The nuclear envelope can form complex structures in physiological and pathological contexts. Current approaches to quantify nuclear envelope structures can be time-consuming or inaccurate. Here, we present a protocol to measure nuclear envelope tubules induced by DNA double-strand breaks using a mid-throughput approach. We describe steps for the induction of these nuclear envelope structures and 3D image analysis using machine-learning-based image segmentation. This protocol can be applied to analyze various nuclear envelope structures in contexts beyond DNA repair. For complete details on the use and execution of this protocol, please refer to Shokrollahi et al.1.

Bioluminescence resonance energy transfer (BRET) is widely employed for real-time monitoring of G protein-coupled receptor activity, interactions, and trafficking in heterologous cell lines, yet its use in neuronal systems remains limited. Here, we present a protocol to apply BRET assays to primary neuronal cultures from mouse embryos. We describe steps and key concepts for generating plasmid constructs and lentivirus preparations, plating and lentiviral transduction of primary cultured neurons in 96-well plates, and BRET data collection and analysis. For complete details on the use and execution of this protocol, please refer to George et al.1.

Here, we present a protocol to quantify interactions among difficult-to-express proteins from Drosophila cells using the select western blot-free tagged-protein interaction (SWFTI) assay. We describe steps for plasmid design, cell plating, protein expression, and immunoprecipitation preparation. We then detail procedures for protein labeling, gel purification, and protein quantification. This protocol offers a fluorescence-based technique for rapid quantification of ectopically expressed proteins that are fused to SNAP and CLIP tags without the need for membrane transfer. For complete details on the use and execution of this protocol, please refer to Lin et al.1.

Sensing is a critical function of artificial cells; however, this is challenging to realize using bottom-up approaches. Here, we present a protocol for building protocell membranes that sense cues important for redox biochemistry and signaling by combining synthetic phospholipids and natural lipids. We detail procedures for building giant unilamellar vesicles as protocell models that fluoresce in response to the biologically significant redox agents peroxynitrite, hydrogen peroxide, and hydrogen sulfide. For complete details on the use and execution of this protocol, please refer to (i) Gutierrez and Aggarwal et al.1 as well as (ii) Erguven and Wang et al.2.

Super-resolution imaging provides unprecedented visualization of sub-cellular structures, but the two main techniques used, single-molecule localization microscopy (SMLM) and stimulated emission depletion (STED), are not easily reconciled. We present a protocol to super-impose nanoscale protein distribution reconstructed with SMLM to sub-cellular morphology obtained in STED. We describe steps for tracking cells on etched coverslips and registering images from two different microscopes with 30-nm accuracy. In this protocol, synaptic proteins are mapped in the dendritic spines of primary neurons. For complete details on the use and execution of this protocol, please refer to Inavalli et al.1.

Tumor acidosis is one of the hallmarks indicating the initiation and progression of various cancers. Here, we present a protocol for preparing a hyperpolarized (HP) 13C-bicarbonate tissue pH MRI imaging contrast agent to detect aggressive tumors. We describe the steps for the formulation and polarization of a precursor molecule 13C-glycerol carbonate (13C-GLC), the post-dissolution reaction, and converting HP 13C-GLC to an injectable HP 13C-bicarbonate solution. We then detail procedures for MRI data acquisition to generate tumor pH maps for assessing tumor aggressiveness. For complete details on the use and execution of this protocol, please refer to Mu et al.1.

A gene-rescue experiment under a mutant background is essential to clarify gene function and the resulting biological potential in vivo. Here, we present a protocol for determining the change in interferon response by microinjecting plasmids into one-cell-stage zebrafish embryos. We describe steps for comparing the resistance potential to virus infection in wild-type and knockout zebrafish larvae following plasmid microinjection. We then detail how to link the enhanced interferon immunity to the improved resistance in knockout zebrafish larvae by gene-rescue experiments. For complete details on the use and execution of this protocol, please refer to Qu et al.1.

Protein-nucleic acid interactions drive some of the most important physiological events in cells. Here, we present a protocol for detecting protein-DNA or protein-RNA interactions in vitro. We describe steps for labeling nucleic acid species and electrophoretic mobility shift assays (EMSAs). This protocol can be used to confirm suspected in vivo interactions using recombinantly expressed/purified proteins of interest and a nucleic acid substrate. It can further be used to investigate mutations that can disrupt interaction or compensatory mutations that restore it. For complete details on the use and execution of this protocol, please refer to Mansouri-Noori et al.1.

Zinc (Zn2+) plays roles in structure, catalysis, and signaling. The majority of cellular Zn2+ is bound by proteins, but a fraction of total Zn2+ exists in a labile form. Here, we present a protocol for measuring labile cytosolic Zn2+ using an in situ calibration of a genetically encoded Förster resonance energy transfer (FRET) sensor. We describe steps for producing buffered Zn2+ solutions for performing an imaging-based calibration and analyzing the imaging data generated to determine labile Zn2+ concentration in single cells. For complete details on the use and execution of this protocol, please refer to Rakshit and Holtzen et al.1.

Comprehensive characterization of small-molecule degraders, including binary and ternary complex formation and degradation efficiency, is critical for bifunctional ligand development and understanding structure-activity relationships. Here, we present a protocol for the biochemical and cellular profiling of small-molecule degraders based on CoraFluor time-resolved fluorescence resonance energy transfer (TR-FRET) technology. We describe steps for labeling antibodies and proteins, tracer saturation binding, binary target engagement, ternary complex profiling, and off-rate determination. We then detail procedures for the quantification of endogenous and GFP fusion proteins in cell lysates. For complete details on the use and execution of this protocol, please refer to Ichikawa et al.1.