植物科学

The target of rapamycin (TOR) is a central hub kinase that promotes growth and development in all eukaryote cells. TOR induces protein synthesis through the phosphorylation of the S6 kinase (S6K), which, in turn, phosphorylates ribosomal S6 protein (RPS6) increasing this anabolic process. Therefore, S6K and RPS6 phosphorylation are generally used as readouts of TOR activity. Protein phosphorylation levels are measured by a western blot (WB) technique using an antibody against one specific phosphosite in cell extracts. However, at the tissue/cell-specific level, there is a huge gap in plants due to the lack of alternative techniques for the evaluation of TOR activity as there are for other organisms such as mammals. Here, we describe an in vivo protocol to detect S6K phosphorylation in tissues/cells of model photosynthetic organisms such as Arabidopsis thaliana and Chlamydomonas reinhardtii. Our proposed method consists of the immunolocalization of a phosphorylated target of TOR kinase using a fluorescent secondary antibody by confocal microscopy. The protocol involves four main steps: tissue/cell fixation, permeabilization, and incubation with primary and secondary antibodies. It is an easy technique that allows handling different samples at the same time. In addition, different ultrastructural cell markers can also be used, such as for nucleus and cell wall detection, allowing a detailed analysis of cell morphology. To our knowledge, this is the first protocol to detect TOR activity in situ in photosynthetic organisms; we consider that it will pave the research on the TOR kinase, opening new possibilities to better understand its complex signaling.

Ribosomes are an archetypal ribonucleoprotein assembly. Due to ribosomal evolution and function, r-proteins share specific physicochemical similarities, making the riboproteome particularly suited for tailored proteome profiling methods. Moreover, the structural proteome of ribonucleoprotein assemblies reflects context-dependent functional features. Thus, characterizing the state of riboproteomes provides insights to uncover the context-dependent functionality of r-protein rearrangements, as they relate to what has been termed the ribosomal code, a concept that parallels that of the histone code, in which chromatin rearrangements influence gene expression. Compared to high-resolution ribosomal structures, omics methods lag when it comes to offering customized solutions to close the knowledge gap between structure and function that currently exists in riboproteomes. Purifying the riboproteome and subsequent shot-gun proteomics typically involves protein denaturation and digestion with proteases. The results are relative abundances of r-proteins at the ribosome population level. We have previously shown that, to gain insight into the stoichiometry of individual proteins, it is necessary to measure by proteomics bound r-proteins and normalize their intensities by the sum of r-protein abundances per ribosomal complex, i.e., 40S or 60S subunits. These calculations ensure that individual r-protein stoichiometries represent the fraction of each family/paralog relative to the complex, effectively revealing which r-proteins become substoichiometric in specific physiological scenarios. Here, we present an optimized method to profile the riboproteome of any organism as well as the synthesis rates of r-proteins determined by stable isotope-assisted mass spectrometry. Our method purifies the r-proteins in a reversibly denatured state, which offers the possibility for combined top-down and bottom-up proteomics. Our method offers a milder native denaturation of the r-proteome via a chaotropic GuHCl solution as compared with previous studies that use irreversible denaturation under highly acidic conditions to dissociate rRNA and r-proteins. As such, our method is better suited to conserve post-translational modifications (PTMs). Subsequently, our method carefully considers the amino acid composition of r-proteins to select an appropriate protease for digestion. We avoid non-specific protease cleavage by increasing the pH of our standardized r-proteome dilutions that enter the digestion pipeline and by using a digestion buffer that ensures an optimal pH for a reliable protease digestion process. Finally, we provide the R package ProtSynthesis to study the fractional synthesis rates of r-proteins. The package uses physiological parameters as input to determine peptide or protein fractional synthesis rates. Once the physiological parameters are measured, our equations allow a fair comparison between treatments that alter the biological equilibrium state of the system under study. Our equations correct peptide enrichment using enrichments in soluble amino acids, growth rates, and total protein accumulation. As a means of validation, our pipeline fails to find “false” enrichments in non-labeled samples while also filtering out proteins with multiple unique peptides that have different enrichment values, which are rare in our datasets. These two aspects reflect the accuracy of our tool. Our method offers the possibility of elucidating individual r-protein family/paralog abundances, PTM status, fractional synthesis rates, and dynamic assembly into ribosomal complexes if top-down and bottom-up proteomic approaches are used concomitantly, taking one step further into mapping the native and dynamic status of the r-proteome onto high-resolution ribosome structures. In addition, our method can be used to study the proteomes of all macromolecular assemblies that can be purified, although purification is the limiting step, and the efficacy and accuracy of the proteases may be limited depending on the digestion requirements.

Biomolecular condensates are membrane-less assemblies of proteins and nucleic acids formed through liquid–liquid phase separation (LLPS). These assemblies are known to temporally and spatially regulate numerous biological activities and cellular processes in plants and animals. In vitro phase separation assay using recombinant proteins represents one of the standard ways to examine the properties of proteins undergoing LLPS. Here, we present a detailed protocol to investigate in vitro LLPS using in vitro expressed and purified recombinant proteins.

Cytochrome P450 reductase (CPR) is a multi-domain protein that acts as a redox partner of cytochrome P450s. The CPR contains a flavin adenine dinucleotide (FAD)–binding domain, a flavin mononucleotide (FMN)-binding domain, and a connecting domain. To achieve catalytic events, the FMN-binding domain needs to move relative to the FAD-binding domain, and this high flexibility complicates structural determination in high-resolution by X-ray crystallography. Here, we demonstrate a seeding technique of sorghum CPR crystals for resolution improvement, which can be applied to other poorly diffracting protein crystals. Protein expression is completed using an E. coli cell line with a high protein yield and purified using chromatography techniques. Crystals are screened using an automated 96-well plating robot. Poorly diffracting crystals are originally grown using a hanging drop method from successful trials observed in sitting drops. A macro seeding technique is applied by transferring crystal clusters to fresh conditions without nucleation to increase crystal size. Prior to diffraction, a dehydration technique is applied by serial transfer to higher precipitant concentrations. Thus, an increase in resolution by 7 Å is achieved by limiting the inopportune effects of the flexibility inherent to the domains of CPR, and secondary structures of SbCPR2c are observed.

Graphical abstract:

A number of molecules, such as secreted peptides, have been shown to mediate root-to-shoot signaling in response to various conditions. The xylem is a pathway for water and molecules that are translocated from roots to shoots. Therefore, collecting and analyzing xylem exudates is an efficient approach to study root-to-shoot long-distance signaling. Here, we describe a step-by-step protocol for the collection of xylem exudate from the model plant Arabidopsis and the crop plant soybean (Glycine max). In this protocol, we can collect xylem exudate from plants cultured under normal growth conditions without using special equipment.

Graphical abstract:

Xylem exudates on the cut surfaces of an Arabidopsis hypocotyl and a soybean internode.

Protein–lipid interactions play important roles in many biological processes, including metabolism, signaling, and transport; however, computational and structural analyses often fail to predict such interactions, and determining which lipids participate in these interactions remains challenging. In vitro assays to assess the physical interaction between a protein of interest and a panel of phospholipids provide crucial information for predicting the functionality of these interactions in vivo. In this protocol, which we developed in the context of evaluating protein–lipid binding of the Arabidopsis thaliana florigen FLOWERING LOCUS T, we describe four independent in vitro experiments to determine the interaction of a protein with phospholipids: lipid–protein overlay assays, liposome binding assays, biotin-phospholipid pull-down assays, and fluorescence polarization assays. These complementary assays allow the researcher to test whether the protein of interest interacts with lipids in the test panel, identify the relevant lipids, and assess the strength of the interaction.

The protein expression and purification process is an essential initial step for biochemical analysis of a protein of interest. Traditionally, heterologous protein expression systems (such as E. coli, yeast, insect cells, and cell-free) are employed for plant protein expression, although a plant expression system is often desirable for plant proteins, to ensure proper post-translational modifications. Here, we describe a method to express and purify the ectodomain of one of the leucine-rich repeat receptor-like kinase called CARD1/HPCA1, from Nicotiana benthamiana apoplastic fluid. First, we express His-tagged CARD1 ectodomain in the apoplastic space of N. benthamiana by the Agroinfiltration method. Then, we collect apoplastic fluids from the leaves and purify the His-tagged protein by Ni²⁺-affinity chromatography. In addition to plant-specific post-translational modifications, protein accumulated in the plant apoplastic space, rather than in the cytosolic space, should be kept under an oxidizing environment. Such an environment will help to maintain the property of intrinsic disulfide bonds in the protein of interest. Further, purification from the apoplastic fluids, rather than the total protein extract, will significantly reduce contaminants (for instance RuBisCO) during protein extraction, and simplify downstream processes. We envisage that our system will be useful for expressing various plant proteins, particularly the apoplastic or extracellular regions of membrane proteins.

The precise regulation of the homeostasis of the cellular proteome is critical for the appropriate growth and development of plants. It also allows the plants to respond to various environmental stresses, by modulating their biochemical and physiological aspects in a timely manner. Ubiquitination of cellular proteins is one of the major protein degradation routes for maintaining cellular protein homeostasis, and ubiquitin E3 ligases, components of ubiquitin ligase complexes, play an important role in the selective degradation of target proteins via substrate-specific interactions. Thus, understanding the role of E3 ligases and their substrate regulation uncovers their specific cellular and physiological functions. Here, we provide protocols for auto- and substrate-ubiquitination analyses that utilize the combination of in vitro purified E3 ubiquitin ligase proteins and immunoprecipitation.

Activity-based protein profiling (ABPP) is a chemoproteomics platform to assess the functional state of enzymes in complex biological systems. Over the two decades, ABPP has emerged from a gel-based to gel-free platform, for in-depth proteome analysis with enhanced resolution, sensitivity for target detection, and discovery of small molecule inhibitors. The gel-free format of ABPP coupled with advanced mass spectrometry is highly sensitive and provides more comprehensive knowledge for the targeted enzyme family than the gel-based method. ABPP strategy is applied across microbe, plant, and animal models. It can be performed both in vitro and in vivo studies, and there is no limitation on sample origin. Here, we report an ultrasensitive, gel-free format of ABPP called active site peptide profiling. This protocol describes the identification of authentic functional proteins, by tagging their active sites in a native biological system. It is high throughput in nature and helps enrich even low abundance functional proteins. Since protein identification is virtually based on a single peptide, the identified peptide should be a unique peptide to identify its parent protein. It can be performed in a facile manner and offers to consolidate identification of protein targets as well as the site of probe modification. We have validated this approach using a fluorophosphonate (FP) serine hydrolase probe in the native proteome of the cereal crop Oryza sativa.

Graphic abstract:

Serine hydrolase active site peptide profiling

Phosphoenolpyruvate carboxylase (PEPC) catalyzes a critical step in carbon metabolism in plants and bacteria, the irreversible reaction between bicarbonate and phosphoenolpyruvate to produce the C₄ compound oxaloacetate. This enzyme is particularly important in the context of C₄ photosynthesis, where it is the initial carbon-fixing enzyme. Many studies have used kinetic approaches to characterize the properties of PEPCs from different species, different post-translational states, and after mutagenesis. Most of these studies have worked at a fixed saturating concentration of bicarbonate. Controlling the concentration of bicarbonate is difficult at low concentrations because of equilibration with atmospheric CO₂. We describe here a simple, repeatable, and gas-tight assay system for PEPC that allows bicarbonate concentrations to be controlled above ca. 50 µM.