分子生物学

Thousands of RNAs are localized to specific subcellular locations, and these localization patterns are often required for optimal cell function. However, the sequences within RNAs that direct their transport are unknown for almost all localized transcripts. Similarly, the RNA content of most subcellular locations remains unknown. To facilitate the study of subcellular transcriptomes, we developed the RNA proximity labeling method OINC-seq. OINC-seq utilizes photoactivatable, spatially restricted RNA oxidation to specifically label RNA in proximity to a subcellularly localized bait protein. After labeling, these oxidative RNA marks are then read out via high-throughput sequencing due to their ability to induce predictable misincorporation events by reverse transcriptase. These induced mutations are then quantitatively assessed for each gene using our software package PIGPEN. The observed mutation rate for a given RNA species is therefore related to its proximity to the localized bait protein. This protocol describes procedures for assaying RNA localization via OINC-seq experiments as well as computational procedures for analyzing the resulting data using PIGPEN.

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.

Science self-efficacy describes the confidence individuals have in their ability to accomplish specific scientific practices. Self-efficacy is one factor linked to success and persistence within STEM fields. The purpose of this protocol is to provide research laboratories with effective methods for teaching and mentoring new students in molecular biology, specifically in the synthesis of virus-like particles (VLPs) derived from bacteriophages. VLPs are multivalent nanoparticle structures that can be utilized in multiple biomedical applications, including platforms for vaccine and drug delivery. Production of bacteriophage VLPs using bacterial expression systems is feasible in most laboratory settings. However, synthesizing and characterizing VLPs can be challenging for new researchers, especially those with minimal laboratory experience or a lack of foundational knowledge in molecular biology. To address this, a multi-phase training protocol was implemented to train new students in VLP synthesis, purification, and characterization. This model was optimized for training numerous high school and undergraduate students. By implementing this multi-phase methodology, the students’ confidence in their abilities to perform specific tasks increased and likely enhanced their persistence in STEM.

Transposon mutagenesis is a powerful tool for investigating gene function in bacteria, particularly in newly discovered species. In this study, we applied the hyperactive EZ-Tn5 transposase system to Pseudomonas argentinensis SA190, an endophytic bacterium known for enhancing plant resilience under drought stress. By leveraging the random amplification of transposon ends (RATE)-PCR method, we successfully mapped the insertion sites of the transposon within the SA190 genome. This approach enabled the precise identification of disrupted genes, offering insights into their roles in bacterial function and interaction with host plants. Our comprehensive protocol, including competent cell preparation, transformation, and insertion site mapping, provides a reliable framework for future studies aiming to explore gene function through mutagenesis.

Transcriptional pausing dynamically regulates spatiotemporal gene expression during cellular differentiation, development, and environmental adaptation. Precise measurement of pausing duration, a critical parameter in transcriptional control, has been challenging due to limitations in resolution and confounding factors. We introduce Fast TV-PRO-seq, an optimized protocol built on time-variant precision run-on sequencing (TV-PRO-seq), which enables genome-wide, single-base resolution mapping of RNA polymerase II pausing times. Unlike standard PRO-seq, Fast TV-PRO-seq employs sarkosyl-free biotin-NTP run-on with time gradients and integrates on-bead enzymatic reactions to streamline workflows. Key improvements include (1) reducing experimental time from 4 to 2 days, (2) reducing cell input requirements, and (3) improved process efficiency and simplified command-line operations through the use of bash scripts.

This manuscript details protocols for the ZnCl₂ precipitation-assisted sample preparation (ZASP) for proteomic analysis. By inducing protein precipitation with ZASP precipitation buffer (ZPB, final concentration of ZnCl ₂ at 100 mM and 50% methanol), ZASP depletes harsh detergents and impurities, such as sodium dodecyl sulfate (SDS), Triton X-100, and urea, at high concentrations in solution from protein solutions prior to trypsin digestion. It is a practical, robust, and cost-effective approach for proteomic sample preparation. It has been observed that 90.2% of the proteins can be recovered from lysates by incubating with an equal volume of ZPB at room temperature for 10 min. In 1 h of data-dependent acquisition (DDA) analysis on an Exploris 480, 4,037 proteins and 25,626 peptides were quantified from 1 μg of mouse small intestine proteins, reaching a peak of 4,500 proteins and up to 30,000 peptides with 5 μg of input. Additionally, ZASP outperformed other common sample preparation methods such as sodium deoxycholate (SDC)-based in-solution digestion, acetone precipitation, filter-aided sample preparation (FASP), and single-pot, solid-phase-enhanced sample preparation (SP3). It demonstrated superior performance in protein (4,456 proteins) and peptide identification (29,871 peptides), lower missing cleavage rates (16.3%), and high reproducibility (Pearson correlation coefficients of 0.96 between replicates) with similar protein distributions and cellular localization patterns. Significantly, the cost of ZASP per sample with 100 μg of protein as input is lower than 30 RMB, including the expense of trypsin.

Cathepsin L (CTSL), a lysosomal cysteine protease belonging to the papain-like protease family, is primarily involved in intracellular protein degradation, antigen processing, and extracellular matrix remodeling. It plays critical roles in pathological conditions, including cancer metastasis, neurodegenerative disorders, and viral infection, due to dysregulated activity or overexpression. Thus, inhibitors targeting CTSL are under investigation for therapeutic applications. Current approaches for identifying CTSL inhibitors predominantly rely on fluorescence-labeled substrates, fluorescence resonance energy transfer (FRET), and cell-based screening assays. Here, we applied the principle of fluorescence polarization (FP) to the detection of substrate cleavage activity by CTSL through changes in millipolarization unit (mp) values and established a cost-effective, quantitative, reagent- and time-saving inhibitor high-throughput screening (HTS) assay. We also provide detailed steps for the expression and purification of highly active CTSL from eukaryotic cells, which lays a solid foundation for the FP-based assay. A key advantage of this assay lies in its reduced susceptibility to fluorescence interference, as the fluorescein isothiocyanate (FITC) fluorophore exhibits high quantum efficiency with an emission peak at 535 nm—a wavelength range distinct from most naturally occurring fluorescent molecules. The assay’s adaptability to reaction time, temperature, and dimethyl sulfoxide (DMSO) concentration minimizes false-positive or false-negative results caused by minor experimental inconsistencies, streamlining the screening process. Furthermore, the protocol requires fewer operational steps, reduced incubation time, and lower quantities of CTSL and substrates compared to conventional methods. This rapid, cost-effective, and scalable approach aligns well with the demands of HTS platforms.

We recently developed an approach for cell type–specific CRISPR/Cas9 editing and transgene expression using a single viral vector. Here, we present a protocol describing how to design and generate plasmids and adeno-associated viruses (AAVs) compatible with this single-vector gene editing approach. This protocol has four components: (1) guide RNA (gRNA) design to target specific genes of interest, (2) ligation and cloning of CRISPR-competent AAV vectors, (3) production of vector-containing AAVs, and (4) viral titer quantification. The resultant vectors are compatible for use with mouse lines expressing the Cas9 protein from Streptococcus pyogenes (SpCas9) and Cre recombinase to enable selective co-expression of standard neuroscience tools in edited cells. This protocol can produce AAVs of any serotype, and the resulting AAVs can be used in the central and peripheral nervous systems. This flexible approach could help identify and test the function of novel genes affecting synaptic transmission, circuit activity, or morphology with a single viral injection.

This protocol provides a step-by-step approach for generating single-gene knockout in hard-to-transfect suspension immune cell lines like THP1, specifically demonstrated by knocking out the GSDMD gene. By employing CRISPR-Cas9 system delivered via lentivirus, this protocol enables precise gene disruption through targeted single-guide RNAs (sgRNAs). Key steps include designing specific sgRNAs, cloning them into a CRISPR vector, viral packaging, and transducing the target cells, followed by selection and validation. This optimized protocol is particularly useful for functional studies in immune cells, allowing researchers to reliably explore gene function in complex cellular pathways.

The cAMP-dependent protein kinase (PKA) is one of the most extensively distributed kinases among intracellular signal cascades, with a pivotal role in the regulation of various processes, including the capacitation of sperm cells. Traditional assessments of PKA activity rely on the utilization of [γ-³²P] ATP and the Kemptide peptide as a substrate. This strategy presents several major drawbacks, including high costs and health risks derived from the manipulation of radioactive isotopes. In this work, we introduce an enhanced non-radioactive assay to quantify PKA activity, termed kinase mobility shift assay (KiMSA), based on the use of a fluorescent-labeled Kemptide (Kemptide-FITC). Once the kinase reaction is terminated, the products can be easily resolved through electrophoresis on an agarose gel and quantified by fluorescence densitometry. We show that KiMSA is suitable for isolated PKA as well as for the enzyme in cell extracts. In addition, it enables quantification of PKA activity during the progression of mouse sperm capacitation. Furthermore, the assay enables monitoring the inhibition of PKA with pharmacological inhibitors in live cells. Therefore, the experimental and optimal assay conditions are set so that KiMSA can be used to assess in vitro as well as in vivo PKA activity in sperm cells. Finally, this method allows for measurement of cAMP concentrations, rendering a versatile technique for the study of cAMP/PKA pathways.