细胞生物学

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.

Sterol regulatory element binding proteins (SREBPs) are transcription factors that reside in the endoplasmic reticulum (ER) membrane as inactive precursors. To be active, SREBPs are translocated to the Golgi where the transcriptionally active N-terminus is cleaved and released to the nucleus to regulate gene expression. Nuclear SREBP levels can be determined by immunoblot analysis; however, this method can only determine the steady-state levels of nuclear SREBPs and does not capture the actual status of activation. The vesicle budding assay provides an alternative way to quantify the activation of SREBPs by monitoring the initiation of SREBP translocation from the ER to the Golgi through vesicles. Microsomal membranes isolated from the liver are incubated in a reaction buffer containing the necessary components to facilitate vesicle formation. Microsomal membranes and vesicles are isolated and SREBPs are quantified in each by immunoblot analysis. The amount of SREBPs found in the budded vesicles provides an assessment of the SREBP activation in the liver.

Cryo-electron microscopy (cryo-EM) is a powerful technique capable of investigating samples in a hydrated state, compared to conventional high-vacuum electron microscopy that requires samples to be completely dry. During the drying process, numerous features and details may be lost due to damage caused by dehydration. Cryo-EM circumvents these problems by cryo-fixing the samples, thereby retaining the intact and original features of hydrated samples. This protocol describes a step-by-step cryo-scanning electron microscopy (cryo-SEM) experimental procedure with Chlorella sorokiniana as the subject. By employing filter paper as the sample substrate, we propose a simple and reliable method for cryo-fixation and freeze-fracture of Chlorella sorokiniana in water suspension. The advantage of using filter paper as a substrate lies in its ability to support a thin film of sample, enabling a cold knife to make a cut effortlessly and produce a clean freeze-fractured surface for SEM investigation. By following the approach described in this protocol, both the internal structure and surface morphology of Chlorella sorokiniana can be easily resolved with high quality. This protocol is highly versatile and can be applied to samples dispersed in water or solvents, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto filter paper.

The human T-lymphotropic virus type-1 (HTLV-1) is an oncogenic retrovirus that predominantly spreads through cell-to-cell contact due to the limited infectivity of cell-free viruses. Among various modes of intercellular transmission, HTLV-1 biofilms emerge as adhesive structures, polarized at the cell surface, which encapsulate virions within a protective matrix. This biofilm is supposed to facilitate simultaneous virion delivery during infection. Yet, the molecular and functional intricacies of viral biofilms remain largely unexplored, despite their pivotal role in understanding retroviral pathogenesis. In this study, we optimized a protocol to isolate HTLV-1 biofilms from chronically infected T cells, facilitating their structural and molecular characterization using proteomic and super-resolution microscopy analyses. This protocol involves cultivating HTLV-1 chronically infected T cells at high density to facilitate the natural detachment of viral biofilms into the supernatant. Then, employing successive centrifugations, the cells are separated from the detached biofilms, and these structures are pelleted at medium speed (10,000× g). This method circumvents the need for mechanical, chemical, or enzymatic biofilm detachment, bypasses the use of ultracentrifugation, and enables us to resuspend the biofilms in the appropriate buffer for subsequent analyses such as western blotting or super-resolution microscopy imaging as presented.

In modern science, the exchange of scientific material between different institutions and collaborating working groups constitutes an indispensable endeavor. For this purpose, bacterial strains are frequently shipped to collaborators to advance joint research projects. Bacterial strains are usually safely shipped as cultures on solid medium, whereas the shipment of liquid cultures requires specific safety measures due to the risk of leakage. Cyanobacterial cultures are frequently maintained as liquid stock cultures, and this problem typically arises. This protocol describes a new method for the shipment of liquid cyanobacterial stock cultures by agarose gel embedding (SCAGE). More specifically, a cyanobacterial culture is mixed with low-melting agarose and cast into sterile plastic bags, resulting in a thin, solid cyanobacterial agarose gel (cyanogel) that can be easily shipped. After delivery, subsequent regeneration of the cyanogel material in liquid media results in full recovery of the examined bacterial strains. Thus, the packaging method devised in the present study comprises an innovative technique to facilitate the shipment of bacterial strains, whilst eliminating previously encountered issues like cell culture leakage.

Two aconitase isoforms are present in mammalian cells: the mitochondrial aconitase (ACO2) that catalyzes the reversible isomerization of citrate to isocitrate in the citric acid cycle, and the bifunctional cytosolic enzyme (ACO1), which also plays a role as an RNA-binding protein in the regulation of intracellular iron metabolism. Aconitase activities in the different subcellular compartments can be selectively inactivated by different genetic defects, iron depletion, and oxidative or nitrative stress. Aconitase contains a [4Fe-4S]²⁺ cluster that is essential for substrate coordination and catalysis. Many Fe-S clusters are sensitive to oxidative stress, nitrative stress, and reduced iron availability, which forms the basis of redox- and iron-mediated regulation of intermediary metabolism via aconitase and other Fe-S cluster-containing metabolic enzymes, such as succinate dehydrogenase. As such, ACO1 and ACO2 activities can serve as compartment-specific surrogate markers of oxygen levels, reactive oxygen species (ROS), reactive nitrogen species (RNS), iron bioavailability, and the status of intermediary and iron metabolism. Here, we provide a protocol describing a non-denaturing polyacrylamide gel electrophoresis (PAGE)-based procedure that has been successfully used to monitor ACO1 and ACO2 aconitase activities simultaneously in human and mouse cells and tissues.

Developing a physiologically relevant in vitro model of the respiratory epithelium is critical for understanding lung development and respiratory diseases. Here, we describe a detailed protocol in which the fetal mouse proximal epithelial progenitors were differentiated into 3D airway organoids, which contain terminal-differentiated ciliated cells and basal stem cells. These differentiated airway organoids could constitute an excellent experimental model to elucidate the molecular mechanisms of airway development and epithelial cell fate determination and offer an important tool for establishing pulmonary dysplasia disease in vitro.

The mammalian kinetochore is a multi-layered protein complex that forms on the centromeric chromatin. The kinetochore serves as the attachment hub for the plus ends of microtubules emanating from the centrosomes during mitosis. For karyokinesis, bipolar kinetochore-microtubule attachment and subsequent microtubule depolymerization lead to the development of inter-kinetochore tension between the sister chromatids. These events are instrumental in initiating a signaling cascade culminating in the segregation of the sister chromatids equally between the new daughter cells. Of the hundreds of conserved proteins that constitute the mammalian kinetochore, many that reside in the outermost layer are loaded during early mitosis and removed around metaphase-anaphase. Dynamically localized kinetochore proteins include those required for kinetochore-microtubule attachment, spindle assembly checkpoint proteins, various kinases, and molecular motors. The abundance of these kinetochore-localized proteins varies at prometaphase, metaphase, and anaphase, and is thus considered diagnostic of the fidelity of progression through these stages of mitosis. Here, we document detailed, state-of-the-art methodologies based on high-resolution fluorescence confocal microscopy followed by quantification of the levels of kinetochore-localized proteins during mitosis. We also document methods to accurately measure distances between sister kinetochores in mammalian cells, a surrogate readout for inter-kinetochore tension, which is essential for chromosome segregation.

Targeted genome editing of human pluripotent stem cells (hPSCs) is critical for basic and translational research and can be achieved with site-specific endonucleases. Cpf1 (CRISPR from Prevotella and Francisella) is a programmable DNA endonuclease with AT-rich PAM sequences. In this protocol, we describe procedures for using a single vector system to deliver Cpf1 and CRISPR RNA (crRNA) for genome editing in hPSCs. This protocol enables indel formation and homologous recombination–mediated precise editing at multiple loci. With the delivery of Cpf1 and a single U6 promoter-driven guide RNA array composed of an AAVS1-targeting and a MAFB-targeting crRNA array, efficient multiplex genome editing at the AAVS1 (knockin) and MAFB (knockout) loci in hPSCs could be achieved in a single experiment. The edited hPSCs expressed pluripotency markers and could differentiate into neurons in vitro. This system also generated INS reporter hPSCs with a 6 kb cassette knockin at the INS locus. The INS reporter cells can differentiate into β-cells that express tdTomato and luciferase, permitting fluorescence-activated cell sorting of hPSC-β-cells. By targeted screening of potential off-target sequences that are most homologous to crRNA sequences, no off-target mutations were detected in any of the tested sequences. This work provides an efficient and flexible system for precise genome editing in mammalian cells including hPSCs with the benefits of less off-target effects.