Plant-derived extracellular vesicles (PDEVs) have emerged as important mediators of intercellular communication and hold growing potential in therapeutic applications. However, standardized methods for their isolation, particularly from Piper betle leaves (PBL), remain unexplored. Existing apoplastic fluid washing (AFW) extraction techniques typically rely on manual syringe infiltration, which often leads to inconsistent pressure control, variable yields, and increased risk of tissue damage. This protocol describes a vacuum-assisted AFW extraction method optimized for the recovery of intact extracellular vesicles (EVs) from PBL. The workflow features controlled negative pressure using a vacuum pump and chamber to achieve more efficient leaf infiltration compared to infiltration using the syringe method and reproducible apoplastic fluid (AF) collection with subsequent low-speed centrifugation steps, to ensure minimal contamination and preservation of vesicle integrity. Piper betle-derived extracellular vesicle (PBdEV) isolation and purification steps are performed using size exclusion chromatography (SEC). The size and concentration of PBdEVs were confirmed using nanoparticle tracking analysis (NTA), whereas the cup-shaped and lipid bilayer morphology of the EVs were confirmed using transmission electron microscopy (TEM). The method is scalable and adaptable to various leaf morphologies and physiological states, making it suitable for both exploratory and high-throughput studies. Overall, this protocol provides a more consistent, efficient, and tissue-preserving alternative to traditional syringe-based AF extraction methods, offering higher-quality EV preparations for plant EV research. Key features • This protocol focuses on the extraction of Piper betle-derived extracellular vesicles (PBdEVs) using a gentle approach to maintain the vesicles' morphology. • This protocol is suitable for large-scale experiments with multiple biological replicates or different leaf samples of similar morphology.

Manganese (Mn) is an essential trace element whose intracellular homeostasis is tightly controlled by specialized membrane transporters. Dysregulation of Mn transport leads to pathological Mn accumulation and severe human disease; however, efficient and quantitative cell-based methods for assessing Mn2+ transporter activity remain limited. Here, we present an optimized cellular Fura-2 manganese extraction assay (CFMEA) that enables robust quantification of cellular Mn content and provides a normalized framework for assessing relative Mn2+ transport activity in a high-throughput format. This protocol integrates Fura-2-based fluorescence detection of Mn2+ at the Ca2+ isosbestic excitation wavelength with dsDNA quantification to normalize dsDNA levels in cell extracts and immunoblotting to account for transporter protein expression levels. Cells expressing Mn2+ transporters are exposed to MnCl2 in 96-well plates, washed to remove extracellular Mn2+, and lysed in a Fura-2-containing extraction buffer. Fluorescence quenched by Mn2+ is quantified and converted to cellular Mn content using a cell-free Mn-Fura-2 standard curve and then normalized to dsDNA content and protein abundance to determine relative transporter activity. This workflow provides a relatively sensitive, reproducible, and low-cost approach for comparative analysis of Mn2+ transporters and their variants across multiple cell types. The protocol is demonstrated using the Mn2+ efflux transporter SLC30A10 in HEK293T cells and is readily adaptable for studying other Mn2+ transport pathways. Key features • High-throughput, cell-based assay for quantifying cellular manganese content and assessing relative Mn2+ transporter function. • Enhanced accuracy and reproducibility by integrating double-stranded DNA quantification and protein normalization into the cellular Fura-2 manganese extraction assay (CFMEA) workflow. • Workflow compatible with diverse cell types and Mn2+ transporters, including systems overexpressing SLC30A10 in HEK293Tcells.

Nanobodies are recombinant single-domain antibodies (VHHs) derived from the heavy chain-only subset of camelid immunoglobulins that can be reverse-engineered into bivalent antibodies by fusion to immunoglobulin Fc constant regions. Mammalian cells are the system of choice to produce VHH-Fcs to ensure authentic folding and post-translation glycosylation of the expressed VHH-Fcs. In a recent project to find neutralising VHH-Fc binders to the spike proteins of SARS-CoV-2 viruses, we identified a need for rapid expression and purification of multiple VHH-Fc fusions from nanobodies selected by phage display. Here, we present a protocol for the construction of expression vectors by parallel ligase-independent cloning, transient small-scale expression in mammalian cells (4 mL culture volume), screening antigen-binding activity, and midi-scale purification (30 mL culture volume) for downstream activity assays. The workflow is completely transferable between different vector formats, of which three are described herein: Fc fusion dimers, monomeric CD4 fusions, and His-tagged monomers. Key features • Miniaturised and parallelised methodology for the screening and production of large numbers of VHHs that bind antigens of interest. • Streamlined and unified, high-throughput cloning strategy for use in multiple modular vectors for monomeric and dimeric VHH production in mammalian culture.

While cell hashing enhances single-cell RNA sequencing (scRNA-seq) efficiency and minimizes batch effects, commercial mouse hashtags often fail in FVB/N and several other strains due to antibody-epitope incompatibility. We describe a robust alternative utilizing biotinylated antibody cocktails and streptavidin-conjugated oligos to enable reliable sample multiplexing. This approach was validated in FVB/N lung tissues, yielding high-quality single-cell libraries. Our protocol offers a practical solution for researchers requiring strain-specific or custom-designed multiplexing strategies for single-cell transcriptomics. Key features • Strain-specific compatibility: Resolves the known H-2q haplotype mismatch in FVB/N mice that fails standard commercial MHC-I hashtag antibodies in cell hashing. • Multi-omic 5' workflow integration: Enables simultaneous sample multiplexing with 10× Genomics 5 chemistry, facilitating joint gene expression and V(D)J repertoire (TCR/BCR) profiling. • Enhanced non-immune cell labeling: Incorporates anti-CD326 (Ep-CAM) to ensure robust hashing of epithelial and tumor cells that may exhibit MHC-I downregulation or lack CD45. • Customizable biotin-streptavidin framework: Provides a modular system using biotinylated antibody cocktails and streptavidin-barcodes, adaptable for any mouse strain or tissue-specific cell markers.

Unsaturated fatty acids (UFAs) play key roles in essential cellular functions such as membrane dynamics, metabolism, and animal development. Disruptions in UFA metabolism are linked to metabolic, cardiovascular, and neurodegenerative disorders. Cellular UFAs composition and quantification are normally determined using methods such as gas chromatography and/or mass spectrometry, which require extraction procedures and prevent analysis of live specimens. Here, we describe a protocol that employs uniform 13C isotope labeling and high-resolution 2D solution-state nuclear magnetic resonance (NMR) spectroscopy to analyze lipid composition and fatty acid unsaturation directly in the model organism Caenorhabditis elegans. The approach enables in vivo assessment of lipid storage compositions with sufficient resolution and sensitivity to distinguish wild-type animals from those with altered fatty acid desaturation. Complementary analysis of total lipid extracts provides information regarding lipid molecules that are not detected in vivo, such as phospholipid molecules organized in biological membranes. Overall, this non-destructive NMR-based method offers a powerful tool for investigating lipid metabolism in C. elegans and other small model systems that can be isotopically enriched. Key features • Solution-state NMR spectroscopy is not destructive and can be used on live cells and multicellular organisms. • 13C isotopic enrichment is required for high-resolution NMR analysis of lipids in live C. elegans. • Lipid signals from live worms arise from the mobile lipid phase in lipid droplets. • NMR provides readouts of lipid compositions in live animals at a highly sensitive rate, enabling precise interpretation of the whole cell lipid metabolism.

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. Key features • Activity measurement of isolated photosynthetic complexes. • Assessing interactions between complexes in the photosynthetic apparatus.

We present a protocol to allow continuous assessment of cell death in Arabidopsis thaliana (L.) seedlings by measuring the release of electrolytes from dying cells upon heat shock. The electrolyte leakage assay is a well-established method to quantify the extent of cell death of plant tissues exposed to pathogen infection, since the activation of the immune response leads to compromised membrane integrity and to the release of ions from the dying cell. This prolonged release of electrolytes is considered a hallmark of regulated cell death in plants. Heat shock in plants induces ferroptosis-like cell death, which can be suppressed either pharmacologically, using inhibitors such as ferrostatin, or genetically through knockout of ferroptosis-related genes. Here, we have adapted the electrolyte leakage assay to quantify cell death in young Arabidopsis seedlings exposed to a heat shock previously shown to induce ferroptosis-like cell death. We also illustrate how this method can be used to assess activation of ferroptosis-like cell death in whole Arabidopsis seedlings using ferrostatin or knockout mutants of potential gene candidates involved in ferroptosis-like cell death. Key features • This protocol does not require any technical experience apart from gentle handling of young seedlings and is less labor-intensive than microscopy-based cell death evaluation. • Builds upon existing methods to quantify the extent of cell death upon immune response in whole seedlings subjected to heat stress. • Only requires a conductivity meter and allows the assessment of continuous cell death using multiple parallel replicates. • The protocol demonstrates how heat shock-induced ferroptosis-like cell death can be inhibited pharmacologically or genetically in whole seedlings, supported with quantitative data.

Structural proteomics methods allow for the proteome-wide interrogation of protein structural differences between two different conditions. Limited proteolysis mass spectrometry (LiP-MS), as originally implemented by the Picotti lab, utilizes a promiscuous protease to cleave at solvent-exposed regions of a protein to encode structural information, which is then read out with mass spectrometry proteomics. Here, we present a protocol that details experimental steps and data analysis for a LiP-MS workflow. First, tissue is homogenized under native conditions and then subjected to limited proteolysis using proteinase K (PK). The samples are prepared for mass spectrometry, and data are acquired using either data-dependent acquisition (DDA) or data-independent acquisition (DIA). Raw data is processed using FragPipe, and raw ion abundances are processed in FragPipe Limited-Proteolysis Processor (FLiPPR). Proteins with structural changes between the two conditions are identified in a proteome-wide manner. Key features • Protocol describes how to perform limited proteolysis mass spectrometry to identify proteins in brain tissue with structural changes proteome-wide between two experimental conditions. • Includes context for how to ensure results are reliable, using permutation analyses. • Utilizes tools (FragPipe and FLiPPR) that are free and open source. • Sample preparation can be performed in two days, not including mass spec acquisition and data analysis.

The placenta is a metabolically active organ whose mitochondrial activity is tightly linked to fetal growth, oxygenation, and nutrient transport, mediating fetal susceptibility to environmental exposures. Accordingly, aberrant mitochondrial function has been implicated in the progression of placental dysfunction. However, existing respirometry platforms require primarily fresh or cryopreserved placental tissue and offer limited throughput, rendering these platforms impractical in the context of large-scale placental dissections. Here, we describe and validate a Seahorse XF approach for measuring mitochondrial respiration in previously frozen placentae, enabling the functional interrogation of placental mitochondria in prenatal studies. Our protocol fundamentally relies on the restoration of matrix substrates that are depleted due to increased mitochondrial membrane permeability following freeze-thaw cycles. We provide a strategy to assess complex I and II-associated respiration adapted for the Seahorse XFe24 Analyzer and further demonstrate comparable oxygen consumption readouts between fresh and frozen placentae. We further demonstrate distinct differences in the magnitude of oxygen consumption between fresh and frozen placentae in the absence of exogenous NADH. Taken together, we present a simplified and convenient protocol for the assessment of respiratory enzyme complex-associated respiration from archived placental tissue. Key features • This protocol is suitable for use with previously frozen mouse placental tissue. • Streamlined protocol for complex-associated respirometry assessments following large-scale placental dissections. • Respirometry data may be acquired in <4 hours.

Calcium ions serve as a universal secondary messenger, integrating diverse external signals, such as light, herbivory, and mechanical stimuli, within plant cells. However, the visualization and mechanistic dissection of calcium signaling specifically in response to mechanical stimulation remain technically challenging and underexplored in most plants. Previous studies have been largely confined to a few model systems, including Arabidopsis; here, we introduce a live-cell imaging approach using the stigmas of Torenia fournieri. This in vitro system enables multiscale observation of calcium signal patterns following controlled mechanical stimulation. This versatile platform not only simplifies the design of calcium imaging assays but also provides a tractable system for functionally validating other key molecular components in this signaling pathway. Key features • Live-cell imaging is employed to monitor calcium signals in response to mechanical stimulation, enabling examination at both the whole-organism and cellular levels. • Stigma vitality is maintained under controlled in vitro conditions throughout the imaging.