Plant Science

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.

The chloroplast lumen contains at least 80 proteins whose function and regulation are not yet fully understood. Isolating the chloroplast lumen enables the characterization of the lumenal proteins. The lumen can be isolated in several ways through thylakoid disruption using a Yeda press or sonication, or through thylakoid solubilization using a detergent. Here, we present a simple procedure to isolate thylakoid lumen by sonication using leaves of the plant Arabidopsis thaliana. The step-by-step procedure is as follows: thylakoids are isolated from chloroplasts, loosely associated thylakoid surface proteins from the stroma are removed, and the lumen fraction is collected in the supernatant following sonication and centrifugation. Compared to other procedures, this method is easy to implement and saves time, plant material, and cost. Lumenal proteins are obtained in high quantity and purity; however, some stromal membrane–associated proteins are released to the lumen fraction, so this method could be further adapted if needed by decreasing sonication power and/or time.

Chlamydomonas reinhardtii is a model organism for various processes, from photosynthesis to cilia biogenesis, and a great chassis to learn more about biofuel production. This is due to the width of molecular tools available, which have recently expanded with the development of a modular cloning system but, most importantly, with CRISPR/Cas9 editing now being possible. This technique has proven to be more efficient in the absence of a cell wall by using specific mutants or by digesting Chlamydomonas cell wall using the mating-specific metalloprotease autolysin (also called gametolysin). Multiple protocols have been used and shared for autolysin production from Chlamydomonas cells; however, they provide very inconsistent results, which hinders the capacity to routinely perform CRISPR mutagenesis. Here, we propose a simple protocol for autolysin production requiring transfer of cells from plates into a dense liquid suspension, gametogenesis by overnight incubation before mixing of gametes, and enzyme harvesting after 2 h. This protocol has shown to be highly efficient for autolysin production regardless of precise control over cell density at any step. Requiring a minimal amount of labor, it will provide a simple, ready-to-go approach to produce an enzyme critical for the generation of targeted mutants.

Graphical overview

Workflow for autolysin production from Chlamydomonas reinhardtii

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.

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.

Lipids in biomembranes can control the structure and, therefore, the functionality of membrane-embedded protein complexes. Unraveling how the lipid composition determines the mode of operation of membrane proteins provides mechanistic insights into their functionality. We applied a proteoliposome technique for studying how proteins function in biomembranes. The incorporation of isolated membrane proteins in preformed liposomes made from a well-defined lipid composition (proteoliposomes) is a powerful tool for studying lipid-protein interactions. Over several decades, the proteoliposome technique was employed for many different membrane proteins. Recently, it was recognized that different lipid compositions control the light-harvesting functionality of the major photosynthetic light-harvesting complex II (LHCII) isolated from plant thylakoid membranes in vitro. This technique allows systematic examination of the role of so-called non-bilayer lipids on light-harvesting characteristics of LHCII. This protocol describes the isolation of LHCII from leaves and details a four-step procedure to incorporate the detergent-solubilized membrane protein in large unilamellar vesicles (LUV). The protocol was optimized to ensure a very high lipid/protein ratio, designed to specifically examine lipid-protein interactions by minimizing LHCII aggregation. The procedure provides structurally and functionally highly intact LHCII in a detergent-free lipid bilayer with a defined composition.

Photosynthesis is the main process by which sunlight is harvested and converted into chemical energy and has been a focal point of fundamental research in plant biology for decades. In higher plants, the process takes place in the thylakoid membranes where the two photosystems (PSI and PSII) are located. In the past few decades, the evolution of biophysical and biochemical techniques allowed detailed studies of the thylakoid organization and the interaction between protein complexes and cofactors. These studies have mainly focused on model plants, such as Arabidopsis, pea, spinach, and tobacco, which are grown in climate chambers even though significant differences between indoor and outdoor growth conditions are present. In this manuscript, we present a new mild-solubilization procedure for use with “fragile” samples such as thylakoids from conifers growing outdoors. Here, the solubilization protocol is optimized with two detergents in two species, namely Norway spruce (Picea abies) and Scots pine (Pinus sylvestris). We have optimized the isolation and characterization of PSI and PSII multimeric mega- and super-complexes in a close-to-native condition by Blue-Native gel electrophoresis. Eventually, our protocol will not only help in the characterization of photosynthetic complexes from conifers but also in understanding winter adaptation.

The majority of cellular proteins are degraded by the 26S proteasome in eukaryotes. However, intrinsically disordered proteins (IDPs), which contain large portions of unstructured regions and are inherently unstable, are degraded via the ubiquitin-independent 20S proteasome. Emerging evidence indicates that plant IDP homeostasis may also be controlled by the 20S proteasome. Relatively little is known about the specific functions of the 20S proteasome and the regulatory mechanisms of IDP degradation in plants compared to other species because there is a lack of systematic protocols for in vitro assembly of this complex to perform in vitro degradation assays. Here, we present a detailed protocol of in vitro reconstitution assay of the 20S proteasome in Arabidopsis by modifying previously reported methods. The main strategy to obtain the 20S core proteasome here is to strip away the 19S regulatory subunits from the 26S proteasome. The protocol has two major parts: 1) Affinity purification of 20S proteasomes from stable transgenic lines expressing epitope-tagged PAG1, an essential component of the 20S proteasome (Procedures A-D) and 2) an in vitro 20S proteasome degradation assay (Procedure E). We anticipate that these protocols will provide simple and effective approaches to study in vitro degradation by the 20S proteasome and advance the study of protein metabolism in plants.

Exploring the structure and function of protein complexes requires their isolation in the native state–a task that is made challenging when studying labile and/or low abundant complexes. The difficulties in preparing membrane-protein complexes are especially notorious. The cyanobacterium Synechocystis sp. PCC 6803 is a widely used model organism for the physiology of oxygenic phototrophs, and the biogenesis of membrane-bound photosynthetic complexes has traditionally been studied using this cyanobacterium. In a typical approach, the protein complexes are purified with a combination of His-affinity chromatography and a size-based fractionation method such as gradient ultracentrifugation and/or native electrophoresis. However, His-affinity purification harbors prominent contaminants and the levels of many proteins are too low for a feasible multi-step purification. Here, we have developed a purification method for the isolation of 3x FLAG-tagged proteins from the membrane and soluble fractions of Synechocystis. Soluble proteins or solubilized thylakoids are subjected to a single affinity purification step that utilizes the highly specific binding of FLAG-affinity resin. After an intensive wash, the captured proteins are released from the resin under native conditions using an excess of synthetic 3x FLAG peptide. The protocol allows fast isolation of low abundant protein complexes with a superb purity.

Laccases are found in cell walls of plants in very low amounts. This protocol provides an efficient method to purify laccases from rice stems. The method involves three steps: 1) Isolation of total protein from rice stems using buffers with high salt concentration to extract protein from cell walls; 2) Purification of laccases using concanavalin-A beads; and, 3) In-gel staining of laccases with 4-hydroxyindole. Concanavalin-A specifically binds to internal or non-reducing terminal α-D-mannosyl and α-D-glucosyl groups found in glycoproteins and glycolipids. Laccases being glycoproteins binds to concanavalin-A during purification process and eluted with mannose.