Molecular Biology

Accurate measurement of protein translation rates is crucial for understanding cellular processes and disease mechanisms. However, existing methods for quantifying translation rates in yeast cells are limited. Here, we present a streamlined protocol for measuring protein translation rates in Saccharomyces cerevisiae using the methionine analog L-azidohomoalanine (AHA), which is the L isoform of this synthetic amino acid, and fluorophore-labeled alkyne dye-based Click chemistry. Our method involves incorporating AHA into newly synthesized proteins, followed by detection using confocal microscopy, flow cytometry, and SDS-PAGE. We validated our protocol by measuring translation rates under various stress conditions, including heat stress, endoplasmic reticulum (ER) stress induced by tunicamycin, and translation inhibition by cycloheximide. Confocal microscopy revealed differential AHA incorporation and fluorescence intensity across conditions. Flow cytometry quantitatively confirmed significant increases in translation rates under heat stress and decreases under ER stress compared to unstressed conditions at 6 and 24 h post-treatment. Imaging of gels under fluorescence detectors following SDS-PAGE further visualized newly synthesized proteins, with no detectable translation after cycloheximide treatment. Our protocol offers enhanced precision and selectivity compared to existing methods for mammalian cells and represents the first standardized approach for measuring translation rates in yeast. Despite limitations in required specialized equipment and expertise, this method holds promise for diverse applications in biotechnology and biomedical research, enabling investigations into protein synthesis regulation in yeast systems.

Immunofluorescence staining is a technique that permits the visualization of components of various cell preparations. Manchette, a transient structure that is only present in elongating spermatids, is involved in intra-manchette transport (IMT) for sperm flagella formation. Sperm flagella are assembled by intra-flagellar transport (IFT). Due to the big complexes formed by IMT and IFT components, it has been challenging to visualize these components in tissue sections. This is because the proteins that make up these complexes overlap with each other. Testicular tissue is digested by a combination of DNase I and Collagenase IV enzymes and fixed by paraformaldehyde and sucrose. After permeabilization with Triton X-100, testicular cells are incubated with specific antibodies to detect the components in the manchette and developing sperm tails. This method allows for cell type–specific resolution without interference from surrounding cells like Sertoli, Leydig, or peritubular myoid cells. Additionally, isolated cells produce cleaner immunofluorescence signals compared to other methods like tissue section/whole mount, making this method the best fit for visualizing protein localization in germ cells when spatial context is not being considered. Hence, this protocol provides the detailed methodology for isolating male mice germ cells for antibody-targeted immunofluorescence assay for confocal/fluorescence microscopy.

Fluorescent protein biosensors (FPBs) that turn on—go from dark to bright upon binding their ligands—enable the detection of targets in living cells with high sensitivity and spatial localization. Several approaches exist for creating turn-on FPBs, most notably the method that gave rise to the GCaMP family of genetically encoded calcium indicators. However, it remains challenging to modify these sensors to recognize new ligands. We recently developed adaptable turn-on maturation (ATOM) biosensors, in which target recognition by a small binding domain triggers chromophore maturation in the fluorescent protein to which it is attached. ATOM sensors are advantageous because they are generalizable (by virtue of the monobody and nanobody binding domains) and modular (binding domains and fluorescent proteins of various colors can be mixed and matched for multiplexed imaging), capable of detecting endogenously expressed proteins, and able to function in subcellular compartments including the cytoplasm, nucleus, endoplasmic reticulum, and mitochondria. The protocols herein detail how to design, clone, and screen new ATOM sensors for detecting targets of choice. The starting materials are the genes encoding for a monobody or nanobody and for a cyan, yellow, or red fluorescent protein. We also present general guidelines for creating ATOM sensors using binding domains other than nanobodies and monobodies.

Candida albicans is the most common human fungal pathogen, able to reside in a broad range of niches within the human body. Even though C. albicans systemic infection is associated with high mortality, the fungus has historically received relatively little attention, resulting in a lack of optimized molecular and fluorescent tools. Over the last decade, some extra focus has been put on the optimization of fluorescent proteins (FPs) of C. albicans. However, as the FPs are GFP-type, they require an aerobic environment and a relatively long period to fully mature. Recently, we have shown the application of a novel type of fluorogen-based FP, with an improved version of fluorescence activating and absorption shifting tag (iFAST), in C. albicans. Due to the dynamic relation between iFAST and its fluorogens, the system has the advantage of being reversible in terms of fluorescence. Furthermore, the combination of iFAST with different fluorogens results in different spectral and cellular properties, allowing customization of the system.

Bottom-up proteomics utilizes sample preparation techniques to enzymatically digest proteins, thereby generating identifiable and quantifiable peptides. Proteomics integrates with other omics methodologies, such as genomics and transcriptomics, to elucidate biomarkers associated with diseases and responses to drug or biologics treatment. The methodologies employed for preparing proteomic samples for mass spectrometry analysis exhibit variability across several factors, including the composition of lysis buffer detergents, homogenization techniques, protein extraction and precipitation methodologies, alkylation strategies, and the selection of digestion enzymes. The general workflow for bottom-up proteomics consists of sample preparation, mass spectrometric data acquisition (LC-MS/MS analysis), and subsequent downstream data analysis including protein quantification and differential expression analysis. Sample preparation poses a persistent challenge due to issues such as low reproducibility and inherent procedure complexities. Herein, we have developed a validated chloroform/methanol sample preparation protocol to obtain reproducible peptide mixtures from both rodent tissue and human cell line samples for bottom-up proteomics analysis. The protocol we established may facilitate the standardization of bottom-up proteomics workflows, thereby enhancing the acquisition of reliable biologically and/or clinically relevant proteomic data.

Membrane proteins play critical roles in cell physiology and pathology. The conventional way to study membrane proteins at protein levels is to use optimal detergents to extract proteins from membranes. Identification of the optimal detergent is tedious, and in some cases, the protein functions are compromised. While this detergent-based approach has produced meaningful results in membrane protein research, a lipid environment should be more suitable to recapture the protein’s native folding and functions. This protocol describes how to prepare amphipathic membrane scaffold-proteins (MSPs)-based nanodiscs of a cation-coupled melibiose symporter of Salmonella enterica serovar Typhimurium (MelB_St), a member of the major facilitator superfamily. MSPs generate nano-assemblies containing membrane proteins surrounded by a patch of native lipids to better preserve their native conformations and functions. This protocol requires purified membrane protein in detergents, purified MSPs in solution, and detergent-destabilized phospholipids. The mixture of all three components at specific ratios is incubated in the presence of Bio-Beads SM-2 resins, which absorb all detergent molecules, allowing the membrane protein to associate with lipids surrounded by the MSPs. By reconstituting the purified membrane proteins back into their native-like lipid environment, these nanodisc-like particles can be directly used in cryo-EM single-particle analysis for structure determination and other biophysical analyses. It is noted that nanodiscs may potentially limit the dynamics of membrane proteins due to suboptimal nanodisc size compared to the native lipid bilayer.

Biological processes are dependent on protein concentration and there is an inherent variability among cells even in environment-controlled conditions. Determining the amount of protein of interest in a cell is relevant to quantitatively relate it with the cells (patho)physiology. Previous studies used either western blot to determine the average amount of protein per cell in a population or fluorescence intensity to provide a relative amount of protein. This method combines both techniques. First, the protein of interest is purified, and its concentration determined. Next, cells containing the protein of interest with a fluorescent tag are sorted into different levels of intensity using fluorescence-activated cell sorting, and the amount of protein for each intensity category is calculated using the purified protein as calibration. Lastly, a calibration curve allows the direct relation of the amount of protein to the intensity levels determined with any instrument able to measure intensity levels. Once a fluorescence-based instrument is calibrated, it is possible to determine protein concentrations based on intensity.

Key features

• This method allows the evaluation and comparison of protein concentration in cells based on fluorescence intensity.

• Requires protein purification and fluorescence-activated cell sorting.

• Once calibrated for one protein, it allows determination of the levels of this protein using any fluorescence-based instrument.

• Allows to determine subcellular local protein concentration based on combining volumetric and intensity measurements.

Graphical overview

Protein level quantification across fluorescence-based platforms