Plant Science

Mal de Río Cuarto disease, caused by a Fijivirus, is a major constraint for maize production in Argentina. The traditional evaluation of resistant hybrids is limited by the low efficiency of natural virus transmission and the lack of standardized field inoculation methods. We developed a protocol that combines laboratory mass-rearing of the planthopper vector Delphacodes kuscheli with a controlled field transmission system. The method involves the synchronized production of large insect populations, acquisition of viruliferous vectors under controlled conditions, and their safe transport to the field using specialized containers. Transmission is achieved through individual cages placed on maize seedlings, ensuring high inoculation pressure under field-like conditions. This protocol enables reliable and reproducible virus transmission, facilitating large-scale screening of maize hybrids and other cereals. Its main advantages are the high throughput of vector production, improved transmission efficiency, and adaptability to diverse experimental designs.

Agrobacterium rhizogenes–mediated hairy root transformation provides a rapid platform for gene function analysis prior to stable whole-plant transformation. However, most existing hairy root transformation methods rely on tissue culture and require chemical or fluorescence-based selection, which increases experimental complexity. Here, we describe a tissue culture–free soybean hairy root transformation protocol incorporating the RUBY visual reporter system. While this work does not introduce a new transformation concept, it presents a streamlined implementation of established soybean hairy root methodologies that emphasizes procedural simplicity, reduced handling, and faster access to functional root material. Transgenic roots expressing RUBY can be directly identified by red pigmentation with the naked eye. In RUBY-positive roots, candidate genes driven by the CaMV 35S promoter showed higher expression levels than those in empty-vector controls, indicating that the system supports effective gene expression. Using this procedure, clearly identifiable transgenic hairy roots can be obtained within 20 days. Overall, this protocol simplifies induction and screening while reducing operational complexity and equipment requirements.

Cellulose synthase complexes (CSCs) play a central role in plant cell wall formation. Their dynamic behavior at the plasma membrane leads to the deposition of cellulose microfibrils into the apoplastic space, thereby shaping the architecture and mechanical properties of the cell wall. Although previous imaging studies have provided important insights into CSC dynamics and localization, standardized and reproducible workflows for quantitative measurements of CSC speed and density remain limited. Here, we present a reproducible live-cell imaging and analysis workflow for quantifying the speed and density of fluorescently labeled CSCs at the plasma membrane in Arabidopsis thaliana. The protocol integrates optimized spinning-disk confocal imaging, surface-based projection of z-stack recordings, automated detection of diffraction-limited CSCs foci, and kymograph-based speed measurements using freely available tools in Fiji. While selected steps, such as region of interest definition and parameter selection for spot detection or trajectory analysis, remain user-guided, these decisions are constrained to well-defined stages within an otherwise standardized pipeline, thereby reducing variability and improving reproducibility across experiments. The workflow has been validated across multiple tissues, reporter lines, genetic backgrounds, and perturbation conditions in Arabidopsis and enables robust comparative analysis of CSC dynamics. Beyond CSCs, this workflow is expected to be adaptable to other fluorescently labeled proteins that appear as diffraction-limited foci at or near the plasma membrane.

In wheat and other crops, some genes display presence/absence variation, and it is occasionally beneficial to select for the absent allele to remove a functional gene. However, current high-throughput genotyping methods used to detect the absence of genes tend to be inconsistent and inconclusive. Kompetitive allele-specific PCR (KASP) and PCR allele competitive extension (PACE) are two well-established methods for allele-specific polymerase chain reaction (AS-PCR) assays, each using fluorescence resonance energy transfer (FRET) to generate a signal for each allele, typically targeting biallelic single-nucleotide polymorphisms. KASP and PACE methods are more difficult to apply to alleles with presence/absence variation because the lack of amplification of the absent allele is indistinguishable from a failed PCR. Here, we present a multiplex fluorescence-based absent allele–specific amplification (FAASA) method using the PACE marker system (compatible with KASP markers) to detect the absence of one particular or all alleles of a target sequence using a primer mix consisting of one target-specific primer pair (TSP) and a second primer set specific to a highly conserved endogenous gene known as a core gene–specific primer pair (CGSP). The forward primer of each pair is tagged with a 5′ terminal tail complementary to dye-labeled oligonucleotides in commercially available FRET cassettes. Lines that amplify only the core gene do not carry the target, while lines that amplify both the core gene and the target carry alleles of both the core gene and the target. The inclusion of the CGSPs allows researchers to confidently distinguish lines with absent alleles of the target from lines with failed PCR reactions, which can happen due to various reasons, including inadequate DNA quality or quantity.

Anthocyanins are specialized flavonoid pigments that play critical roles in plant coloration, photoprotection, and responses to environmental stress. Arabidopsis thaliana serves as a valuable genetic model for dissecting anthocyanin biosynthesis and regulatory networks. Conventional methods for anthocyanin quantification, such as crude spectrophotometric assays, often compromise pigment integrity, yield inconsistent results, and provide limited information on compound composition. Here, we describe a simple, reproducible, and high-fidelity protocol for the induction, extraction, quantification, and chromatographic profiling of anthocyanins in Arabidopsis thaliana seedlings. The workflow employs well-defined anthocyanin-inductive conditions (AIC), methanol/formic acid extraction, lyophilization for dry-weight normalization, and dual quantification via spectrophotometry and High-performance liquid chromatography with diode-array detection (HPLC-DAD) analysis. This protocol enables accurate comparison between wild-type and mutant genotypes, facilitating both mutant screening and metabolic pathway analysis. The approach minimizes pigment degradation, enhances reproducibility across replicates, and offers a robust tool for research in plant metabolism, stress physiology, and flavonoid biochemistry.

Protoplast systems are widely used in plant research as versatile platforms for studying cellular processes and validating gene editing tools. In maize, they are particularly valuable because stable transformation in immature embryos is slow and labor-intensive, often requiring months to regenerate plants. However, existing protocols often yield inconsistent results in protoplast recovery, transfection efficiency, and viability. We present an optimized protocol for maize mesophyll protoplast isolation and PEG-mediated transfection. Two-week-old etiolated seedlings are processed using vertical cutting, improving the yield and viability of protoplasts. Protoplasts are then immediately transformed with a CRISPR/Cas9 construct after isolation, via PEG4000 with only 10 μg of plasmid DNA, reducing the resource demands of standard methods. Modified washing and storage conditions extend transformed protoplast viability to seven days, enabling longer-term monitoring and expanded downstream analyses. Editing outcomes are quantified by sequencing target sites and calculating efficiency with Cas-Analyzer. This protocol provides a rapid, efficient, and reproducible method for the rapid evaluation of gene editing in maize. This protocol offers a methodology to accelerate agricultural crop studies and broader plant molecular biology.

Biomolecular condensates organize cellular processes through liquid–liquid phase separation, creating membrane-less compartments enriched in specific proteins and RNAs. Understanding their RNA composition is essential for elucidating plant stress responses, yet capturing these transiently associated RNAs remains technically challenging. We present Turbo-RIP (TurboID-based proximity labeling with RNA immunopurification), a comprehensive protocol for identifying condensate-associated RNAs in plants. Turbo-RIP employs the biotin ligase TurboID to label proximal proteins at 22 °C, followed by formaldehyde crosslinking and streptavidin-based capture of protein–RNA complexes. We provide detailed procedures for three cloning strategies, transformation of Nicotiana benthamiana and Arabidopsis thaliana, validation of TurboID activity, and RNA recovery. The protocol successfully captured processing body–associated RNAs with minimal background. Turbo-RIP enables systematic mapping of RNA populations within plant condensates under diverse conditions. The protocol requires 3–5 days from sample preparation to RNA isolation, with construct validation taking 2–4 weeks. All procedures use standard laboratory equipment, making Turbo-RIP accessible for plant molecular biology laboratories.

The plant cell wall is a dynamic and complex extracellular matrix that not only provides structural integrity and determines cell shape but also mediates intercellular communication. Among its major components, pectins play essential roles in cell adhesion, wall porosity, hydration, and flexibility. Rhamnogalacturonan-I (RG-I), a structurally diverse pectic polysaccharide, remains one of the least understood components of the plant cell wall. Its backbone is substituted with arabinan, galactan, and arabinogalactan side chains that vary in length, branching, and composition across tissues, species, and developmental stages. In addition, RG-I can undergo modifications such as backbone acetylation, further contributing to its structural complexity and functional diversity. To advance understanding of RG-I, we present a detailed method for isolating RG-I from the model plant Arabidopsis thaliana. Leveraging Arabidopsis as a model system provides major advantages owing to its well-characterized genome and powerful molecular toolkit, enabling deeper investigation into the roles of RG-I in plant development and responses to environmental stress. Our method consists of two major steps: an initial chemical extraction using oxalate, followed by endo-polygalacturonase (EPG) digestion to fragment the pectic domains. An advantage of this approach is that it produces a dry material that can be stored at room temperature without special handling and does not introduce chemicals that may interfere with downstream analyses. The purified RG-I can be used for detailed compositional and structural analyses, as well as for functional studies of enzymes involved in pectin biosynthesis, modification, and degradation. Although this protocol was developed for isolating RG-I from Arabidopsis rosette leaves, it is also applicable to other Arabidopsis organs and other plant species.

Pinpointing causal genes for complex traits from genome-wide association studies (GWAS) remains a central challenge in crop genetics, particularly in species with extensive linkage disequilibrium (LD) such as rice. Here, we present CisTrans-ECAS, a computational protocol that overcomes this limitation by integrating population genomics and transcriptomics. The method’s core principle is the decomposition of gene expression into two distinct components: a cis-expression component (cis-EC), regulated by local genetic variants, and a trans-expression component (trans-EC), influenced by distal genetic factors. By testing the association of both components with a phenotype, CisTrans-ECAS establishes a dual-evidence framework that substantially improves the reliability of causal inference. This protocol details the complete workflow, demonstrating its power not only to identify causal genes at loci with weak GWAS signals but also to systematically reconstruct gene regulatory networks. It provides a robust and powerful tool for advancing crop functional genomics and molecular breeding.

Accurate profiling of soil and root-associated bacterial communities is essential for understanding ecosystem functions and improving sustainable agricultural practices. Here, a comprehensive, modular workflow is presented for the analysis of full-length 16S rRNA gene amplicons generated with Oxford Nanopore long-read sequencing. The protocol integrates four standardized steps: (i) quality assessment and filtering of raw reads with NanoPlot and NanoFilt, (ii) removal of plant organelle contamination using a curated Viridiplantae Kraken2 database, (iii) species-level taxonomic assignment with Emu, and (iv) downstream ecological analyses, including rarefaction, diversity metrics, and functional inference. Leveraging high-performance computing resources, the workflow enables parallel processing of large datasets, rigorous contamination control, and reproducible execution across environments. The pipeline’s efficiency is demonstrated on full-length 16S rRNA gene datasets from yellow pea rhizosphere and root samples, with high post-filter read retention and high-resolution community profiles. Automated SLURM scripts and detailed documentation are provided in a public GitHub repository (https://github.com/henrimdias/emu-microbiome-HPC; release v1.0.2, emu-pipeline-revised) and archived on Zenodo (DOI: 10.5281/zenodo.17764933).