Plant Science

Iron (Fe) is an indispensable micronutrient for plant growth and development. Since both deficiency, as well as a surplus of Fe, can be detrimental to plant health, plants need to constantly tune uptake rates to maintain an optimum level of Fe. Quantification of Fe serves as an important parameter for analyzing the fitness of plants from different accessions, or mutants and transgenic lines with altered expression of specific genes. To quantify metals in plant samples, methods based on inductively coupled plasma-optical emission spectrometry (ICP-OES) or inductively coupled plasma-mass spectrometry (ICP-MS) have been widely employed. Although these methods are highly accurate, these methodologies rely on sophisticated equipment which is not always available. Moreover, ICP-OES and ICP-MS allow for surveying several metals in the same sample, which may not be necessary if only the Fe status is to be determined. Here, we outline a simple and cost-efficient protocol to quantify Fe concentrations in roots and shoots of Arabidopsis seedlings, by using a spectroscopy-based assay to quantify Fe²⁺-BPDS₃ complexes against a set of standards. This protocol provides a fast and reproducible method to determine Fe levels in plant samples with high precision and low costs, which does not depend on expensive equipment and expertise to operate such equipment.

The Xenopus oocyte is a powerful system for the exogenous expression and functional characterization of plant membrane transport proteins. Until now, a number of potassium transporters and channels have been identified in oocytes expression system by the two-electrode voltage clamp technology. It is difficult to characterize K⁺/H⁺ anti-transporters, especially, electroneutral transporter. The K⁺ efflux assay system enables easy, fast, large-scale measurement of the transporters activity without two-electrode voltage clamp technology. This protocol describes a technique to measure the efflux activity of potassium transporter in oocytes expressing system.

Nitric oxide (NO), is a redox-active, endogenous signalling molecule involved in the regulation of numerous processes. It plays a crucial role in adaptation and tolerance to various abiotic and biotic stresses. In higher plants, NO is produced either by enzymatic or non-enzymatic reduction of nitrite and an oxidative pathway requiring a putative nitric oxide synthase (NOS)-like enzyme. There are several methods to measure NO production: mass spectrometry, tissue localization by DAF-FM dye. Electron paramagnetic resonance (EPR) also known as electron spin resonance (ESR) and spectrophotometric assays. The activity of NOS can be measured by L-citrulline based assay and spectroscopic method (NADPH utilization method). A major route for the transfer of NO bioactivity is S-nitrosylation, the addition of a NO moiety to a protein cysteine thiol forming an S-nitrosothiol (SNO). This experimental method describes visualization of NO using DAF-FM dye by fluorescence microscopy (Zeiss AXIOSKOP 2). The whole procedure is simplified, so it is easy to perform but has a high sensitivity for NO detection. In addition, spectrophotometry based protocols for assay of NOS, Nitrate Reductase (NR) and the content of S-nitrosothiols are also described. These spectrophotometric protocols are easy to perform, less expensive and sufficiently sensitive assays which provide adequate information on NO based regulation of physiological processes depending on the treatments of interest.

This protocol provides a detailed description of how to fabricate and use the dual-flow-RootChip (dfRootChip), a novel microfluidic platform for investigating root nutrition, root-microbe interactions and signaling and development in controlled asymmetric conditions. The dfRootChip was developed primarily to investigate how plants roots interact with their environment by simulating environmental heterogeneity. The goal of this protocol is to provide a detailed resource for researchers in the biological sciences wishing to employ the dfRootChip in particular, or microfluidic devices in general, in their laboratory.

Boron (B) is essential for plant growth and taken up by plant roots as boric acid. Under B limitation, B uptake and translocation in plants are dependent on the boric acid channels located in the plasma membrane. Xenopus leavis oocyte is a reliable heterologous expression system to characterize transport activities of boric acid channels and related major intrinsic proteins (aquaporins). Here, we outline the protocols for expression of boric acid channels and boric acid uptake assay in Xenopus leavis oocytes.

To determine boron quantity in soil, water and biological samples, several protocols are available. Colorimetric assays are the simplest and cheapest methods which can be used to determine boron concentration. However, published protocols do not give straightforward guidance for beginners to adopt these protocols for routine use in the laboratory. Based on a previously published available procedure, we present a detailed and modified version of a curcumin based colorimetric protocol to determine boron concentration extracted from any sample. Our modified protocol is able to determine up to 0.2 nmole of Boron in a sample volume of 300 µl.

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.

In ectomycorrhizal plants, the fungal cells colonize the roots of their host plant to create new organs called ectomycorrhizae. In these new organs, the fungal cells colonize the walls of the cortical cells, bathing in the same apoplasm as the plant cells in a space named the ‘Hartig net’, where exchanges between the two partners take place. Finally, the efficiency of ectomycorrhizal fungi to improve the phosphorus nutrition of their host plants will depend on the regulation of phosphate transfer from the fungal cells to plant cells in the Hartig net through as yet unknown mechanisms. In order to investigate these mechanisms, we developed an in vitro experimental device mimicking the common apoplasm of the ectomycorrhizae (the Hartig net) to study the phosphorus metabolism in the ectomycorrhizal fungus Hebeloma cylindrosporum when the fungal cells are associated or not with the plant cells of the host plant Pinus pinaster. This device can be used to monitor ³²Phosphate efflux from the fungus previously incubated with ³²P-orthophosphate.