More Like this

1. Development, Screening, and Validation of Camelid‐Derived Nanobodies for Neuroscience Research

   https://doi.org/10.1002/cpns.107  

   Gavira‐O'Neill, Clara E. ; Dong, Jie‐Xian ; Trimmer, James S. ( November 2020 , Current Protocols in Neuroscience)

   Nanobodies (nAbs) are recombinant antigen‐binding variable domain fragments obtained from heavy‐chain‐only immunoglobulins. Among mammals, these are unique to camelids (camels, llamas, alpacas, etc.). Nanobodies are of great use in biomedical research due to their efficient folding and stability under a variety of conditions, as well as their small size. The latter characteristic is particularly important for nAbs used as immunolabeling reagents, since this can improve penetration of cell and tissue samples compared to conventional antibodies, and also reduce the gap distance between signal and target, thereby improving imaging resolution. In addition, their recombinant nature allows for unambiguous definition and permanent archiving in the form of DNA sequence, enhanced distribution in the form of sequences or plasmids, and easy and inexpensive production using well‐established bacterial expression systems, such as the IPTG induction method described here. This article will review the basic workflow and process for developing, screening, and validating novel nAbs against neuronal target proteins. The protocols described make use of the most common nAb development method, wherein an immune repertoire from an immunized llama is screened via phage display technology. Selected nAbs can then be taken through validation assays for use as immunolabels or as intrabodies in neurons. © 2020 Wiley Periodicals LLC.

   This article was corrected on 26 June 2021. See the end of the full text for details.

   Basic Protocol 1: Total RNA isolation from camelid leukocytes

   Basic Protocol 2: First‐strand cDNA synthesis; V_HH and V_Hrepertoire PCR

   Basic Protocol 3: Preparation of the phage display library

   Basic Protocol 4: Panning of the phage display library

   Basic Protocol 5: Small‐scale nAb expression

   Basic Protocol 6: Sequence analysis of selected nAb clones

   Basic Protocol 7: Nanobody validation as immunolabels

   Basic Protocol 8: Generation of nAb‐pEGFP mammalian expression constructs

   Basic Protocol 9: Nanobody validation as intrabodies

   Support Protocol 1: ELISA for llama serum testing, phage titer, and screening of selected clones

   Support Protocol 2: Amplification of helper phage stock

   Support Protocol 3: nAb expression in amber suppressorE. colibacterial strains

    

   more » « less
2. Imaging Spontaneous Neuronal Activity with Voltage‐Sensitive Dyes

   https://doi.org/10.1002/cpz1.48  

   Raliski, Benjamin K. ; Kirk, Molly J. ; Miller, Evan W. ( March 2021 , Current Protocols)

   Accurately mapping changes in cellular membrane potential across large groups of neurons is crucial for understanding the organization and maintenance of neural circuits. Measuring cellular voltage changes by optical means allows greater spatial resolution than traditional electrophysiology methods and is adaptable to high‐throughput imaging experiments. VoltageFluors, a class of voltage‐sensitive dyes, have recently been used to optically study the spontaneous activity of many neurons simultaneously in dissociated culture. VoltageFluors are particularly useful for experiments investigating differences in excitability and connectivity between neurons at different stages of development and in different disease models. The protocols in this article describe general procedures for preparing dissociated cultures, imaging spontaneous activity in dissociated cultures with VoltageFluors, and analyzing optical spontaneous activity data. © 2021 Wiley Periodicals LLC.

   This article was corrected on 20 July 2022. See the end of the full text for details.

   Basic Protocol 1: Preparation of dissociated rat hippocampal or cortical cultures

   Alternate Protocol: Preparation of microisland dissociated cultures

   Basic Protocol 2: Imaging of spontaneous activity in dissociated cultures using voltage‐sensitive dyes

   Basic Protocol 3: Analysis of spontaneous activity imaging data

    

   more » « less
3. CUT&RUN for Chromatin Profiling in Caenorhabditis elegans

   https://doi.org/10.1002/cpz1.445  

   Emerson, Felicity J. ; Lee, Siu Sylvia ( June 2022 , Current Protocols)

   Cleavage under targets and release using nuclease (CUT&RUN) is a recently developed chromatin profiling technique that uses a targeted micrococcal nuclease cleavage strategy to obtain high‐resolution binding profiles of protein factors or to map histones with specific post‐translational modifications. Due to its high sensitivity, CUT&RUN allows quality binding profiles to be obtained with only a fraction of the starting material and sequencing depth typically required for other chromatin profiling techniques such as chromatin immunoprecipitation. Although CUT&RUN has been widely adopted in multiple model systems, it has rarely been utilized inCaenorhabditis elegans, a model system of great importance to genomic research. Cell dissociation techniques, which are required for this approach, can be challenging inC. elegansdue to the toughness of the worm's cuticle and the sensitivity of the cells themselves. Here, we describe a robust CUT&RUN protocol for use inC. elegansto determine the genome‐wide localization of protein factors and specific histone marks. With a simple protocol utilizing live, uncrosslinked tissue as the starting material, performing CUT&RUN in worms has the potential to produce physiologically relevant data at a higher resolution than chromatin immunoprecipitation. This protocol involves a simple dissociation step to uniformly permeabilize worms while avoiding sample loss or cell damage, resulting in high‐quality CUT&RUN profiles with as few as 100 worms and detectable signal with as few as 10 worms. This represents a significant advancement over chromatin immunoprecipitation, which typically uses thousands or hundreds of thousands of worms for a single experiment. The protocols presented here provide a detailed description of worm growth, sample preparation, CUT&RUN workflow, library preparation for high‐throughput sequencing, and a basic overview of data analysis, making CUT&RUN simple and accessible for any worm lab. © 2022 Wiley Periodicals LLC.

   Basic Protocol 1: Growth and synchronization ofC. elegans

   Basic Protocol 2: Worm dissociation, sample preparation, and optimization

   Basic Protocol 3: CUT&RUN chromatin profiling

   Alternate Protocol: Improving CUT&RUN signal using a secondary antibody

   Basic Protocol 4: CUT&RUN library preparation for Illumina high‐throughput sequencing

   Basic Protocol 5: Basic data analysis using Linux

    

   more » « less
4. Programmable Gene Knockdown in Diverse Bacteria Using Mobile‐CRISPRi

   https://doi.org/10.1002/cpmc.130  

   Banta, Amy B. ; Ward, Ryan D. ; Tran, Jennifer S. ; Bacon, Emily E. ; Peters, Jason M. ( December 2020 , Current Protocols in Microbiology)

   Facile bacterial genome sequencing has unlocked a veritable treasure trove of novel genes awaiting functional exploration. To make the most of this opportunity requires powerful genetic tools that can target all genes in diverse bacteria. CRISPR interference (CRISPRi) is a programmable gene‐knockdown tool that uses an RNA‐protein complex comprised of a single guide RNA (sgRNA) and a catalytically inactive Cas9 nuclease (dCas9) to sterically block transcription of target genes. We previously developed a suite of modular CRISPRi systems that transfer by conjugation and integrate into the genomes of diverse bacteria, which we call Mobile‐CRISPRi. Here, we provide detailed protocols for the modification and transfer of Mobile‐CRISPRi vectors for the purpose of knocking down target genes in bacteria of interest. We further discuss strategies for optimizing Mobile‐CRISPRi knockdown, transfer, and integration. We cover the following basic protocols: sgRNA design, cloning new sgRNA spacers into Mobile‐CRISPRi vectors, Tn7transfer of Mobile‐CRISPRi to Gram‐negative bacteria, and ICEBs1transfer of Mobile‐CRISPRi to Bacillales. © 2020 The Authors.

   Basic Protocol 1: sgRNA design

   Basic Protocol 2: Cloning of new sgRNA spacers into Mobile‐CRISPRi vectors

   Basic Protocol 3: Tn7transfer of Mobile‐CRISPRi to Gram‐negative bacteria

   Basic Protocol 4: ICEBs1transfer of Mobile‐CRISPRi to Bacillales

   Support Protocol 1: Quantification of CRISPRi repression using fluorescent reporters

   Support Protocol 2: Testing for gene essentiality using CRISPRi spot assays on plates

   Support Protocol 3: Transformation ofE. coliby electroporation

   Support Protocol 4: Transformation of CaCl₂‐competentE. coli

    

   more » « less
5. Detection and Quantification of RNA Phosphorothioate Modifications Using Mass Spectrometry

   https://doi.org/10.1002/cpnc.113  

   Wu, Ying ; Zheng, Ya Ying ; Lin, Qishan ; Sheng, Jia ( August 2020 , Current Protocols in Nucleic Acid Chemistry)

   This article describes a protocol for detecting and quantifying RNA phosphorothioate modifications in cellular RNA samples. Starting from solid‐phase synthesis of phosphorothioate RNA dinucleotides, followed by purification with reversed‐phase HPLC, phosphorothioate RNA dinucleotide standards are prepared for UPLC‐MS and LC‐MS/MS methods. RNA samples are extracted from cells using TRIzol reagent, then digested with a nuclease mixture and analyzed by mass spectrometry. UPLC‐MS is employed first to identify RNA phosphorothioate modifications. An optimized LC‐MS/MS method is then employed to quantify the frequency of RNA phosphorothioate modifications in a series of model cells. © 2020 Wiley Periodicals LLC.

   Basic Protocol 1: Synthesis, purification, and characterization of RNA phosphorothioate dinucleotides

   Basic Protocol 2: Digestion of RNA samples extracted from cells

   Basic Protocol 3: Detection and quantification of RNA phosphorothioate modifications by mass spectrometry

    

   more » « less