Aminoacyl-tRNA synthetases (aaRSs) provide an essential functional link between an mRNA sequence and the protein it encodes. aaRS enzymes catalyze a two-step chemical reaction that acylates specific tRNAs with a cognate α-amino acid. In addition to their role in translation, acylated tRNAs contribute to non-ribosomal natural product biosynthesis and are implicated in multiple human diseases. In synthetic biology, the acylation of tRNAs with a non-canonical α-amino acid or, more recently, a non-α-amino acid monomer is a critical first step in the incorporation of these monomers into proteins, where they can be used for fundamental and applied science. These endeavors all demand an understanding of aaRS activity and specificity. Here, we describe a liquid chromatography-mass spectrometry assay that directly monitors aaRS activity by detecting the intact acyl-tRNA product. After a simple tRNA acylation reaction workup, acyl- and non-acyl-tRNA molecules are resolved by using ion-pairing reverse-phase chromatography, and their exact masses are determined by using high-resolution time-of-flight mass spectrometry. Our assay is fast and simple, quantifies reaction yields as low as 0.23% and can also be used on tRNAs acylated with flexizyme to detect products that are undetectable by using standard techniques. The protocol requires basic expertise in molecular biology, liquid chromatography-mass spectrometry and RNase-free techniques. This protocol takes ≥5 h to complete, depending on the number of samples.

Premetastatic cancer cells often spread from the primary lesion through the lymphatic vasculature and, clinically, the presence or absence of lymph node metastases impacts treatment decisions. However, little is known about cancer progression via the lymphatic system or of the effect that the lymphatic environment has on cancer progression. This is due, in part, to the technical challenge of studying lymphatic vessels and collecting lymph fluid. Here we provide a step-by-step procedure to collect both lymph and tumor-draining lymph in mouse models of cancer metastasis. This protocol has been adapted from established methods of lymph collection and was developed specifically for the collection of lymph from tumors. The approach involves the use of mice bearing melanoma or breast cancer orthotopic tumors. After euthanasia, the cisterna chyli and the tumor are exposed and viewed using a stereo microscope. Then, a glass cannula connected to a 1 mL syringe is inserted directly into the cisterna chyli or the tumor-draining lymphatics for collection of pure lymph. These lymph samples can be used to analyze the lymph-derived cancer cells using highly sensitive multiomics approaches to investigate the impact of the lymph environment during cancer metastasis. The procedure requires 2 h per mouse to complete and is suitable for users with minimal expertise in small animal handling and use of microsurgical tools under a stereo microscope.

Cell-matrix interactions, mediated by cellular force and matrix remodeling, result in dynamic reciprocity that drives numerous biological processes and disease progression. Currently, there is no available method for directly quantifying cell traction force and matrix remodeling in three-dimensional matrices as a function of time. To address this long-standing need, we developed a high-resolution microfabricated device that enables longitudinal measurement of cell force, matrix stiffness and the application of mechanical stimulation (tension or compression) to cells. Here a specimen comprising of cells and matrix self-assembles and self-integrates with the sensor. With primary fibroblasts, cancer cells and neurons we have demonstrated the feasibility of the sensor by measuring single or multiple cell force with a resolution of 1 nN and changes in tissue stiffness due to matrix remodeling by the cells. The sensor can also potentially be translated into a high-throughput system for clinical assays such as patient-specific drug and phenotypic screening. We present the detailed protocol for manufacturing the sensors, preparing experimental setup, developing assays with different tissues and for imaging and analyzing the data. Apart from microfabrication of the molds in a cleanroom (one time operation), this protocol does not require any specialized skillset and can be completed within 4-5 h.

Metal halide perovskite semiconductors have attracted considerable attention because they enable the development of devices with exceptional optoelectronic and electronic properties via cost-effective and high-throughput chemical solution processes. However, challenges persist in the solution processing of perovskite films, including limited control over crystallization and the formation of defective deposits, leading to suboptimal device performance and reproducibility. Tin (Sn) halide perovskite holds promise for achieving high-performance thin-film transistors (TFTs) due to its intrinsic high hole mobility. Nevertheless, reliable production of high-quality Sn perovskite films remains challenging due to the rapid crystallization compared with more extensively studied lead (Pb)-based materials. Recently, composition engineering has emerged as a mature and effective strategy for realizing the high-yield fabrication of Sn halide perovskite thin films. This approach cannot only achieve improved TFT performance with high hole mobilities and current ratios, but also enable reliable device operation with hysteresis-free character and long-term stability. Here we provide the experimental procedure for precursor preparation, film and device fabrication and characterization. The entire process typically takes 20-24 h. This protocol requires a basic understanding of metal halide perovskites, perovskite film coating process, standard TFT fabrication and measurement techniques.

Sensitive, rapid and label-free biochemical sensors are needed for many applications. In this protocol, we describe biochemical detection using FLOWER (frequency locked optical whispering evanescent resonator)-a technique that we have used to detect single protein molecules in aqueous solution as well as exosomes, ribosomes and low part-per-trillion concentrations of volatile organic compounds. Whispering gallery mode microtoroid resonators confine light for extended time periods (hundreds of nanoseconds). When light circulates within the resonator, a portion of the electromagnetic field extends beyond the cavity, forming an evanescent field. This field interacts with bound analytes resulting in a change in the cavity's effective refractive index, which can be tracked by monitoring shifts in the resonance wavelength. The surface of the microtoroid can be functionalized to respond specifically to an analyte or biochemical interaction of interest. The frequency-locking feature of frequency locked optical whispering evanescent resonator means that the instruments respond to perturbations in the surface by very rapidly finding the new resonant frequency. Here we describe microtoroid fabrication (4-6 h), how to couple light into these devices using tapered optical fibers (20-40 min) and procedures for coupling antibodies as well as G-protein coupled receptors to the microtoroid's surface (from 1 h to 1 d depending on the target analyte). In addition, we describe our liquid handling perfusion system as well as the use of a rotary selector valve and custom fluidic chamber to optimize sample delivery. Step-by-step details on how to perform biosensing experiments and analyze the data are described; this takes 1-2 d.

The clinical potential of current chimeric antigen receptor-engineered T (CAR-T) cell therapy is hampered by its autologous nature that poses considerable challenges in manufacturing, costs and patient selection. This spurs demand for off-the-shelf therapies. Here we introduce an ex vivo feeder-free culture method to differentiate gene-engineered hematopoietic stem and progenitor (HSP) cells into allogeneic invariant natural killer T (NKT) cells and their CAR-armed derivatives (CAR-NKT cells). We include detailed information on lentivirus generation and titration, as well as the five stages of ex vivo culture required to generate CAR-NKT cells, including HSP cell engineering, HSP cell expansion, NKT cell differentiation, NKT cell deep differentiation and NKT cell expansion. In addition, we describe procedures for evaluating the pharmacology, antitumor efficacy and mechanism of action of CAR-NKT cells. It takes ~2 weeks to generate and titrate lentiviruses and ~6 weeks to generate mature CAR-NKT cells. Competence with human stem cell and T cell culture, gene engineering and flow cytometry is required for optimal results.

Top-down analysis of intact proteins and middle-down analysis of proteins subjected to limited digestion require efficient detection of traces of proteoforms in samples, necessitating the reduction of sample complexity by thorough pre-fractionation of the proteome components in the sample. SDS-PAGE is a simple and inexpensive high-resolution protein-separation technique widely used in biochemical and molecular biology experiments. Although its effectiveness for sample preparation in bottom-up proteomics has been proven, establishing a method for highly efficient recovery of intact proteins from the gel matrix has long been a challenge for its implementation in top-down and middle-down proteomics. As a much-awaited solution to this problem, we present an experimental protocol for efficient proteoform fractionation from complex biological samples using passively eluting proteins from polyacrylamide gels as intact species for mass spectrometry (PEPPI-MS), a rapid method for extraction of intact proteins separated by SDS-PAGE. PEPPI-MS allows recovery of proteins below 100 kDa separated by SDS-PAGE in solution with a median efficiency of 68% within 10 min and, unlike conventional electroelution methods, requires no special equipment, contributing to a remarkably economical implementation. The entire protocol from electrophoresis to protein purification can be performed in <5 h. By combining the resulting PEPPI fraction with other protein-separation techniques, such as reversed-phase liquid chromatography and ion mobility techniques, multidimensional proteome separations for in-depth proteoform analysis can be easily achieved.

Deep and accurate proteome analysis is crucial for understanding cellular processes and disease mechanisms; however, it is challenging to implement in routine settings. In this protocol, we combine a robust chromatographic platform with a high-performance mass spectrometric setup to enable routine yet in-depth proteome coverage for a broad community. This entails tip-based sample preparation and pre-formed gradients (Evosep One) combined with a trapped ion mobility time-of-flight mass spectrometer (timsTOF, Bruker). The timsTOF enables parallel accumulation-serial fragmentation (PASEF), in which ions are accumulated and separated by their ion mobility, maximizing ion usage and simplifying spectra. Combined with data-independent acquisition (DIA), it offers high peak sampling rates and near-complete ion coverage. Here, we explain how to balance quantitative accuracy, specificity, proteome coverage and sensitivity by choosing the best PASEF and DIA method parameters. The protocol describes how to set up the liquid chromatography-mass spectrometry system and enables PASEF method generation and evaluation for varied samples by using the py_diAID tool to optimally position isolation windows in the mass-to-charge and ion mobility space. Biological projects (e.g., triplicate proteome analysis in two conditions) can be performed in 3 d with ~3 h of hands-on time and minimal marginal cost. This results in reproducible quantification of 7,000 proteins in a human cancer cell line in quadruplicate 21-min injections and 29,000 phosphosites for phospho-enriched quadruplicates. Synchro-PASEF, a highly efficient, specific and novel scan mode, can be analyzed by Spectronaut or AlphaDIA, resulting in superior quantitative reproducibility because of its high sampling efficiency.

Photoporation with free photothermal nanoparticles (NPs) is a promising technology for gentle delivery of functional biomacromolecules into living cells, offering great flexibility in terms of cell types and payload molecules. However, the translational use of photoporation, such as for transfecting patient-derived cells for cell therapies, is hampered by safety and regulatory concerns as it relies on direct contact between cells and photothermal NPs. A solution is to embed the photothermal NPs in electrospun nanofibers, which form a substrate for cell culture. Here we present a protocol for photothermal electrospun nanofiber (PEN)-mediated photoporation that induces membrane permeabilization by photothermal effects and enables efficient intracellular delivery of payload molecules into various cell types. By incorporating photothermal NPs within biocompatible electrospun nanofibers, direct cellular contact with NPs is avoided, thus largely mitigating safety or regulatory issues. Importantly, PEN photoporation is gentler to cells compared with electroporation, the most commonly used physical transfection method, resulting in higher-quality genetically engineered cells with better therapeutic potential. According to this protocol, it takes 2-3 d to prepare PEN culture wells with the desired cells, 3-4 d to optimize PEN photoporation parameters for intracellular delivery of payload molecules into different cell types in vitro and 4-5 weeks to evaluate the in vivo therapeutic efficacy of PEN-photoporated T cells. The protocol also provides details on how to construct the laser-based setup for performing photoporation experiments.

Implantable multifunctional probes have transformed neuroscience research, offering access to multifaceted brain activity that was previously unattainable. Typically, simultaneous access to both optical and electrical signals requires separate probes, while their integration into a single device can result in the emergence of photogenerated electrical artifacts, affecting the quality of high-frequency neural recordings. Among the nontrivial strategies aimed at the realization of an implantable multifunctional interface, the integration of optical and electrical capabilities on a single, minimally invasive, tapered optical fiber probe has been recently demonstrated using fibertrodes. Fibertrodes require the application of a set of planar microfabrication techniques to a nonplanar system with low and nonconstant curvature radius. Here we develop a process based on multiple conformal depositions, nonplanar two-photon lithography and chemical wet etching steps to obtain metallic patterns on the highly curved surface of the fiber taper. We detail the manufacturing, encapsulation and back end of the fibertrodes. The design of the probe can be adapted for different experimental requirements. Using the optical setup design, it is possible to perform angle selective light coupling with the fibertrodes and their implantation and use in vivo. The fabrication of fibertrodes is estimated to require 5-9 d. Nonetheless, due to the high scalability of a large part of the protocol, the manufacture of multiple fibertrodes simultaneously substantially reduces the required time for each probe. The procedure is suitable for users with expertise in microfabrication of electronics and neural recordings.

Individual ion mass spectrometry (IMS) is the Orbitrap-based extension of the niche mass spectrometry technique known as charge detection mass spectrometry (CDMS). While traditional CDMS analysis is performed on in-house-built instruments such as the electrostatic linear ion trap, IMS extends CDMS analysis to Orbitrap analyzers, allowing charge detection analysis to be available to the scientific community at large. IMS simultaneously measures the mass-to-charge ratios (m/z) and charges (z) of hundreds to thousands of individual ions within one acquisition event, creating a spectral output directly into the mass domain without the need for further spectral deconvolution. A mass distribution or 'profile' can be created for any desired sample regardless of composition or heterogeneity. To assist in reducing IMS analysis to practice, we developed this workflow for data acquisition and subsequent data analysis, which includes (i) protein sample preparation, (ii) attenuation of ion signals to obtain individual ions, (iii) the creation of a charge-calibration curve from standard proteins with known charge states and finally (iv) producing a meaningful mass spectral output from a complex or unknown sample by using the STORIboard software. This protocol is suitable for users with prior experience in mass spectrometry and bioanalytical chemistry. First, the analysis of protein standards in native and denaturing mode is presented, setting the foundation for the analysis of complex mixtures that are intractable via traditional mass spectrometry techniques. Examples of complex mixtures included here demonstrate the relevant analysis of an intact human monoclonal antibody and its intricate glycosylation patterns.

Glycosylated RNAs (glycoRNAs) have recently emerged as a new class of molecules of substantial interest owing to their potential roles in cellular processes and diseases. However, studying glycoRNAs is challenging owing to the lack of effective research tools including, but not limited to, imaging techniques to study the spatial distribution of glycoRNAs. Recently, we reported the development of a glycoRNA imaging technique, called sialic acid aptamer and RNA in situ hybridization-mediated proximity ligation assay (ARPLA), to visualize sialic acid-containing glycoRNAs with high sensitivity and specificity. Here we describe the experimental design principles and detailed step-by-step procedures for ARPLA-assisted glycoRNA imaging across multiple cell types. The procedure includes details for target selection, oligo design and preparation, optimized steps for RNA in situ hybridization, glycan recognition, proximity ligation, rolling circle amplification and a guideline for image acquisition and analysis. With properly designed probe sets and cells prepared, ARPLA-based glycoRNA imaging can typically be completed within 1 d by users with expertise in biochemistry and fluorescence microscopy. The ARPLA approach enables researchers to explore the spatial distribution, trafficking and functional contributions of glycoRNAs in various cellular processes.

Templates for the acquisition of large datasets such as the Human Connectome Project guide the neuroimaging community to reproducible data acquisition and scientific rigor. By contrast, small animal neuroimaging often relies on laboratory-specific protocols, which limit cross-study comparisons. The establishment of broadly validated protocols may facilitate the acquisition of large datasets, which are essential for uncovering potentially small effects often seen in functional MRI (fMRI) studies. Here, we outline a procedure for the acquisition of fMRI datasets in rats and describe animal handling, MRI sequence parameters, data conversion, preprocessing, quality control and data analysis. The procedure is designed to be generalizable across laboratories, has been validated by using datasets across 20 research centers with different scanners and field strengths ranging from 4.7 to 17.2 T and can be used in studies in which it is useful to compare functional connectivity measures across an extensive range of datasets. The MRI procedure requires 1 h per rat to complete and can be carried out by users with limited expertise in rat handling, MRI and data processing.

Programmable gene integration technologies are an emerging modality with exciting applications in both basic research and therapeutic development. Programmable addition via site-specific targeting elements (PASTE) is a programmable gene integration approach for precise and efficient programmable integration of large DNA sequences into the genome. PASTE offers improved editing efficiency, purity and programmability compared with previous methods for long insertions into the mammalian genome. By combining the specificity and cargo size capabilities of site-specific integrases with the programmability of prime editing, PASTE can precisely insert cargoes of at least 36 kb with efficiencies of up to 60%. Here we outline best practices for design, execution and analysis of PASTE experiments, with protocols for integration of EGFP at the human NOLC1 and ACTB genomic loci and for readout by next generation sequencing and droplet digital PCR. We provide guidelines for designing and optimizing a custom PASTE experiment for integration of desired payloads at alternative genomic loci, as well as example applications for in-frame protein tagging and multiplexed insertions. To facilitate experimental setup, we include the necessary sequences and plasmids for the delivery of PASTE components to cells via plasmid transfection or in vitro transcribed RNA. Most experiments in this protocol can be performed in as little as 2 weeks, allowing for precise and versatile programmable gene insertion.

Antibody-based research applications are critical for biological discovery. Yet there are no industry standards for comparing the performance of antibodies in various applications. We describe a knockout cell line-based antibody characterization platform, developed and approved jointly by industry and academic researchers, that enables the systematic comparison of antibody performance in western blot, immunoprecipitation and immunofluorescence. The scalable protocols, which require minimal technological resources, consist of (1) the identification of appropriate cell lines for antibody characterization studies, (2) development/contribution of isogenic knockout controls, and (3) a series of antibody characterization procedures focused on the most common applications of antibodies in research. We provide examples of expected outcomes to guide antibody users in evaluating antibody performance. Central to our approach is advocating for transparent and open data sharing, enabling a community effort to identify specific antibodies for all human proteins. Mid-level graduate students with training in biochemistry and prior experience in cell culture and microscopy can complete the protocols for a specific protein within 1 month while working part-time on this effort. Antibody characterization is needed to meet standards for resource validation and data reproducibility, which are increasingly required by journals and funding agencies.

Microcrystal electron diffraction (MicroED) has advanced structural methods across a range of sample types, from small molecules to proteins. This cryogenic electron microscopy (cryo-EM) technique involves the continuous rotation of small 3D crystals in the electron beam, while a high-speed camera captures diffraction data in the form of a movie. The crystal structure is subsequently determined by using established X-ray crystallographic software. MicroED is a technique still under development, and hands-on expertise in sample preparation, data acquisition and processing is not always readily accessible. This comprehensive guide on MicroED sample preparation addresses commonly used methods for various sample categories, including room temperature solid-state small molecules and soluble and membrane protein crystals. Beyond detailing the steps of sample preparation for new users, and because every crystal requires unique growth and sample-preparation conditions, this resource provides instructions and optimization strategies for MicroED sample preparation. The protocol is suitable for users with expertise in biochemistry, crystallography, general cryo-EM and crystallography data processing. MicroED experiments, from sample vitrification to final structure, can take anywhere from one workday to multiple weeks, especially when cryogenic focused ion beam milling is involved.

The ability to apply controlled forces to individual molecules or molecular complexes and observe their behaviors has led to many important discoveries in biology. Instruments capable of probing single-molecule forces typically cost >US$100,000, limiting the use of these techniques. The centrifuge force microscope (CFM) is a low-cost and easy-to-use instrument that enables high-throughput single-molecule studies. By combining the imaging capabilities of a microscope with the force application of a centrifuge, the CFM enables the simultaneous probing of hundreds to thousands of single-molecule interactions using tethered particles. Here we present a comprehensive set of instructions for building a CFM module that fits within a commercial benchtop centrifuge. The CFM module uses a 3D-printed housing, relies on off-the-shelf optical and electrical components, and can be built for less than US$1,000 in about 1 day. We also provide detailed instructions for setting up and running an experiment to measure force-dependent shearing of a short DNA duplex, as well as the software for CFM control and data analysis. The protocol is suitable for users with basic experience in analytical biochemistry and biophysics. The protocol enables the use of CFM-based experiments and may facilitate access to the single-molecule research field.