the relative rates of carbon unloading into heterotrophic sink organs, specifically roots, after long-distance transport (Yadav et al., 2017b). We propose that conducting all three protocols on experimental and control plants provides a reliable comparison of whole-plant carbon partitioning, and minimizes ambiguities associated with a single protocol conducted in isolation (Dasgupta et al., 2014; Khadilkar et al., 2016). In this protocol, Arabidopsis leaf disks isolated from mature rosette leaves are infiltrated with a buffered solution containing [¹⁴C]Suc. Suc transporters (SUCs or SUTs) load Suc into the phloem and excess, unloaded Suc in the leaf disk is then washed away. Loading of labeled Suc into the veins is visualized by autoradiography of lyophilized leaf disks and quantified by scintillation counting. Results are expressed as disintegration per minute per unit of leaf disk fresh weight or area.

Phloem loading and transport of photoassimilate from photoautotrophic source leaves to heterotrophic sink organs are essential physiological processes that help the disparate organs of a plant function as a single, unified organism. We present three protocols we routinely use in combination with each other to assess (1) the relative rates of sucrose (Suc) loading into the phloem vascular system of mature leaves (Yadav et al., 2017a), (2) the relative rates of carbon loading and transport through the phloem (Yadav et al., 2017b), and (3) the relative rates of carbon unloading into heterotrophic sink organs, specifically roots, after long-distance transport (this protocol). We propose that conducting all three protocols on experimental and control plants provides a reliable comparison of whole-plant carbon partitioning, and minimizes ambiguities associated with a single protocol conducted in isolation (Dasgupta et al., 2014; Khadilkar et al., 2016). In this protocol, [¹⁴C]CO₂ is photoassimilated in source leaves and phloem loading and transport of the ¹⁴C label to heterotrophic sink organs, particularly roots, is quantified by scintillation counting. Using this protocol, we demonstrated that overexpression of sucrose transporters and a vacuolar proton pumping pyrophosphatase in the companion cells of Arabidopsis enhanced transport of ¹⁴C label photoassimilates to sink organs (Dasgupta et al., 2014; Khadilkar et al., 2016). This method can be adapted to quantify long-distance transport in other plant species.

Phloem loading and transport of photoassimilate from photoautotrophic source leaves to heterotrophic sink organs are essential physiological processes that help the disparate organs of a plant function as a single, unified organism. We present three protocols we routinely use in combination with each other to assess (1) the relative rates of sucrose (Suc) loading into the phloem vascular system of mature leaves (Yadav et al., 2017a), (2) the relative rates of carbon loading and transport through the phloem (this protocol), and (3) the relative rates of carbon unloading into heterotrophic sink organs, specifically roots, after long-distance transport (Yadav et al., 2017b), We propose that conducting all three protocols on experimental and control plants provides a reliable comparison of whole-plant carbon partitioning, and minimizes ambiguities associated with a single protocol conducted in isolation (Dasgupta et al., 2014; Khadilkar et al., 2016). In this protocol, [¹⁴C]CO₂ is photoassimilated in source leaves and phloem loading and transport of photoassimilate is quantified by collecting phloem exudates into an EDTA solution followed by scintillation counting.

This protocol describes a method to extract ribulose-1,5-bisphosphate carboxylase oxygenase (Rubisco) from diatoms (Bacillariophyta) to determine catalytic performance. This protocol has been adapted from use in cyanobacteria and higher plants (Andrews, 1988; Whitney and Sharwood, 2007). First part (steps A1-A3) of the extraction provides a crude extract of Rubisco that is sufficient for carboxylation assays to measure the Michaelis constant for CO₂ (K_C) and the catalytic turnover rate (k_cat^c). However, the further purification steps outlined (steps B1-B4) are needed for measurements of Rubisco CO₂/O₂ Specificity (S_C/O, [Kane et al., 1994]).

Premature leaf senescence induced by drought stress is a main factor for yield losses in barley. Research in drought stress tolerance has become more important as due to climate change the number of drought periods will increase and tolerance to drought stress has become a goal of high interest in barley breeding. However, reliable screening for drought stress tolerance is still a difficult task. This protocol describes the experimental design for the phenotyping for drought stress tolerance and early leaf senescence in the juvenile stage of barley (A) and the determination of six physiological parameters involved in drought tolerance and leaf senescence (B to G) according to Wehner et al., (2015).

Chloroplasts accumulate to weak light and escape from strong light. These light-induced responses have been known from the 19th century (Böhm, 1856). Up to now, many scientists have developed different methods to investigate these dynamic phenomena in a variety of plant species including the model plant Arabidopsis thaliana, a terrestrial dicot (Wada, 2013). Especially, a serial recording to trace the position of individual chloroplast for the analysis of its mode of movement is critical to understand the underlying mechanism. An aquatic monocot Vallisneria (Alismatales: Hydrocharitaceae, Figure 1A) has contributed over a century to such investigation (Senn, 1908; Zurzycki, 1955; Seitz, 1967), because Vallisneria leaves have rectangular parallelepiped-shaped epidermal cells aligned orderly in a monolayer (Figure 1B), providing an excellent experimental system for microscopic studies. Here we describe a protocol for the up-to-date time-lapse imaging procedures to analyze Vallisneria chloroplast movement. Using this and prototype procedures, the relevant photoreceptor systems (Izutani et al., 1990; Dong et al., 1995; Sakai et al., 2015), association with actin cytoskeleton (Dong et al., 1996; Dong et al., 1998; Sakai and Takagi 2005; Sakurai et al., 2005), and regulatory roles of Ca²⁺ (Sakai et al., 2015) have been strenuously investigated.


Figure 1. Vallisneria plant. A. Whole plant body; B. A bright-field image of adaxial epidermal cells containing a large number of chloroplasts; C. Culture facilities.

Chlorophyll fluorescence measurement is a widely used technique to determine photosynthetic performance. Light energy absorbed by a chlorophyll molecule can be dissipated by driving photochemical energy conversion, as heat in non-photochemical quenching processes, or it is re-emitted as fluorescence. The loss of light energy as chlorophyll fluorescence is primarily derived from photosystem II. Photosystem II is a thylakoid-embedded multiprotein complex which provides the high redox potential needed to oxidize water. Within photosystem II photons of light are captured and used to energize electrons. Energized electrons are fed into the linear electron transport chain and photosystem II replenishes lost electrons with electrons from splitting of water. Chlorophyll fluorescence yield can be quantified using a modulated fluorometer device. In such a device, a modulated measuring light beam (switched on and off at a high frequency) and the parallel detection of fluorescence exclusively excited by the measuring light allows chlorophyll fluorescence measurements in the presence of photosynthetic (actinic) light. In addition, high intensity, but short duration light flashes (saturating pulses) are used to determine maximum fluorescence yields in dark and light adapted leaves. In this protocol the procedure to receive a typical fluorescence graph of Arabidopsis wild-type leaves is given. Furthermore, this procedure can be used to identify Arabidopsis mutant plants affecting photosystem II, on the basis of the respective fluorescence graphs and values.