skip to main content

Explore Research Products in the NSF-PAR

Title: Targeted Genetic Changes in Candida albicans Using Transient CRISPR‐Cas9 Expression
Abstract

Candida albicansis an opportunistic fungal pathogen responsible for significant disease and mortality. Absent complete mating and other convenient methods, dissection of its virulence factors relies on robust tools to delete, complement, and otherwise modify genes of interest in this diploid organism. Here we describe the design principles and use of CRISPR associated nuclease 9 (Cas9) and single‐guide RNAs transiently expressed from PCR cassettes to modify genes of interest, generating homozygous mutants in a single transformation step. © 2021 Wiley Periodicals LLC.

This article was corrected on 27 February 2023. See the end of the full text for details.

Basic Protocol 1: PCR amplification of CRISPR components

Basic Protocol 2: Transformation ofCandida albicans

Basic Protocol 3: Selecting and genotyping transformants

Alternate Protocol 1: Deletion with recyclable markers by CRISPR induced marker excision (CRIME)

Alternate Protocol 2: Knock‐in and combining multiple cassettes with overlapping homology

  more » « less
NSF-PAR ID:
10238656
Author(s) / Creator(s):
 ;  ;  
Publisher / Repository:
Wiley Blackwell (John Wiley & Sons)
Date Published:
Journal Name:
Current Protocols
Volume:
1
Issue:
1
ISSN:
2691-1299
Format(s):
Medium: X
Sponsoring Org:
National Science Foundation
More Like this
  1. ; ; ; ; ; ; ; ; ( , mSphere)
    Mitchell, Aaron P. (Ed.)
    ABSTRACT

    The World Health Organization recently published the first list of priority fungal pathogens highlighting multipleCandidaspecies, includingCandida glabrata,Candida albicans, andCandida auris. However, prior studies in these pathogens have been mainly limited to the use of two drug resistance cassettes,NatMXandHphMX, limiting genetic manipulation capabilities in prototrophic laboratory strains and clinical isolates. In this study, we expanded the toolkit forC. glabrata,C. auris, andC. albicansto includeKanMXandBleMXwhen coupled with anin vitroassembled CRISPR-Cas9 ribonucleoprotein (RNP)-based system. Repurposing these drug resistance cassettes forCandida, we were able to make single gene deletions, sequential and simultaneous double gene deletions, epitope tags, and rescue constructs. We applied these drug resistance cassettes to interrogate the ergosterol pathway, a critical pathway for both the azole and polyene antifungal drug classes. Using our approach, we determined for the first time that the deletion ofERG3inC. glabrata,C. auris,andC. albicansprototrophic strains results in azole drug resistance, which further supports the conservation of the Erg3-dependent toxic sterol model. Furthermore, we show that anERG5deletion inC. glabratais azole susceptible at subinhibitory concentrations, suggesting that Erg5 could act as an azole buffer for Erg11. Finally, we identified a synthetic growth defect when bothERG3andERG5are deleted inC. glabrata,which suggests the possibility of another toxic sterol impacting growth. Overall, we have expanded the genetic tools available to interrogate complex pathways in prototrophic strains and clinical isolates.

    IMPORTANCE

    The increasing problem of drug resistance and emerging pathogens is an urgent global health problem that necessitates the development and expansion of tools for studying fungal drug resistance and pathogenesis. Prior studies inCandida glabrata,Candida auris, andCandida albicanshave been mainly limited to the use ofNatMX/SAT1andHphMX/CaHygfor genetic manipulation in prototrophic strains and clinical isolates. In this study, we demonstrated thatNatMX/SAT1, HphMX, KanMX,and/orBleMXdrug resistance cassettes when coupled with a CRISPR-ribonucleoprotein (RNP)-based system can be efficiently utilized for deleting or modifying genes in the ergosterol pathway ofC. glabrata,C. auris, andC. albicans. Moreover, the utility of these tools has provided new insights intoERGgenes and their relationship to azole resistance inCandida. Overall, we have expanded the toolkit forCandidapathogens to increase the versatility of genetically modifying complex pathways involved in drug resistance and pathogenesis.

     
    more » « less
  2. ; ; ( , Current Protocols in Microbiology)
    Abstract

    Vibrio fischeriis a nonpathogenic organism related to pathogenicVibriospecies. The bacterium has been used as a model organism to study symbiosis in the context of its association with its host, the Hawaiian bobtail squidEuprymna scolopes. The genetic tractability of this bacterium has facilitated the mapping of pathways that mediate interactions between these organisms. The protocols included here describe methods for genetic manipulation ofV. fischeri. Following these protocols, the researcher will be able to introduce linear DNA via transformation to make chromosomal mutations, to introduce plasmid DNA via conjugation and subsequently eliminate unstable plasmids, to eliminate antibiotic resistance cassettes from the chromosome, and to randomly or specifically mutagenizeV. fischeriwith transposons. © 2020 Wiley Periodicals LLC.

    Basic Protocol 1: Transformation ofV. fischeriwith linear DNA

    Basic Protocol 2: Plasmid transfer intoV. fischerivia conjugation

    Support Protocol 1: Removing FRT‐flanked antibiotic resistance cassettes from theV. fischerigenome

    Support Protocol 2: Eliminating unstable plasmids fromV. fischeri

    Alternate Protocol 1: Introduction of exogenous DNA using a suicide plasmid

    Alternate Protocol 2: Site‐specific transposon insertion using a suicide plasmid

    Alternate Protocol 3: Random transposon mutagenesis using a suicide plasmid

     
    more » « less
  3. ; ; ; ; ( , Current Protocols in Microbiology)
    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, 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
  4. ; ; ( , Current Protocols)
    Abstract

    Until recently, precise genome editing has been limited to a few organisms. The ability of Cas9 to generate double stranded DNA breaks at specific genomic sites has greatly expanded molecular toolkits in many organisms and cell types. Before CRISPR‐Cas9 mediated genome editing,P. patenswas unique among plants in its ability to integrate DNA via homologous recombination. However, selection for homologous recombination events was required to obtain edited plants, limiting the types of editing that were possible. Now with CRISPR‐Cas9, molecular manipulations inP. patenshave greatly expanded. This protocol describes a method to generate a variety of different genome edits. The protocol describes a streamlined method to generate the Cas9/sgRNA expression constructs, design homology templates, transform, and quickly genotype plants. © 2023 Wiley Periodicals LLC.

    Basic Protocol 1: Constructing the Cas9/sgRNA transient expression vector

    Alternate Protocol 1: Shortcut to generating single and pooled Cas9/sgRNA expression vectors

    Basic Protocol 2: Designing the oligonucleotide‐based homology‐directed repair (HDR) template

    Alternate Protocol 2: Designing the plasmid‐based HDR template

    Basic Protocol 3: Inducing genome editing by transforming CRISPR vector intoP. patensprotoplasts

    Basic Protocol 4: Identifying edited plants.

     
    more » « less
  5. ; ; ; ; ( , Current Protocols)
    Abstract

    Visualization of gene products inCaenorhabditis eleganshas provided insights into the molecular and biological functions of many novel genes in their native contexts. Single‐molecule fluorescencein situhybridization (smFISH) and immunofluorescence (IF) enable the visualization of the abundance and localization of mRNAs and proteins, respectively, allowing researchers to ultimately elucidate the localization, dynamics, and functions of the corresponding genes. Whereas both smFISH and immunofluorescence have been foundational techniques in molecular biology, each protocol poses challenges for use in theC. elegansembryo. smFISH protocols suffer from high initial costs and can photobleach rapidly, and immunofluorescence requires technically challenging permeabilization steps and slide preparation. Most importantly, published smFISH and IF protocols have predominantly been mutually exclusive, preventing the exploration of relationships between an mRNA and a relevant protein in the same sample. Here, we describe protocols to perform immunofluorescence and smFISH inC. elegansembryos either in sequence or simultaneously. We also outline the steps to perform smFISH or immunofluorescence alone, including several improvements and optimizations to existing approaches. These protocols feature improved fixation and permeabilization steps to preserve cellular morphology while maintaining probe and antibody accessibility in the embryo, a streamlined, in‐tube approach for antibody staining that negates freeze‐cracking, a validated method to perform the cost‐reducing single molecule inexpensive FISH (smiFISH) adaptation, slide preparation using empirically determined optimal antifade products, and straightforward quantification and data analysis methods. Finally, we discuss tricks and tips to help the reader optimize and troubleshoot individual steps in each protocol. Together, these protocols simplify existing workflows for single‐molecule RNA and protein detection. Moreover, simultaneous, high‐resolution imaging of proteins and RNAs of interest will permit analysis, quantification, and comparison of protein and RNA distributions, furthering our understanding of the relationship between RNAs and their protein products or cellular markers in early development. © 2021 Wiley Periodicals LLC.

    Basic Protocol 1: Sequential immunofluorescence and single‐molecule fluorescencein situhybridization

    Alternate Protocol: Abbreviated protocol for simultaneous immunofluorescence and single‐molecule fluorescencein situhybridization

    Basic Protocol 2: Simplified immunofluorescence inC. elegansembryos

    Basic Protocol 3: Single‐molecule fluorescencein situhybridization or single‐molecule inexpensive fluorescencein situhybridization

     
    more » « less
  • Citation Formats
  • MLA
  • APA
  • Chicago
  • BibTeX
  • Save / Share this Record
  • Send to Email