生物化学

Cytosolic peptide: N-glycanase (PNGase/NGLY1 in mammals), an amidase classified under EC:3.5.1.52, is a highly conserved enzyme across eukaryotes that catalyzes the removal of N-glycans from glycoproteins, converting N-glycosylated asparagine residues into aspartic acid. This enzyme also plays a role in the quality control system for nascent glycoproteins. Despite the development of non-radioisotope-based assay systems such as those using S-alkylated RNase or fluorescent-labeled glycopeptides as substrates, these methods are incompatible with crude enzyme sources, primarily due to the degradation of reaction products by contaminating endogenous proteases. We previously developed an assay system using a 5-carboxyfluorescein-labeled glycosylated cyclo-heptapeptide (5FAM-GCP), a substrate remarkably resistant to endogenous peptidase activity. This system enables the accurate measurement of endogenous NGLY1 activity in various samples, including cell lines, tissues, peripheral blood mononuclear cells, and NGLY1-deficient patient-derived cells, without the interference of proteolytic degradation. We recently advanced this approach by producing a novel fluorescence resonance energy transfer (FRET)-based GCP probe (fGCP) and demonstrated its ability to detect endogenous NGLY1 activity across diverse enzyme sources via fluorescence on multiarray plates. This innovative and straightforward assay now offers reliable disease diagnostics and also allows the measurement of endogenous PNGase/NGLY1 activities across various organisms.

The motile parameters of kinesin superfamily proteins are fundamental to intracellular transport. Single-molecule motility assays using total internal reflection fluorescence (TIRF) microscopy are a gold standard technique for measuring the motile parameters of kinesin motors. With this technique, one can evaluate the velocity, run length, and binding frequency of kinesins on microtubules by directly observing their motility. This protocol provides a comprehensive procedure for single molecule assays of kinesins, including the preparation of labeled microtubules, the measurement of kinesin motility via TIRF microscopy, and the quantification of kinesin motor parameters.

Proteomics analysis is crucial for understanding the molecular mechanisms underlying muscle adaptations to different types of exercise, such as concentric and eccentric training. Traditional methods like two-dimensional gel electrophoresis and standard mass spectrometry have been used to analyze muscle protein content and modifications. This protocol details the preparation of muscle samples for proteomics analysis using ultra-high-performance liquid chromatography (UHPLC). It includes steps for muscle biopsy collection, protein extraction, digestion, and UHPLC-based analysis. The UHPLC method offers high-resolution separation of complex protein mixtures, providing more detailed and accurate proteomic profiles compared to conventional techniques. This protocol significantly enhances sensitivity, reproducibility, and efficiency, making it ideal for comprehensive muscle proteomics studies.

Cyclic diadenosine monophosphate (c-di-AMP) is a recently discovered second messenger that modulates several signal transduction pathways in bacterial and host cells. Besides the bacterial system, c-di-AMP signaling is also connected with the host cytoplasmic surveillance pathways (CSP) that induce type-I IFN responses through STING-mediated pathways. Additionally, c-di-AMP demonstrates potent adjuvant properties, particularly when administered alongside the Bacillus Calmette–Guérin (BCG) vaccine through mucosal routes. Because of its pivotal role in bacterial signaling and host immune response, this molecule has garnered significant interest from the pharmaceutical industry. This protocol outlines the quantification of c-di-AMP by an HPLC-based assay to enumerate the activity of c-di-AMP synthase from Mycobacterium smegmatis. The following protocol is designed to be generic, enabling the study of c-di-AMP synthase activity from other bacterial species. However, modifications may be required depending on the specific activity of c-di-AMP synthase from different bacterial sources.

The target of rapamycin (TOR) is a central hub kinase that promotes growth and development in all eukaryote cells. TOR induces protein synthesis through the phosphorylation of the S6 kinase (S6K), which, in turn, phosphorylates ribosomal S6 protein (RPS6) increasing this anabolic process. Therefore, S6K and RPS6 phosphorylation are generally used as readouts of TOR activity. Protein phosphorylation levels are measured by a western blot (WB) technique using an antibody against one specific phosphosite in cell extracts. However, at the tissue/cell-specific level, there is a huge gap in plants due to the lack of alternative techniques for the evaluation of TOR activity as there are for other organisms such as mammals. Here, we describe an in vivo protocol to detect S6K phosphorylation in tissues/cells of model photosynthetic organisms such as Arabidopsis thaliana and Chlamydomonas reinhardtii. Our proposed method consists of the immunolocalization of a phosphorylated target of TOR kinase using a fluorescent secondary antibody by confocal microscopy. The protocol involves four main steps: tissue/cell fixation, permeabilization, and incubation with primary and secondary antibodies. It is an easy technique that allows handling different samples at the same time. In addition, different ultrastructural cell markers can also be used, such as for nucleus and cell wall detection, allowing a detailed analysis of cell morphology. To our knowledge, this is the first protocol to detect TOR activity in situ in photosynthetic organisms; we consider that it will pave the research on the TOR kinase, opening new possibilities to better understand its complex signaling.

Carbohydrates serve crucial functions in most living cells, encompassing structural and metabolic roles. Within the realms of plant and algal biology, carbohydrate biosynthesis and partitioning play pivotal roles in growth, development, stress physiology, and various practical applications. These applications span diverse fields, including the food and feed industry, bioenergetics (biofuels), and environmental management. However, existing methods for carbohydrate determination tend to be costly and time-intensive. In response to that, we propose a novel approach to assess carbohydrate partitioning from small samples. This method leverages the differential solubility of various fractions, including soluble sugars, starch, and structural polymers (such as cellulose). After fractionation, a straightforward spectrophotometric analysis allows for the quantification of sugars.

The extracellular matrix (ECM) is a complex network of proteins that provides structural support and biochemical cues to cells within tissues. Characterizing ECM composition is critical for understanding this tissue component’s roles in development, homeostasis, and disease processes. This protocol describes an integrated pipeline for profiling both cellular and ECM proteins across varied tissue types using mass spectrometry–based proteomics. The workflow covers stepwise extraction of cellular and extracellular proteins, enzymatic digestion into peptides, peptide cleanup, mass spectrometry analysis, and bioinformatic data processing. The key advantages include unbiased coverage of cellular, ECM-associated, and core-ECM proteins, including the fraction of ECM that cannot be solubilized using strong chaotropic agents such as urea or guanidine hydrochloride. Additionally, the method has been optimized for reproducible ECM enrichment and quantification across diverse tissue samples. This protocol enables systematic mapping of the ECM at a proteome-wide scale.

This protocol outlines the use of the previously described sodium hypochlorite extraction method for estimating the accumulation of polyhydroxybutyrate (PHB) in bacteria. Sodium hypochlorite (NaClO) is widely used for PHB extraction as it oxidizes most components of the cells except PHB. We assessed the feasibility of using NaClO extraction for the estimation of PHB accumulation in bacterial cells (expressed as a percentage w/w). This allowed us to use a simple spectrophotometric measurement of the turbidity of the PHB extracted by NaClO as a semiquantitative estimation of PHB accumulation in the marine microorganisms Halomonas titanicae KHS3, Alteromonas sp., and Cobetia sp. However, this fast and easy protocol could be used for any bacterial species as long as some details are considered. This estimation exhibited a good correlation with the accumulation measured as dry cell weight or even with the accumulation measured by crotonic acid and HPLC quantifications. The key advantage of this protocol is how fast it allows an estimation of PHB accumulation in Halomonas, Alteromonas, and Cobetia cultures (results are available in 50 min), enabling the identification of the appropriate moment to harvest cells for further extraction, polymer characterization, and accurate quantification using more reliable and time-consuming methods. This protocol is very useful during bacterial cultivation for a quick evaluation of PHA accumulation without requiring (i) large volumes of cultures, (ii) a long time for analysis compared to dry cell weight, (iii) preparation of standard curves with sulfuric acid hydrolysis for crotonic acid quantification, or (iv) specific equipment and/or technical services for HPLC quantification.

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.

The eukaryotic cytoskeleton is formed in part by microtubules, which are relatively rigid filaments with inherent structural polarity. One consequence of this polarity is that the two ends of a microtubule have different properties with important consequences for their cellular roles. These differences are often challenging to probe within the crowded environment of the cell. Fluorescence microscopy–based in vitro assays with purified proteins and stabilized microtubules have been used to characterize polarity-dependent and end-specific behaviors. These assays require ways to visualize the polarity of the microtubules, which has previously been achieved either by the addition of fluorescently tagged motor proteins with known directionality or by fluorescently polarity marking the microtubules themselves. However, classical polarity-marking protocols require a particular chemically modified tubulin and generate microtubules with chemically different plus and minus segments. These chemical differences in the segments may affect the behavior of interacting proteins of interest in an undesirable manner. We present here a new protocol that uses a previously characterized, reversibly binding microtubule plus-end capping protein, a designed ankyrin repeat protein (DARPin), to efficiently produce polarity-marked microtubules with different fluorescently labeled, but otherwise biochemically identical, plus- and minus-end segments.