Molecular Biology


Protocols in Current Issue
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0 Q&A 495 Views Aug 5, 2023

Generation of zebrafish (Danio rerio) models with targeted insertion of epitope tags and point mutations is highly desirable for functional genomics and disease modeling studies. Currently, CRISPR/Cas9-mediated knock-in is the method of choice for insertion of exogeneous sequences by providing a repair template for homology-directed repair (HDR). A major hurdle in generating knock-in models is the labor and cost involved in screening of injected fish to identify the precise knock-in events due to low efficiency of the HDR pathway in zebrafish. Thus, we developed fluorescent PCR–based high-throughput screening methods for precise knock-in of epitope tags and point mutations in zebrafish. Here, we provide a step-by-step guide that describes selection of an active sgRNA near the intended knock-in site, design of single-stranded oligonucleotide (ssODN) templates for HDR, quick validation of somatic knock-in using injected embryos, and screening for germline transmission of precise knock-in events to establish stable lines. Our screening method relies on the size-based separation of all fragments in an amplicon by fluorescent PCR and capillary electrophoresis, thus providing a robust and cost-effective strategy. Although we present the use of this protocol for insertion of epitope tags and point mutations, it can be used for insertion of any small DNA fragments (e.g., LoxP sites, in-frame codons). Furthermore, the screening strategy described here can be used to screen for precise knock-in of small DNA sequences in any model system, as PCR amplification of the target region is its only requirement.

Key features

• This protocol expands the use of fluorescent PCR and CRISPR-STAT for screening of precise knock-in of small insertions and point mutations in zebrafish.

• Allows validation of selected sgRNA and HDR template within two weeks by somatic knock-in screening.

• Allows robust screening of point mutations by combining restriction digest with CRISPR-STAT.

Graphical overview

Overview of the three-phase knock-in pipeline in zebrafish (created with

0 Q&A 1849 Views Oct 20, 2022

Directed evolution is a powerful technique for identifying beneficial mutations in defined DNA sequences with the goal of improving desired phenotypes. Recent methodological advances have made the evolution of short DNA sequences quick and easy. However, the evolution of DNA sequences >5kb in length, notably gene clusters, is still a challenge for most existing methods. Since many important microbial phenotypes are encoded by multigene pathways, they are usually improved via adaptive laboratory evolution (ALE), which while straightforward to implement can suffer from off-target and hitchhiker mutations that can adversely affect the fitness of the evolved strain. We have therefore developed a new directed evolution method (Inducible Directed Evolution, IDE) that combines the specificity and throughput of recent continuous directed evolution methods with the ease of ALE. Here, we present detailed methods for operating Inducible Directed Evolution (IDE), which enables long (up to 85kb) DNA sequences to be mutated in a high throughput manner via a simple series of incubation steps. In IDE, an intracellular mutagenesis plasmid (MP) tunably mutagenizes the pathway of interest, located on the phagemid (PM). MP contains a mutagenic operon (danQ926, dam, seqA, emrR, ugi, and cda1) that can be expressed via the addition of a chemical inducer. Expression of the mutagenic operon during a cell cycle represses DNA repair mechanisms such as proofreading, translesion synthesis, mismatch repair, and base excision and selection, which leads to a higher mutation rate. Induction of the P1 lytic cycle results in packaging of the mutagenized phagemid, and the pathway-bearing phage particles infect naïve cells, generating a mutant library that can be screened or selected for improved variants. Successive rounds of IDE enable optimization of complex phenotypes encoded by large pathways (as of this writing up to 36 kb), without requiring inefficient transformation steps. Additionally, IDE avoids off-target genomic mutations and enables decoupling of mutagenesis and screening steps, establishing it as a powerful tool for optimizing complex phenotypes in E. coli.

Graphical abstract:

Figure 1. Overview of Inducible Directed Evolution (IDE). Pathways of interest are cloned into a P1 phagemid (PM) backbone and transformed into a strain of E. coli containing MP (diversification strain). The mutagenesis plasmid is induced to generate mutations. Phage lysate is produced and used to infect a strain that expresses the phenotype of interest (screening/selection strain). The resulting strain library is screened to identify those with improved properties. Narrowed-down libraries can then go through another IDE cycle by infecting a fresh diversification strain.

0 Q&A 4572 Views Jul 5, 2022

Profiling the specificities of antibodies can reveal a wealth of information about humoral immune responses and the antigens they target. Here, we present a protocol for VirScan, an application of the phage immunoprecipitation sequencing (PhIP-Seq) method for profiling the specificities of human antiviral antibodies. Accompanying this protocol is a video of the experimental procedure. VirScan and, more generally, PhIP-Seq are techniques that enable high-throughput antibody profiling by combining high-throughput DNA oligo synthesis and bacteriophage display with next-generation sequencing. In the VirScan method, human sera samples are screened against a library of peptides spanning the entire human viral proteome. Bound phage are immunoprecipitated and sequenced, identifying the viral peptides recognized by the antibodies. VirScan Is a powerful tool for uncovering individual viral exposure histories, mapping the epitope landscape of viruses of interest, and studying fundamental mechanisms of viral immunity.

Graphical abstract:

0 Q&A 1962 Views Mar 20, 2022

Phytophthora sojae is a model species for the study of plant pathogenic oomycetes. The initial research on gene function using Phytophthora was mainly based on gene silencing technology. Recently, the CRISPR/Cas9-mediated genome editing technology was successfully established in P. sojae and widely used in oomycetes. In this protocol, we describe the operating procedures for the use of CRISPR/Cas9-based genome editing technology and PEG-mediated stable transformation of P. sojae protoplasts. Two plasmids were co-transformed into P. sojae: pYF515 expressing Cas9 and the single guide RNA, and the homologous replacement vector of the candidate gene. Finally, the ORF of candidate gene were replaced with the ORF of the entire hygromycin B phosphotransferase gene (HPH), to achieve precise knockout.

1 Q&A 8661 Views Jul 20, 2021

Efficient precision genome engineering requires high frequency and specificity of integration at the genomic target site. Multiple design strategies for zebrafish gene targeting have previously been reported with widely varying frequencies for germline recovery of integration alleles. The GeneWeld protocol and pGTag (plasmids for Gene Tagging) vector series provide a set of resources to streamline precision gene targeting in zebrafish. Our approach uses short homology of 24-48 bp to drive targeted integration of DNA reporter cassettes by homology-mediated end joining (HMEJ) at a CRISPR/Cas induced DNA double-strand break. The pGTag vectors contain reporters flanked by a universal CRISPR sgRNA sequence to liberate the targeting cassette in vivo and expose homology arms for homology-driven integration. Germline transmission rates for precision-targeted integration alleles range 22-100%. Our system provides a streamlined, straightforward, and cost-effective approach for high-efficiency gene targeting applications in zebrafish.

Graphic Abstract:

GeneWeld method for CRISPR/Cas9 targeted integration.

0 Q&A 3315 Views Mar 5, 2021

Transposon insertion sequencing (TIS) is an emerging technique which utilizes a massive transposon mutant library to screen specific phenotype and determine the conditional essential genetic requirements for bacterial fitness under distinct conditions combined with high-throughput parallel sequencing technology. Compared with a massive mutant library in traditional TIS, the defined mutant library sequencing (DML-Seq) has advantages as: 1) efficient mutagenesis; 2) low bottleneck effects; 3) avoid hotpots caused by screening; 4) can be directly used in the following experiments. Here, we described an optimized procedure of DML-Seq for fitness screen to supply classical TIS using the marine pathogenic bacterium Edwardsiella piscicida as an example.

0 Q&A 5296 Views Feb 20, 2021

Precise genome engineering has become a commonplace technique for metabolic engineering. Also, insertion, deletion and alteration of genes and other functional DNA sequences are essential for understanding and engineering cells. Several techniques have been developed to this end (e.g., CRISPR/Cas-assisted methods, homologous recombination, or λ Red recombineering), yet most of them rely on the use of auxiliary plasmids, which have to be cured after the editing procedure. Temperature-sensitive replicons, counter-selectable markers or repeated passaging of plasmid-bearing cells have been traditionally employed to circumvent this hurdle. While these protocols work reasonably well in some bacteria, they are not applicable for other species or are time consuming and laborious. Here, we present a fast and versatile protocol of fluorescent marker-assisted genome editing in Pseudomonas putida, followed by clean curing of auxiliary plasmids through user-controlled plasmid replication. One fluorescent marker facilitates identification of genome-edited colonies, while the second reporter enables detection of plasmid-free bacterial clones. Not only is this protocol the fastest available for Pseudomonas species, but it can be easily adapted to any type of genome modifications, including sequence deletions, insertions, and replacements.

Graphical abstract:

Rapid genome engineering of Pseudomonas with curable plasmids

0 Q&A 4967 Views Sep 5, 2020
The efficiency of cleavage of individual CRISPR/Cas9-sgRNAs remains difficult to predict based on the CRISPR target sequence alone. Different intracellular environments (dependent on cell type or cell cycle state for example) may affect sgRNA efficiency by altering accessibility of genomic DNA through DNA modifications such as epigenetic marks and DNA-binding proteins (e.g., histones) as well as alteration of the chromatin state of genomic DNA within the nucleus.

We recently reported a multi-step screening method for the identification of efficient sgRNAs targeting the Herpes simplex virus (HSV-1) genome and reported a differential mechanism for viral inhibition by CRISPR-Cas9 in the latent versus lytic phase. The screening platform detailed in this protocol allows step-by-step testing of the efficiency of cleavage in a cell-free system and in the context of viral target cells such as human foreskin fibroblasts followed by functional testing of the effects of CRISPR/sgRNA on viral protein expression, replication, and reactivation. This strategy could be readily applied to other target cells such as pluripotent stem cell-derived human sensory neurons or other human DNA viruses.
0 Q&A 4385 Views Feb 20, 2020
Anabaena sp. PCC 7120 (hereafter Anabaena) is a model cyanobacterium to study nitrogen fixation, cellular differentiation and several other key biological functions that are analogous in plants. As with any other organism, many genes in Anabaena encode an essential life function and hence cannot be deleted, causing a bottleneck in the elucidation of its genomic function. Antisense RNA (asRNA) mediated approach renders the study of essential genes possible by suppressing (but not completely eliminating) expression of the target gene, thus allowing them to function to some extent. Recently, we have successfully implemented this approach using the strong endogenous promoter of the psbA1 gene (D1 subunit of Photosystem II) introduced into a high-copy replicative plasmid (pAM1956) to suppress the transcript level of the target gene alr0277 (encoding a sigma factor, SigJ/Alr0277) in Anabaena. This protocol represents an efficient and easy procedure to further explore the functional genomics, expanding the scope of basic and applied research in these ecologically important cyanobacteria.
0 Q&A 5049 Views Jan 5, 2020
Site-directed scanning mutagenesis is a useful tool applied in studying protein function and designing proteins with new properties, such as increased stability or enzymatic activity. Creating a systematic library of hundreds of site-directed mutants is still a demanding and expensive task. The established protocols for making such libraries include PCR amplification of the recombinant DNA using a pair of primers carrying a target mutation in the same PCR. Unfortunately, this approach is very often coupled with PCR artifacts which compromise overall efficiency of site-directed mutagenesis. To reduce the failure rate due to the PCR artifacts, we have set up a high-throughput mutagenesis protocol based on a two-fragment PCR approach. To this end, each mutation is introduced in two separate PCRs resulting in two linear fragments of the mutated plasmid. In the next steps, the PCR template is digested and the two matching plasmid fragments are joined together using Gibson assembly. Separating the corresponding mutagenic primers into two different PCRs decreases a number of artifacts and thus increases overall cloning efficiency. Furthermore, free software that we developed facilitates both high-throughput primer design and analysis of sequencing results. Overall, this protocol enabled us to efficiently produce several alanine-scanning libraries of 400 single-point mutations with complete coverage of the protein sequence.

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