Selection Cassette Research Articles

Markerless gene editing in Neisseria gonorrhoeae.

Neisseria gonorrhoeae, the gonococcus, is a pathogen of major public health concern, but sophisticated approaches to gene manipulation are limited for this species. For example, there are few methods for generating markerless mutations, which allow the generation of precise point mutations and deletions without introducing additional DNA sequence. Markerless mutations are central to studying pathogenesis, the spread of antimicrobial resistance (AMR) and for vaccine development. Here we describe the use of galK as a counter-selectable marker that can be used for markerless mutagenesis in N. gonorrhoeae. galK encodes galactokinase, an enzyme that metabolizes galactose in bacteria that can utilize it as a sole carbon source. GalK can also phosphorylate a galactose analogue, 2-deoxy-galactose (2-DOG), into a toxic, non-metabolisable intermediate, 2-deoxy-galactose-1-phosphate. We utilized this property of GalK to develop a markerless approach for mutagenesis in N. gonorrhoeae. We successfully deleted both chromosomally and plasmid-encoded genes, that are important for gonococcal vaccine development and studies of AMR spread. We designed a positive-negative selection cassette, based on an antibiotic resistance marker and galK, that efficiently rendered N. gonorrhoeae susceptible to growth on 2-DOG. We then adapted the galK-based counter-selection and the use of 2-DOG for markerless mutagenesis, and applied biochemical and phenotypic analyses to confirm the absence of target genes. We show that our markerless mutagenesis method for N. gonorrhoeae has a high success rate, and should be a valuable gene editing tool in the future.

Accelerated embryonic stem cell screening with a highly efficient genotyping pipeline.

Gene targeting in mouse ES cells replaces or modifies genes of interest; conditional alleles, reporter knock-ins, and amino acid changes are common examples of how gene targeting is used. For example, enhanced green fluorescent protein or Cre recombinase is placed under the control of endogenous genes to define promoter expression patterns. The most important step in the process is to demonstrate that a gene targeting vector is correctly integrated in the genome at the desired chromosomal location. The rapid identification of correctly targeted ES cell clones is facilitated by proper targeting vector construction, rapid screening procedures, and advances in cell culture. Here, we optimized and functionally linked magnetic activated cell sorting (MACS) technology as well as multiplex droplet digital PCR (ddPCR) to our ES cell screening process to achieve a greater than 60% assurance that ES clones are correctly targeted. In a further refinement of the process, drug selection cassettes are removed from ES cells with adenovirus technology. We describe this improved workflow and illustrate the reduction in time between therapeutic target identification and experimental validation. In sum, we describe a novel and effective implementation of ddPCR, multiMACS, and adenovirus recombinase into a streamlined screening workflow that significantly reduces timelines for gene targeting in mouse ES cells.

Generation of human induced pluripotent stem cell lines encoding for genetically encoded calcium indicators RCaMP1h and GCaMP6f

Calcium plays a key role in cardiomyocytes (CMs) for the translation of the electrical impulse of an action potential into contraction forces. A rapid, not-invasive fluorescence imaging technology allows for the monitoring of calcium transients in human induced pluripotent stem cell derived-cardiomyocytes (hiPSC-CMs) to investigate the cardiac electrophysiology in vitro and after cell transplantation in vivo. The genetically encoded calcium indicators (GECIs) GCaMP6f or RCaMP1h were successfully transfected in the previously established hiPSC line MHHi001-A, together with a cardiac specific antibiotic selection cassette facilitating the monitoring of the calcium handling in highly pure populations of hiPSC-CMs.

A high-efficiency and versatile CRISPR/Cas9-mediated HDR-based biallelic editing system.

Clustered regulatory interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 nuclease (Cas9), the third-generation genome editing tool, has been favored because of its high efficiency and clear system composition. In this technology, the introduced double-strand breaks (DSBs) are mainly repaired by non-homologous end joining (NHEJ) or homology-directed repair (HDR) pathways. The high-fidelity HDR pathway is used for genome modification, which can introduce artificially controllable insertions, deletions, or substitutions carried by the donor templates. Although high-level knock-out can be easily achieved by NHEJ, accurate HDR-mediated knock-in remains a technical challenge. In most circumstances, although both alleles are broken by endonucleases, only one can be repaired by HDR, and the other one is usually recombined by NHEJ. For gene function studies or disease model establishment, biallelic editing to generate homozygous cell lines and homozygotes is needed to ensure consistent phenotypes. Thus, there is an urgent need for an efficient biallelic editing system. Here, we developed three pairs of integrated selection systems, where each of the two selection cassettes contained one drug-screening gene and one fluorescent marker. Flanked by homologous arms containing the mutated sequences, the selection cassettes were integrated into the target site, mediated by CRISPR/Cas9-induced HDR. Positively targeted cell clones were massively enriched by fluorescent microscopy after screening for drug resistance. We tested this novel method on the amyloid precursor protein (APP) and presenilin 1 (PSEN1) loci and demonstrated up to 82.0% biallelic editing efficiency after optimization. Our results indicate that this strategy can provide a new efficient approach for biallelic editing and lay a foundation for establishment of an easier and more efficient disease model.

CelloSelect – A synthetic cellobiose metabolic pathway for selection of stable transgenic CHO cell lines

Current protocols for generating stable transgenic cell lines mostly rely on antibiotic selection or the use of specialized cell lines lacking an essential part of their metabolic machinery, but these approaches require working with either toxic chemicals or knockout cell lines, which can reduce productivity. Since most mammalian cells cannot utilize cellobiose, a disaccharide consisting of two β-1,4-linked glucose molecules, we designed an antibiotic-free selection system, CelloSelect, which consists of a selection cassette encoding Neurospora crassa cellodextrin transporter CDT1 and β-glucosidase GH1-1. When cultivated in glucose-free culture medium containing cellobiose, CelloSelect-transfected cells proliferate by metabolizing cellobiose as a primary energy source, and are protected from glucose starvation. We show that the combination of CelloSelect with a PiggyBac transposase-based integration strategy provides a platform for the swift and efficient generation of stable transgenic cell lines. Growth rate analysis of metabolically engineered cells in cellobiose medium confirmed the expansion of cells stably expressing high levels of a cargo fluorescent marker protein. We further validated this strategy by applying the CelloSelect system for stable integration of sequences encoding two biopharmaceutical proteins, erythropoietin and the monoclonal antibody rituximab, and confirmed that the proteins are efficiently produced in either cellobiose- or glucose-containing medium in suspension-adapted CHO cells cultured in chemically defined media. We believe coupling heterologous metabolic pathways additively to the endogenous metabolism of mammalian cells has the potential to complement or to replace current cell-line selection systems.

Combined fluorescent seed selection and multiplex CRISPR/Cas9 assembly for fast generation of multiple Arabidopsis mutants

BackgroundMultiplex CRISPR-Cas9-based genome editing is an efficient method for targeted disruption of gene function in plants. Use of CRISPR-Cas9 has increased rapidly in recent years and is becoming a routine method for generating single and higher order Arabidopsis thaliana mutants. Low entry, reliable assembly of CRISPR/Cas9 vectors and efficient mutagenesis is necessary to enable a maximum of researchers to break through the genetic redundancy within plant multi-gene families and allow for a plethora of gene function studies that have been previously unachievable. It will also allow routine de novo generation of mutations in ever more complex genetic backgrounds that make introgression of pre-existing alleles highly cumbersome.ResultsTo facilitate rapid and efficient use of CRISPR/Cas9 for Arabidopsis research, we developed a CRISPR/Cas9-based toolbox for generating mutations at multiple genomic loci, using two-color fluorescent seed selection. In our system, up-to eight gRNAs can be routinely introduced into a binary vector carrying either a FastRed, FastGreen or FastCyan fluorescent seed selection cassette. FastRed and FastGreen binary vectors can be co-transformed as a cocktail via floral dip to introduce sixteen gRNAs at the same time. The seeds can be screened either for red or green fluorescence, or for the presence of both colors. Importantly, in the second generation after transformation, Cas9 free plants are identified simply by screening the non-fluorescent seeds. Our collection of binary vectors allows to choose between two widely-used promoters to drive Cas enzymes, either the egg cell-specific (pEC1.2) from A. thaliana or the constitutive promoter from Petroselinum crispum (PcUBi4-2). Available enzymes are “classical” Cas9 codon-optimized for A. thaliana and a recently reported, intron-containing version of Cas9 codon-optimized for Zea mays, zCas9i. We observed the highest efficiency in producing knockout phenotypes by using intron-containing zCas9i driven under egg-cell specific pEC1.2 promoter. Finally, we introduced convenient restriction sites flanking promoter, Cas9 and fluorescent selection cassette in some of the T-DNA vectors, thus allowing straightforward swapping of all three elements for further adaptation and improvement of the system.ConclusionA rapid, simple and flexible CISPR/Cas9 cloning system was established that allows assembly of multi-guide RNA constructs in a robust and reproducible fashion, by avoiding generation of very big constructs. The system enables a flexible, fast and efficient screening of single or higher order A. thaliana mutants.

Fast and Efficient Generation of Isogenic Induced Pluripotent Stem Cell Lines Using Adenine Base Editing.

Isogenic induced pluripotent stem cell (iPSC) lines are currently mostly created by homology directed repair evoked by a double-strand break (DSB) generated by CRISPR-Cas9. However, this process is in general lengthy and inefficient. This problem can be overcome, specifically for correction or insertion of transition mutations, by using base editing (BE). BE does not require DSB formation, hence avoiding creation of genomic off-target breaks and insertions and deletions, and as it is highly efficient, it also does not require integration of selection cassettes in the genome to enrich for edited cells. BE has been successfully used in many cell types as well as in some in vivo settings to correct or insert mutations, but very few studies have reported generation of isogenic iPSC lines using BE. Here, we describe a simple and fast workflow to generate isogenic iPSCs efficiently with a compound heterozygous or a homozygous Wolfram syndrome 1 (WFS1) mutation using adenine BE, without the need to include a genomic selection cassette and without off-target modifications. We demonstrated that correctly base-edited clones can be generated by screening only five cell clones in less than a month, provided that the mutation is positioned in a correct place with regards to the protospacer adjacent motif sequence and no putative bystander bases exist.

Screening and validation of genome-edited animals.

The emergence of an array of genome-editing tools in recent years has facilitated the introduction of genetic modifications directly into the embryo, increasing the ease, efficiency and catalogue of alleles accessible to researchers across a range of species. Bypassing the requirement for a selection cassette and resulting in a broad range of outcomes besides the desired allele, genome editing has altered the allele validation process both temporally and technically. Whereas traditional gene targeting relies upon selection and allows allele validation at the embryonic stem cell modification stage, screening for the presence of the intended allele now occurs in the (frequently mosaic) founder animals. Final confirmation of the edited allele can only take place at the subsequent G1 generation and the validation strategy must differentiate the desired allele from a range of unintended outcomes. Here we present some of the challenges posed by gene editing, strategies for validation and considerations for animal colony management.

VEGF receptor 2 (KDR) protects airways from mucus metaplasia through a Sox9-dependent pathway.

Mucus-secreting goblet cells are the dominant cell type in pulmonary diseases, e.g., asthma and cystic fibrosis (CF), leading to pathologic mucus metaplasia and airway obstruction. Cytokines including IL-13 are the major players in the transdifferentiation of club cells into goblet cells. Unexpectedly, we have uncovered a previously undescribed pathway promoting mucous metaplasia that involves VEGFa and its receptor KDR. Single-cell RNA sequencing analysis coupled with genetic mouse modeling demonstrates that loss of epithelial VEGFa, KDR, or MEK/ERK kinase promotes excessive club-to-goblet transdifferentiation during development and regeneration. Sox9 is required for goblet cell differentiation following Kdr inhibition in both mouse and human club cells. Significantly, airway mucous metaplasia in asthmatic and CF patients is also associated with reduced KDR signaling and increased SOX9 expression. Together, these findings reveal an unexpected role for VEGFa/KDR signaling in the defense against mucous metaplasia, offering a potential therapeutic target for this common airway pathology.

Efficient Targeted Mutagenesis Mediated by CRISPR-Cas12a Ribonucleoprotein Complexes in Maize.

Recent advances in the development of CRISPR-Cas genome editing technologies have made it possible to perform targeted mutagenesis and precise gene replacement in crop plants. CRISPR-Cas9 and CRISPR-Cas12a are two main types of widely used genome editing systems. However, when CRISPR-Cas12a editing machinery is expressed from a transgene, some chromosomal targets encountered low editing frequency in important crops like maize and soybean. Here, we report efficient methods to directly generate genome edited lines by delivering Cas12a-gRNA ribonucleoprotein complex (RNP) to immature maize embryos through particle bombardment in an elite maize variety. Genome edited lines were obtained at ~7% frequency without any selection during regeneration via biolistic delivery of Cas12a RNP into immature embryos. Strikingly, the gene editing rate was increased to 60% on average and up to 100% in some experiments when the Cas12a RNP was co-delivered with a PMI selectable marker gene cassette and the induced callus cultures were selected with mannose. We also show that use of higher activity Cas12a mutants resulted in improved editing efficiency in more recalcitrant target sequence. The advances described here provide useful tools for genetic improvement of maize.

Rapid Genome Modification in Serratia marcescens Through Red Homologous Recombination.

Despite the great potential of Serratia marcescens in industrial applications, lack of powerful genetic modification tools limits understanding of the regulatory networks of the useful metabolites and therefore restricts their mass production. To meet the urgent demand, we established a genome-editing strategy for S. marcescens based on Red recombineering in this study. Without host modification in advance, nucA and pigA were substituted by PCR-amplified resistance genes. No long homologous arms were required at the two sides of resistance genes. Using this procedure, the fragment at the S. marcescens as large as 20 kb was easily deleted. Then we constructed a counter-selection gene kil constructed under the control of inducible PBAD operon, which demonstrates obvious lethality to S. marcescens. Subsequently, GmR-kil double selection cassette was inserted into the CDS of pigA gene. Using single-stranded DNA-mediated recombination, this insertion mutation was efficiently repaired through kil counter-selection. A powerful genetic modification platform based on Red recombineering system was successfully established for S. marcescens. Multiple types of modification and multiple recombination strategies can all be performed easily in this species. We hope this study will be useful for the theoretical research and the research of metabolic engineering in S. marcescens.

H3.3-K27M drives neural stem cell-specific gliomagenesis in a human iPSC-derived model.

Diffuse intrinsic pontine glioma (DIPG) is an aggressive childhood tumor of the brainstem with currently no curative treatment available. The vast majority of DIPGs carry a histone H3 mutation leading to a lysine 27-to-methionine exchange (H3K27M). We engineered human induced pluripotent stem cells (iPSCs) to carry an inducible H3.3-K27M allele in the endogenous locus and studied the effects of the mutation in different disease-relevant neural cell types. H3.3-K27M upregulated bivalent promoter-associated developmental genes, producing diverse outcomes in different cell types. While being fatal for iPSCs, H3.3-K27M increased proliferation in neural stem cells (NSCs) and to a lesser extent in oligodendrocyte progenitor cells (OPCs). Only NSCs gave rise to tumors upon induction of H3.3-K27M and TP53 inactivation in an orthotopic xenograft model recapitulating human DIPGs. In NSCs, H3.3-K27M leads to maintained expression of stemness and proliferative genes and a premature activation of OPC programs that together may cause tumor initiation.

A CRISPR-Cas9, Cre-lox, and Flp-FRT Cascade Strategy for the Precise and Efficient Integration of Exogenous DNA into Cellular Genomes.

We describe a protocol for the precise integration of exogenous DNA into user-defined genomic loci in cultured cells. This strategy first introduces a promoter and a lox site to a specific location via a Cas9-induced double-strand break. Second, a gene of interest (GOI) is inserted into the lox site via Cre-lox recombination. Upon correct insertion, a cis-linked antibiotic resistance gene will be expressed from a promoter introduced into the genome in the first step assuring selection for correct integrants. Last, the selection cassette is excised via a Flp-FRT recombination event, leaving a precisely targeted GOI. This method is broadly applicable to any exogenous DNA to be integrated, choice of integration site, and choice of cell type. The most remarkable aspect of this versatile approach, termed "CasPi" (cascaded precise integration), is that it allows for precise genome targeting with large, frequently complex, and repetitive DNA sequences that do not integrate efficiently or at all with current genome targeting methods.

Peel-1 negative selection promotes screening-free CRISPR-Cas9 genome editing in Caenorhabditis elegans.

Improved genome engineering methods that enable automation of large and precise edits are essential for systematic investigations of genome function. We adapted peel-1 negative selection to an optimized Dual-Marker Selection (DMS) cassette protocol for CRISPR-Cas9 genome engineering in Caenorhabditis elegans and observed robust increases in multiple measures of efficiency that were consistent across injectors and four genomic loci. The use of Peel-1-DMS selection killed animals harboring transgenes as extrachromosomal arrays and spared genome-edited integrants, often circumventing the need for visual screening to identify genome-edited animals. To demonstrate the applicability of the approach, we created deletion alleles in the putative proteasomal subunit pbs-1 and the uncharacterized gene K04F10.3 and used machine vision to automatically characterize their phenotypic profiles, revealing homozygous essential and heterozygous behavioral phenotypes. These results provide a robust and scalable approach to rapidly generate and phenotype genome-edited animals without the need for screening or scoring by eye.

Sizing, stabilising, and cloning repeat-expansions for gene targeting constructs.

Aberrant microsatellite repeat-expansions at specific loci within the human genome cause several distinct, heritable, and predominantly neurological, disorders. Creating models for these diseases poses a challenge, due to the instability of such repeats in bacterial vectors, especially with large repeat expansions. Designing constructs for more precise genome engineering projects, such as engineering knock-in mice, proves a greater challenge still, since these unstable repeats require numerous cloning steps in order to introduce homology arms or selection cassettes. Here, we report our efforts to clone a large hexanucleotide repeat in the C9orf72 gene, originating from within a BAC construct, derived from a C9orf72-ALS patient. We provide detailed methods for efficient repeat sizing and growth conditions in bacteria to facilitate repeat retention during growth and sub-culturing. We report that sub-cloning into a linear vector dramatically improves stability, but is dependent on the relative orientation of DNA replication through the repeat, consistent with previous studies. We envisage the findings presented here provide a relatively straightforward route to maintaining large-range microsatellite repeat-expansions, for efficient cloning into vectors.

Footprint-free gene mutation correction in induced pluripotent stem cell (iPSC) derived from recessive dystrophic epidermolysis bullosa (RDEB) using the CRISPR/Cas9 and piggyBac transposon system

BackgroundRecessive dystrophic epidermolysis bullosa (RDEB) is a monogenic skin blistering disorder caused by mutations in the type VII collagen gene. A combination of biological technologies, including induced pluripotent stem cells (iPSCs) and several gene-editing tools, allows us to develop gene and cell therapies for such inherited diseases. However, the methodologies for gene and cell therapies must be continuously innovated for safe clinical use. ObjectiveIn this study, we used the clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 technology to correct the pathogenic mutation in RDEB-specific iPSCs, and the piggyBac transposon system so that no residual gene fragments remained in the genome of iPSCs after correcting the mutation. MethodsFor homologous recombination (HR)-based gene editing using CRISPR/Cas9, we designed guide RNA and template DNA including homologous sequences with drug-mediated selection cassette flanked by inverted repeat sequences of the transposon. HR reaction using CRISPR/Cas9 was induced in RDEB-specific iPSCs, and mutation-corrected iPSCs (MC-iPSCs) was obtained. Consequently, the selection cassette in the genome of MC-iPSCs was removed by transposase expression. ResultsAfter CRISPR/Cas9-induced gene editing, we confirmed that the pathogenic mutation in RDEB-specific iPSCs was properly corrected. In addition, MC-iPSCs had no genetic footprint after removing the selection cassette by transposon system, and maintained their “stemness”. When differentiating MC-iPSCs into keratinocytes, the expression of type VII collagen was restored. ConclusionsOur study demonstrated one of the safer approaches to establish gene and cell therapies for skin hereditary disorders for future clinical use.

In Vitro Directed Evolution of a Restriction Endonuclease with More Stringent Specificity

Restriction endonuclease (REase) specificity engineering is extremely difficult. Here we describe a multistep protocol that helps to produce REase variants that have more stringent specificity than the parental enzyme. The protocol requires the creation of a library of expression selection cassettes (ESCs) for variants of the REase, ideally with variability in positions likely to affect DNA binding. The ESC is flanked on one side by a sequence for the restriction site activity desired and a biotin tag and on the other side by a restriction site for the undesired activity and a primer annealing site. The ESCs are transcribed and translated in a water-in-oil emulsion, in conditions that make the presence of more than one DNA molecule per droplet unlikely. Therefore, the DNA in each cassette molecule is subjected only to the activity of the translated, encoded enzyme. REase variants of the desired specificity remove the biotin tag but not the primer annealing site. After breaking the emulsion, the DNA molecules are subjected to a biotin pulldown, and only those in the supernatant are retained. This step assures that only ESCs for variants that have not lost the desired activity are retained. These DNA molecules are then subjected to a first PCR reaction. Cleavage in the undesired sequence cuts off the primer binding site for one of the primers. Therefore, PCR amplifies only ESCs from droplets without the undesired activity. A second PCR reaction is then carried out to reintroduce the restriction site for the desired specificity and the biotin tag, so that the selection step can be reiterated. Selected open reading frames can be overexpressed in bacterial cells that also express the cognate methyltransferase of the parental REase, because the newly evolved REase targets only a subset of the methyltransferase target sites.

In Vitro Directed Evolution of a Restriction Endonuclease with More Stringent Specificity.

Restriction endonuclease (REase) specificity engineering is extremely difficult. Here we describe a multistep protocol that helps to produce REase variants that have more stringent specificity than the parental enzyme. The protocol requires the creation of a library of expression selection cassettes (ESCs) for variants of the REase, ideally with variability in positions likely to affect DNA binding. The ESC is flanked on one side by a sequence for the restriction site activity desired and a biotin tag and on the other side by a restriction site for the undesired activity and a primer annealing site. The ESCs are transcribed and translated in a water-in-oil emulsion, in conditions that make the presence of more than one DNA molecule per droplet unlikely. Therefore, the DNA in each cassette molecule is subjected only to the activity of the translated, encoded enzyme. REase variants of the desired specificity remove the biotin tag but not the primer annealing site. After breaking the emulsion, the DNA molecules are subjected to a biotin pulldown, and only those in the supernatant are retained. This step assures that only ESCs for variants that have not lost the desired activity are retained. These DNA molecules are then subjected to a first PCR reaction. Cleavage in the undesired sequence cuts off the primer binding site for one of the primers. Therefore, PCR amplifies only ESCs from droplets without the undesired activity. A second PCR reaction is then carried out to reintroduce the restriction site for the desired specificity and the biotin tag, so that the selection step can be reiterated. Selected open reading frames can be overexpressed in bacterial cells that also express the cognate methyltransferase of the parental REase, because the newly evolved REase targets only a subset of the methyltransferase target sites.

Correction of ATM mutations in iPS cells from two ataxia-telangiectasia patients restores DNA damage and oxidative stress responses.

Patients with ataxia-telangiectasia (A-T) lack a functional ATM kinase protein and exhibit defective repair of DNA double-stranded breaks and response to oxidative stress. We show that CRISPR/Cas9-assisted gene correction combined with piggyBac (PB) transposon-mediated excision of the selection cassette enables seamless restoration of functional ATM alleles in induced pluripotent stem cells from an A-T patient carrying compound heterozygous exonic missense/frameshift mutations, and from a patient with a homozygous splicing acceptor mutation of an internal coding exon. We show that the correction of one allele restores expression of ~ 50% of full-length ATM protein and ameliorates DNA damage-induced activation (auto-phosphorylation) of ATM and phosphorylation of its downstream targets, KAP-1 and H2AX. Restoration of ATM function also normalizes radiosensitivity, mitochondrial ROS production and oxidative-stress-induced apoptosis levels in A-T iPSC lines, demonstrating that restoration of a single ATM allele is sufficient to rescue key ATM functions. Our data further show that despite the absence of a functional ATM kinase, homology-directed repair and seamless correction of a pathogenic ATM mutation is possible. The isogenic pairs of A-T and gene-corrected iPSCs described here constitute valuable tools for elucidating the role of ATM in ageing and A-T pathogenesis.

Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in <em>Chlamydia trachomatis</em>

Chlamydia trachomatis is an obligate intracellular pathogen that has been historically difficult to genetically manipulate. Definitive progress in elucidating the mechanisms that C. trachomatis use to create and maintain a privileged intracellular niche has been limited due to a lack of genetic tools. Fortunately, there have recently been several new advances in genetic manipulation techniques. Among these is the development of fluorescence-reported allelic exchange mutagenesis (FRAEM). This method allows targeted gene deletion coupled with insertion of a selection cassette encoding antibiotic resistance and green fluorescent protein (GFP). Reliance on this strategy can be complicated when targeting genes within polycistronic operons due to the potential of polar effects on downstream genes. Floxed cassette allelic exchange mutagenesis (FLAEM), the protocol for which is described here, was developed to alleviate cassette-induced polar effects. FLAEM utilizes Cre-loxP genome editing to remove the selection cassette after targeted deletion by allelic exchange. The resulting strains contain markerless gene deletions of one or more coding sequences. This technique facilitates direct assessment of gene function and expands the repertoire of tools for genetic manipulation in C. trachomatis.