Cell Biology

Protein phosphorylation is a dynamic post-translational modification that regulates fundamental processes, including signal transduction, cell proliferation, differentiation, and effector function of immune cells. The Janus Kinase/Signal Transducer and Activator of Transcription (JAK/STAT) pathway is a key mediator of cytokine responses, essential for maintaining immune cell homeostasis and determining cell fate across diverse immune subsets. Dysregulation of JAK/STAT signaling has been linked to a broad spectrum of pathologies, including monogenic immune disorders, autoimmunity, and cancer. Platforms facilitating single-cell analysis of protein phosphorylation offer the ability to reveal subtle signaling defects and dissect the pleiotropy in cellular composition and phosphorylation status, providing insights into immune phenotype and function, while identifying potential therapeutic targets. While an application of cytometry-by-time-of-flight, termed phospho-CyTOF, has proven invaluable for studying protein phosphorylation in cryopreserved peripheral blood mononuclear cells (cPBMCs), its application is limited by cell loss and signaling artifacts stemming from isolation and cryopreservation. Conversely, whole blood (WB) approaches, preserving the native immune cell composition and signaling context, offer a more physiological representation but necessitate robust and consistent protocols for broad application. Herein, we present optimized dual phospho-CyTOF workflows tailored for both cPBMCs and whole blood, building upon established protocols for cytokine stimulation of both samples. These workflows facilitate comprehensive, high-dimensional profiling of JAK/STAT signaling in response to pleiotropic cytokines such as Type I interferons (IFN-α), Type II interferons (IFN-γ), and Interleukin-21 (IL-21). By leveraging CyTOF's capacity for high-dimensional profiling using pure heavy metal–labeled antibodies, these protocols aim to identify pathway-specific alterations in STAT phosphorylation across major immune subsets that may be overlooked by traditional flow cytometry. Together, these optimized dual workflows provide scalable, translationally relevant tools for dissecting the subtle and differential JAK/STAT-driven immune responses in both clinical and research settings, while also being compatible with the simultaneous assessment of crosstalk with alternative immune cell signaling pathways.

Accurate labeling of excitatory postsynaptic sites remains a major challenge for high-resolution imaging due to the dense and sterically restricted environment of the postsynaptic density (PSD). Here, we present a protocol utilizing Sylites, 3 kDa synthetic peptide probes that bind with nanomolar affinity to key postsynaptic markers, PSD-95 and Gephyrin. eSylites (excitatory Sylites) specifically target the PDZ1 and PDZ2 domains of PSD-95, enabling precise and efficient labeling of excitatory postsynaptic density (ePSD). In contrast, iSylites (inhibitory Sylites) bind to the dimerizing E-domain of the Gephyrin C-terminus, allowing selective visualization of inhibitory postsynaptic density (iPSD). Their small size reduces linkage error and enhances accessibility compared to conventional antibodies, enabling clear separation of PSD-95 nanodomains in super-resolution microscopy. The protocol is compatible with co-labeling using standard antibodies and integrates seamlessly into multichannel immunocytochemistry workflows for primary neurons and brain tissue. This method enables robust, reproducible labeling of excitatory synapses with enhanced spatial resolution and can be readily adapted for expansion microscopy or live-cell applications.

Studying G protein-coupled receptor (GPCR) activation of heterotrimeric G proteins is crucial for understanding diverse physiological processes and developing novel therapeutics. Traditional methods to assay GPCR activation of G proteins, including assays of second messengers and biosensors, involve complex or indirect procedures. However, second messengers like cAMP and calcium are not direct readouts of GPCR activity due to signaling crosstalk, while biosensors can have undesired consequences due to structural alteration caused by fluorescent protein insertion. Here, we present a streamlined protocol employing GST-tagged bait proteins and epitope-embedded Gα subunits to achieve direct monitoring of Gα activity within cells. This method involves purification of GST-tagged bait constructs from bacteria and subsequent direct interaction studies with GluGlu-tagged Gα proteins expressed in any human cells of interest by including GST-tagged bait proteins in the cell lysis buffer. The approach enables sensitive detection of activated Gα within cells following extracellular stimulation. Advantages of this protocol include high sensitivity, enhanced monitoring of GPCR signaling dynamics under physiologically relevant conditions with minimum alteration in Gα, and the ability to distinguish between highly homologous isoforms within the same Gα family.

The target of rapamycin complex 1 (TORC1) is a highly conserved protein complex whose primary function is to link nutrient availability to cell growth in eukaryotes, particularly nitrogen sources. It was originally identified during the screening of Saccharomyces cerevisiae strains resistant to rapamycin treatment. For its part, S. cerevisiae is well known for being a key model organism in biological research and an essential microorganism for the fermentation of food and beverages. This yeast is widely distributed in nature, with domesticated and wild strains existing. However, little is known about what effects domestication has had on its different phenotypes; for example, how nitrogen sources are sensed for TORC1 activation and what impact domestication has had on TORC1 activation are questions that still have no complete answer. To study the genetic basis of TORC1 activation associated with domestication through approaches such as quantitative trait loci (QTL) mapping or genome-wide association studies (GWAS), and more generally for any study requiring TORC1 activity as a readout for a large number of individuals, it is necessary to have a high-throughput methodology that allows monitoring the activation of this pathway in numerous yeast strains. In this context, the present protocol was designed to assess phenotypical differences in TORC1 activation using a new reporter plasmid, the pTOMAN-G plasmid, specifically designed to monitor TORC1 activation. As a proof of concept, this methodology allowed phenotyping a large population of yeast strains derived from the 1002 Yeast Genomes Project, the most complete catalog of genetic variation in yeasts. This protocol proved to be an efficient alternative to assess TORC1 pathway activation compared to techniques based on immunoblot detection, which, although effective, are considerably more laborious. Briefly, the protocol involves the design and construction of the pTOMAN-G plasmid, which carries a construct containing the firefly luciferase gene (Luc) under the control of the TORC1-regulated RPL26A gene promoter (P_RPL26A). The protocol then details the process for selecting subgroups of yeasts based on their ability to grow under nutrient-limited conditions, using proline as the sole nitrogen source. These yeasts are then transformed with the TOMAN-G plasmid, using two alternative transformation methods. Finally, those yeasts that emit luminescence are selected, whose phenotype for TORC1 activation is measured by a nitrogen-upshift experiment in microculture. This approach, using the pTOMAN-G plasmid, offers a rapid and consistent method for assessing TORC1 signaling pathway activation in a large number of yeast strains, highlighting its usefulness to study the activation of the TORC1 pathway and the domestication process associated with it. In the future, a redesign of the plasmid could extend its use as a reporter tool to monitor the activation of the TORC1 pathway, or other pathways, in other yeast species.

Long-term depression (LTD), a key form of synaptic plasticity, is typically induced through regulated Ca²⁺ entry via NMDA receptors and achieved by prolonged (up to hundreds of seconds) low-frequency presynaptic stimulation or bath application of NMDA receptor agonists. Electrophysiological approach to LTD induction requires specialized equipment, while bath applications limit productivity, as only one neuron per sample may be recorded. Here, we present a simple and effective protocol for pharmacological modeling of LTD in primary cultured neurons. This approach relies on highly localized iontophoretic application of NMDA, which induces LTD in individual cells, enhancing experimental throughput. We have analyzed spatio-temporal patterns of iontophoretic drug delivery and demonstrated how this technique may be combined with electrophysiological and live-cell imaging approaches to investigate LTD-related changes in synaptic strength and Ca²⁺-dependent signaling of neuronal Ca²⁺ sensor proteins.

Recordings of electric potential changes on plant surfaces have been utilized to identify the components and mechanisms involved in the formation and transmission of systemic signals elicited by stimuli such as herbivory, wounding, or burning. The recorded responses, commonly referred to as slow wave or variation potentials, exhibit striking variability in their waveform. The extent to which this variability is due to differences in experimental procedures or plant biological variability remains unclear. Here, we provide a detailed and robust protocol refined from years of experience in conducting leaf surface potential recordings of Arabidopsis thaliana in response to mechanical wounding. This protocol serves as a comprehensive tutorial covering plant growth, procedures for reproducible mechanical wounding, critical aspects of electrophysiological recordings, and statistical analysis of surface potential recordings. It particularly emphasizes the construction and maintenance of electrodes, placement of the reference or ground electrode, mechanisms for wounding, and data analysis. This protocol aims to promote and facilitate the adoption, standardization, and interoperability of plant surface potential recordings among research groups, thereby increasing the reproducibility and comparability of data within the field.

ALPK1 is an atypical protein kinase that is activated during bacterial infection by ADP-heptose and phosphorylates TIFA to activate a cell signaling pathway. In contrast, specific mutations in ALPK1 allow it to also be activated by endogenous human nucleotide sugars such as UDP-mannose, leading to the phosphorylation of TIFA in the absence of infection. This protocol describes a quantitative, cell-free phosphorylation assay that can directly measure the catalytic activity of wildtype and disease-causing ALPK1 in the presence of different nucleotide sugars. In this method, overexpressed ALPK1 is first immunoprecipitated from the extracts of ALPK1 knockout HEK-Blue cells transfected with plasmids encoding either FLAG-tagged wildtype or mutant ALPK1, and then subjected to a radioactive phosphorylation assay in which the phosphorylation of purified GST-tagged TIFA by ALPK1 is quantified by measuring the incorporation of radioactivity derived from radiolabeled ATP.

Chlamydomonas (Chlamydomonas reinhardtii) is a unicellular model alga that has been shown to undergo programmed cell death (PCD) that can be triggered in response to different stresses. We have recently shown that Chlamydomonas is particularly well suited to the study and quantification of PCD. We have shown for the first time that S-nitrosoglutathione (GSNO), a nitric oxide (NO) donor, is able to induce PCD and can be used as a study system in Chlamydomonas. In this article, we provide a simple and robust protocol for quantifying GSNO-induced PCD, which can be adapted to any other treatment. We explain how to detect NO production in the cell following GSNO treatment. We show how PCD can be identified simply by analyzing the degradation profile of genomic DNA. We also provide an easy and reproducible cell death quantification protocol, which makes it possible to follow the course of PCD over time and highlight very fine differences in the number of affected cells between different samples.

Stomata are pores surrounded by a pair of specialized cells, called guard cells, that play a central role in plant physiology through the regulation of gas exchange between plants and the environment. Guard cells have features like cell-autonomous responses and easily measurable readouts that have turned them into a model system to study signal transduction mechanisms in plants. Here, we provide a detailed protocol to analyze different physiological responses specifically in guard cells. We describe, in detail, the steps and conditions to isolate epidermal peels with tweezers and to analyze i) stomatal aperture in response to different stimuli, ii) cytosolic parameters such as hydrogen peroxide (H₂O₂), glutathione redox potential (E_GSH), and MgATP^-2 in vivo dynamics using fluorescent biosensors, and iii) gene expression in guard cell–enriched samples. The importance of this protocol lies in the fact that most living cells on epidermal peels are guard cells, enabling the preparation of guard cell–enriched samples.

Plasma membrane proteins mediate important aspects of physiology, including nutrient acquisition, cell–cell interactions, and monitoring homeostasis. The trafficking of these proteins, involving internalisation from and/or recycling back to the cell surface, is often critical to their functions. These processes can vary among different proteins and cell types and states and are still being elucidated. Current strategies to measure surface protein internalisation and recycling are typically microscopy or biochemical assays; these are accurate but generally limited to analysing a homogenous cell population and are often low throughput. Here, we present flow cytometry–based methods involving probe-conjugated antibodies that enable quantification of internalisation or recycling rates at the single-cell level in complex samples. To measure internalisation, we detail an assay where the protein of interest is labelled with a specific antibody conjugated to a fluorescent oligonucleotide-labelled probe. To measure recycling, a specific antibody conjugated to a cleavable biotin group is employed. These probes permit the differentiation of molecules that have been internalised or recycled from those that have not. When combined with cell-specific marker panels, these methods allow the quantitative study of plasma membrane protein trafficking dynamics in a heterogenous cell mixture at the single-cell level.