Serum is a complex mixture of proteins that originate from a wide range of cells and tissues. At present, it is difficult to know what set of proteins any given tissue contributes to the circulating proteome. Here, we describe a method of using proximity biotinylation of proteins that normally transit through the endoplasmic reticulum to profile secreted proteins from cells in culture. We also describe a mouse model that enables the elucidation of the in vivo tissue-specific secretome. As examples, we demonstrate how we can readily identify in vivo endothelial-specific secretion, and how this model allows for the characterization of muscle-derived serum proteins that either increase or decrease with exercise. Our "Secretome Mouse" model is broadly applicable to other cell and tissue types.

The use of antibodies as research tools has contributed to the characterization of specific biomolecules and the understanding of their role in biological processes and disease. However, the usefulness of antibodies is only enhanced by their specificity for the target molecule because off-target and non-specific reactivity confounds results. Using accessible websites and standard hybridoma technology, we describe the development of targeted antibodies with high specificity and antibody validation using genetically deficient animals.

Gene expression is frequently regulated with cell type specificity and at multiple levels, including through tightly-controlled protein synthesis. While existing methods of protein synthesis profiling with non-canonical amino acid or isotope-containing amino acid labeling are effective and efficient for use in cultured cells, they are often costly to carry out in vivo. We previously developed a method to visualize and capture nascent proteomes in a cell type-specific manner in Drosophila brain explants using phenylacetyl-OPP (PhAc-OPP), a modified form of the puromycin analog O-propargyl-puromycin (OPP) that can incorporate into growing polypeptide chains. Targeted expression of Penicillin G acylase (PGA) in a cell population of interest removes an enzyme-labile blocking group on PhAc-OPP, thus permitting OPP-dependent labeling of nascent proteins in that cell population. This method, which we call POPPi (PGA-dependent OPP incorporation), provides a versatile approach to visualize or identify proteins synthesized in cell populations of interest in vivo. Here, we provide detailed protocols for labeling newly-synthesized protein in Drosophila brain cell populations by POPPi followed by detection via immunofluorescence or by capture and protein identification.

Combining MetRS*-based cell-selective protein labeling with mass spectrometry-based proteomics is a powerful approach for investigating intercellular communication within tissues. Cell-selective labeling overcomes limitations of cell sorting techniques and facilitates cell type-specific proteome and secretome analyses in vivo. Our recent work has showcased the application of this method for the comprehensive proteomic characterization of cellular proteins in tissues, as well as released proteins in the bloodstream. Here, we present experimental guidelines for MetRS*-based cell-selective proteomics experiments in vivo and a detailed sample preparation protocol for tissues and body fluids.

High-throughput screening (HTS) is a cornerstone of modern drug discovery and particularly important for the development of novel anticancer therapeutics as it allows to identify agents with specific biological activities among a large number of chemical compounds. So far, most anticancer HTS approaches focused either on specifically targeting pathways critical for tumor development or on identifying drugs capable of increasing the immunogenicity of cancer cells, thus directly decreasing tumor cell viability or rendering them detectable to the immune system, respectively. Here, we employ an in vitro antigen presentation system for HTS with the purpose to identify agents that exhibit immunostimulatory potential in the context of dendritic cell (DC)-mediated antigen presentation. To this aim DCs derived from immortalized precursors are treated with an array of compounds and are then confronted with the model antigen ovalbumin (OVA). Coculture of these DCs with T-cell hybridoma cells expressing an OVA-specific T cell receptor (TCR) that triggers the secretion of IL-2 then allows the ELISA-quantifiable assessment of antigen presentation. Altogether, this system can be used to identify immunomodulatory drugs that affect the intricate crosstalk between innate and adaptive immunity.

Cell type-specific proteome labeling provides enhanced understanding of cellular function and structure within tissues by tagging proteins during translation, while cells and tissues are in their "native" states. TurboID is a methodology to enable rapid and efficient biotinylation of proteins. New TurboID viral constructs and a Cre-mediated TurboID transgenic mouse line enable cell type-specific proteomic investigations in cell culture and in vivo settings. Together, these new tools enable diverse studies designed to interrogate individual cell type contributions within complex multi-cellular systems. While biotin-based labeling enables enrichment of the labeled proteome via immunoprecipitation, it is also compatible with biotin/streptavidin-based immunoassays, including the Luminex xMAP multiplexed immunoassay platform. Here, we detail protocols to utilize existing commercially available Luminex kits to directly detect TurboID-biotinylated (and therefore cell type-specific) proteins of interest from cell culture and bulk tissue samples. Luminex immunoassays have multiple advantages compared to other methodologies, including (1) requiring small sample volumes/masses, (2) direct immuno-reaction-based quantification from a target sample without intermediate processing steps, (3) reduced costs and (4) direct readout. Below, we describe an adapted Luminex xMAP protocol to quantify phospho-proteins and cytokines from TurboID-labeled cells or tissues with cell type-specificity.

Inter-organ communication, including the release of secreted proteins, plays a key role in synchronized physiological responses and organismal homeostasis. Recent studies have emphasized functions of muscle-secreted proteins (i.e., myokines), in regulating metabolic pathways and improving metabolic dysfunction distally in the liver. Thus, experimental workflows to study myokines and their impact on target cell types are of scientific value. Here, we describe a cell culture-based method to investigate communication from muscle to liver mediated by secreted proteins. Briefly, C2C12 myoblasts are differentiated into myotubes, myotube-conditioned media is collected, and myotube-secreted proteins are isolated and stored. To demonstrate the utility of this method, AML12 hepatocytes were treated with myotube-secreted proteins and effects on bioenergetics were assessed. This method can be useful as a proof of principle tool, for mechanistic studies, or paired with proteomic or biochemical analyses to identify novel myokines. We also envision it is adaptable in terms of cell type, downstream application, and signaling direction.

Extracellular ATP (eATP) serves as a crucial signaling molecule in diverse physiological and pathological processes, including neurotransmission, inflammation, and cancer. Despite its importance, accurate measuring eATP dynamics in vivo has remained a significant technical challenge. Traditional methods, such as soluble luciferase systems, fluorescent probes, microelectrode biosensors, and high-performance liquid chromatography (HPLC), exhibit limitations in spatial resolution, tissue permeability, and real-time monitoring. Fluorescent probes offer high spatial resolution but are hindered by poor tissue penetration and the need for excitation light. Microelectrode biosensors provide localized detection but require invasive procedures, while HPLC, though highly sensitive, is restricted to ex vivo applications.To address these limitations, the plasma-membrane-targeted luciferase (pmeLUC) probe was developed. This bioluminescent system allows real-time, quantitative monitoring of eATP levels in living cells and animal models without the need for external excitation light. The pmeLUC is anchored on the outer surface of the plasma membrane, positioning its catalytic site extracellularly for direct eATP sensing. Its high sensitivity, tissue permeability, and adaptability for both in vitro and in vivo studies have enabled significant advancements in understanding eATP dynamics across different pathological contexts, including tumor microenvironments, immune responses, and brain injury models. Furthermore, the creation of pmeLUC-transgenic mice and of AAV-mediated delivery systems, has expanded the applications of this tool for longitudinal and systemic monitoring of eATP in living organisms. This review highlights the rationale behind choosing pmeLUC over other methodologies, emphasizing its superior capabilities in overcoming existing technical barriers and advancing eATP research in both basic and translational sciences.

S-palmitoylation of cysteine residues is the only lipid-based posttranslational modification of proteins that is reversible and therefore has important implications in cellular function. S-palmitoylation has been associated with several cellular processes (e.g., cell signaling, protein transport, cell cycle, immune response, lipid metabolism, host-pathogen interaction) and human diseases, including neurological disorders, cancer, and infectious diseases. However, S-palmitoylation research has been hampered by the cumbersome experimental protocols necessary for its study. Currently, there are two main methodologies that, coupled with mass spectrometry (MS), allow the study of S-palmitoylated proteins proteome-wide. They mainly differ in the way of labeling palmitoylated proteins: one relies on "metabolic labeling" with a palmitic acid analog in living cells, while the other is based on "chemical labeling" of thiol groups derived from palmitoylated sites in extracted proteins. Although metabolic labeling is restricted to cultured cells, we will focus on this technique as it is more sensitive and specific than others. Here, we describe the protocol to measure palmitoylation in cancer cells using metabolic labeling coupled to SILAC-based mass spectrometry quantification, which can be applied to other mammalian cell models. Facilitating the use of this methodology will extend the knowledge of palmitoylation signaling and unravel potential therapeutic avenues for diseases in which this unexplored modification is implicated.

Several studies have shown that extracellular vesicles (EVs) are biosynthesized in all cells and then transported to other cells in an endocrine or paracrine manner through the bloodstream. They are considered to be taken up via the EV receptor in the destination cell membrane, resulting in a subsequent intracellular signal transduction cascade induced by the contents inside EV particles (e.g., DNA, RNA, miRNA protein, etc.). Since EVs are synthesized and secreted by all cells, it is important to "classify" them for EV research. However, challenges such as the numerous types and large number of EVs in biological samples and the complexity of target antigens for classification necessitate the development of more effective experimental methods. This chapter discusses a novel strategy to identify specific EV surface markers for classifying EVs using the "proximity labeling" analysis, which labels proximal molecular groups typically used for protein-protein interaction analysis in recent years.

One important strategy of cells to communicate with their environment is the release of proteins which can serve as signals for other cells nearby or at distant places in a complex organism. A first step in the characterization and investigation of cell-cell communication in this context is to figure out which proteins are released from cells under defined experimental conditions. Here, we present an approach that will give rise to a high-quality secretome and detects proteins that will be confidently released by cultured cells. This approach is based on the separate preparation of proteins from conditioned medium and corresponding cell lysates. After protein digestion and quantitative mass spectrometric analysis, protein abundances are compared and proteins showing a significantly higher abundance in the secretome are identified. We assume that these proteins have a higher probability of being released by well-directed processes and not simply by contamination of (dead) cells. We show an optimized protocol in which samples from primary human normal dermal fibroblasts (NHDF) are prepared with the single-pot solid-phase-enhanced sample preparation (SP3) method from only 450μL conditioned medium along with one-hour gradient separations and data-independent mass spectrometric data acquisition.

In the heart, interaction among cells is essential for homeostasis and response to injury. Identifying paracrine/autocrine factors released by cardiomyocytes is fundamental to understand this complex network. However, limitations in the tools available to culture and manipulate isolated adult cardiomyocytes have hindered the study of these released factors. Here, we outline a protocol for profiling the secretome from isolated and in vitro cultured adult cardiomyocytes, using the well-established proximity ligation system.

Proteome-level investigations of distinct cell types while retaining their native states in tissue can provide key insights into disease mechanisms that may not be captured by transcriptomic studies. We describe protocols to achieve cell type-specific in vivo biotinylation of proteins (CIBOP) in mouse brain. CIBOP uses a proximity labeling approach in which biotin ligase TurboID is expressed in specific cell types, leading to broad proteomic biotinylation. Subsequently, biotinylated proteins can be enriched from bulk tissue homogenates without requiring cell type isolation, followed by mass spectrometry-based quantitative proteomics of biotinylated proteins, yielding cell type-specific proteomes. We showcase CIBOP to label neurons and astrocytes, using adenovirus-based as well as transgenic approaches. This versatile pipeline may be readily applicable to various cell types as well as to neurological and non-neurological disease model systems.

Mouse blood extraction is a crucial technique in a wide range of scientific research, including immunological studies. Various blood sampling methods have been outlined and categorized based on factors such as sample volume, sampling frequency, and the necessity of anesthesia. However, the complexity of these techniques arises from the anatomy and size of the mouse, potentially impacting sample quality, endangering the mouse's well-being, and influencing its behavior and the accuracy of analytical results. In this context, the protocol for blood extraction from the saphenous vein of the leg is presented with the goal of minimizing stress in animals and ensuring the safe collection of samples of suitable quality and quantity for different immunological analyses. Thus, this presented technique aims to reduce analytical variations resulting from distress, inflammation, or the use of anesthesia, which might affect subsequent immunological studies.

Communication between tissues or different cells within a tissue is often a result of secreted molecules such as metabolites, lipids, nucleic acids, or proteins (referred to as the secretome). These enter the extracellular space and may subsequently pass into the circulation. Depending on their nature, concentration and context, these molecules initiate specific responses in their target cells. Environmental stimuli such as exercise and cold exposure, but also different diseases, are known to significantly alter the secretome and thereby affect whole body homeostasis. Thus, identifying these factors is of great interest. The analysis of secreted proteins, however, represents a unique challenge for the field. This is mainly because mass spectrometry can be limited by the dynamic range problem, whereby the detection of low abundance polypeptides can be masked by the presence of high abundance proteins. Plasma, muscle, and fat all contain specific proteins of very high abundance, making it tremendously challenging to detect low abundance proteins in these biological samples. Thus, secreted, hormone-like polypeptides frequently remain undetected. Because muscle and fat are known to communicate by secretion of myokines and adipokines, respectively, we have sought to develop methods that can circumvent these issues through the isolation of extracellular fluids (EF) which surround these tissues. EFs had previously been isolated for analysis of metabolites; however, whether this method could be made useful for in depth proteomics analysis was not known. Recently, we have developed a method that modifies these procedures and makes it applicable for the study of EF proteins. We have applied this to muscle and fat EFs, but in principle, it can be used to study secreted proteins from almost any tissue in any species, including humans. A step-by-step protocol and methods of quality control are given below.

The tumor microenvironment (TME) plays a pivotal role in tumor development, influencing interactions with immune cells, tumor progression, and responses to treatment. Understanding the heterogeneity of the TME is essential for uncovering the mechanisms underlying cell-to-cell interactions and their contribution to tumor dynamics. Recent advancements in high-dimensional imaging technologies, such as multiplex immunofluorescence imaging, imaging mass cytometry (IMC), and multiplexed ion beam imaging (MIBI), have provided powerful tools to investigate the complexity of the TME. These technologies allow for the simultaneous analysis of multiple cellular markers and spatial organization of cells within tissue samples, offering detailed insights into the composition and dynamics of the TME. Specifically, IMC offers a unique advantage by enabling the detection of over 40 proteins in a single tissue slide, facilitating a deeper understanding of cellular interactions in situ, at high resolution, and with minimal interchannel crosstalk. Here, we present the analysis approaches and tools developed for imaging mass cytometry data to unravel cellular composition, spatial organization, and interactions within the TME. These methods aim to enhance our understanding of the intricate interplay between cancer cells, immune cells, and stromal components, ultimately supporting the development of more effective therapeutic strategies.

Multiple myeloma is a hematologic malignancy characterized by clonal plasma cell proliferation, with recent therapeutic advances focusing on immunotherapies targeting cell surface antigens. While several surface markers are well-characterized, there remains a critical need to identify additional specific targets for relapsed cases. Comprehensive surface proteome analysis is challenging due to the low abundance of surface proteins and limited cell numbers available from patient samples.

Tertiary lymphoid structures (TLS) are ectopic lymphoid aggregates that are correlated with improved patient outcomes in several solid cancers, including melanoma. Multiplex immunofluorescent histology (mIFH) has been used in numerous studies to identify and characterize TLS. However, detailed studies evaluating immune cell subsets and markers of immune activity at TLS sites have been limited. Here, we introduce multiplex immunofluorescence histology methods to identify TLS, their associated immune cell components, and markers of immune activity. We outline two mIFH panels for evaluating and quantifying TLS, and markers of immune activity, offering methodologies that can be used to gain a more nuanced understanding of the role and immunological activity of TLS in cancer prognosis and therapy.

Accurate identification of the locations of endogenous proteins is crucial for understanding their functions in tissues and cells. However, achieving precise cell-type-specific labeling of proteins has been challenging in vivo. A notable solution to this challenge is the self-complementing split green fluorescent protein (GFP) system. In this paper, we present a detailed protocol for labeling endogenous proteins in a cell-type-specific manner using the GFP system in fruit flies. This approach depends on the reconstitution of the GFP and GFP fragments, creating a fluorescence signal. We insert the GFP fragment into a specific genomic locus while expressing its counterpart, GFP, through an available Gal4 driver line. The unique aspect of this system is that neither GFP nor GFP alone emits fluorescence, enabling the precise detection of protein localization only in Gal4-positive cells expressing the GFP tagged endogenous protein. We illustrate this technique using the adhesion molecule gene teneurin-m (Ten-m) as a model, highlighting the generation and validation of GFP protein trap lines via Minos-mediated integration cassette (MiMIC) insertion. Furthermore, we demonstrate the cell-type-specific labeling of Ten-m proteins in the larval brains of fruit flies. This method significantly enhances our ability to image endogenous protein localization patterns in a cell-type-specific manner and is adaptable to various model organisms beyond fruit flies.

Eosinophils are a rare immune cell subset with important roles in Th2 immunity and cancer. Interleukin IL-33 (IL-33) plays important functions in the survival and activation of eosinophils. Recent evidence unraveled the role of IL-33 in activating the anti-tumor activities of eosinophils enhancing their degranulation and tumor cell killing in a contact-dependent manner. We propose a dual approach methodology to extrapolate the physical interactions of eosinophils with tumor cells as a result of eosinophil stimulation. Human eosinophils isolated from the blood of healthy donors by dextran sedimentation followed by magnetic sorting are exposed to either IL-33 (Eos33) or IL-5 (Eos5) for 18 h. These pre-conditioned cells are then co-cultured with A375P melanoma cells to monitor cell-cell interactions. Acoustic focusing flow cytometry analysis is employed to evaluate the presence of eosinophil-tumor cell conjugates after 1 h incubation of human eosinophils and A375P melanoma cells. Moreover, a 24 h time-lapse video recording approach is employed to obtain single cell tracking eosinophil profiles. This allows to quantitatively determine the interaction extent of Eos33, as opposed to Eos5, with tumor cells. In conclusion, our protocols easily and quickly allow the extrapolation of relevant kinematic and biologically relevant parameters for tumor reactive eosinophils. Furthermore, these methods are adaptable to various models with other types of immune cell subsets and cancer cells and can be implemented on different video microscopy platforms and advanced flow cytometry systems.