The development of new technologies for the efficient expression of recombinant hemoglobin (rHb) is of interest for experimental studies of protein biochemistry and the development of cell‐free blood substitutes in transfusion medicine. Expression of rHb in
Transmembrane proteins are responsible for many critical cellular functions and represent one of the largest families of drug targets. However, these proteins, especially multipass transmembrane proteins, are difficult to study because they must be embedded in a lipid bilayer to maintain their native conformations. The development of the virion display (VirD) technology enables transmembrane proteins to be integrated into the viral envelope of herpes simplex virus 1 (HSV‐1). Combining high‐throughput cloning, expression, and purification techniques, VirD technology has been applied to the largest set of human transmembrane proteins, namely G‐protein‐coupled receptors, and has allowed the identification of interactions that are both specific and functional. This article describes the procedures to integrate an open reading frame for any transmembrane protein into the HSV‐1 genome and produce recombinant HSV‐1 virus to ultimately generate pure VirD virions for biological and pharmaceutical studies. © 2020 Wiley Periodicals LLC.
- NSF-PAR ID:
- 10238058
- Publisher / Repository:
- Wiley Blackwell (John Wiley & Sons)
- Date Published:
- Journal Name:
- Current Protocols in Molecular Biology
- Volume:
- 132
- Issue:
- 1
- ISSN:
- 1934-3639
- Format(s):
- Medium: X
- Sponsoring Org:
- National Science Foundation
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Abstract Escherichia coli host cells has numerous advantages, but one disadvantage of using prokaryotic systems to express eukaryotic proteins is that they are incapable of performing post‐translational modifications such as NH2‐terminal acetylation. One possible solution is to coexpress additional enzymes that can perform the necessary modifications in the host cells. Here, we report a new method for synthesizing human rHb with proper NH2‐terminal acetylation. Mass spectrometry experiments involving native and recombinant human Hb confirmed the efficacy of the new technique in producing correctly acetylated globin chains. Finally, functional experiments provided insights into the effects of NH2‐terminal acetylation on O2binding properties. © 2020 Wiley Periodicals LLC.Basic Protocol 1 : Gene synthesis and cloning the cassette to the expression plasmidBasic Protocol 2 : Selection ofE. coli expression strains for coexpressionBasic Protocol 3 : Large‐scale recombinant hemoglobin expression and purificationSupport Protocol 1 : Measuring O2equilibration curvesSupport Protocol 2 : Mass spectrometry to confirm NH2‐terminal acetylation -
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Abstract Intracellular signaling processes are frequently based on direct interactions between proteins and organelles. A fundamental strategy to elucidate the physiological significance of such interactions is to utilize optical dimerization tools. These tools are based on the use of small proteins or domains that interact with each other upon light illumination. Optical dimerizers are particularly suitable for reproducing and interrogating a given protein‐protein interaction and for investigating a protein's intracellular role in a spatially and temporally precise manner. Described in this article are genetic engineering strategies for the generation of modular light‐activatable protein dimerization units and instructions for the preparation of optogenetic applications in mammalian cells. Detailed protocols are provided for the use of light‐tunable switches to regulate protein recruitment to intracellular compartments, induce intracellular organellar membrane tethering, and reconstitute protein function using enhanced Magnets (eMags), a recently engineered optical dimerization system. © 2021 Wiley Periodicals LLC.
This article was corrected on 25 July 2022. See the end of the full text for details.
Basic Protocol 1 : Genetic engineering strategy for the generation of modular light‐activated protein dimerization unitsSupport Protocol 1 : Molecular cloningBasic Protocol 2 : Cell culture and transfectionSupport Protocol 2 : Production of dark containers for optogenetic samplesBasic Protocol 3 : Confocal microscopy and light‐dependent activation of the dimerization systemAlternate Protocol 1 : Protein recruitment to intracellular compartmentsAlternate Protocol 2 : Induction of organelles’ membrane tetheringAlternate Protocol 3 : Optogenetic reconstitution of protein functionBasic Protocol 4 : Image analysisSupport Protocol 3 : Analysis of apparent on‐ and off‐kineticsSupport Protocol 4 : Analysis of changes in organelle overlap over time -
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Abstract 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 leukocytesBasic Protocol 2 : First‐strand cDNA synthesis; VHH and VHrepertoire PCRBasic Protocol 3 : Preparation of the phage display libraryBasic Protocol 4 : Panning of the phage display libraryBasic Protocol 5 : Small‐scale nAb expressionBasic Protocol 6 : Sequence analysis of selected nAb clonesBasic Protocol 7 : Nanobody validation as immunolabelsBasic Protocol 8 : Generation of nAb‐pEGFP mammalian expression constructsBasic Protocol 9 : Nanobody validation as intrabodiesSupport Protocol 1 : ELISA for llama serum testing, phage titer, and screening of selected clonesSupport Protocol 2 : Amplification of helper phage stockSupport Protocol 3 : nAb expression in amber suppressorE. coli bacterial strains -
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Abstract 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, Tn
7 transfer of Mobile‐CRISPRi to Gram‐negative bacteria, and ICEBs1 transfer of Mobile‐CRISPRi to Bacillales. © 2020 The Authors.Basic Protocol 1 : sgRNA designBasic Protocol 2 : Cloning of new sgRNA spacers into Mobile‐CRISPRi vectorsBasic Protocol 3 : Tn7 transfer of Mobile‐CRISPRi to Gram‐negative bacteriaBasic Protocol 4 : ICEBs1 transfer of Mobile‐CRISPRi to BacillalesSupport Protocol 1 : Quantification of CRISPRi repression using fluorescent reportersSupport Protocol 2 : Testing for gene essentiality using CRISPRi spot assays on platesSupport Protocol 3 : Transformation ofE. coli by electroporationSupport Protocol 4 : Transformation of CaCl2‐competentE. coli -
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Abstract Protein activity is generally functionally integrated and spatially restricted to key locations within the cell. Knocksideways experiments allow researchers to rapidly move proteins to alternate or ectopic regions of the cell and assess the resultant cellular response. Briefly, individual proteins to be tested using this approach must be modified with moieties that dimerize under treatment with rapamycin to promote the experimental spatial relocalizations. CRISPR technology enables researchers to engineer modified protein directly in cells while preserving proper protein levels because the engineered protein will be expressed from endogenous promoters. Here we provide straightforward instructions to engineer tagged, rapamycin‐relocalizable proteins in cells. The protocol is described in the context of our work with the microtubule depolymerizer MCAK/Kif2C, but it is easily adaptable to other genes and alternate tags such as degrons, optogenetic constructs, and other experimentally useful modifications. Off‐target effects are minimized by testing for the most efficient target site using a split‐GFP construct. This protocol involves no proprietary kits, only plasmids available from repositories (such as addgene.org). Validation, relocalization, and some example novel discoveries obtained working with endogenous protein levels are described. A graduate student with access to a fluorescence microscope should be able to prepare engineered cells with spatially controllable endogenous protein using this protocol. © 2023 The Authors. Current Protocols published by Wiley Periodicals LLC.
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