植物科学

Obtaining homozygous mutant and transgenic lines is a critical yet time-consuming step in Arabidopsis thaliana research. Conventional breeding procedures require seeds to undergo complete maturation and natural desiccation before harvest, followed by cold stratification to overcome seed dormancy. This process substantially prolongs generation turnover and delays genetic screening. Here, we describe a rapid germination strategy based on the use of partially dehydrated seeds collected approximately 15 days after pollination (DAP). At this developmental stage, embryos have reached physiological maturity, while the seeds have not yet entered deep desiccation-induced dormancy. After surface sterilization and short-term cold treatment (2–3 days at 4 °C), these seeds readily germinate on 1/2 MS medium and develop into normal seedlings. By bypassing the prolonged maturation and dormancy phases associated with naturally dried seeds, this protocol shortens each generation cycle by approximately 1–2 weeks. The method is particularly useful for accelerating the propagation of transgenic materials and the identification of homozygous mutant lines in Arabidopsis.

Identifying substrates of protein phosphatases has been technically challenging and has hampered progress in the field of plant sciences. Small molecule inhibitors of protein phosphatases have aided in uncovering classes of phosphatases that target substrates, but that too has severe limitations. Here, we describe a method that enriches phosphorylated substrates using TiO₂ and phospho-tyrosine antibodies in phosphatase knockout lines of Arabidopsis thaliana. When compared to wild-type plants, this approach permits identification of putative substrates and specific phosphorylation sites by mass spectrometry, allowing for further in vitro or functional validation. The key to the approach described here is the use of phosphatase knockout lines to maintain substrates in a phosphorylated state and using phospho-tyrosine antibodies to enrich for tyrosine phosphorylated peptides.

Plant roots dynamically respond to environmental changes and serve as an ideal system for studying cell development and gene regulation. Recent advances in imaging-based spatial transcriptomics have enabled high-resolution mapping of gene expression while preserving spatial context. However, existing sample preparation techniques remain inadequate for handling rigid plant tissues such as crop roots. Here, we present a detailed and practical protocol for preparing rigid plant tissue samples for imaging-based spatial transcriptomics. The workflow ensures effective tissue handling while maintaining RNA integrity and spatial organization. Within approximately eight days, samples can be processed and mounted onto commercial slides, making them ready for subsequent probe hybridization and imaging. This protocol also includes an integrated sample attachment test performed to assess slide quality. It has been optimized to produce consistent and reliable results across experiments. Overall, our method provides a robust solution for spatial transcriptomic analysis in rigid plant tissues, facilitating broader application of these technologies in plant research.

In the model alga Chlamydomonas reinhardtii, CRISPR (clustered regularly interspaced short palindromic repeat)-based gene editing using Cas (CRISPR-associated) enzymes enables both (a) insertion of large gene cassettes and (b) the creation of knockouts based on the introduction of indels, and specific mutations via mutation-directing oligonucleotides. Owing to the relatively low efficiency of this process, selection markers are frequently used to enrich the candidate pool prior to screening, which typically employs PCR. Unfortunately, few selection markers are available for Chlamydomonas. Furthermore, each marker requires different selection media, and deletion of the selectable marker can be difficult. When multiple successive gene editing steps are required, the use of these markers becomes onerous. The SCREAM (sequential CRISPR via recycling endogenous auxotrophic markers) technique employs an endogenous gene as a marker, the mutation of which can be selected both in the forward (loss of function) and reverse (gain of function) directions. During the first gene editing step, crRNA and mutation-directing oligonucleotides are provided for both the marker and the first target gene (Target 1). Candidates with edited marker genes are selected by loss of marker function, prior to screening for the desired modification of the first target gene. Using a successful candidate, a subsequent gene editing step directs reversion of the mutant marker gene to wild-type status, with candidates being selected on auxotrophic media to detect the regain of function of the auxotrophic marker to wild type (i.e., reversion). Simultaneously, a second target gene modification is produced using Target 2–specific crRNA and oligonucleotides. Revertants, now with a wild-type auxotrophic marker, are then screened for the specific mutation of Target 2. This reversion strategy enables a single selectable marker to be reused indefinitely, facilitating the creation of many successive mutations in a single cell line. As the marker can be completely reconstituted, strains can be created in which only the target gene is altered. Employment of homology-directed repair, using single-stranded oligonucleotides for mutation creation, enables the creation of site-directed mutants, tag insertion, and gene knockouts or reversion, rather than the insertion of large gene cassettes. In this implementation, nitrate reductase is used as the endogenous auxotrophic marker, and the adenine phosphoribosyltransferase gene is used as an example of a target gene.

CRISPR/Cas9-based genome editing is a powerful approach for functional genomics and bioenergy research in woody plants. However, conventional single guide RNA (gRNA) strategies predominantly generate small insertions or deletions that may not fully disrupt gene function and often require extensive sequencing for mutation identification. Here, we present an optimized protocol for the efficient generation of large-fragment deletion mutants in Populus tremula × P. alba clone INRA 717-1B4 using a dual-gRNA CRISPR/Cas9 system. Co-expression of two gRNAs flanking the target region induces double-strand breaks at both sites, enabling the deletion of the intervening genomic fragment, typically larger than 50 bp. This protocol describes step-by-step procedures for gRNA design, vector construction, Agrobacterium-mediated transformation, plant regeneration, and molecular validation. Using the PtFBX230 gene as a representative target, large deletions are readily identified by conventional PCR and agarose gel electrophoresis, enabling rapid and cost-effective genotyping. This protocol can be readily adopted to other loci in poplar and related woody species and provides a robust framework for generating null alleles to support functional genomics and bioenergy-related trait engineering in woody plants.

The canonical role of transfer RNAs (tRNAs) in protein synthesis has been extensively characterized; however, recent studies have uncovered novel functions for tRNA as a mediator of long-distance signaling in plants. Several studies have identified dicistronic tRNA-mRNA transcripts that contain a tRNA gene and an adjacent protein-coding gene (PCG) that are transcribed as a single unit. These transcripts are associated with RNA systemic mobility through the plant’s vascular tissues, potentially acting as non-cell-autonomous signaling messengers in coordinating development and stress responses. Here, we report a computational pipeline to detect dicistronic tRNA-mRNA transcripts from short-read next-generation RNA-sequencing datasets; to our knowledge, this is the only established pipeline for the systematic identification of such candidates in plants. The dicistronic RNA transcript version 2 (v2) described here improves on the earlier version DiRT v1 by expanding the repertoire of dicistronic transcripts detected to include tRNA-like structures (TLS) as well as functional tRNAs, which were already supported in the pipeline. The updated protocol also includes detection of dicistronic tRNA or TLS sequences within genomic features such as untranslated regions (UTRs). The accurate detection of both tRNAs and UTR-embedded tRNA-like sequences (TLS) is critical, as these RNA structures have been reported to function as mediators of long-distance RNA mobility. Furthermore, as NGS datasets are prone to sequencing artifacts and potential DNA contamination, we improved the pipeline’s statistical robustness by including read coverage of flanking intronic regions as a baseline control. To account for potential DNA contamination during RNA-seq library preparation, detected tRNA-mRNA transcripts are deemed as putatively dicistronic only if the coverage of their intergenic region is significantly higher (Student’s t-test, FDR < 0.05) than flanking intronic regions. Furthermore, the updated pipeline allows this statistical test to be applied to intronless and single-intron genes. Using this updated protocol, we identified novel tRNA and TLS dicistronic transcripts in both grapevine (Vitis spp. Ruggeri 140) and Arabidopsis thaliana datasets and validated in vitro using RT-PCR. We provide a fast and reliable method to detect dicistronic transcripts that can be applied to any short-read RNA-sequencing dataset, fast-tracking the functional characterization of these newly emerging transcripts.

While traditional kinetic studies of the cytochrome b₆f complex have frequently relied on measurements within the complex environment of intact leaves or whole-organism systems, such approaches can be limited by overlapping signals and physiological variables. This protocol advances existing frameworks by introducing a streamlined, multi-wavelength spectroscopic approach utilizing a reconstituted in vitro system to elucidate the inter-complex electron transfer kinetics between photosystem I and cytochrome b₆f. Utilizing the JTS-150 pulsed spectrometer, supplied with a Smart Lamp, we monitored the redox transitions of P700⁺ and Cytf by simultaneously measuring the absorbance changes of our isolated complexes system in six different wavelengths (546, 554, 563, 574, 705, and 740 nm). Kinetic analysis was divided into two phases: laser-induced flash kinetics and steady-state actinic induction. We resolved the second-order re-reduction of P700⁺ by plastocyanin, accounting for detector saturation constraints with a 2 ms post-flash delay. Steady-state measurements under actinic light revealed complex Cytf turnover, characterized by a double-exponential decay. Furthermore, dark relaxation kinetics were used to quantify ferredoxin-mediated re-reduction of the cytochrome pool. By allowing the incorporation of specific regulatory and inhibitory factors, this methodology sets the ground for the deconvolution of competing electron pathways. It can therefore be used as a robust framework for assessing the mechanism of regulatory processes on photosynthetic flux.

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.

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.

A simple and effective method to identify genetic markers of yield response to nitrogen (N) fertilizer among maize hybrids is urgently needed. In this article, we describe a detailed methodology to identify genetic markers and develop associated assays for the prediction of yield N-response in maize. We first outline an in silico workflow to identify high-priority single-nucleotide polymorphism (SNP) markers from genome-wide association studies (GWAS). We then describe a detailed methodology to develop cleaved amplified polymorphic sequences (CAPS) and derived CAPS (dCAPS)-based assays to quickly and effectively test genetic marker subsets. This protocol is expected to provide a robust approach to determine N-response type among maize germplasm, including elite commercial varieties, allowing more appropriate on-farm N application rates, minimizing N fertilizer waste.