Hydrogels are widely used as injectable biomaterials for minimally invasive stem cell delivery, providing a hydrated three-dimensional matrix that supports cell survival and retention. Nanocomposite hydrogels, formed by combining hydrogels with nanoscale fillers, offer additional opportunities to tailor rheological behavior and mechanical performance for injectable cell-delivery applications. However, their translational potential depends on the ability to rationally characterize both material behavior and cell compatibility under injection-relevant conditions. This chapter presents a structured experimental protocol for the preparation and initial characterization of collagen-Laponite nanocomposite hydrogels containing human adipose-derived mesenchymal stem cells as a representative model for injectable therapeutic applications. The protocol describes hydrogel formulation, nanoparticle dispersion, stem cell encapsulation, rheological characterization, injectability testing, post-injection viability assessment, and short-term metabolic activity analysis. Although a collagen-Laponite formulation and human adipose-derived mesenchymal stem cells are used as the working example, the protocol provides a practical and systematic framework for evaluating injectable stem cell-laden nanocomposite hydrogels, with particular focus on balancing injectability, structural integrity, and preservation of cellular function.

Mass spectrometry-based proteomics allows the unbiased identification and quantification of proteins and phosphopeptides in biological materials. The nature of walled plant cells requires specific protocols for effective and efficient protein isolation, and, in general, the plant sciences can benefit from more accessible, optimized proteomics workflows. Advances in MS instrumentation now allow the measurement of large numbers of samples, shifting constraints in proteomics toward the accurate, high-throughput preparation of samples. Here, we describe a high-throughput (phospho)proteomics protocol that enables processing of samples using different filter types in a 96-well format.

The plasma membrane (PM) is the primary interface between plant cells and their environment, and its resident proteins mediate key processes such as extracellular signal perception and downstream cellular reprogramming. Yet, PM proteins are typically underrepresented in total protein extracts, and existing enrichment strategies are often laborious and require extensive optimization. Here, a simple and robust workflow is described for enriching PM proteins from Arabidopsis thaliana seedlings using total microsomal membranes obtained by differential centrifugation as starting material. Sequential low- and high-speed spins are used to isolate total microsomal membranes and progressively deplete contaminating organelles, thereby increasing the relative abundance of PM proteins. Coupled with the rich genetic toolkit available in Arabidopsis, this protocol provides an accessible platform for systematic characterization of the PM proteome.

This section details the optimized methods for single-cell RNA sequencing (scRNA-seq) in Medicago truncatula roots and nodules. We describe a detailed experimental workflow, comprising plant growth and bacteria application, protoplast isolation, quality control, and library processing according to 10× Genomics®, suitable for both long- and short-read sequencing. This protocol can serve as a starting point for establishing robust, high-quality scRNA-seq workflows to study nodulation and root development in legumes.

In plant roots, the diversity of tissues along the radial axis and of developmental stages along the longitudinal axis implies a highly dynamic and heterogeneous gene expression. Many master regulators of gene expression exhibit distinctive temporal expression patterns or are restricted to specific cell types, making whole-root molecular analyses insufficient to resolve their roles in developmental and physiological processes. Here, we report a protocol for fluorescence-activated cell sorting (FACS)-coupled RNA sequencing to analyze, reconstruct, and ultimately dissect the gene regulatory networks (GRNs) involved in root development and physiology, particularly well-suited for genes that are expressed transiently or in rare or restricted cell types. The method relies on selecting an appropriate cell-type-specific reporter line, which enables the isolation of labeled cells with high specificity and minimal contamination. Furthermore, we discuss the relative advantages of FACS-coupled RNA-seq compared with alternative gene-expression methodologies, including quantitative PCR (qPCR), bulk root RNA-seq, single-cell RNA-seq (scRNA-seq), and spatial transcriptomics, considering sensitivity, resolution, input requirements, and cost.

In situ mRNA hybridization, which relies on the complementarity between antisense riboprobes and specific RNA populations, was originally used on thin sections with colorimetric dyes to visualize mRNA distribution. The use of fluorescent probes and confocal imaging enabled whole-mount in situ hybridization (WISH) and 3D reconstruction of gene expression domains. Here, we present a versatile protocol based on microwave treatment that results in high and homogeneous hybridization yields. When used with a cell wall marker and tissue clearing, it enables simultaneous visualization of gene expression and cell segmentation across various Arabidopsis organs, including leaves, shoots, roots, and embryos enclosed within integuments. This easy, rapid, robust, and low-cost method also accommodates different detection kits and cytological dyes, and it can be extended to Double WISH for the simultaneous detection of two distinct mRNA targets.

Iron (Fe) is involved in numerous key physiological processes due to its transfer or gain electron capacity. However, in excess, Fe is toxic to cells. The understanding of mechanisms governing its homeostasis requires imaging techniques capable of localizing and quantifying Fe and its redox state in situ. The Perls-DAB histochemical staining is a widely used method for visualizing Fe at a cellular level. Other advanced approaches, such as micro X-ray fluorescence and X-ray absorption near-edge structure spectroscopy, also provide information on the spatial distribution and oxidation state of Fe. However, all of these methods require sample fixation before observing Fe localization, which can introduce biases in the results. Recent development in synthetic fluorescent probes now enables the real-time visualization of ferrous (Fe2+) and ferric (Fe3+) forms in living roots, offering a robust and user-friendly approach to study Fe redox distribution in planta. This live imaging approach provides access to Fe dynamics and enables semi-quantitative analyses, thereby opening new avenues for Fe homeostasis research. This chapter provides detailed protocols for applying these methods in Arabidopsis roots.

Quantitative methods using imaging and spectroscopy offer valuable insights into cell biology and molecular biophysics by elucidating complex mechanisms across various cellular environments. Fluorescence correlation spectroscopy (FCS) is particularly advantageous for quantifying the dynamics of fluorescent particles in low-concentration samples. As a high-resolution spectroscopic method with single-molecule sensitivity, FCS is well-suited to investigating membrane proteins, molecular interactions, aggregation, and conformational changes in living systems, including plant cells. However, challenges such as background fluorescence and artifacts arising from biological sample properties require careful experimental design, thereby limiting the routine application of FCS in plant tissues. Recent advancements in technology, such as confocal optics and highly efficient photon detectors, have substantially enhanced FCS's sensitivity and expanded its capacity for single-molecule detection. These advancements now empower plant biologists to explore the dynamics of plant proteins under various conditions. This chapter focuses on methodologies in fluorescence autocorrelation spectroscopy (FACS/FCS) and their application to plant research, with a particular emphasis on plasma membrane proteins in Arabidopsis root cells.

The division of a mother cell into two daughter cells is a fundamental process in biology, involving the transfer of genomic information to the next generation of cells. In plant roots, cell divisions occur in the root apical meristem, a specialized tissue located at the plant's root tip. Cell proliferation rates in the root apical meristem shape root growth, thereby contributing to general development as well as the acclimatization to stressful environmental conditions through phenotypic plasticity. Here, we present a protocol for the microscopical analysis of cell proliferation in Arabidopsis thaliana root meristems by 5-ethynyl-2'-deoxyuridine (EdU) staining. EdU, a thymine-analog, is incorporated into newly synthesized DNA of proliferative cells during the S-phase of the cell cycle. Subsequently, EdU-containing nuclei can be labeled with a fluorophore and detected by confocal laser scanning microscopy. The combination of the EdU assay with cell wall staining enables the quantification of root meristem architecture parameters and cell division activity from microscopic images. Together, this can elucidate how root meristems respond to changes in environmental conditions or address fundamental questions of developmental plant biology.

The polar transport of the phytohormone auxin regulates plant organogenesis, morphogenesis, and gravitropism, which are coordinately regulated by auxin influx and efflux carriers. To measure the basipetal (shootward) transport of auxin in roots, we applied radiolabeled IAA to the root tips of 7-day-old seedlings in the model plant Arabidopsis thaliana. A scintillation counter was then used to measure the radioactivity of the samples. Here, we describe a method for measuring auxin transport in A. thaliana roots via 3H-indole-3-acetic acid (3H-IAA).

The capillary-based Organic Electronic Ion Pump (cOEIP) is an emerging bioelectronic delivery tool that enables precise and electronically controlled delivery of charged biomolecules to biological targets without inducing bulk fluid flow. Traditional treatment methods often induce mechanical disturbances or secondary signaling that obscure the direct physiological effects of the delivered compounds. In contrast, the iontronic devices enable precise, flow-free ionic delivery, allowing the isolation of responses specifically attributable to the applied biomolecule. Here, we present a detailed protocol for integrating the cOEIP with live confocal imaging of Arabidopsis thaliana roots, enabling spatiotemporally resolved studies of root physiology under controlled ionic delivery. As a model application, we demonstrate the localized delivery of protons (H⁺) to the root microenvironment, providing a framework for investigating pH-dependent signaling processes. The protocol includes step-by-step instructions for device assembly, preparation, calibration, and experimental setup, accompanied by representative images illustrating the mounting and operational procedures. Importantly, this approach can be adapted for the delivery of a wide range of charged biomolecules, expanding the toolkit for dynamic in vivo studies of plant-environment interactions.

Water potential is the driving force behind the movement of water within the soil-plant-atmosphere continuum. Measuring water potential and its components provides valuable information about plant- and root water relations, including osmoregulation. Here, we describe a straightforward method for determining water and osmotic potential, as well as cell turgor in roots, using dew point hygrometry. The chapter begins with a brief overview of issues related to plant and root water potential, and then proceeds to describe the principles and measurement procedures of dew point hygrometry in more detail, including calibration, equilibration, and sample preparation. Practical recommendations are provided to minimize common error sources. This methodology may be useful for researchers studying plant-water balance, root water uptake, plant stress physiology, and osmoregulation.

Water is distributed unevenly in soil, creating fine-scale patterns that shift with soil structure and drying. To navigate this complexity, roots deploy localized adaptive responses that allow them to sense and respond to water availability with high spatial precision. These include alterations in root branching (hydropatterning and xerobranching) and directional growth responses (hydrotropism and xerotropism), all of which collectively optimize water capture. Despite their importance, reproducing these heterogeneous water cues in the laboratory is challenging because soil systems are structurally and hydraulically complex. To address this, a suite of agar-based model systems has been developed to provide reproducible, tractable, and quantifiable platforms for studying root water sensing. These approaches rely on manipulating water potential using solutes or air interfaces to mimic microscale environmental heterogeneity and trigger adaptive responses. In this chapter, we outline the conceptual principles underlying water-driven root adaptive responses and provide detailed protocols for five agar-based experimental systems designed to study hydrotropism, xerotropism, xerobranching, and hydropatterning. These methods support robust mechanistic investigation and can be adapted for both Arabidopsis and crop species.

Root hairs play pivotal roles in nutrient and water acquisition and in plant-microbe interactions. Consequently, understanding the mechanisms underlying root hair development and their regulatory pathways is an important aspect of plant physiology research. Quantifying root hairs and root hair length is often essential in such studies, but is labor-intensive and prone to subjectivity. The availability of straightforward tools for automated root hair measurements is limited, and existing options are often tailored for specific images or do not measure individual root hairs. To address this, we developed RootHairML, a flexible and simple Python-based machine learning tool designed for efficient quantification of root hair lengths. Based on labeled images and pixel features, RootHairML trains a Random Forest model to enable the detection of root hairs in new images. It provides measurements of individual root hair lengths per image and generates annotated images showing all detected root hairs, allowing for manual verification and adjustments. Here, we describe and showcase the use of RootHairML. Overall, RootHairML offers a valuable tool for root hair analysis, enabling researchers to increase data collection and enhance the reproducibility of root hair studies.

Imaging has revolutionized phenotyping analysis by providing accurate documentation and records of observed phenotypes. Analyzing imaging data remains a major bottleneck in root phenotyping analysis. Advances in technology offer a plethora of analysis options at varying automation levels and price points, providing a reliable open-source pipeline for image analysis that delivers stable, reproducible results. Here, we discuss a maize seedling image analysis pipeline powered by open-source tools that enables rapid analysis of seedling root phenotypes. This image analysis pipeline uses Root Painter, an optional AI cleanup step, and RhizoVision Explorer to extract phenotypic data from maize seedlings grown on rolled towel assays, which are commonly used, enabling rapid analysis of these images.